The Liver: Biology and Pathobiology 5th Edition [5 ed.] 9780470723135, 0470723130

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The Liver

The Liver Biology and Pathobiology Fifth Edition

Editor Irwin M. Arias Senior Scientist, National Institutes of Health, Bethesda, MD, USA; Emeritus Professor of Physiology, Tufts University School of Medicine, Boston, MA, USA; Visiting Professor of Medicine, Albert Einstein College of Medicine, Bronx, NY, USA

Associate Editors Harvey J. Alter Distinguished NIH Investigator, Department of Transfusion Medicine, Clinical Center, National Institutes of Health, Bethesda, MD, USA; Clinical Professor of Medicine, Georgetown University, Washington, DC, USA; Adjunct Scientist, Southwest Foundation for Biomedical Research, San Antonio, TX, USA

James L. Boyer Department of Internal Medicine and Liver Center Yale University School of Medicine, New Haven, CT, USA

David E. Cohen Department of Medicine, Gastroenterology Division Brigham and Women’s Hospital, Harvard Medical School and Harvard—Massachusetts Institute of Technology Division of Health Sciences and Technology, Boston, MA, USA

Nelson Fausto Department of Pathology, School of Medicine, University of Washington, Seattle, WA, USA

David A. Shafritz Marion Bessin Liver Research Center Albert Einstein College of Medicine, Bronx, New York, NY, USA

Allan W. Wolkoff Division of Hepatology, Marion Bessin Liver Research Center, and Department of Anatomy and Structural Biology, Albert Einstein College of Medicine, Bronx, NY, USA

A John Wiley & Sons, Ltd., Publication

This edition first published 2009,  2009, John Wiley & Sons Ltd. Wiley-Blackwell is an imprint of John Wiley & Sons, formed by the merger of Wiley’s global Scientific, Technical and Medical business with Blackwell Publishing. Registered office: John Wiley & Sons Ltd, The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, UK Other Editorial Offices 9600 Garsington Road, Oxford, OX4 2DQ, UK 111 River Street, Hoboken, NJ 07030-5774, USA For details of our global editorial offices, for customer services and for information about how to apply for permission to reuse the copyright material in this book please see our website at www.wiley.com/wiley-blackwell The right of the author to be identified as the author of this work has been asserted in accordance with the Copyright, Designs and Patents Act 1988. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, except as permitted by the UK Copyright, Designs and Patents Act 1988, without the prior permission of the publisher. Wiley also publishes its books in a variety of electronic formats. Some content that appears in print may not be available in electronic books. Designations used by companies to distinguish their products are often claimed as trademarks. All brand names and product names used in this book are trade names, service marks, trademarks or registered trademarks of their respective owners. The publisher is not associated with any product or vendor mentioned in this book. This publication is designed to provide accurate and authoritative information in regard to the subject matter covered. It is sold on the understanding that the publisher is not engaged in rendering professional services. If professional advice or other expert assistance is required, the services of a competent professional should be sought. The contents of this work are intended to further general scientific research, understanding, and discussion only and are not intended and should not be relied upon as recommending or promoting a specific method, diagnosis, or treatment by physicians for any particular patient. The publisher and the author make no representations or warranties with respect to the accuracy or completeness of the contents of this work and specifically disclaim all warranties, including without limitation any implied warranties of fitness for a particular purpose. In view of ongoing research, equipment modifications, changes in governmental regulations, and the constant flow of information relating to the use of medicines, equipment, and devices, the reader is urged to review and evaluate the information provided in the package insert or instructions for each medicine, equipment, or device for, among other things, any changes in the instructions or indication of usage and for added warnings and precautions. Readers should consult with a specialist where appropriate. The fact that an organization or Website is referred to in this work as a citation and/or a potential source of further information does not mean that the author or the publisher endorses the information the organization or Website may provide or recommendations it may make. Further, readers should be aware that Internet Websites listed in this work may have changed or disappeared between when this work was written and when it is read. No warranty may be created or extended by any promotional statements for this work. Neither the publisher nor the author shall be liable for any damages arising herefrom. Library of Congress Cataloguing-in-Publication Data The liver : biology and pathobiology / editors, Irwin M. Arias . . . [et al.]. – 5th ed. p. ; cm. Includes bibliographical references and index. ISBN 978-0-470-72313-5 1. Liver–Physiology. 2. Liver–Pathophysiology. I. Arias, Irwin M. [DNLM: 1. Liver–pathology. 2. Liver–physiology. WI 700 L7825 2009] QP185.L54 2009 612.3′ 5–dc22 2009025380 ISBN: 978-0-470-72313-5 A catalogue record for this book is available from the British Library. Typeset in 9.75/11.75 Times Roman by Laserwords Private Limited, Chennai, India Printed in Singapore by Fabulous Printers Pte Ltd. First Impression 2009

Dedication We dedicate this edition to the National Institutes of Health (NIH) of the Department of Health and Human Services of the United States Government. It was the vision of James Shannon, Director from 1955 to 1968, that the creation of extramural and intramural research programs would generate a strong basis for accelerating post-war biologic science and improving human health. Since the creation of these programs, the NIH has been the major worldwide source of public funds supporting biomedical research, and the peer review system proved to be the most effective mechanism for allocation of public funds. The Intramural and Extramural programs have been catalytic for virtually every major advance in biology and medicine in the United States, including better understanding of liver structure, function, and disease.

Major advances in hepatology would not have been possible without the support of public funds administered through the NIH. The Intramural program played a particularly major role in the study, detection, and analysis of hepatitis viruses, and also prevention of post-transfusion hepatitis and other infections. NIH support has been global, as have the resulting advances and benefits. At least three generations of American and other basic and clinical investigators owe their careers to support from the NIH. We hope that this investment by the American public in science and world health will be sustained, thereby permitting effective bridging of the amazing advances in the biological sciences with human health.

Contents List of Contributors

xiii

Preface

xxiii

Acknowledgements

xxiv

PART ONE

INTRODUCTION

1

1 Organizational Principles of the Liver Joe W. Grisham

3

2 Embryonic Development of the Liver Roque Bort and Kenneth S. Zaret

17

PART TWO THE CELLS

27

3 Microtubules, Actin Filaments and Motor-mediated Vesicular Transport Ronald R. Marchelletta and Sarah F. Hamm-Alvarez

29

4 Molecular Motors Peter Satir

45

5 Ion Pumps and Molecular Motors: P-, F-, and V-type ATPases Sarah Bond, Daniel J. Cipriano, and Michael Forgac

57

6 Hepatocyte Surface Polarity: Its Dynamic Maintenance and Establishment Lelita T. Braiterman and Ann L. Hubbard

73

7 Endocytosis as an Essential Process in Liver Function and Pathology Barbara Schroeder and Mark McNiven

107

8 Membrane Transport in Hepatocellular Secretion Susan Chi and Mark McNiven

125

9 Mitochondria Kasturi Mitra

137

10 Nuclear Pore Complex Joseph S. Glavy

147

11 Protein Maturation and Processing at the Endoplasmic Reticulum Ramanujan S. Hegde

157

12 Protein Degradation and the Lysosomal System Susmita Kaushik and Ana Maria Cuervo

173

viii

CONTENTS

13 Peroxisome Assembly, Degradation, and Disease Peter K. Kim

191

14 Gap and Tight Junctions in Liver: Composition, Regulation, and Function Takashi Kojima, Norimasa Sawada, Hiroshi Yamaguchi, Alfredo G. Fort, and David C. Spray

201

SECTION A HEPATOCYTE

221

15 Copper Metabolism and the Liver Michael L. Schilsky and Dennis J. Thiele

223

16 The Central Role of the Liver in Iron Storage and Regulation of Systemic Iron Homeostasis Tracey A. Rouault, Victor Gordeuk, and Gregory Anderson

235

17 Disorders of Bilirubin Metabolism Namita Roy Chowdhury and Jayanta Roy Chowdhury

251

4.21. The porphyrias Peter N. Meissner, Richard J. Hift, Ralph E. Kirsch 18 Hepatic Fatty Acid Metabolism and Dysfunction David L. Silver

257

19 Lipoprotein Metabolism and Cholesterol Balance David E. Cohen

271

SECTION B

287

BILE SECRETION

20 Bile Acids and the Enterohepatic Circulation Alan F. Hofmann

289

21 Hepatocyte Basolateral Membrane Organic Anion Transporters Jo H. Choi, John W. Murray, and Allan W. Wolkoff

305

22 Nuclear Receptors Regulate Bile Acid Synthesis Guorong Xu and Gerald Salen

323

4.27. Hormonal regulation of bile secretion Francis R. Simon 4.28. Nucleotide transport and regulation of bile formation Richard M. Roman and J. Gregory Fitz 23 The Function of the Canalicular Membrane in Bile Formation and Secretion Ronald P.J. Oude Elferink and Coen C. Paulusma

339

24 Apical Recycling of Canalicular ABC Transporters Yoshiyuki Wakabayashi and Irwin M. Arias

349

25 Cholangiocyte Functions in Health and Disease: The Ciliary Connection Anatoliy I. Masyuk, Tatyana V. Masyuk, and Nicholas F. LaRusso

359

SECTION C SINUSOIDAL CELLS

371

26 The Hepatic Sinusoidal Endothelial Cell: Morphology, Function, and Pathobiology Laurie D. DeLeve

373

www.GastroHep.com

CONTENTS

ix

4.30. Endothelial and pit cells Filip Braet, Dianzhong Luo, Ilan Spector, David Vermijlen, Eddie Wisse 27 Fenestrations in the Liver Sinusoidal Endothelial Cell Victoria C. Cogger and David G. Le Couteur

389

28 Hepatic Stellate Cells Marcos Rojkind and Karina Reyes-Gordillo

407

29 Hepatic Fibrosis Ram´on Bataller and David A. Brenner

433

30 Matrix Giuliano Ramadori and Jozsef Dudas

453

PART THREE

469

INTERRELATED CELL FUNCTIONS

31 Insulin Resistance Varman T. Samuel, Kitt F. Petersen, and Gerald I. Shulman

471

32 Ca2+ Signaling in the Liver Fatima M. Leite, Mateus T. Guerra, and Michael H. Nathanson

485

4.35. Synthesis of and signalling through D-3 phosphoinositides Rial A Christensen, Isabel De Aos Scherpenseel, Lyuba Varticovski 4.39. Nitric oxide in the liver Mark G. Clemens 33 Role of Intracellular Iron Movement and Oxidant Stress in Hepatocellular Injury John J. Lemasters, Akira Uchiyama, Jae-Sung Kim, Kazuyoshi Kon, and Hartmut Jaeschke

511

4.18. Hypoxic, Ischemic and reperfusion injury to liver John J. Lemasters 4.19. Protective mechanisms against reactive oxygen species Masayasu Inoue 34 Regulatory Pathways of Liver Gene Expression: The Central Role of Cyclic AMP Giuseppe Servillo, Maria Agnese Della Fazia, and Paolo Sassone-Corsi

521

35 AMPK: Central Regulator of Glucose and Lipid Metabolism Maria M. Mihaylova and Reuben J. Shaw

535

3.16. Energy Metabolism Sam Seifter and Sasha Englard 36 Liver Regeneration Nelson Fausto and Jean S. Campbell

549

4.40. Interleukin-6 signaling during the acute-phase response of the liver Johannes G. Bode, Peter C. Heinrich 4.43. Hepatocyte growth factor: its role in hepatic growth and pathobiology Reza Zarnegar, Marie C. Defrances, George K. Michalopoulos 37 Ribosome Biogenesis and its Role in Cell Growth and Proliferation in the Liver Stefano Fumagalli and George Thomas

www.GastroHep.com

567

x

CONTENTS

4.5. Gene regulation and in vivo function of liver transcription factors Robert H. Costa, Al-Xuan Le Holternan, Francisco M. Rausa, Guy R. Adami 38 Liver Repopulation by Cell Transplantation and the Role of Stem Cells David A. Shafritz, Michael Oertel, Mariana D. Dabeva, and Markus Grompe

577

PART FOUR

597

RELATION TO OTHER ORGANS

39 Hepatic Encephalopathy Roger F. Butterworth and Javier Vaquero

599

40 The Kidney in Liver Disease Moshe Levi

619

41 Critical Role of the Liver in Coagulation Robert Fathke, Ze Peng, Basil Golding, and Chava Kimchi-Sarfaty

639

PART FIVE

659

PATHOBIOLOGICAL ANALYSIS

42 Inheritable Cholestatic Disorders Paul Gissen and A. S. Knisely

661

4.48. Alpha 1-Antitrypsin deficiency David H. Perlmutter 43 Adaptive Regulation of Hepatocyte Transporters in Cholestasis James L. Boyer

681

44 Pathogenesis of Portal Hypertension Roberto J. Groszmann and Juan G. Abraldes

703

45 Non-alcoholic Fatty Liver Disease: A Pathophysiological Perspective Michael Fuchs and Arun J. Sanyal

719

46 Pathophysiology of Alcoholic Liver Disease Natalia Nieto and Marcos Rojkind

743

47 Inflammation and Drug-induced Liver Injury Robert A. Roth and Patricia E. Ganey

773

48 Hepatocyte Apoptosis Cynthia R.L. Webster

783

49 Back to the Future: A Backward Glance at the Forward Progress of Hepatitis Virus Research Harvey J. Alter

803

50 Molecular Biology of Hepatitis Viruses Christoph Seeger, Michael M.C. Lai, and William S. Mason

807

51 Immune Mechanisms of Viral Clearance and Disease Pathogenesis During Viral Hepatitis Carlo Ferrari and Mario Mondelli

835

4.41. Innate immune sensing and the toll-like receptors Bruce Beutler 52 Clinical Implications of the Molecular Biology of Hepatitis B Virus Timothy M. Block, Ju-Tao Guo, and W. Thomas London

859

53 Viral Escape Mechanisms in Hepatitis C and the Clinical Consequences of Persistent Infection Stanley M. Lemon, Patrizia Farci, and Marc G. Ghany

877

www.GastroHep.com

CONTENTS

54 Current and Future Therapy for Hepatitis B and C Gary L. Davis and Jean-Michel Pawlotsky

xi

899

55 Biological Principles and Clinical Issues Underlying Liver Transplantation for Virus-induced End-stage Liver Disease James R. Burton Jr, Hugo R. Rosen, and Paul Martin

921

PART SIX

933

HORIZONS

56 Tissue Engineering of the Liver Gregory H. Underhill, Salman R. Khetani, Alice A. Chen, and Sangeeta N. Bhatia

935

57 Aging and the Uncertain Roles of Sirtuins J. Fred Dice

955

58 The Liver Proteome Laura Beretta

961

59 Liver-directed Gene Therapy Betsy T. Kren, Clifford J. Steer, Namita Roy Chowdhury, and Jayanta Roy Chowdhury

965

4.63. Novel strategies for manipulating hepatic gene expression in vivo Marxa L. Figueiredo, Eric P. Sandgren 60 Decoding the Liver Cancer Genome Ju-Seog Lee and Snorri S. Thorgeirsson

991

61 Genome-wide Expression Profiling of Human Hepatocellular Carcinoma Anuradha Budhu and Xin Wei Wang

999

62 Cell Cycle Control in the Liver Jeffrey H. Albrecht and Lisa K. Mullany

1015

63 miRNAs and Liver Biology Charles E. Rogler and Leslie E. Rogler

1029

64 Imaging Cellular Proteins and Structures: Smaller, Brighter, and Faster Erik Snapp

1053

65 Zebrafish as a Model System for the Study of Liver Development and Disease Randolph P. Matthews

1067

66 The Hepatocyte and the Cancer Cell: Dr Jekyll and Mr Hyde Jean-Pierre Gillet, Michael M. Gottesman, and Mitsunori Okabe

1075

67 The Role of Endocannabinoids and Their Receptors in the Control of Hepatic Functions George Kunos, Douglas Osei-Hyiaman, S´andor B´atkai, P´al Pacher, Bin Gao, Won-Il Jeong, Jie Liu, and Gregorz Godlewski

1091

68 Telomeres and Aging, Cancer, and Hepatic Fibrosis Hans L. Tillmann, Ruben R. Plentz, Yvonne Begus-Nahrmann, Andr´e Lechel, and Lenhard K. Rudolph

1105

69 Treatment of Cirrhosis with Vitamin A-coupled Liposomes Carrying siRNA against Heat Shock Protein 47 Yoshiro Niitsu, Yasushi Sato, Kazuyuki Murase, and Junji Kato

1121

70 The “Green Liver” and Transcriptional Regulation of Phase II Detoxification Genes Christopher Johnson and Jonathan Arias

1131

Index

1139

www.GastroHep.com

List of Contributors Juan G. Abraldes Hepatic Hemodynamic Laboratory Liver Unit Hospital Clinic University of Barcelona IDIBAPS and CiberEHD Barcelona Spain Jeffrey H. Albrecht Division of Gastroenterology University of Minnesota and Hennepin County Medical Center Minneapolis, MN USA Harvey J. Alter Distinguished NIH Investigator Department of Transfusion Medicine Clinical Center National Institutes of Health Bethesda, MD USA Clinical Professor of Medicine Georgetown University Washington, DC USA and Adjunct Scientist Southwest Foundation for Biomedical Research San Antonio, TX USA Gregory Anderson Queensland Institute of Medical Research PO Royal Brisbane Hospital Brisbane, Queensland Australia Irwin M. Arias Senior Scientist National Institutes of Health Bethesda, MD USA

Emeritus Professor of Physiology Tufts University School of Medicine Boston, MA USA and Visiting Professor of Medicine Albert Einstein College of Medicine Bronx, NY USA Jonathan Arias Department of Biological Sciences University of Maryland Baltimore County Baltimore, MD USA Ram´on Bataller Liver Unit Institut de Malalties Digestives i Metab`oliques Hospital Cl´ınic IDIBAPS Barcelona Spain S´andor B´atkai National Institute on Alcohol Abuse and Alcoholism National Institutes of Health Bethesda, MD USA Yvonne Begus-Nahrmann Institute of Molecular Medicine and Max Planck Research Group on Stem Cell Aging Ulm University Ulm Germany Laura Beretta Molecular Diagnostics Program Public Health Sciences Division Fred Hutchinson Cancer Research Center Seattle, WA USA

xiv

LIST OF CONTRIBUTORS

Sangeeta N. Bhatia Harvard-MIT Division of Health Sciences and Technology/Electrical Engineering and Computer Science Massachusetts Institute of Technology Boston, MA USA and Division of Medicine Brigham & Women’s Hospital Boston, MA USA Timothy M. Block Drexel University College of Medicine and the Hepatitis B Foundation Pennsylvania Biotechnology Center Doylestown, PA USA Sarah Bond Department of Physiology Tufts University School of Medicine Boston, MA USA Roque Bort Unidad de Hepatolog´ıa Experimental CIBEREHD Centro de Investigaci´on Hospital Universitario La Fe Valencia Spain James L. Boyer Department of Internal Medicine and Liver Center Yale University School of Medicine New Haven, CT USA Lelita T. Braiterman Department of Cell Biology Johns Hopkins University School of Medicine Baltimore, MD USA David A. Brenner UCSD School of Medicine University of California San Diego, CA USA Anuradha Budhu Liver Carcinogenesis Section Laboratory of Human Carcinogenesis Center for Cancer Research National Cancer Institute National Institutes of Health Bethesda, MD USA

James R. Burton Jr University of Colorado Denver, CO USA Roger F. Butterworth Neuroscience Research Unit Hˆopital Saint-Luc University of Montreal Montreal Canada Jean S. Campbell Department of Pathology School of Medicine University of Washington Seattle, WA USA Alice A. Chen Harvard-MIT Division of Health Sciences and Technology/Electrical Engineering and Computer Science Massachusetts Institute of Technology Boston, MA USA Susan Chi Department of Biochemistry and Molecular Biology Miles and Shirley Fiterman Center for Digestive Diseases Mayo Clinic & Foundation Rochester, MN USA Jo H. Choi Division of Hepatology Marion Bessin Liver Research Center and Department of Anatomy and Structural Biology Albert Einstein College of Medicine Bronx, New York, NY USA Daniel J. Cipriano Department of Physiology Tufts University School of Medicine Boston, MA USA Victoria C. Cogger Centre for Education and Research on Ageing University of Sydney and Concord RG Hospital Sydney Australia

LIST OF CONTRIBUTORS

David E. Cohen Department of Medicine Gastroenterology Division Brigham and Women’s Hospital Harvard Medical School and Harvard-Massachusetts Institute of Technology Division of Health Sciences and Technology Boston, MA USA David G. Le Couteur Centre for Education and Research on Ageing University of Sydney and Concord RG Hospital Sydney Australia Ana Maria Cuervo Department of Developmental and Molecular Biology Marion Bessin Liver Research Center Albert Einstein College of Medicine Bronx New York, NY USA Mariana D. Dabeva Marion Bessin Liver Research Center Albert Einstein College of Medicine Bronx NY USA Gary L. Davis Department of Hepatology Baylor University Medical Center Dallas, TX USA Laurie D. DeLeve Division of Gastrointestinal and Liver Diseases University of Southern California Keck School of Medicine Los Angeles, CA USA J. Fred Dice Department of Physiology Tufts University School of Medicine Boston, MA USA Jozsef Dudas Department of Internal Medicine Section of Gastroenterology and Endocrinology Georg-August-University Goettingen Goettingen Germany

Patrizia Farci Hepatic Pathogenesis Unit Laboratory of Infectious Diseases National Institute of Allergy and Infectious Diseases National Institutes of Health Bethesda, MD USA Robert Fathke Division of Hematology Center for Biologics Evaluation and Research US Food and Drug Administration Bethesda, MD USA Nelson Fausto Department of Pathology School of Medicine University of Washington Seattle, WA USA Maria Agnese Della Fazia Department of Clinical and Experimental Medicine School of Medicine University of Perugia Perugia Italy Carlo Ferrari Unit of Infectious Diseases and Hepatology Laboratory of Viral Immunopathology Azienda Ospedaliero–Universitaria di Parma Parma Italy Michael Forgac Department of Physiology Tufts University School of Medicine Boston, MA USA Alfredo G. Fort Department of Neuroscience Albert Einstein College of Medicine Bronx New York, NY USA Michael Fuchs Division of Gastroenterology, Hepatology and Nutrition Department of Internal Medicine Virginia Commonwealth University School of Medicine Richmond, VA USA

xv

xvi

LIST OF CONTRIBUTORS

Stefano Fumagalli Department of Cancer and Cell Biology Genome Research Institute University of Cincinnati Cincinnati, OH USA Patricia E. Ganey Department of Pharmacology and Toxicology Center for Integrative Toxicology Michigan State University East Lansing, MI USA Bin Gao National Institute on Alcohol Abuse and Alcoholism National Institutes of Health Bethesda, MD USA Marc G. Ghany Liver Diseases Branch National Institute of Diabetes and Digestive and Kidney Diseases National Institutes of Health Bethesda, MD USA Jean-Pierre Gillet Laboratory of Cell Biology Center for Cancer Research National Cancer Institute National Institutes of Health Bethesda, MD USA Paul Gissen Inherited Metabolic Diseases Unit Birmingham Children’s Hospital Birmingham UK Joseph S. Glavy Department of Chemistry, Chemical Biology & Biomedical Engineering Stevens Institute of Technology Hoboken, NJ USA Gregorz Godlewski National Institute on Alcohol Abuse and Alcoholism National Institutes of Health Bethesda, MD USA Basil Golding Division of Hematology Center for Biologics Evaluation and Research US Food and Drug Administration Bethesda, MD USA

Victor Gordeuk Howard University Medical Center Washington, D. C. USA Michael M. Gottesman Laboratory of Cell Biology Center for Cancer Research National Cancer Institute National Institutes of Health Bethesda, MD USA Joe W. Grisham Department of Pathology and Laboratory Medicine University of North Carolina at Chapel Hill Chapel Hill, NC USA Markus Grompe Oregon Health Sciences University Portland, OR USA Roberto J. Groszmann Veterans Administration Medical Center West Haven, CT USA and Yale University School of Medicine New Haven, CT USA Mateus T. Guerra Department of Physiology and Biophysics UFMG Belo Horizonte Brazil Ju-Tao Guo Drexel University College of Medicine and the Hepatitis B Foundation Pennsylvania Biotechnology Center Doylestown, PA USA Sarah F. Hamm-Alvarez Department of Pharmacology and Pharmaceutical Sciences USC School of Pharmacy Los Angeles, CA USA Ramanujan S. Hegde Cell Biology and Metabolism Program National Institutes of Child Health and Human Development National Institutes of Health Bethesda, MD USA

LIST OF CONTRIBUTORS

Alan F. Hofmann Division of Gastroenterology Department of Medicine University of California San Diego, CA USA

Peter K. Kim Program in Cell Biology The Hospital for Sick Children Toronto, Ontario Canada

Ann L. Hubbard Department of Cell Biology Johns Hopkins University School of Medicine Baltimore, MD USA

Chava Kimchi-Sarfaty Division of Hematology Center for Biologics Evaluation and Research US Food and Drug Administration Bethesda, MD USA

Hartmut Jaeschke Department of Pharmacology, Toxicology & Therapeutics University of Kansas Medical Center Kansas City, KS USA

A. S. Knisely Institute of Liver Studies/Histopathology King’s College Hospital London UK

Won-Il Jeong National Institute on Alcohol Abuse and Alcoholism National Institutes of Health Bethesda, MD USA

Takashi Kojima Department of Pathology Sapporo Medical University School of Medicine Sapporo Japan

Christopher Johnson Department of Biological Sciences University of Maryland Baltimore County Baltimore, MD USA

Kazuyoshi Kon Department of Gastroenterology Juntendo University School of Medicine Tokyo Japan

Junji Kato Fourth Department of Internal Medicine Sapporo Medical University Sapporo Japan Susmita Kaushik Department of Developmental and Molecular Biology Marion Bessin Liver Research Center Albert Einstein College of Medicine Bronx, New York, NY USA Salman R. Khetani Harvard-MIT Division of Health Sciences and Technology/Electrical Engineering and Computer Science Massachusetts Institute of Technology Boston, MA USA Jae-Sung Kim Department of Surgery University of Florida Gainesville, FL USA

Betsy T. Kren Department of Medicine University of Minnesota School of Medicine Minnesota, MN USA George Kunos National Institute on Alcohol Abuse and Alcoholism National Institutes of Health Bethesda, MD USA Michael M.C. Lai Howard Hughes Medical Institute Department of Molecular Microbiology and Immunology University of Southern California Los Angeles, CA USA Nicholas F. LaRusso Gasteroenterology Research Unit Mayo Graduate School of Medicine Mayo Clinic College of Medicine Rochester, MN USA

xvii

xviii

LIST OF CONTRIBUTORS

Andr´e Lechel Institute of Molecular Medicine and Max Planck Research Group on Stem Cell Aging Ulm University Ulm Germany Ju-Seog Lee Department of Systems Biology The University of Texas M. D. Anderson Cancer Center Houston, TX USA Fatima M. Leite Department of Physiology and Biophysics UFMG Belo Horizonte Brazil Stanley M. Lemon Center for Hepatitis Research Institute for Human Infections and Immunity Galveston, TX USA John J. Lemasters Center for Cell Death, Injury and Regeneration Departments of Pharmaceutical & Biomedical Sciences and Biochemistry & Molecular Biology Medical University of South Carolina Charleston, SC USA Moshe Levi Division of Renal Diseases and Hypertension University of Colorado Denver, CO USA Jie Liu National Institute on Alcohol Abuse and Alcoholism National Institutes of Health Bethesda, MD USA W. Thomas London Fox Chase Cancer Center Philadelphia, PA USA Ronald R. Marchelletta Department of Pharmacology and Pharmaceutical Sciences USC School of Pharmacy Los Angeles, CA USA

Paul Martin Division of Hepatology University of Miami Miami, FL USA William S. Mason Fox Chase Cancer Center Philadelphia, PA USA Anatoliy I. Masyuk Mayo Graduate School of Medicine Mayo Clinic College of Medicine Rochester, MN USA Tatyana V. Masyuk Mayo Graduate School of Medicine Mayo Clinic College of Medicine Rochester, MN USA Randolph P. Matthews Division of Gastroenterology, Hepatology and Nutrition The Children’s Hospital of Philadelphia and Department of Pediatrics University of Pennsylvania School of Medicine Philadelphia, PA USA Mark McNiven Department of Biochemistry & Molecular Biology Miles and Shirley Fiterman Center for Digestive Diseases Mayo Clinic & Foundation Rochester, MN USA Maria M. Mihaylova Molecular and Cell Biology Laboratory The Salk Institute for Biological Studies La Jolla, CA USA Kasturi Mitra Cell Biology and Metabolism Program National Institute of Child Health and Human Development National Institutes of Health Bethesda, MD USA

LIST OF CONTRIBUTORS

Mario Mondelli Laboratori di Ricerca –Area Infettivologica IRCCS Policlinico S. Matteo Universit`a degli Studi di Pavia Pavia Italy

Ronald P.J. Oude Elferink AMC Liver Center Academic Medical Center Amsterdam The Netherlands

Lisa K. Mullany Minneapolis Medical Research Foundation Minneapolis, MN USA

P´al Pacher National Institute on Alcohol Abuse and Alcoholism National Institutes of Health Bethesda, MD USA

Kazuyuki Murase Fourth Department of Internal Medicine Sapporo Medical University Sapporo Japan

Coen C. Paulusma AMC Liver Center Academic Medical Center Amsterdam The Netherlands

John W. Murray Division of Hepatology Marion Bessin Liver Research Center and Department of Anatomy and Structural Biology Albert Einstein College of Medicine Bronx, New York, NY USA

Jean-Michel Pawlotsky Department of Virology and INSERM U955 Henri Mondor Hospital University of Paris 12 Cr´eteil France

Michael H. Nathanson Departments of Medicine and Cell Biology Yale University School of Medicine New Haven, CT USA Natalia Nieto Mount Sinai School of Medicine Department of Medicine/Division of Liver Diseases New York, NY USA Yoshiro Niitsu Department of Molecular Target Exploration Sapporo Medical University Sapporo Japan Michael Oertel Marion Bessin Liver Research Center Albert Einstein College of Medicine Bronx, NY USA Mitsunori Okabe Tohoku University School of Medicine Sendai Japan Douglas Osei-Hyiaman National Institute on Alcohol Abuse and Alcoholism National Institutes of Health Bethesda, MD USA

Ze Peng Division of Hematology Center for Biologics Evaluation and Research US Food and Drug Administration Bethesda, MD USA Kitt F. Petersen Department of Internal Medicine Section of Endocrinology Yale University School of Medicine New Haven, CT USA Ruben R. Plentz Department of Gastroenterology, Hepatology and Endocrinology Medical School Hannover Hannover Germany Giuliano Ramadori Department of Internal Medicine Section of Gastroenterology and Endocrinology Georg-August-University Goettingen Goettingen Germany Karina Reyes-Gordillo Department of Biochemistry and Molecular Biology The George Washington University Medical Center Washington, DC USA

xix

xx

LIST OF CONTRIBUTORS

Charles E. Rogler Marion Bessin Liver Research Center Division of Hepatology Department of Medicine Albert Einstein College of Medicine Bronx New York, NY USA Leslie E. Rogler Marion Bessin Liver Research Center Division of Hepatology Department of Medicine Albert Einstein College of Medicine Bronx New York, NY USA Marcos Rojkind Department of Biochemistry and Molecular Biology The George Washington University Medical Center Washington, DC USA Hugo R. Rosen University of Colorado Denver, CO USA Robert A. Roth Department of Pharmacology and Toxicology Center for Integrative Toxicology Michigan State University East Lansing, MI USA Tracey A. Rouault National Institute of Child Health and Human Development National Institutes of Health Bethesda, MD USA Jayanta Roy Chowdhury Departments of Medicine and Genetics and Marion Bessin Liver Research Center Albert Einstein College of Medicine Bronx New York, NY USA Namita Roy Chowdhury Departments of Medicine and Genetics and Marion Bessin Liver Research Center Albert Einstein College of Medicine Bronx New York, NY USA

Lenhard K. Rudolph Institute of Molecular Medicine and Max Planck Research Group on Stem Cell Aging Ulm University Ulm Germany Gerald Salen University of Medicine and Dentistry of New Jersey New Jersey Medical School NJ USA Varman T. Samuel Department of Internal Medicine Section of Endocrinology Yale University School of Medicine New Haven, CT USA and Veterans Affairs Medical Center West Haven, CT USA Arun J. Sanyal Division of Gastroenterology, Hepatology and Nutrition Department of Internal Medicine Virginia Commonwealth University School of Medicine Richmond, VA USA Paolo Sassone-Corsi Department of Pharmacology School of Medicine University of California Irvine, CA USA Peter Satir Department of Anatomy and Structural Biology Albert Einstein College of Medicine Bronx, New York, NY USA Yasushi Sato Fourth Department of Internal Medicine Sapporo Medical University Sapporo Japan Norimasa Sawada Department of Pathology Sapporo Medical University School of Medicine Sapporo Japan

LIST OF CONTRIBUTORS

Michael L. Schilsky Department of Medicine Yale University Medical Center New Haven, CT USA Barbara Schroeder Department of Biochemistry & Molecular Biology Miles and Shirley Fiterman Center for Digestive Diseases Mayo Clinic & Foundation Rochester, MN USA Christoph Seeger Fox Chase Cancer Center Philadelphia, PA USA Giuseppe Servillo Department of Clinical and Experimental Medicine School of Medicine University of Perugia Perugia Italy David A. Shafritz Marion Bessin Liver Research Center Albert Einstein College of Medicine Bronx, NY USA Reuben J. Shaw Molecular and Cell Biology Laboratory The Salk Institute for Biological Studies La Jolla, CA USA Gerald I. Shulman Department of Internal Medicine Section of Endocrinology Yale University School of Medicine New Haven, CT USA Department of Molecular and Cellular Physiology and Howard Hughes Medical Institute and Yale University School of Medicine New Haven, CT USA David L. Silver Department of Biochemistry Albert Einstein College of Medicine Bronx, New York, NY USA

xxi

Erik Snapp Department of Anatomy and Structural Biology Albert Einstein College of Medicine Bronx New York, NY USA David C. Spray Department of Neuroscience Albert Einstein College of Medicine Bronx, New York, NY USA Clifford J. Steer Departments of Medicine and Genetics, Cell Biology and Development University of Minnesota School of Medicine Minnesota, MN USA Dennis J. Thiele Department of Pharmacology and Cancer Biology Duke University Medical Center Durham, NC USA George Thomas Department of Cancer and Cell Biology Genome Research Institute University of Cincinnati Cincinnati, OH USA Snorri S. Thorgeirsson Laboratory of Experimental Carcinogenesis Center for Cancer Research National Cancer Institute National Institutes of Health Bethesda, MD USA Hans L. Tillmann Duke Clinical Research Institute GI/Hepatology Research Program Division of Gastroenterology Durham, NC USA Akira Uchiyama Department of Gastroenterology Juntendo University School of Medicine Tokyo Japan

xxii

Gregory H. Underhill Harvard-MIT Division of Health Sciences and Technology/Electrical Engineering and Computer Science Massachusetts Institute of Technology Boston, MA USA Javier Vaquero Neuroscience Research Unit Hˆopital Saint-Luc University of Montreal Montreal Canada Yoshiyuki Wakabayashi Unit of Cellular Polarity Cell Biology and Metabolism Program National Institute of Child Health and Human Development National Institutes of Health Bethesda, MD USA Xin Wei Wang Liver Carcinogenesis Section Laboratory of Human Carcinogenesis Center for Cancer Research National Cancer Institute National Institutes of Health Bethesda, MD USA Cynthia R.L. Webster Cummings School of Veterinary Medicine at Tufts University North Grafton, MA USA

LIST OF CONTRIBUTORS

Allan W. Wolkoff Division of Hepatology Marion Bessin Liver Research Center and Department of Anatomy and Structural Biology Albert Einstein College of Medicine Bronx, New York, NY USA Guorong Xu University of Medicine and Dentistry of New Jersey New Jersey Medical School NJ USA and VA Medical Center East Orange, NJ USA Hiroshi Yamaguchi Department of Pathology Sapporo Medical University School of Medicine Sapporo Japan Kenneth S. Zaret Cell and Developmental Biology Program Fox Chase Cancer Center Philadelphia, PA USA

Preface Predictions stated in previous editions of The Liver: Biology and Pathobiology (1982, 1988, 1994, and 1999) have consistently come to pass at a faster rate than had been anticipated. Major advances in genetics, immunology, virology, chemistry, biophysics, and structural, molecular, and cellular biology increasingly affect our understanding of liver function and disease. The near future promises additional developments resulting from the sequencing of the human genome, proteomics, small RNAs, and advances in combinatorial chemistry, micro-imaging, stem cell biology, and other areas of research. The challenge addressed by this book has not changed since the Preface to the First Edition was written in 1982: The amazing advances in fundamental biology that have occurred within the past two decades have brought hepatology and other disciplines into new, uncharted and exciting waters. The dramatic changes in biology will profoundly influence our ability to diagnose, treat, and prevent liver disease. How can a student of the liver and its disease maintain a link to these exciting advances? Most physicians lack the time to take post-graduate courses in basic biology; most basic researchers lack an understanding of liver physiology and disease. This book strives to bridge the ever-increasing gap between the amazing advances in basic biology and their application to liver structure, function, and disease.

A problem arises. How can a new edition remain reasonable in cost and size and still present essential background information, which has not changed since the last edition, as well as the panoply of major new contributions to our understanding of liver disease? In the Fourth Edition, we introduced a novel solution which has been expanded in the current Fifth Edition. The Fourth Edition emphasized exciting important new concepts and discoveries of the previous five years, and background information and references from the previous edition were provided on an open web site. Now, in collaboration with our new publisher, Wiley-Blackwell, we have provided 17 chapters from the

Fourth Edition on the web site GastroHep.com. These are freely available and we have cross-referenced the new text to these chapters. They are designated by [insert computer screen symbol] and web site chapter numbers W-1 through W-17 when cited in the text. Therefore, readers can obtain new information in the printed Fifth Edition and background information through GastroHep.com. The Fifth Edition includes 60 NEW chapters and 14 chapters are on the web site that present major progress as achieved in the laboratory and clinic. All other chapters either have been completely revised or appear on the web site. Harvey Alter, David Cohen and Allan Wolkoff have become Associate Editors. Previous editions included a section entitled “Horizons”, which presented advances of extraordinary nature in areas of potentially major importance to the liver. Virtually all of these fields have rapidly expanded and become topics for future chapters. Fifteen new “Horizons” are presented in this edition. One may safely predict that their impact on hepatology will also be considerable. As stated in the Preface to the Third and Fourth Editions: The amazing advance in science proceeds at an ever-increasing pace. The implications for students of liver disease are considerable. The authors and editors will have achieved our goals if the reader finds within this volume glimpses into the current state and future direction of our discipline and perspectives that lead to better understanding of liver function and disease.

Irwin M. Arias, Editor Harvey J. Alter, Associate Editor James L. Boyer, Associate Editor David E. Cohen, Associate Editor Nelson Fausto, Associate Editor David A. Shafritz, Associate Editor Allan W. Wolkoff, Associate Editor

Acknowledgments We thank the distinguished authors for their expertise, enthusiastic participation, and patience in responding to editorial suggestions. Appreciation is also acknowledged for the administrative assistance of Katrina Matthews

at the National Institutes of Health, and to the staff at Wiley-Blackwell, particularly Gill Whitley and Joan Marsh.

PART ONE : INTRODUCTION

1

Organizational Principles of the Liver Joe W. Grisham Department of Pathology and Laboratory Medicine, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA

PRINCIPLES OF LIVER ORGANIZATION The liver provides functions required to maintain homeostasis in the organism. To accomplish this the liver synthesizes numerous essential molecules of diverse sort; extracts and metabolizes a plethora of nutrients and xenobiotics brought into the body through the alimentary tract (and substances entering by other routes), as well as worn-out molecules and cells; stores, exports and/or excretes the metabolic products; and neutralizes numerous foreign antigens and microbes from the gut. These varied functions take place in a structurally complex, multicellular tissue with a unique angioarchitecture that has slowly evolved to its present form. Major features of liver structure are a functional tissue (parenchyma) composed of at least seven distinct types of cell—hepatocytes, cholangiocytes, sinusoidal endothelial cells, macrophages, lymphocytes of several different phenotypes, dendritic cells, and stellate cells—that conjointly possess the capacities to synthesize, metabolize and eliminate a wide range of complex molecules and to carry out immune fnctions, all arranged in a matrix that facilitates their cooperative interaction. This cellular matrix is perfused with blood at low pressure and flow rate through uniquely structured capillary-size blood vessels, which are supplied by two sources of blood: (i) venous blood that has already circulated through the gut, pancreas, and spleen, is reduced in oxygen and pressure, and is enriched

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

in nutrients and toxins absorbed from the gut and in viscerally generated hormones and growth factors; and (ii) arterial blood at systemic levels of oxygen, pressure, and composition. The most fundamental feature of liver organization is a unique vascular pattern in which afferent (supplying) and efferent (draining) blood vessels of all sizes interdigitate uniformly, always maximally separated by parenchymal tissue and connected almost exclusively by the smallest capillary-size vessels (the sinusoids). Afferent blood vessels branch to form up to 8–10 orders of diminishing size from their entrance at the liver hilum; terminal portal veins, which supply blood to sinusoids, arise from the smallest two or three orders of preterminal portal veins. Sinusoids are interposed between afferent terminal portal veins and small efferent hepatic (“central”) veins, which collect sinusoidal blood and merge to form larger hepatic veins. This vascular pattern provides a large volume of blood at a high flow rate through large vessels with high compliance and capacity to supply the sinusoids at a low flow rate and pressure. Total liver blood flow is large only because there are myriad sinusoids. Hemodynamic conditions resulting from this vascular pattern create “watersheds” or flow currents that segment the continuous mass of liver parenchyma into separate afferent and efferent vascular units at both macroscopic and microscopic levels. The liver parenchyma is not divided by connective tissue into identical capsule-delimited lobules, as is characteristic of all other glands, but is

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

4

THE LIVER: EMBRYOLOGY OF THE LIVER

segmented only by hemodynamic patterns of afferent and efferent blood flow (see below). The smallest hemodynamic segments of parenchyma perfused with blood by sinusoids supplied by terminal afferent vessels and drained by small efferent vessels (the afferent and efferent microvascular segments often termed “lobules,” acini, etc.) are the parenchymal units in which the metabolic activity of the liver parenchyma is focused (see below). The human liver contains millions of virtually identical microvascular segments. Large afferent and efferent vessels hemodynamically divide the parenchyma into at least eight macrovascular segments, which enable the surgical resection of large portions (segments) of the blood-filled, sponge-like liver through hemodynamic fissures between afferent and efferent blood flows (see below).

PHYLOGENESIS AND EVOLUTION OF THE LIVER The multiple functions of the mammalian liver are carried out by a combination of exocrine, endocrine, and paracrine/juxtacrine mechanisms involving the several types of cell listed above, as well as stromal cells (fibroblasts), and cells forming nerves and large blood vessels. The liver is the largest visceral organ in humans and other vertebrates, comprising generally 2–5% of the body weight in most species. It is a comparatively late evolutionary development, emerging more or less in concert with the vertebral column. Although the functions carried out by the various cellular components of the liver parenchyma are not limited to vertebrates, the aggregation of these different cells into the tissue of a single organ is unique. In invertebrates the liver’s functions are distributed among various cells and tissues dispersed throughout the body; glandular appendages to the gut (“midgut or digestive glands”) provide a few (but not all) of the metabolic functions of the vertebrate liver and pancreas [1]. A more liver-like gut appendage, associated with a vestigial portal circulation and some hepatocyte-like functions, occurs in the cephalochordate, amphioxus or lancelet (Branchiostoma sp.) [2, 3], a primitive chordate which occupies a proximate position in the evolution of vertebrates [4]. Development of a portal circulation may be the crucial event that enabled the formation of the liver in vertebrates [1]. The livers of mammals, birds, amphibians, and fish pass through similar developmental sequences, but vary structurally in adults [5]. In some cartilaginous fish, for example, the pancreatic and hepatic elements derived from the primitive foregut are incompletely separated; in primitive cartilaginous fish the biliary ducts regress in adults [6, 7]; and in bony fish there is a simpler ramification of blood vessels (and, consequently, vascular segments) and a different arrangement of hepatocytes and terminal bile ducts [8, 9].

EMBRYOLOGY OF THE LIVER The evolutionary steps that eventuated in the emergence of the mammalian liver containing multiple types of functioning cell are obscure. To the extent that the ontogenesis of the liver mirrors its phylogenesis, the embryonic development of the mammalian liver suggests the way in which the aggregation of multiple types of cell into the hepatic parenchyma may have evolved (Figure 1.1). Liver embryogenesis in mammals (see Chapter 2) begins in a few endodermal cells of the ventral foregut (the “liver bud”) located adjacent to the mesoderm of the septum transversum, which lies between the developing heart and the yolk sac and contains several types of mesenchymal cells and small blood vessels [10]. Interactions between epithelial cells of the foregut endoderm and mesenchymal cells of adjacent mesoderm direct the early development of the liver through the mediation of growth factors and cytokines [10, 11]. Hepatic commitment of endodermal cells of the ventral foregut is first signaled by the neoexpression of a group of genes for transcription factors and for liver-enriched or specific proteins, including α-fetoprotein and albumin [10]. Committed endodermal cells progressively invade the septum transversum, directed by competent endothelial cells in the latter [12], to form the first vestiges of the plate-like structure of the adult liver parenchyma, in which endodermal cells (hepatoblasts) that eventually become hepatocytes are separated by primordial sinusoids. Bipotent hepatoblasts associated with endothelial cells in the septum transversum differentiate into both hepatocytes and cholangiocytes of the adult liver [10, 11, 13]. Hepatoblasts of humans express cytokeratins (CK) 8, 18, and 19 [14], and most of them amplify expression of CK 8 and 18 and mature to form hepatocytes in conjunction with cessation of CK 19 expression [14]. The relatively few hepatoblasts that touch the mesenchyme of the primitive portal tracts form a double-layer of cells termed the “ductal plate,” acquire the expression of CK 7 and amplify expression of CK 19, while continuing to express CK 8 and 18 [15], gradually gaining the form and function of cholangiocytes. Early cholangiocytes in ductal plates remodel to form separate ducts, while intervening ductal plate cells regress. Larger ducts are incorporated into the developing portal tracts and the smallest ducts connect to bile canaliculi in hepatic plates. Development of cholangiocytes and bile ducts from hepatoblasts is regulated by cytokines produced by portal mesenchyme [10, 13]. With the early migration of primitive hematopoietic tissue from the yolk sac and the embryonic aortagonad-mesonephros, the developing liver becomes the main hematopoietic organ of the mammalian embryo [16]. Cytokines produced by hematopoietic tissue promote the further development of the liver parenchyma and vice versa [10, 17]. Hematopoiesis migrates from the mammalian liver to the bone marrow during later stages of embryogenesis, but several cells derived from

1: ORGANIZATIONAL PRINCIPLES OF THE LIVER

YOLK SAC/ AGM AREA

VENTRAL FOREGUT

BONE MARROW

Endodermal Cells Hematopoiesis

SINUSOIDAL ENDOTHELIUM MACROPHAGES IMMUNOCYTES STELLATE CELLS

HEPATOCYTES

HEPATOBLASTS

CARDIAC AREA

5

LIVER STEM CELLS (Canals of Hering) Ductal Plate Perivenous Mesenchyme

Venous Endothelium

SEPTUM TRANSVERSUM

CHOLANGIOCYTES

FETAL LIVER

POSTNATAL LIVER

Figure 1.1 Schematic depiction of some of the major events and cell–tissue interactions during the embryogenesis of the liver. Under influences (growth factors, etc.) from the developing heart (cardiac area) and from endogenous endothelial cells of the septum traversum, endordermal cells (the liver bud) migrate from the ventral foregut into the septum traversum, forming the primordial hepatoblasts. Hematopoietic tissue also migrates to the septum traversum from sites in the fetal yolk sac and the aortic-gonadal-mesonephros (AGM) region of the embryo. Hematopoietic tissue and hepatoblasts interact to further the development of both. Hepatoblasts that contact the mesenchyme around the emerging portal veins progressively change to form the ductal plates and, subsequently, cholangiocytes. A few hepatoblasts become liver stem cells. Late in the development of the fetal liver, in conjunction with the maturation of hepatocytes, hematopoietic tissue migrates from the liver to bone, but a few representatives of several hematopoietic cells—sinusoid endothelial cells, macrophages, immunocytes, and stellate cells—remain in the liver. Development of the liver is completed postnatally

hematopoietic tissue—including macrophages, immunocytes, some sinusoidal endothelial cells, and possibly stellate cells—become permanent residents of the liver [18–20]. Small vessels in the septum transversum are the source of endothelial cells required for emergence of endodermal cells from the foregut [10, 12], but some endothelial cells originate from angioblasts in hematopoietic tissue [21]. Stellate cells have been posited to originate from such widely disparate sources as gut endoderm, mesoderm of the septum transversum, the neural crest, and/or hematopoietic tissue [22], but recent studies exclude a neural crest origin [23] and support an origin from hematopoietic tissue [24–26]. In fact, after transplantation of either bone marrow ([27], references S3, S8, S64 therein) or liver ([27], references S67–S69 therein), all of the liver’s non-epithelial cells are partially replenished from hematopoietic tissue in mammals, but they can all also proliferate locally (see below). The pattern of emergence of the liver in fish, birds, and mammals is similar [4, 28, 29], but endothelial cells do not appear to direct the emergence of endodermal cells from the gut during development of the liver in zebrafish [29]. Furthermore, endothelial cells with scavenger activity are located in the gills and kidneys

of cartilaginous and bony fish, and not in the liver as in all terrestrial animals [30]; the location of scavenger endothelial cells in the liver reflects a late step in the evolution of the mammalian liver. Nevertheless, the general pattern of expression of transcription factors and genes involved in liver development is conserved in all of these species [10], suggesting a common transcriptional strategy for assembling the liver. When this strategy first emerged awaits further genetic analysis of gut appendages in chordate ancestors of vertebrates.

ANATOMY OF THE LIVER The liver of adult humans weighs from 1300 to 1700 g, depending on sex and body size. It is a continuous sponge-like parenchymal mass penetrated by tunnels (lacunae) that contain the interdigitating networks of afferent and efferent vessels [31]. The blood vessels and their investments of connective tissue provide the soft, spongy liver with its major structural support, or “skeleton.” Larger afferent vessels, portal veins, and hepatic arteries, are contained together in connective tissue—the portal tracts—which are

6

THE LIVER: LIVER HEMODYNAMICS

continuous with the mesenchymal components of the liver’s mesothelium-covered surface capsule (Glisson’s capsule). Portal tract connective tissue is progressively diminished as the portal veins decrease in size. In humans and many other mammals, the smallest (terminal) portal vein is almost bare of connective tissue. (In contrast, connective tissue accompanies the terminal portal veins of adult swine and a few other animals, giving the false appearance of capsule-encased lobules.) Portal tracts also contain bile ducts, lymphatic vessels, nerves, and varying populations of other types of cells, such as macrophages, immunocytes, stellate cells, and possibly hematopoietic stem cells ([32] and references therein). The collagenous investment of the efferent vessels is less robust and lacks large numbers of adventitious cells. The hepatic artery is distributed to the tissues of portal tracts, the liver capsule, and the walls of large vessels [33–36]. In portal tracts arterial branches form a capillary network (the peribiliary plexus) arborized around bile ducts [36, 37]. Efferent twigs from the peribiliary plexus empty into adjacent portal veins and sinusoids, forming an intrahepatic portal system [36, 37]. The portal vein supplies blood to the parenchymal mass only through its terminal branches [36–38]. Portal and arterial blood appear to be well mixed before entering sinusoids [38], but the direct supply of arterial blood to sinusoids by small branches of the hepatic artery is uncertain [33–37]. Glisson’s capsule, including portal tracts, contains extraparenchymal cells and tissues that make important contributions to liver function, including bile metabolism (cholangiocytes in bile ducts), vascular regulation (sympathetic nerves), pain perception (parasympathetic nerves), immune function (immunocytes), and lymph formation (lymph vessels). A large volume of lymph (up to half of all lymph) is produced in the liver, mostly in the ramifications of Glisson’s capsule [39]. Lymph vessels are not found in the parenchyma [40] and no satisfactory mechanism has been proposed that could separate the countercurrent flow of lymph in spaces of Disse and flow of blood in sinusoids with large fenestrations, although tracer studies suggest that lymph originates within the parenchyma [41]. In most mammalian species the liver is multilobed, the individual lobes reflecting the distribution of the major branches of afferent and efferent blood vessels. In contrast, the human liver parenchyma is fused into a continuous parenchymal mass with two major lobes, right and left, delineated only by being supplied and drained by separate first- and second-order branches of the portal and hepatic veins. Right and left lobes are topographically separated by the remnants of the embryonic umbilical vein (the falciform ligament), but this landmark does not locate the true anatomic division. Anatomically, the medial segment of the left lobe is located to the right side of the falciform ligament, centered on the anterior branches of the left portal vein. Interdigitation of first- and second-order branches of the portal and hepatic veins produces eight or more macrovascular parenchymal segments centered

on large portal veins and separated by large hepatic veins [42]. Hemodynamic watersheds or fissures separating afferent and efferent macrovascular segments permit the surgical resection of individual or adjacent segments.

LIVER HEMODYNAMICS The hepatic vasculature is characterized by high capacity, high compliance, and low resistance [43]. Blood vessels encompass about 22% of the liver’s mass/volume [44] and the liver contains about 12% of the total blood volume under physiological conditions [43], a sizeable fraction of which can be expelled by contraction of larger vessels by sympathetic nerve stimulation: the liver is a blood reservoir. The pressure of portal venous blood is reduced as the major afferent vessels dichotomize through the parenchyma, from about 130 mm of water in the extrahepatic portal vein to about 60 mm of water in the preterminal portal veins of the exteriorized liver of the anesthetized rat, amounting to about 60% of the total transhepatic pressure gradient [44]. A similar portal pressure gradient has been found in humans [43]. Blood flow through the liver amounts to about 1500–2000 ml minute−1 in adult humans, about 25% of the resting cardiac output [43]. About 25% of the total liver blood flow is derived from the hepatic artery at prevailing arterial pressure and oxygenation. The portal venous blood (about 75% of total liver blood flow) arrives at the liver partially depleted of oxygen and at a reduced pressure as a consequence of having already perfused the splanchnic viscera. In aggregate, sinusoids comprise about 60% of the liver’s vascular volume, or about 13% of the total liver mass/volume [44]. A significant decrement in blood pressure occurs in sinusoids (about 40% of the transhepatic pressure gradient), the pressure declining to about 25 mm of water in terminal hepatic veins of exteriorized liver of anesthetized rats [45]; the pressure gradient in the short sinusoids is especially steep. Blood pressure in the inferior vena cava approximates that in the terminal hepatic vein [43]. Consequently, although flow of blood through sinusoids faces little resistance, it is slow and somewhat intermittent and is assisted by negative pressure produced by respiratory expiration [43]. Possible mechanisms of regulating blood flow within sinusoids are controversial. Sinusoids appear to have limited contractile ability, possibly produced by contraction of encircling stellate cells (pericytes) [46, 47]. Studies of the exteriorized liver of rodents suggest that sinusoidal flow may be regulated at both inlet and outlet levels [46], but other similar studies have not detected sphincters at either point [45]. However, sinusoidal flow is strongly affected by postsinusoidal resistance [43]. Unlike capillaries elsewhere, liver sinusoids are composed of endothelial cells that are penetrated by holes (fenestrae) and lack a basal membrane [48], features that allow free egress of the fluid components and solutes of

1: ORGANIZATIONAL PRINCIPLES OF THE LIVER

the perfusing blood. For example, tagged albumin has access to a space in the liver that is about 48% larger than the sinusoidal volume, in contrast to other tissues in which capillary space and albumin space are nearly the same [44].

HEMODYNAMIC MICROSEGMENTATION OF THE LIVER PARENCHYMA Profiles of portal tracts and hepatic veins of various sizes are a prominent feature of liver histology [33–35]. Profiles of smaller branches of the afferent and efferent vessels (together with their stromal components) predominate in tissue sections taken from peripheral, subcapsular locations, whereas tissue sections taken from more proximal areas nearer to the hilum also contain larger vascular structures [35]. These vascular/stromal elements are contained in tunnels (lacunae) that penetrate the parenchymal mass [33]. While hepatic plates have been analogized as brick-like walls (muralia) of hepatocytes one cell (one brick) thick, this description somewhat oversimplifies the situation. Hepatic plates merge and branch frequently [41], and at branch points they are focally more than one cell thick [33, 49]. Furthermore, in livers undergoing growth or repair, hepatic plates are focally several cells thick at sites of hepatocyte proliferation before they are remodeled to a thinner structure [50, 51]. In histological sections of mammalian liver afferent and efferent vessels interdigitate regularly in an approximate ratio of 2–3 portal tract profiles for each profile of a hepatic vein, to form a pattern of cross-sections of portal tracts and hepatic veins separated by parenchyma [34, 35]. Most of the cross-sectioned portal tracts contain preterminal (penultimate) portal veins that represent the 7th–10th-order branches from the hilar portal vein in large mammals, such as humans. These small portal tracts and hepatic (central) veins penetrate the parenchyma in nearly parallel orientations about 0.5–1.0 mm apart. The terminal portal veins (septal or inlet venules) branch from preterminal portal veins at points on the circumference of the latter separated by about 120 radial degrees (triradial branching) and penetrate the parenchyma approximately perpendicular to and midway between two adjacent terminal hepatic veins [34, 35]. (Afferent liver blood vessels branch one order more than do efferent vessels.) The disposition of terminal portal veins in two dimensions yields a roughly hexagonal pattern, forming the edges of the more-or-less hexagonal “Kiernan lobule” (see below); the portal tracts that contain the parental preterminal portal veins are positioned at alternating corners of the hexagons. This description of the distribution pattern of preterminal and terminal portal veins and of terminal hepatic veins is, again, somewhat idealized; the distribution of terminal portal veins in two dimensions actually forms patterns not limited to

7

regular hexagons since not every terminal hepatic vein is bordered by exactly three portal tracts containing preterminal portal veins and not every terminal portal vein is of the same length. During their course through the parenchyma terminal portal (inlet) veins break up completely into sinusoids, which are oriented more or less perpendicularly to the veins (Figure 1.2). Because they are hardly larger than sinusoids, terminal portal veins are not conspicuous in humans and other mammals that lack a connective sheath around them. However, in adult swine their course through the parenchyma is clearly marked by connective tissue. Capillary-size sinusoids occupy the smallest and most numerous tunnels (lacunae) in the parenchymal mass [33]. In favorably oriented histological sections more or less parallel, longitudinal profiles of sinusoids alternate with hepatic plates [49]. In other orientations, sinusoids may appear in histological sections as circular cross-sections located in a slightly larger circular cross-section of a small tunnel in the parenchymal mass. A narrow cleft, termed the space of Disse, separates sinusoids from hepatocytes located in adjacent hepatic plates [48, 52]. At their proximal (portal venous) ends, sinusoids are narrow and somewhat tortuous, whereas their middle and distal (hepatic venous) portions are larger and straighter [37, 53, 54]. Sinusoids and hepatic plates are disposed radially around the draining hepatic veins and extend more or less directly to the supplying terminal portal veins [54]. Microvascular segments (often incorrectly termed “lobules”) of the liver parenchyma are delineated by this pattern of distribution of microvessels and by the resulting hemodynamic properties of blood flow through them (alternating “sources” and “sinks”), which separate the flow of blood to individual microsegments. In humans and other mammals that lack connective tissue along the terminal portal veins, the parenchyma is segmented only by the hemodynamic barrier (“vascular septum” or “watershed”) formed by currents of blood flowing from the terminal portal veins into the sinusoids. These currents effectively separate the flow of blood in adjacent microvascular segments. Concepts of the smallest structural and functional units of the liver have generated controversy for more than 400 years [55]. To a large extent this controversy has centered on efforts to determine the “true” lobular structure of the liver, when, in fact, the liver is a continuous parenchymal mass that lacks lobules, even in swine! Whether the correct equivalent of the “liver lobule” should be centered on the draining efferent vessels (as in the “Kiernan lobule”) [56] or on the supplying afferent vessels (as in the “Mall lobule”) [57] has consumed much argument [55]. The need for this controversy is eliminated when one realizes that the liver lacks true lobules; this focuses concepts and terminology on hemodynamically determined parenchymal microsegments. In hemodynamic terms the classic hexagonal “Kiernan lobule” [56] is the efferent microvascular segment,

8

THE LIVER: HEMODYNAMIC MICROSEGMENTATION OF THE LIVER PARENCHYMA

Sinusoids

Terminal portal veins

Terminal hepatic vein

Portal tract containing branches of portal vein, hepatic artery, and bile duct

Source Sink

Source

Figure 1.2 Schematic diagram of the distribution of the terminal branches of the portal and hepatic veins and of the flow of blood through sinusoids that connect them. Terminal hepatic veins and the portal veins of several orders from which terminal portal veins arise penetrate the parenchyma more or less in parallel. Terminal portal veins branch from the parental vessels at points around the circumference about 120◦ apart and at somewhat different points along the length of the latter, to form a quasi-hexagonal pattern of distribution. Terminal portal veins branch dichotomously along their entire circumferences and lengths to form sinusoids that merge with terminal hepatic veins, which, in turn, connect with larger (sublobular) hepatic veins. Not depicted here, terminal twigs of the hepatic artery and intrahepatic bile duct (the latter as Canals of Hering) accompany the terminal portal veins without a connective tissue sheath in most mammals, including humans. Peripheral sinusoids near terminal portal veins are tortuous, becoming straighter in the remainder of their course toward terminal hepatic veins. Sinusoids branch and merge freely throughout their lengths to form a vascular “sponge.” Plates of hepatocytes (not shown) fill the spaces between sinusoids. Arrows depict the prevailing directions of blood flow from terminal portal veins through sinusoids into terminal hepatic veins. The relatively high blood pressure near terminal portal veins forms a hemodynamic barrier that directs sinusoidal flow into the nearest terminal hepatic vein where the pressure is at its lowest (see text). The sinusoidal network between adjacent terminal portal veins is the locus of the hemodynamic barrier that delimits microsegments of parenchyma. The segmentation of the parenchymal mass into microscopic units is a hemodynamic phenomenon produced by the in-flow sources located at the junctions of terminal portal veins and sinusoids, and sinks at the junctions of sinusoids and terminal hepatic veins

being the smallest unit of parenchyma that is drained of blood by a single efferent (terminal hepatic or “central”) vein. The nature of the afferent microvascular segment, the smallest unit of parenchyma supplied with blood by an individual afferent (terminal portal) vein, has continued to be controversial, but recent studies have greatly calmed the controversy. The afferent microvascular segment, which is supplied with blood by a single terminal portal vein, was defined by Rappaport, who called this unit of parenchyma the “liver acinus” [34, 58]. More recent studies by Matsumoto and Kawakami [35, 59] and by Ekataksin and Wake [60] have refined the structure of the afferent microvascular segment of liver

parenchyma that is supplied with blood by a single terminal portal vein. In their terminologies, the approximate unit of parenchyma encompassed by the “Rappaport acinus” is called the “primary lobule” [38, 59] or the “hepatic microcirculatory subunit” [60]. Matsumoto et al. defined the basis of the “vascular septum,” caused by the flow of blood from adjacent terminal portal veins into intervening sinusoids [35], which produces a hemodynamic barrier that separates the flow of blood in adjacent microvascular segments. An afferent microvascular segment (acinus, primary lobule, microcirculatory subunit) overlaps into the two neighboring efferent microvascular segments (“hexagonal lobules”) that center on adjacent terminal

1: ORGANIZATIONAL PRINCIPLES OF THE LIVER

hepatic veins; and each efferent microvascular segment is composed of cone-shaped bits of parenchyma from six (on average) afferent microvascular segments. Ekataksin and Wake showed that the afferent microvascular segment is also the smallest unit of parenchyma drained of bile by terminal bile ducts [60], demonstrating that this hemodynamic segment is also the smallest excretory unit of parenchyma. Terminal bile ducts accompany terminal portal veins through the parenchyma, but the biliary epithelial cells in these tiny ducts are difficult to see in histological sections unless stained with an antibody to CK 19 [61]. Terminal bile ducts accompanying the terminal portal veins merge with hepatic plates to form the Canals of Hering, tiny tubules composed of both biliary epithelial cells and small hepatocytes [61].

THE CELLS OF THE LIVER PARENCHYMA Hepatocytes are commonly denoted as “parenchymal cells” and the other cells of the liver tissue matrix as “non-parenchymal cells.” This convention is somewhat artificial since hepatocytes alone are not competent to perform all essential hepatic functions, and the several types of cell in the liver tissue matrix function as an integrated community to carry out conjointly the multiplicity of hepatic functions. Functional integration of this cellular community is accomplished by several communication mechanisms, including signaling networks involving numerous cytokines and chemokines, and by direct transfer of small molecules through gap junctions [62]. Hepatocytes, responsible for most of the synthetic and many of the metabolic functions of the liver (see Chapters 3–25), are large polygonal cells (averaging about 25–30 µm in cross section and 5000–6000 µm3 in volume [63]). They are the most numerous cells in the liver parenchyma; the adult human liver probably contains about 10−11 hepatocytes, representing about 60% of all cells in the parenchymal matrix and composing about 80% of its mass/volume [52]. Hepatocytes are shaped as complex rhomboids with several distinct surfaces [34] that comprise functionally distinct domains (see Chapter 6). About 35% of the total hepatocyte surface faces sinusoids, and the area of this surface is greatly amplified by the folding of the plasma membrane to form innumerable microvilli that extend into the space of Disse [63]. About 50% of the total hepatocyte surface faces adjacent hepatocytes [63]. The plasma membrane of these intercellular surfaces is mostly flat except where it is infolded to form bile canaliculi, which comprise about 13% of the total hepatocyte surface [63], also amplified by microvilli. The flat intercellular surface membranes contain intercellular adhesion complexes (tight junctions, intermediate junctions, and desmosomes) that pin together the adjacent hepatocytes and form a permeability barrier between the perisinusoidal space of Disse and bile canaliculi. Bile

9

canaliculi form a belt-like extracellular space (about 1 µm in diameter) that is continuous along the lengths of hepatic plates, connecting at the portal ends with bile ducts. The flat intercellular hepatocyte surface also contains gap junctions that allow communication between adjacent hepatocytes by transfer of small molecules. Hepatocytes are polarized by molecular specializations of their various surface membranes in the forms of receptors, pumps, transport channels, and carrier proteins (see Chapters 6, 14, 21, 23 and 24). The canalicular membrane is modified for bile excretion, whereas the sinusoidal surface is equipped for extraction of a great variety of molecules from the blood and for the simultaneous secretion into the blood of other molecules that have been modified or synthesized by hepatocytes. As befits their numerous metabolic functions, hepatocytes contain a complex array of mitochondria (∼1700 per cell on average), peroxisomes (∼370 per cell), lysosomes (∼250 per cell), Golgi complexes (∼50 per cell), aggregates of rough and smooth endoplasmic reticulum (∼15% of cell volume), and numerous microtubules/microfilaments [63]. Cholangiocytes comprise much less than 1% of the total number of cells in the liver parenchyma, since most are located in bile ducts in portal tracts [64]. Only the smallest bile ducts penetrate the parenchymal mass in the company of terminal portal veins, where they connect with bile canaliculi in hepatic plates. The points of connection of ducts with hepatic plates are defined by tubular structures, the Canals of Hering, composed of both small cholangiocytes and small hepatocytes [61], which are the location of liver epithelial stem cells that can differentiate into both hepatocytes and cholangiocytes [65, 66]. Larger bile ducts contain cholangiocytes that rest on a basal membrane and vary in number and size in proportion to duct size [64]. Their luminal surface membranes are expanded by microvilli and contain stereocilia, which signal from bile flow/content to cell cytoplasm [67]. Although they contain fewer mitochondria and sparser endoplasmic reticulum than do hepatocytes, cholangiocytes in intrahepatic bile ducts, together with the network of capillaries that surrounds them (the peribiliary plexus), form a metabolic unit that modifies the composition of canalicular bile by altering the content of water and solutes [68]. Endothelial cells of sinusoids comprise about 3% of the parenchymal mass/volume [52] and probably number about 3 × 1010 in an adult human liver. The thin cytoplasm of these flattened cells is penetrated by holes (fenestrations), each about 150–170 nm in diameter, that form groups termed sieve plates [48, 69]. Unlike other capillaries, sinusoids lack a basal membrane, but they are surrounded by a complex mixture of molecules, including collagens I, III, IV, V, and VI, laminin, heparin sulfate and dermatan sulfate proteoglycans, fibronectin, and chondroitin sulfate [70]. The unique structure of liver endothelium and sinusoids enables the free escape of fluid components of blood, but the matrix molecules located in the space of Disse can bind some molecules ([71] and

10

THE LIVER: FUNCTIONAL AND STRUCTURAL HETEROGENEITY ALONG HEPATIC PLATES AND SINUSOIDS

references therein) and may retard the access of some solutes to hepatocyte surfaces. Sinusoidal endothelial cells are actively pinocytic and avidly clear effete proteins and colloids from the perfusing blood through several scavenger receptors, providing the main pathway for clearance of effete molecules from the circulation [69, 72]. (see Chapters 26–28). Macrophages (Kupffer cells) comprise about 2% of the parenchymal volume/mass [51] and perhaps number about 2 × 1010 /adult human liver. They are located within the lumens of sinusoids, most numerous in the portal regions, and are held in place by loose attachments to the sinusoidal endothelium [69]. Liver macrophages are avidly phagocytic through C3 and Fc receptors, clearing the sinusoidal blood of relatively large particulate materials including bacteria, effete cells (worn-out erythrocytes, dead or damaged hepatocytes, etc.) [69, 72]. Together with sinusoidal endothelial cells they form the organism’s major system for removing worn-out cells and proteins from perfusing blood. Activated macrophages produce a large number of chemokines and cytokines that have a fundamental role in the implementation of the liver’s acute phase reaction, coordinating the responses to injury of all of the parenchymal cells [73]. Immunocytes of the liver—T, NK, and NKT (and a few B) lymphocytes and dendritic cells—are components of a liver-centered immune system, largely segregated from the rest of the body’s immune system [19, 73]. The human liver is estimated to contain about 1010 lymphocytes of different phenotypes, located along sinusoids and in portal tracts [73]. The liver-centered immune system includes a major fraction of the body’s innate (native) immune capacity, as well as a small component of its acquired (adaptive) immune capacity [19, 73]. In addition to lymphocytes and dendritic cells, liver macrophages (Kupffer cells) and sinusoidal endothelial cells are also essential components of the liver-centered immune system [73]. Both liver macrophages and sinusoidal endothelial cells function as antigen presenters and both types of cell when activated secrete chemokines/cytokines that help stimulate the acute phase reaction and thymus-independent maturation of antigen-specific clones of T lymphocytes [19, 69, 73]. Liver immunity is involved in the removal/neutralization of numerous foreign antigens that reach the liver from the gut, including bacteria, particularly by the mechanisms of innate (native) immunity. In addition to the clearance of foreign antigens, the liver’s native immune system has a major regulatory role in the repair of the liver after cell injury and loss (see below). The small population of liver T lymphocytes, part of the body’s acquired (adaptive) immune capacity, are involved in virus elimination, clearance of activated T lymphocytes, and development of antigen tolerance [19, 73]. Stellate cells, located in the space of Disse outside of and partly encircling sinusoids (pericytes), comprise about 1.5% of the parenchymal volume/mass [52]. Stellate cells

are multifunctional (see Chapters 28–29); together with hepatocytes they participate in the metabolism of vitamin A and store this fat-soluble vitamin (as retinyl esters) in lipid inclusions; this gave origin to previous designations as “lipocytes, fat-storing, and/or vitamin A-storing cells” [74]. Stellate cells synthesize, secrete, and degrade components of the perisinusoidal extracellular matrix [74]. They respond to several cytokines by becoming actively migratory and by acquiring a myofibroblastic phenotype, signaled by their expression of desmin, α-smooth muscle actin, and several neuroendocrine proteins [74]. As myofibroblasts, stellate cells have a major role in the fibrotic responses of the liver to injury of various sorts.

FUNCTIONAL AND STRUCTURAL HETEROGENEITY ALONG HEPATIC PLATES AND SINUSOIDS Hepatic plates and adjacent sinusoids form associations that are structurally similar in all parts of the liver. Various liver cells show numerical, structural, and functional heterogeneities related to their location along the afferent–efferent axis of hepatic plates and sinusoids. Among the structural differences are ploidy variations in hepatocytes; in adult mammals hepatocytes located at the portal ends of hepatic plates are diploid, while cells of higher ploidy are located further downstream [75]. Gap junctions containing connexin 26 are more numerous on portal hepatocytes, whereas junctions containing connexin 32 are distributed on hepatocytes in all parts of hepatic plates [76]. In the portal regions, hepatic plates merge and branch frequently, accompanied by narrow, tortuous sinusoids [53, 54]. A larger fraction of stellate cells of the portal regions express desmin [74] and macrophages are larger and more numerous in this part of the parenchyma [73], whereas endothelial cell fenestrations are smaller and less numerous [48, 69]. Distribution of the molecular components of the perisinusoidal matrix also varies along the afferent–efferent axis of the space of Disse [50]. These variations in structure and cellular composition are associated with functional differences among hepatocytes located at different points along the afferent– efferent axis of plates–sinusoids. Rappaport divided the portal-hepatic (afferent–efferent) lengths of hepatic plates into three arbitrary zones (termed I, II, and III) and cited published studies documenting that hepatocytes located in these zones differ in their functional capabilities and susceptibilities to pathological damage [58]. Metabolic zonation of hepatocytes is strikingly exemplified by regional differences in carbohydrate metabolism (gluconeogenesis and glycogen storage by periportal hepatocytes; glycolysis by perihepatic vein hepatocytes), the enzymes

1: ORGANIZATIONAL PRINCIPLES OF THE LIVER

of ammonia metabolism (carbamoyl phosphate synthetase is concentrated in periportal vein hepatocytes; glutamine synthetase is confined to a single ring of hepatocytes bordering terminal hepatic veins). Recent research has shown that many liver functions are dispersed heterogeneously, with dispersed functions often acting as integrated parts of coordinate metabolic systems [77]. Zonation of liver functions is related to sinusoidal hemodynamics, which produces gradients in blood-borne substances available to hepatocytes and other cells of the parenchymal matrix [44]. Hepatocytes and other cells located at afferent and efferent ends of hepatic plates are subjected to different microenvironmental conditions. Certain molecules are largely extracted by the first hepatocytes that encounter the perfusing blood, lowering their concentration downstream. For example, oxygen levels in the blood at afferent and efferent ends of sinusoids differ greatly because oxygen is efficiently extracted by hepatocytes located at the afferent ends of hepatic plates, exposing downstream hepatocytes to relatively hypoxic conditions; the oxygen gradient alone can explain much of the heterogeneity of hepatocyte function related to position in plates [78]. Other molecules modified or produced by upstream hepatocytes are excreted into the sinusoidal blood and may be removed by hepatocytes located further downstream. The complex interplay of metabolite concentration in the perfusing blood, coupled with extraction, modification, secretion, re-extraction, and further modification, influence the metabolic events that occur in individual cells and define unequal parenchymal territories that produce zonal variations in different physiological capabilities and pathological susceptibilities [44]. Differential concentration of certain growth factors (termed “morphogens”) by hepatocytes in various parts of the hepatic plates may be especially important in producing functional heterogeneity [79]. A countervailing hypothesis holds that functional heterogeneity of hepatocytes results from age-dependent differentiation, producing cells with “hard-wired” differences in metabolic capabilities. This hypothesis is based on the concept of the “streaming liver” [80], which posits that hepatocytes are born at the afferent ends of hepatic plates and mature functionally as they migrate proximodistally along the length of the plates, becoming functionally variant as they mature, losing the capacity to proliferate and senescing/dying at the efferent ends of the plates [75, 81], analogous to the lifespan sequence of enterocytes migrating along the length of the small intestinal cryptvillus. Although this hypothesis is superficially attractive, many studies of the kinetics of hepatocyte proliferation provide overwhelming evidence that hepatic plates do not constitute tracks along which hepatocytes regularly migrate during their life cycles [82]. All hepatocytes (including polyploid cells) can proliferate, obviating the possibility that functional heterogeneity results from their age-dependent differentiation and ultimate senescence.

11

TURNOVER OF LIVER CELLS All liver parenchymal cells have a finite lifespan which can be shortened by physiological or pathological conditions that increase cell death and cell birth. Physiological turnover of hepatocytes occurs slowly with a lifespan of about 400 days in an adult steady-state hepatocyte population, about 0.025% of which typically will be engaged in DNA synthesis [83]. Cholangiocytes undergo turnover at similar rates. Analysis of the specific lifespans of the other cells of the liver parenchyma is complicated by the fact that their number can be augmented both by local proliferation of cells resident in the liver populations [84] and by repletion from the bone marrow. Repletion of these hematopoietically-derived cells from bone marrow is evident in recipients of bone marrow transplants, in which they are replaced by cells of the new bone marrow genotype [27], and in recipients of liver transplants, in which these types of liver cell are replaced with cells of the host genotype [27]. In contrast, hepatocytes are not generated from bone marrow cells in significant number under either circumstance [27].

LIVER PARENCHYMAL REPAIR Three distinct processes have evolved to produce the new hepatocytes needed to meet physiologically increased functional demand or to replace hepatocytes that are lost pathologically to trauma and toxicity. These processes center either on the temporary reactivation of cell cycle transit in fully differentiated, mitotically quiescent hepatocytes, with or without the coordinate proliferation and integration of other types of cells into a parenchymal matrix, and/or on the generation of entirely new hepatocyte lineages from adult liver stem cells (see Chapters 36, 38). The most direct and rapid of these parenchymal augmentation/replacement processes involves the upregulation of hepatocyte birth in the absence of a preceding increase in hepatocyte death, often associated with increased hepatic functional demand due to physiological need [85]. Hyperplasia of hepatocytes by this mechanism enlarges the parenchymal mass and increases hepatocyte functional capacity. This process is regulated by the binding of ligands to hepatocyte nuclear receptors, of which nearly 50 have been identified [86]. Nuclear receptors are transcription factors that, when bound to ligands, directly up-regulate the combination of genes required to drive hepatocytes through the cell cycle [85, 86]. Several ligands for nuclear receptors (termed “primary hepatocyte mitogens”), including adrenal corticoids, bile acids, sex steroids, thyroid hormone, peroxisome proliferators, and 9-cis-retinoic acid directly stimulate the proliferation of hepatocytes and increase liver mass after binding to nuclear receptors [85]. Although it would seem that new endothelial cells would be needed to support the additional

12

THE LIVER: REFERENCES

hepatocytes, no documentation of coordinate endothelial cell proliferation has been presented; it is possible that needed endothelial cells are derived from bone marrow. Next in process complexity and in rapidity of response is the replacement of cellularly diverse liver parenchyma by the sequential proliferation of all of the component cells (hepatocytes, cholangiocytes, endothelial cells, macrophages, stellate cells, and immunocytes), and the melding of the new cells into tissue that closely reiterates the functional units of the undamaged liver [87]. This process, which is capable of replacing up to 70% of the parenchymal mass in mammals, is often called “liver regeneration”—a misnomer since in mammals the part of the liver removed surgically does not “regenerate” in the same way as body parts in certain lower animal species. Instead, the liver remaining after resection is enlarged by the formation of new tissue microunits that accurately correct deficient liver functions. (In contrast to the process in mammals, liver repair after partial hepatectomy in fish most intensively involves cells at the resection margin [88, 89] and may culminate in the regrowth (regeneration) of the resected tissue [88].) The cell proliferation phase of this reparative process in mammals has been subjected to intensive kinetic and regulatory analyses ([87] and references therein). After tissue loss, residual hepatocytes are activated to proliferate within a few hours; hepatocyte proliferation begins at the portal ends of plates [83], and successive waves of hepatocyte proliferation ultimately involve virtually all residual hepatocytes [83, 90]. Hepatocyte replacement is followed sequentially by proliferation of sinusoidal endothelial cells and macrophages [83, 84], and the other cells of the parenchymal matrix. To the extent that it has been elucidated (see Chapters 36, 38), regulation of hepatocyte proliferation is effected by a complex mixture of cytokines and growth factors [87]. Most of the regulatory molecules are produced by various liver cells or released from storage sites within the liver [87], and many are components of the acute-phase reaction [91] and other elements of the liver’s native immune system [92, 93]. The less completely analyzed remodeling phase centrally involves endothelial cells and likely the other cells of the liver parenchyma. For example, proliferating hepatocytes initially form focal multicellular clumps [50, 51], which are cleaved into one-hepatocyte-wide plates by signaling from and separation by endothelial and stellate cells [50, 51]. The mechanism of liver tissue reformation at higher organizational levels (development of new afferent and efferent microvessels and establishment of new parenchymal microunits) is yet to be elucidated. Although known regulatory mechanisms drive the reparative process, the mechanism that “triggers” the onset of repair after loss of liver tissue is still obscure. Since the liver vasculature must accept the entire portal blood volume, it has long been suspected that the trigger may be the massive increase in portal blood flow per unit of residual mass that follows loss of liver tissue [94]. Increased

portal blood flow and pressure cause shear stress in sinusoids [94], which produces a burst of nitric oxide and prostaglandin production by sinusoidal endothelial cells, possibly providing the molecular trigger [95, 96]. Alternatively (or in concert), early activation of the nuclear receptor mechanism of hepatocyte proliferation may function as a trigger [97], and it is possible that multiple alterations in the physiological status of the liver remaining after tissue loss may converge to comprise a “mass action” trigger. The mechanistically most complex (and slowest) process of liver parenchymal repair involves the generation of new hepatocyte and biliary epithelial cell lineages from adult liver stem cells (located in and around the Canals of Hering) [61, 65, 66], and the subsequent neoformation of liver functional units in a process that resembles aspects of liver embryogenesis. In this process, progeny of liver stem cells proliferate to form populations of poorly differentiated cells (termed “oval cells” in rodents) that have developmental analogies to embryonic hepatoblasts. In rodents, activation of hepatocyte formation through the “oval cell” reaction appears to occur only when liver repair from mature hepatocytes is blocked [65]. However, parenchymal repair after surgical resection of liver in teleost fish includes both hepatocyte proliferation and the generation of new hepatocyte lineages from stem cells [89]. The “oval cell” reaction in rodents is also regulated by elements of hepatic immunomodulation centered on the acute-phase reaction [98]. Although the subject of intensive scrutiny recently, there is no substantial evidence that hematopoietic stem cells are a significant source for the generation of hepatocytes or biliary epithelial cells in either humans or experimental animals [27]. This situation contrasts with the replenishment from hematopoietic sources of other cells of the liver parenchyma [27].

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Embryonic Development of the Liver Roque Bort1 and Kenneth S. Zaret2 1 Unidad

de Hepatolog´ıa Experimental, CIBEREHD, Centro de Investigaci´on, Hospital Universitario La Fe, Valencia, Spain 2 Cell and Developmental Biology Program, Fox Chase Cancer Center, Philadelphia, PA, USA

INTRODUCTION The liver is one of the first organs to develop in the embryo and it rapidly becomes one of the largest organs in the fetus. The most essential function of the mammalian fetal liver is to provide a site for hematopoiesis. The early dependence of the fetus on its own blood cell supply makes embryonic liver growth and viability a sensitive phenotypic indicator for gene inactivation studies. From an experimental perspective, the large size and small number of cell types in the developing liver make it is easy to study, and new insights have emerged from recent work on genetically-modified mice, explants of embryonic tissues, and other vertebrate models such as zebra fish and frogs. Much is being learned about how gene function is orchestrated to control tissue morphogenesis, helping to establish liver development as a paradigm for the genesis of other gut-derived tissues. Understanding the mechanisms that govern liver development should also provide insight into future therapies for liver diseases. Examples of such applications include activating stem cells in the adult liver, replenishing diseased livers with cells from embryonic stem cells, trans-differentiating cells from different organs, and reconstituting proper liver morphology and function. This review will describe the early stages of liver development from the initial specification to hematopoietic cell invasion, which covers the competence of progenitor cells to become hepatocytes, the formation of the liver bud, and the early morphogenesis and differentiation The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

of the liver. The review will also present the key effectors of hepatocyte maturation and discuss the control of liver size and regeneration in the embryo and in the adult. The reader is referred to other reviews for summaries of the mid- and late-fetal stages of liver development [1–6].

ACQUISITION OF HEPATIC COMPETENCE WITHIN THE ENDODERM The liver, lung, pancreas, thyroid, and gastrointestinal tract are derived from the anterior-ventral definitive endoderm, the latter being one of the three germ layers that arise during gastrulation. Initially, the endoderm is an epithelial sheet that lines the ventral surface of the embryo. Infolding of the sheet at the anterior and posterior of the embryo generates the foregut and hindgut (Figure 2.1). When these morphogenetic movements reach the middle of the embryo, the gut tube closes off. During gut tube formation, different tissues are specified along the anterior–posterior and dorsal–ventral axes of the embryo, with the liver arising from the prospective ventral endoderm domain of the foregut (Figure 2.1). Are there “pre-patterns” or local domains of endoderm that are competent to differentiate into the liver? LeDouarin demonstrated, using heterotypic grafts of quail embryo segments into recipient chick embryos, that only the prospective anterior-ventral domain of endoderm had the capacity to

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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THE LIVER: FROM DEFINITIVE TO HEPATIC ENDODERM

Figure 2.1 Parasagittal diagram of a mammalian embryo at the time of hepatic specification. Relevant tissues and signaling molecules are indicated. The time of development corresponds to about 8.25 days gestation in the mouse and about three weeks in the human

develop into the liver [7, 8]. More recent studies with mouse embryos and more sensitive assays of gene expression revealed that while the prospective dorsal endoderm (Figure 2.1) does not normally activate liver genes or become liver, in a tissue explant assay the dorsal endoderm cells could initiate liver gene expression when isolated from the adjacent mesodermal tissues [9]. Only recently has the molecular mechanism underlying these transplantation studies been deciphered, using Xenopus (frog) embryos; similar mechanisms likely operate in mammalian embryos. It was found that due to the anterior expression of Wnt inhibitors such as Dkk, sFrp-1, or sFrp-5, the function of the Wntβ-catenin signaling pathway is repressed upon gastrulation in the anterior endoderm [10], but it is active posteriorly, where the Wnt inhibitors are absent. Wnt downregulation in the anterior endoderm was shown to be crucial for liver and pancreas specification; in fact, posterior endoderm that was forced to activate GSK-3β, a Wnt inhibitor, possessed hepatogenic competence [11]. Wnt signaling induces the transcription factor Vent in the endoderm, which in turn represses the homeobox gene Hex that is required in the endoderm for liver and pancreas development [12–17]. In summary, Wnt repression in the anterior endoderm is required for liver and pancreas specification, whereas active Wnt signaling in the posterior endoderm suppresses those fates. These findings provide a molecular basis to help explain the patterning of the endoderm. Another approach to understanding how the endoderm gains the competence to develop into liver is to identify the transcription factors that directly enable the process. Regulatory factors that are known to be important for liver differentiation are expressed in pre-hepatic endoderm, prior to hepatic specification. Such factors could operate by helping to open up chromatin structure for genes that need to be transcribed during liver differentiation [18]. Two transcription factors that are expressed in the pre-hepatic domain of ventral foregut endoderm, and later in the liver, are FoxA2 [19–22] and GATA4 [23–26]. The roles of these transcription factors can be discerned by

Figure 2.2 Transcriptional competence factors in the endoderm. DNA binding sites for GATA and FOXA transcription factors are occupied on the alb1 gene prior to alb1 expression or hepatic commitment. During hepatic specification, other transcription factors bind DNA at adjacent sites and the albumin gene becomes active

understanding how the factors function in a chromatin context. DNA binding by both FoxA and GATA factors is required for the activity of a transcriptional enhancer of alb1 , the gene encoding serum albumin [27, 28]; alb1 is one of the earliest genes to be activated in hepatic development [9, 29]. An analysis of the DNA binding sites for FoxA2 and GATA4 [30] showed that both sites are occupied on the alb1 enhancer in the endoderm (Figure 2.2), prior to alb1 transcriptional activation or hepatic commitment [9, 27]. Once hepatic specification has occurred, adjacent binding sites for a variety of other transcription factors become occupied at the enhancer and alb1 becomes active (Figure 2.2). Recently, genetic experiments have confirmed the essential role of FoxA and GATA factors in hepatogenesis. A transgenic mouse with a conditional ablation of both FoxA1 and FoxA2 in the foregut endoderm completely lacked the induction of hepatic markers of specification, such as alb1 , afp, or ttr [31]. Similarly, while the liver is specified in GATA6–/– or GATA4–/– mouse embryos (though subsequent development is blocked), a double knockout of both genes in zebra fish revealed a complete failure in liver induction [32]. These findings provide genetic evidence supporting the concept that FoxA and GATA factors serve as “pioneer” factors in the endoderm, being among the first to bind a target gene in development and endow competence for the gene to be activated, under the influence of cell-type inductive signals.

FROM DEFINITIVE TO HEPATIC ENDODERM Once the hepatogenic competence in the definitive endoderm is acquired, what causes the liver to be specified from the ventral foregut endoderm? The mesoderm secretes patterning signals that instruct the differentiation of

2: EMBRYONIC DEVELOPMENT OF THE LIVER

the underlying endoderm. Several lines of research conducted in chicken and mouse demonstrated that cardiac mesoderm helps instruct the underlying ventral foregut endoderm to become hepatic endoderm (Figure 2.1). In vitro culture of mouse embryonic explants together with the use of zebra fish and xenopus embryos have identified some of the relevant molecular signals. There are over 20 genes for fibroblast growth factors (FGFs) and 4 genes for FGF receptors [33, 34]. Each of the FGF receptors has different binding specificities for FGFs, and the existence of multiple spliced isoforms of the receptors provides further complexity to the receptor–ligand relationships. Cardiogenic mesoderm expresses FGF1, FGF2, FGF8, and FGF10 in the period prior to hepatic induction in the endoderm. In an embryonic tissue explant system, purified FGF1 or FGF2 can efficiently activate liver gene expression in ventral foregut explants where cardiogenic mesoderm has been removed, and an FGF antagonist can inhibit liver gene induction in foregut explants where cardiogenic mesoderm has been retained [35]. FGF binding induces tyrosine phosphorylation activity by the cytoplasmic domain of the receptors, resulting in activation of mitogen-activated protein kinase signaling pathways within cells [34, 36]. A combination of in vivo-genetic, whole-embryo culture and tissue explant approaches revealed that a transient activation of the MAPK pathway by FGF signaling in foregut endoderm explants is necessary for the initiation and stabilization of the hepatic program [37]. Although the PI3K/AKT pathway is activated in the endoderm shortly after the MAPK pathway, PI3K/AKT pathway activation appears not to be downstream of FGF signaling and the pathways do not cross-regulate in the hepatic endoderm [37]. BMP4 is a member of the TGFβ superfamily. BMP2, BMP4, BMP5, and BMP7 are highly expressed in the septum transversum mesenchyme (STM), the latter consisting of loose mesenchyme cells that surround the cardiac and ventral endoderm domains [38–42]. The BMP receptors BMPRIA, BMPRII, and ActRIIA are expressed in the endoderm [43, 44]. The mouse foregut explant system described above was used to reveal that BMP signaling from STM cells is also crucial for hepatic gene induction in the endoderm [45]. Thus, hepatic induction requires positive signals (FGF and BMP) from two different cell sources (cardiac mesoderm and STM), indicating the importance of combinatorial signaling. The general relevance of FGFs and BMPs for hepatic induction has been highlighted by genetic experiments in zebra fish [46]. The role of Wnt signaling during hepatic induction appears dynamic. The above step of FGF–BMP induction of the liver occurs when Wnt signaling is being suppressed in the foregut. But immediately after the induction of the hepatic program in the endoderm, Wnt signaling appears to be required for further outgrowth of the endoderm into a liver bud [11]. In zebra fish, expression of Wnt2b in the lateral plate mesoderm, acting through the β-catenin canonical pathway, appears essential for liver specification

19

in the endoderm and bud induction [47]. Differences in the development of the liver in fish vs amniotes may explain an earlier positive role for Wnt signaling in hepatic induction [5].

FROM HEPATIC ENDODERM TO LIVER BUD The progress from hepatic endoderm to liver bud has been divided in three morphogenetic stages [13]: stage I, the formation of a thickened, columnar hepatic epithelium; stage II, the formation of a pseudo-stratified epithelium (Figure 2.3c); and stage III, laminin breakdown and hepatic cell migration from the epithelium into the STM. Hepatoblast migration is accompanied by major remodeling of the extracellular matrix surrounding the hepatic cells [48]. During this period, the entire ventral foregut domain extends toward the midgut, bringing the liver region with it (Figure 2.3a,b). The mass of cells emerging from the endodermal epithelium and concentrating in the septum transversum is referred to as the liver bud, and the cells within the liver bud are referred to as hepatoblasts. Hepatoblasts will later differentiate into hepatocytes and cholangiocytes (biliary cells); see below. At stage I of liver bud formation in mammals, endothelial cells, not yet assembled into a vasculature, are adjacent to the hepatoblast epithelium [49]. Genetic depletion of endothelial cells at this stage has led to the discovery that endothelial cells are an essential stimulus to further liver bud development: the absence of endothelial cells halted liver bud development at stage II, with hepatoblasts remaining within the limits of the basal epithelial membrane. A similar organogenic stimulatory role has been attributed to endothelial cells in the developing pancreas [50]. In the pancreas, vascular endothelial growth factor A (VEGF-A) secreted by beta cells attracts endothelial cells to the developing islet [51]. Endothelial cells, in turn, form a basal membrane rich in laminins, collagen IV, and fibronectin required for the correct function of β-cells. The nature of the signals originating from the endothelial cells that trigger morphogenesis and cell differentiation in the emerging liver bud is not known. As already stated, the formation of the liver bud requires the degradation of the basal membrane to allow migration of hepatocytes into the STM. One of the genes controlling this step is Prox1 [48]. The Prox1–/– mouse embryonic hepatic endoderm is unable to degrade the basal epithelial membrane, resulting in the clustering of hepatoblasts into a smaller liver. It is not known whether this activity is caused by a slow degradation or production of excess basal membrane components by the embryonic hepatoblasts. As noted above, the homeodomain factor Hex is crucial for liver development [16] and plays an essential role in transformation from stage I to stage II [13]. Hex seems to control different aspects of liver bud formation, including

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THE LIVER: BEYOND THE LIVER BUD: CHOLANGIOCYTE DIFFERENTIATION

(a)

(b)

(c)

Figure 2.3 The beginning of liver development. (a) Mouse embryo, nine days gestation. Tail is removed; arrow indicates plane of section in (b). (b) 100× magnification of transverse section. Note the thickening of the endodermal epithelium of the gut tube at the region of the liver bud. Boxed area is magnified to 400× in (c). Arrows depict the liver bud and other landmarks of the embryo. Photographs courtesy of J. Rossi

the adequate proliferation of hepatic endoderm cells, the formation of a pseudostratified epithelium, and the stable maintenance of the hepatic cell type [12–14]. How Hex exerts its role is not known, but it has been suggested that it represses sonic hedgehog signaling within the ventral foregut endoderm, contributing to the exclusion of the intestinal fate [13].

BEYOND THE LIVER BUD: CHOLANGIOCYTE DIFFERENTIATION The complex signals that cause the emergence of the liver bud are followed closely by distinct signals required to grow the bud into the liver organ. The hepatoblasts differentiate into hepatocytes and cholangiocytes at about embryonic day of gestation 13.5 (E13.5) in the mouse and seven weeks in humans [52–55]. Cells that are slightly caudal to the liver bud, or within the caudal portion of the liver bud itself, give rise to cells of the gall bladder [55], while cells located in the cranial portion of the liver will give rise to duct cells (intrahepatic,

hepatic, cystic, and common). Similar studies in the spf ash mouse suggested that there are three lineages, hepatoblasts giving rise to cholangiocytes (biliary cells), hepatoblasts giving rise to hepatocytes, and hepatoblasts giving rise to both [56]; but genetic lineage marking studies will be needed to confirm this complexity. The molecular effectors involved in the fate decisions involve the Onecut transcription factors OC1 (HNF6) and OC2. Double homozygous OC1/2 mutant embryos present a disturbance of the TGFβ gradient across the axis of the portal vein to the hepatic parenchyma that causes the differentiation of hepatoblasts to hybrid cells with characteristics of both hepatocytes and cholangiocytes [57] (Figure 2.4). How OC factors control the TGFβ gradient in the developing liver is currently unknown. Together with TGFβ, notch signaling is required for normal biliary tract morphogenesis. Alagille syndrome, a developmental disorder characterized by a paucity of intrahepatic bile ducts (IHBDs), is caused predominantly by mutations in the Jagged1 (JAG1) gene, which encodes a ligand for notch family receptors [58]. Recent studies using notch loss-of-function mice and zebra fish models have indicated that notch regulates bile duct paucity rather than cell fate specification [59–61].

2: EMBRYONIC DEVELOPMENT OF THE LIVER

21

Figure 2.4 Schema of steps of liver development, and relevant signals and transcription factors that help mediate the steps. See text for details

ROLE OF MESENCHYMAL CELLS IN HEPATOCYTE DIFFERENTIATION Hepatocyte differentiation and liver morphogenesis are dependent upon the cellular microenvironment. During early stages of liver bud development, as discussed above, the microenvironment is provided by endothelial and mesenchymal cells in the STM, and, after E10.5 in the mouse, when liver becomes a hematopoietic organ, hematopoietic stem cells (HSCs) also contribute. The relevance of these mesenchymal cell types is highlighted by genetic loss-of-function studies in mice, where genes expressed in mesenchymal cells surrounding the nascent liver but not in hepatoblasts are found to be essential for correct liver formation. One of these cases, Lhx2 , is a LIM-homeobox gene expressed in the STM and mesenchymal components in the adult liver (presumably stellate cells). Lhx2 –/– livers display a disrupted cellular organization with increased deposition of extracellular matrix (ECM) and an altered gene expression pattern of the early hepatocytes [62]. Besides the early role of individual endothelial cells in promoting liver bud growth, blood vessels develop de novo within the liver bud (Figure 2.3b), forming a capillary bed that becomes interspersed within the expanding hepatoblast population [63]. These transitions establish the liver’s sinusoidal architecture, which is critical for organ function and sets the stage for the fetal liver to support hematopoiesis. Hematopoietic cells migrate to the early liver first from the yolk sac [64] and later from the aorta-gonad-mesonephros region [65]. Proper intermediate filament expression is critical for blood-cell homing, as erythrocytes accumulate excessively in the fetal liver of keratin 8 mutants [66]. Embryos deficient in the heavy metal-responsive transcription factor MTF1 exhibit reduced cytokeratin expression as well as enlarged sinusoids and dissociated epithelial cells, but not anemia [67]. The primary defect in MTF1-deficient embryos appears to be in a failure to control metal homeostasis and the oxidation-reduction state in hepatocytes at mid-gestation.

Both hematopoietic and endothelial cells provide differentiating signals to the hepatoblasts, because heterogeneity in hepatic gene expression has been found to be related to the vascular architecture of the fetal liver [68]. More specifically, mouse gene inactivation studies have shown that oncostatin M signaling from hematopoietic cells to nascent hepatocytes is critical for liver growth [69]. Impaired hematopoietic cell proliferation in c-myb mutant embryos [70], or impaired erythrocytic cell proliferation in retinoblastoma gene (Rb) mutant embryos [71, 72], also result in impaired liver growth. The failure of hematopoietic cells to migrate to the liver in β1 integrin-null embryos is therefore also thought to contribute to the liver developmental defect in the β1 mutants [73, 74]. As the fetal liver matures and grows, a capsule forms around it. N-myc gene expression in the liver capsule and jumonji gene expression in the hepatic stromal cells help promote growth during mid-gestation [75–77]. Although many of the genes that affect hepatoblast proliferation are probably most important during the initial transition from the liver bud to the organ stage, inactivation of these genes usually manifests itself as a hematopoietic defect well after the organ is formed. In these cases, a liver capsule develops and hematopoietic cells migrate to the liver, but the paucity of hepatoblasts leads to a defect in the hematopoietic environment and, consequently, embryonic lethality. Furthermore, mutations of certain liver regulatory factors yield a fetal liver growth defect due to apoptosis of the hepatoblasts. These proteins include c-jun [78, 79], IKK2 [80], RelA [81], and XBP1 [82].

EMBRYOLOGIC CONTROL OF LIVER REGENERATION The liver is among the few internal organs that can rapidly regenerate after removal of tissue in the adult. Recent studies indicate that the regenerative capacity of the liver is attained as early as the hepatoblast stage in embryos. Tissue complementation experiments, where Hex–/– mouse embryonic stem cells were injected into Hex+/+ blastocyts, discovered that at the onset of liver

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THE LIVER: FUTURE AND PERSPECTIVES

morphogenesis (E9.0), wild type hepatic progenitor cells can increase their proliferation rate to compensate for the failure of Hex–/– cells in the liver bud to survive [13]. In an independent study, when two-thirds of liver progenitors are genetically ablated between E9.5 and E13.5, the remaining cells are still able to engage in a compensatory growth, generating a normal-sized fetal liver within four days [83]. The pancreas, also an endoderm-derived organ, is unable to grow in this fashion, resulting in major losses of pancreatic mass after partial ablation of its embryonic progenitors [83]. It seems that liver, but not pancreas, has a full regenerating potential at nearly all developmental stages, and while the growth limit of the liver is set by the organ size, in the case of the pancreas it is set by the cell division number. It is noteworthy that even when the pancreas does not recover its original size after 50% pancreatectomy, it does recover nearly 100% of its beta cell mass after four weeks [84], suggesting different regenerating capabilities between the endocrine and exocrine compartments. Another interesting aspect is the difference between humans and animals, since recent experiments pointed out that humans, unlike rodents, do not have a significant increase in beta cell proliferation after partial pancreatectomy [85]. Identifying the genetic programs that cause the marked difference in hepatic and pancreatic regeneration at the embryonic stage, when the cells are otherwise quite immature, could be a convenient way to reveal the basis for adult liver regeneration.

HEPATOCYTE DIFFERENTIATION Hepatocyte differentiation spans from liver specification in ventral foregut endoderm until the postnatal maturation of hepatocytes (Figure 2.4). Downstream of signaling molecules that induce liver differentiation are the transcription factors that execute the liver program, including HNF1, HNF4, HNF6, FoxA, and C/EBP [4–6]. These liver-specific genes act, in a cross-regulatory fashion, on each other’s promoters. HNF4 is essential for neither liver specification nor early liver morphogenesis [86]. Extensive RT-PCR analyses revealed that the expression of the primary liver-enriched transcription factors was unchanged in HNF4–/– livers at E12, except for PXR and HNF1α, direct target genes of HNF4 [86]. However, a large number of tissue-specific genes involved in the maturation of hepatoblasts to hepatocytes failed to be induced in HNF4e–/– embryos. The extensive effects were explained by a genome-wide chromatin analysis, whereby the location of HNF4 protein bound to promoter sequences of nearly 10 000 genes was assessed simultaneously, using microarray technology [87]. Such analysis was also performed for HNF6, HNF1a [87]. HNF1a was bound to 222 target genes in human hepatocytes, or 1.6% of the genes on the array; HNF6 bound 227 genes, or 1.7%; and HNF4a bound to 1575 gene promoters, a

striking 12% on the array. The genes bound by HNF4a corresponded to ∼42% of the genes bound by RNA polymerase II. That is, nearly half of the active genes tested in the liver are bound by HNF4α. In summary, HNF4 controls the expression of many hepatic genes directly, rather than indirectly by controlling the expression of other transcription factors. Subsequent studies found that HNF4 regulates genes involved in cell junction assembly and adhesion in the developing liver [88], consequently promoting epithelial maturation of the liver parenchyma [89]. The limited apparent role of relevant transcription factors like HNF1α in liver development, based on gene inactivation studies in animals, contrasts sharply with ectopic and overexpression studies in cultured hepatic cell lines, which suggest that HNF1α is critical for the expression of a wide variety of liver-specific genes [90]. Such distinctions indicate how strongly the mechanisms of gene regulation are influenced by whole-animal physiology, beyond simple notions of gene redundancy, e.g. [91, 92]. Indeed, for future studies, identification of target genes is perhaps best guided by phenotypic expression changes in gene inactivation studies. For example, gene expression array analysis of embryos that were homozygous mutants for the basic leucine zipper (bZIP) transcription factor XBP1 revealed hepatic defects in α1-antitrypsin expression, and the XBP1 protein was then found to bind and activate the α1-antitrypsin gene promoter [82]. While many other liver enriched transcription factors are expressed early in liver development, their inactivation does not yield an embryonic defect [93]. Compound gene inactivation studies need to be performed in order to uncover potential redundancies. Ultimately, it is crucial to determine how inductive signaling pathways converge on regulatory transcription factor genes to coordinately promote early liver differentiation and morphogenesis.

FUTURE AND PERSPECTIVES This review has highlighted ways that our understanding of embryonic liver development has expanded greatly in the past 10 years or so. The regulatory molecules that control liver development are now being used successfully to program hepatic cells from embryonic stem cells and other sources [94]. Given the large number of proteins that were discovered in studies of adult livers and yet give embryonic liver phenotypes when deleted in mice, there is high confidence that many of these proteins’ functions in the adult are a recapitulation of activities in the embryo. Thus, continued investigation of embryonic liver development is certain to be instructive about the function, regeneration, and repair of the adult liver, and seems likely to provide new sources of cells and molecules to combat liver disease. Further refinements in the ability to inactivate genes in the early liver and better methods of embryo tissue culture are needed to advance the analysis of liver development. Such advances may come from adaptations

2: EMBRYONIC DEVELOPMENT OF THE LIVER

of methodology for other endoderm-derived organ systems [95]. In addition, relevant new genes will emerge from studies of model organisms where genetic screens can be coupled with knowledge of genomic sequences and interrelationships of protein function. Considering that our mechanistic understanding of liver development has emerged so recently, the prospects are bright for much deeper knowledge and new applications for liver therapies in the future.

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ACKNOWLEDGMENTS Thanks to Eileen Pytko for help in preparing the manuscript. R.B. is supported by a grant from the Spanish Ministry of Science (SAF-64414). CIBEREHD is funded by the Instituto de Salud Carlos III, Spain. K.Z.’s research on liver development is supported by a grant from the NIH (GM36477).

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44. Roelen, B.A., Goumans, M.J., van Rooijen, M.A. et al. (1997) Differential expression of BMP receptors in early mouse development. Int J Dev Biol , 41, 541–49. 45. Rossi, J.M., Dunn, N.R., Hogan, B.L.M. et al. (2001) Distinct mesodermal signals, including BMP’s from the septum transversum mesenchyme, are required in combination for hepatogenesis from the endoderm. Genes Dev , 15, 1998–2009. 46. Shin, D., Shin, C.H., Tucker, J. et al. (2007) Bmp and Fgf signaling are essential for liver specification in zebrafish. Development , 134, 2041–50. 47. Ober, E.A., Verkade, H., Field, H.A. et al. (2006) Mesodermal Wnt2b signalling positively regulates liver specification. Nature, 442, 688–91. 48. Sosa-Pineda, B., Wigle, J.T. and Oliver, G. (2000) Hepatocyte migration during liver development requires Prox1. Nat Genet , 25, 254–55. 49. Matsumoto, K., Yoshitomi, H., Rossant, J. et al. (2001) Liver organogenesis promoted by endothelial cells prior to vascular function. Science, 294, 559–63. 50. Lammert, E., Cleaver, O. and Melton, D. (2001) Induction of pancreatic differentiation by signals from blood vessels. Science, 294, 564–67. 51. Nikolova, G., Jabs, N., Konstantinova, I. et al. (2006) The vascular basement membrane: a niche for insulin gene expression and Beta cell proliferation. Dev Cell , 10, 397–405. 52. Germain, L., Blouin, M.J. and Marceau, N. (1988) Biliary epithelial and hepatocytic cell lineage relationships in embryonic rat liver as determined by the differential expression of cytokeratins, α-fetoprotein, albumin, and cell surface-exposed components. Cancer Res, 48, 4909–18. 53. Shiojiri, N. (1981) Enzymo- and immunocytochemical analyses of the differentiation of liver cells in the prenatal mouse. J Embryol Exp Morphol, 62, 139–52. 54. Shiojiri, N. (1984) Analysis of differentiation of hepatocytes and bile duct cells in developing mouse liver by albumin immunofluorescence. Dev Growth Differ, 26, 555–61. 55. Shiojiri, N. (1997) Development and differentiation of bile ducts in the mammalian liver. Microsc Res Tech, 39, 328–35. 56. Shiojiri, N., Inujima, S., Ishikawa, K. et al. (2001) Cell lineage analysis during liver development using the spf(ash)-heterozygous mouse. Lab Invest , 81, 17–25. 57. Clotman, F., Jacquemin, P., Plumb-Rudewiez, N. et al. (2005) Control of liver cell fate decision by a gradient of TGF beta signaling modulated by Onecut transcription factors. Genes Dev , 19, 1849–54. 58. Oda, T., Elkahloun, A.G., Pike, B.L. et al. (1997) Mutations in the human Jagged1 gene are responsible for Alagille syndrome. Nat Genet , 16, 235–42. 59. Kodama, Y., Hijikata, M., Kageyama, R. et al. (2004) The role of notch signaling in the development of intrahepatic bile ducts. Gastroenterology, 127, 1775–86. 60. Lorent, K., Yeo, S.Y., Oda, T. et al. (2004) Inhibition of Jagged-mediated Notch signaling disrupts zebrafish biliary development and generates multi-organ defects

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79. Hilberg, F., Aguzzi, A., Howells, N. et al. (1993) c-Jun is essential for normal mouse development and hepatogenesis. Nature, 365, 179–81. 80. Li, Q., Antwerp, D.V., Mercurio, F. et al. (1999) Severe liver degeneration in mice lacking the lκB kinase 2 gene. Science, 284, 321–25. 81. Beg, A.A., Sha, W.C., Bronson, R.T. et al. (1995) Embryonic lethality and liver degeneration in mice lacking the RelA component of NF-KB. Nature, 376, 167–70. 82. Reimold, A.M., Etkin, A., Clauss, I. et al. (2000) An essential role in liver development for transcription factor XBP-1. Genes Dev , 14, 152–57. 83. Stanger, B.Z., Tanaka, A.J. and Melton, D.A. (2007) Organ size is limited by the number of embryonic progenitor cells in the pancreas but not the liver. Nature, 445, 886–91. 84. Lee, C.S., De Leon, D.D., Kaestner, K.H. et al. (2006) Regeneration of pancreatic islets after partial pancreatectomy in mice does not involve the reactivation of neurogenin-3. Diabetes, 55, 269–72. 85. Menge, B.A., Tannapfel, A., Belyaev, O. et al. (2008) Partial pancreatectomy in adult humans does not provoke beta-cell regeneration. Diabetes, 57, 142–49. 86. Li, J., Ning, G. and Duncan, S.A. (2000) Mammalian hepatocyte differentiation requires the transcription factor HNF-4alpha. Genes Dev , 14, 464–74. 87. Odom, D.T., Zizlsperger, N., Gordon, D.B. et al. (2004) Control of pancreas and liver gene expression by HNF transcription factors. Science, 303, 1378–81. 88. Battle, M.A., Konopka, G., Parviz, F. et al. (2006) Hepatocyte nuclear factor 4alpha orchestrates expression of cell adhesion proteins during the epithelial transformation of the developing liver. Proc Natl Acad Sci USA, 103, 8419–24. 89. Parviz, F., Matullo, C., Garrison, W.D. et al. (2003) Hepatocyte nuclear factor 4alpha controls the development of a hepatic epithelium and liver morphogenesis. Nat Genet , 34, 292–96. 90. Tronche, F., Bach, I., Chouard, T. et al. (1994) Hepatocyte nuclear factor 1(HNF1) and liver gene expression, in Liver Gene Expression (eds F. Tronche and M. Yaniv), R.G. Landes Company, pp. 155–82. 91. Barbacci, E., Reber, M., Ott, M.O. et al. (1999) Variant hepatocyte nuclear factor 1 is required for visceral endoderm specification. Development , 126, 4795– 805. 92. Cereghini, S., Ott, M.-O., Power, S. et al. (1992) Expression patterns of vHNF1 and HNF1 homeoproteins in early postimplantation embryos suggest distinct and sequential developmental roles. Development , 116, 783–97. 93. Cereghini, S. (1996) Liver-enriched transcription factors and hepatocyte differentiation. FASEB J , 10, 267–82. 94. Gouon-Evans, V., Boussemart, L., Gadue, P. et al. (2006) BMP-4 is required for hepatic specification of mouse embryonic stem cell-derived definitive endoderm. Nat Biotechnol , 24, 1402–11. 95. Wells, J.M. and Melton, D.A. (1999) Vertebrate endoderm development. Annu Rev Cell Dev Biol , 15, 393–410.

PART TWO : THE CELLS

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Microtubules, Actin Filaments and Motor-mediated Vesicular Transport Ronald R. Marchelletta and Sarah F. Hamm-Alvarez Department of Pharmacology and Pharmaceutical Sciences, USC School of Pharmacy, Los Angeles, CA, USA

INTRODUCTION Microtubules, actin filaments, and structural filaments called intermediate filaments have long been observed in non-muscle cells in classical ultrastructural studies [1, 2]. Since these filaments appeared to function primarily in the maintenance of cellular structure in a wide variety of eukaryotic cell types, they were originally all thought of as the “cell’s skeleton,” hence, they were collectively labeled the cytoskeleton. Over the past several decades, it has begun to emerge that these structures are not only scaffolds that maintain the structural integrity of the cell, but are also dynamic structures that mediate vital cellular functions such as secretion, endocytosis, and transcytosis. In particular, studies in the intracellular role of microtubules, actin filaments, and their associated proteins called motor proteins have led to improved understanding of many events in vesicular transport. In this chapter, the structure and the integral role of microtubules, actin filaments and their associated motor proteins in membrane trafficking will be examined.

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

MICROTUBULES AND ACTIN FILAMENTS: HISTORICAL CONTEXT As early as 1954, electron microscopy revealed rod-like structures in rat sperm cell flagella [3]. These rod-like structures appeared as “filaments.” The filamentous structures were hypothesized to have a role in the beating and maintenance of the structural integrity of sperm tails and epithelial cilia [3, 4]. Advancing electron microscopy technology improved the resolution of these rod-like structures. These structures appeared to consist of multiple subunits [5]. Along with rapidly improving electron microscopes, improved techniques in sample fixation enabled direct observation of the tubular architecture of these fibrils [6]. Work done by Ledbetter and Porter enabled them to observe, in their planar thin sections, 13 subunits arranged in a ring. They extrapolated that these multiple subunits were arranged as a long tube consisting of 13 interacting chains or

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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THE LIVER: MICROTUBULE COMPOSITION AND POLYMERIZATION DYNAMICS

protofilaments organized around a hollow center [6, 7]. Because of this tube-like appearance, Ledbetter and Porter named these structures microtubules. In addition, they found that microtubules were conserved across a wide range of eukaryotic cells, from plants to vertebrates. One year later, Gary Borisy and Edwin Taylor were able to find a colchicine-binding protein [8, 9]. They observed that when cells were exposed to colchicine, the microtubules disappeared, which led them to conclude that this colchicine-binding protein was a microtubule subunit. Soon after this discovery, the name of “tubulin” was applied to this now-characterized subunit of microtubules [10]. Later, Borisy and Klugman revealed that tubulin assembly into microtubules is dependent on the presence of guanosine triphosphate (GTP), inorganic salts and temperature [11]. Actin filaments were first isolated from rabbit muscle in 1942 [12]. Soon after the isolation of filamentous actin, it became apparent that filamentous actin was in fact a polymer made up of subunits of globular actin monomers [13–15]. To distinguish polymeric from monomeric actin, filamentous actin was given the name of F-actin and its monomer, globular actin, was given the name G-actin [15, 16]. It was discovered that the polymerization of F-actin was dependent ATP, and inorganic salts. Actin in non-muscle cells was discovered biochemically in Physarum polycephalum slime mold [17]. Two years later, using electron microscopy, Abercrombie, Heaysman, and Pegrum were the first to observe unambiguously the presence of non-muscle actin filaments in fibroblasts [18]. While the function of non-muscle actin in the maintenance of cellular structure was apparent, almost immediately non-muscle actin was also implicated in the ruffling of lamellipodia and in the motility of fibroblast cells [18, 19]. Soon after electron microscopic visualization, actin filaments were observed with fluorescence in non-muscle cells at the light microscope level [20].

MICROTUBULE COMPOSITION AND POLYMERIZATION DYNAMICS The observation that microtubules disassemble in the presence of colchicine gave scientists the opportunity to investigate the role of microtubules in cell shape and function. When primary mesenchymal cells from Arbacia punctulata were exposed to colchicine, the axopodia disassembled and cell shape was disrupted [21, 22]. Curiously, this observation of disassembly of microtubules could be replicated with incubation of Actinosphaerium nucleofilum at 4 ◦ C [23]. Microtubules, as well as cell shape, were therefore shown to be temperature- and colchicine-sensitive. This was the first direct evidence that microtubules were involved in cell shape. Not only was cell shape disrupted when cells were exposed to colchicine or 4 ◦ C,

but mitosis also was inhibited because the mitotic spindle responsible for chromosomal segregation was disassembled. Thus, microtubules were shown to be involved not only in maintenance of cell shape, but also in cellular events such as mitosis and cell motility [24]. Cellular microtubules are largely assembled from two types of tubulin, α-tubulin and β-tubulin: α-tubulin and β-tubulin form a dimer, which is the repeating subunit of the microtubule protofilament [25]. Like actin, tubulin polymerization is energy-dependent. However, unlike actin, the energy for polymerization is harnessed through GTP binding [26]. Both α- and β-tubulin bind GTP prior to assembly into the microtubule polymer. Dimers assemble in a head-to-tail fashion [27]. Because of the polar structure of the dimer subunits consisting of α/β-tubulin, microtubule ends are structurally distinct, with one end exposing α-tubulin and the other exposing β-tubulin [25]. α/β-tubulin prefers to polymerize to the exposed β-tubulin pole rather than the exposed α-tubulin pole. Because of this, on the microtubules pole with the exposing β-tubulin is called the “+” pole, while the pole with the exposing α-tubulin is called the “−” pole. These structural differences and α/β-tubulin assembly preferences lead to distinct rate constants of polymerization and depolymerization at each pole. Furthermore, the rate of polymerization, or subunit addition, is α/β-tubulin concentration dependent; depolymerization, or subunit removal, is GTP hydrolysis dependent, with hydrolysis occurring on the β-tubulin subunit of the dimer [28]. Since microtubule polymerization is α/β-tubulin concentration dependent, the concentration in which net addition of α/β-tubulin dimers to the microtubule poles ceases is called the critical concentration [28]. Therefore, there is net addition of α/β-tubulin dimer to the microtubule poles at concentrations above their critical concentrations, and net removal of α/β-tubulin subunits below their critical concentrations. The “−” end requires a higher concentration of α/β-tubulin for net dimer addition relative to the “+” end and thus a higher critical concentration for assembly. Since there are two poles, there are two critical concentrations. The weighted average of both these concentrations is called the bulk critical concentration. If the bulk tubulin concentration is above the critical concentration at the “+” end but below the critical concentration at the “−” end then there is a net addition of α/β-tubulin dimers to the “+” end and net removal from the “−” end. This behavior, observed largely in vitro, is known as “treadmilling.” In cells, the “−” ends are typically anchored on stabilizing centers called microtubule-organizing centers or MTOCs and only the “+” ends are exposed. The polar nature of microtubules confers unique properties on the polymer. One characteristic of microtubules is their ability to exhibit a behavior called dynamic instability [29, 30] in vitro and in vivo. Dynamic instability results in the detection of simultaneous growth and shrinkage of individual microtubules within a cell. This behavior

3: MICROTUBULES, ACTIN FILAMENTS AND MOTOR-MEDIATED VESICULAR TRANSPORT

is due to the balance between the growth of the polymer due to subunit addition to the microtubule “+” end and the shrinkage associated with GTP hydrolysis by β-tubulin within the newly added dimers. After incorporation of the tubulin dimer into the microtubule, the β-tubulin with its bound GTP is exposed at the “+” end. The exposed GTP-bound β-tubulin encourages further polymerization at the “+” end, which is energetically favorable [28, 29]. The kinetics of GTP hydrolysis on β-tubulin within the polymer is slow enough that the addition to the “+” end is faster than hydrolysis. This leads to the accumulation of the “GTP cap,” which encourages more dimer addition [29, 30]. As the subunit concentration decreases, the rate of dimer addition slows and GTP hydrolysis at the microtubule “+” end catches up [29]. Once GTP is hydrolyzed to GDP, a sudden depolymerization occurs at the “+” end [26, 29, 30]. This results in a shrinking microtubule. This phenomenon is called catastrophe [28]. The shrinking microtubule undergoing catastrophe can be rescued by the new addition of GTP-bound tubulin dimers to the “+” end if the local concentration of α/β-tubulin returns to the critical concentration [26, 28]. At any one time, subpopulations of microtubules will be undergoing catastrophe or polymerization when subunit concentrations are near critical concentration [28, 30]. Microtubule-binding proteins present in different cells can regulate microtubule dynamics; for instance, the MT-associated proteins or MAPs enhance growth and stability, while the severing proteins such as katanin enhance disassembly [31, 32]. α/β-tubulin assembly into microtubules requires a template or scaffold to begin polymerization. This scaffold includes, among other accessory proteins, another tubulin isoform called γ-tubulin [29]. Once a scaffold of γ-tubulin is established, α/β-tubulin dimers can add to the template, thereby polymerizing into a microtubule [29]. MTOCs which anchor the “−” ends of microtubules in vivo are enriched in γ-tubulin. The location of the MTOCs denotes the position of the “−” end of the microtubule [33]. The number of cellular MTOCs and the positioning of these structures within the cell differs with different cell types [33–35].

F-ACTIN COMPOSITION AND POLYMERIZATION DYNAMICS Early X-ray crystallography and electron microscopy analysis were able to resolve the structure of F-actin as consisting of two polymers interacting in a helical arrangement [36, 37]. Further electron microscopy analysis later confirmed this helical polymer arrangement [36, 38, 39]. Electron microscopy and X-ray crystallography also revealed that actin filaments were asymmetric [40, 41]. This asymmetry was intrinsic to F-actin because of the inherent asymmetry of G-actin, which when polymerized yielded F-actin with distinct ends. These polar ends became apparent when F-actin was incubated with the

31

S1 fragment of muscle myosin 2 [40]. Because of the appearance of F-actin incubated with S1 fragments under electron microscopy, researchers labeled one pole the “barbed” end and the other the “pointed” end [42]. At the same time, biochemists and biophysicists studying the kinetics of G-actin monomer addition to F-actin found that G-actin monomers added to the polymer at both poles, but preferred to add more to one pole and less to the other [43]. Researchers labeled the preferred end of monomer addition the “+” end and the other the “−” end. It was later revealed that the barbed end is the “+” end and the pointed is the “−” end [40, 43]. Thus, the asymmetry of the polar ends of F-actin is both a structural and a kinetic feature that arises from the intrinsic asymmetry of the G-actin monomer. F-actin polymerization is dependent on the concentration of ATP-bound G-actin. The concentration of ATP-bound G-actin at which net addition to F-actin ceases is called the critical concentration. Net ATP-bound G-actin addition to the poles of the F-actin polymer occurs at concentrations above the critical concentration, while net removal occurs at concentrations below the critical concentration [28]. As for microtubules, there are two critical concentrations for the two poles of F-actin, a lower concentration and a higher concentration, for the “+” ends and “−” ends, respectively. The bulk critical concentration is the weighted average of the two critical concentrations [26, 28]. However, unlike microtubules, there is not a GTP cap for actin filaments, so polymerization is entirely reliant on the bulk critical concentration of G-actin [26]. It is therefore unsurprising that a host of actin-binding proteins exist which can regulate the critical concentration of G-actin near the ends of the F-actin, including effectors such as profilin and thymosin β4 [28, 44, 45]. F-actin dynamics are characterized largely by a process called “treadmilling,” where the rate of polymerization at the “+” end equals the rate of depolymerization at the “−” end without significant change in the length of the filament. While filamentous actin and microtubules undergo both treadmilling and dynamic instability behavior, dynamic instability is the predominant feature of microtubules while treadmilling is the predominant feature of actin filaments. De novo polymerization of actin is energetically unfavorable [46]. Cells can circumvent the energy barrier through the use of accessory proteins. Three main proteins aid in the initiation of actin filament polymerization: N-WASP, Arp 2, and Arp 3 [47]. N-WASP and Arp 2/3 bind to each other. This binding establishes the scaffold in which G-actin subunits can associate, thereby initiating polymerization. N-WASP and Arp 2/3 can also initiate actin branching from a central filament. N-WASP is the main regulator of actin polymerization and is sensitive to signaling molecules such as Rho, Rac, and Cdc42, which enables the cell to control actin assembly and dynamics in structures including stress fibers, lamellipodia, membrane

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THE LIVER: MICROTUBULE-BASED MOTOR PROTEINS

ruffling, and filopodia, in response to external stimuli [46, 47].

In conclusion, the orientation of microtubules in epithelial cells differs depending on the cell type. Microtubule and actin filament organization in different cell models is shown in Plates 3.1 and 3.2.

MICROTUBULE AND ACTIN FILAMENT ORGANIZATION IN EPITHELIAL CELLS

MICROTUBULE-BASED MOTOR PROTEINS

The cytoskeletal organization in epithelial cells is such that actin filaments and microtubules occupy distinct regions and domains in the cell. In epithelial cells such as hepatocytes, lacrimal gland or pancreatic acinar cells, and intestinal enterocytes, actin filaments are densely enriched in the cell cortex, specifically the region underneath the apical plasma membrane (APM) [42, 48–51]. In hepatocytes, this region of APM and the juxtaposed actin-rich subapical region are called the canalicular membrane and the pericanalicular region, respectively. Analysis of cortical actin in this region of hepatocytes and intestinal enterocytes by electron microscopy has revealed that actin filaments are oriented so that the “+” ends are directed toward the APM and the “−” ends toward the cell interior [50, 52, 53]. In addition, the actin filaments form a branched network where new filaments emerge from existing filaments to generate an arboreal network with the “+” ends pointed toward the APM [28]. Several of these actin filament networks layer on top of each other to form an actin meshwork called the terminal web or, in the case of hepatocytes, the pericanalicular web [52, 54]. Since the terminal/pericanalicular web occupies only the subapical cortex of polarized epithelial cells, this actin is sometimes referred to as the cortical actin [54]. Actin filaments form parallel bundles in the cores of the microvilli of intestinal enterocytes, hepatocytes and acinar cells [42, 52, 54]. The filaments that make up these parallel bundles all have their “−” ends pointing toward the cell body [42, 52]. In hepatocytes, as these bundles enter the pericanalicular web, the bundles fray and merge with the web, where filaments from adjacent microvilli cross each other bidirectionally, parallel to the canalicular membrane [52]. Below the cell cortex, microtubules are mostly localized to the cell interior [48, 50]. In many epithelial cells, including enterocytes and acinar cells, microtubules have their “−” ends associated with the MTOCs beneath the APM cell cortex and their “+” ends extending throughout the cell toward the APM (Plate 3.1) [35, 48]. However, in hepatocytes, the MTOCs are located perinuclearly (Plate 3.1) [50]. This suggests that in hepatocytes, the microtubules direct their “+” ends toward the APM/canalicular membrane as well as the basolateral or sinusoidal membrane, and their “−” ends to the perinuclear MTOCs. Both orientations differ from the radial array of microtubules emanating from the perinuclear MTOCs in simple cultured cell models such as the HeLa cell (Plate 3.2).

In 1965, Rowe and Gibbons isolated an ATPase from the ciliary protozoan Tetrahymena pyriformis [55, 56]. Ultrastructural studies showed that this ATPase formed “arms,” and it was thought that these structures were responsible for the ATP-dependent beating of the cilia surrounding the protozoa. They called this protein “dynein”, a composite name from the Greek term “dyne”, meaning “force”, and “-in”, the suffix meaning “protein”. Soon after, Gibbons discovered dynein in the flagella of sea urchin spermatozoa. Again, dynein was implicated in having a role in the chemo-mechanical beating of both sea urchin sperm flagella and protozoan cilia [56, 57]. Both flagellar and ciliary structures are enriched in microtubules, and it soon became apparent that dynein-microtubule associations were responsible for the beating of flagella and cilia. This focus on the role of microtubules and their associated motor proteins in cellular structure and motility expanded when the microtubule subunit tubulin was extracted in high amounts from brain [58, 59]. Since neuronal cells are non-motile, the presence of tubulin suggested that microtubules might have additional roles other than participation in structural or mechanical functions [55]. An ATPase motor protein very similar to flagellar and ciliary dynein was isolated from brain [60]. It was soon discovered that this ATPase shared similar structure and amino acid sequence [58, 61, 62]. It became apparent that this protein was another dynein, called “cytoplasmic dynein”. The term “cytoplasmic dynein” was given to this new protein to distinguish it from its close flagellar and ciliary cousin, axonemal dynein [62]. To date there have been only two cytoplasmic dyneins discovered, cytoplasmic dynein 1 and cytoplasmic dynein 2 [63]. Cytoplasmic dynein 2 is localized in the cytoplasm of the flagella and cilia. Cytoplasmic dynein 1 is present throughout all microtubule-containing cells. Because of the ubiquitous expression of cytoplasmic dynein 1, the general term “cytoplasmic dynein” is assumed to refer to cytoplasmic dynein 1 unless otherwise specified. Evidence supports a role for cytoplasmic dynein 1 in vesicle and organelle transport by mediating the “walking” of cargo-laden vesicles along microtubules, a function first observed in neuronal cells and then in other cell types such as epithelial cells [51, 62]. The second type of microtubule-based motor protein to be discovered was kinesin [64]. Kinesin was discovered when glass beads were observed to move on microtubules

3: MICROTUBULES, ACTIN FILAMENTS AND MOTOR-MEDIATED VESICULAR TRANSPORT

after their immersion in squid axoplasm [65]. Other kinesins were later discovered in non-neuronal cells such as epithelial cells [66]. There are now upwards of 600 kinesins, including 45 kinesins identified in humans and mice; the first kinesin identified is now labeled “conventional kinesin” [67]. Kinesins are implicated in a wide variety of cellular functions, such as the maintenance of cellular polarity, chromosome motility during mitosis, and organelle and vesicle trafficking. Similar to cytoplasmic dynein 1, kinesins can associate with vesicles and walk them along microtubules in an ATP-dependent manner [64, 67].

Cytoplasmic dynein Cytoplasmic dynein is a large multiprotein complex composed of two conventional dynein heavy chains, three intermediate chains, and four light chains [58]. The two heavy chains comprise the two motor domains that interact with microtubules. In addition to the microtubule binding site, the motor domains also contain six AAA+ domains, in which one or two of these domains are thought to have ATPase activity [63, 68]. The hydrolysis of ATP enables the motor domain to “walk” along the microtubules, although the way ATP hydrolysis drives cytoplasmic dynein and its cargo along the microtubule at the molecular level has not been clearly ascertained [68]. Current models show that cytoplasmic dynein associates with vesicles through a multiprotein complex called the dynactin complex, however some recent work shows that cytoplasmic dynein can bind directly to vesicles [58, 69]. The dynactin complex’s role in cytoplasmic dynein-driven transport can be shown by overexpressing exogenous dynamitin/p50, a component of the dynactin complex, which inhibits cytoplasmic dynein vesicle movement [70]. However, evidence is emerging that the dynactin complex may function by regulating cytoplasmic dynein [69]. Furthermore, the overexpression of an exogenous dominant negative mutant 150Glued , another dynactin complex constituent, does not inhibit cytoplasmic dynein processivity [69]. The exact interaction of dynactin and its role in cytoplasmic dynein function is still a work in progress. Once cytoplasmic dynein associates with vesicles, the motor assembly utilizes ATP to drive microtubule-mediated transport [68]. This transport is observed to be directed toward the “−” end of microtubules, therefore cytoplasmic dynein is a “−” end motor protein [58]. Since the orientation of microtubules in epithelial cells differs depending on the cell type, the direction of cytoplasmic dynein-driven vesicle movements (i.e. toward the apical versus basolateral membranes) may differ between cell types. For instance, hepatocytes have the “−” ends oriented toward the perinuclear region

33

and acinar cells have the “−” ends oriented toward the APM, therefore cytoplasmic dynein likely functions as a motor for movement of materials to the APM in acinar cells but not in hepatocytes [71, 72].

Kinesins Since the discovery of conventional kinesin, discoveries of other kinesins have greatly expanded the superfamily. To date, 45 kinesins in humans have been discovered, and these have systematically been categorized into 14 classes [64]. Under the new system of classification, conventional kinesin is called KIF5B and all members of the kinesin superfamily are labeled as KIFs [73]. Kinesins have a tripartite structure consisting of a semiconserved catalytic head domain (also called the catalytic core) that both binds to microtubules and hydrolyzes ATP for force generation; a neck region that is susceptible to cellular regulation; and a stalk or tail that can interact with organelles such as vesicles or nucleic acids [64]. The structural diversity of the differing classes of kinesins is notable even between members of the same class. The highly divergent tail/stalk structures are thought to be reflective of the highly diverse functions of kinesins, which mediate the binding and microtubule-based movement of organelles ranging from the synaptic vesicles in neurons to the chromosomes during meiosis and mitosis. In most kinesins the catalytic domains are located at the N-terminus, but in some classes the catalytic domains are in the C-terminus or in the middle of the primary amino acid sequence. The location of the motor domain within the amino acid sequence has a major impact on the directionality of kinesins. For instance, the conventional kinesin class and other members of the KIF5B class have their catalytic domains at the N-terminal, which enables them to walk along microtubules toward the “+” end. However, kinesins in the Kinesin-14 class walk toward the “−” end of microtubules because their catalytic domains are at the C-terminus, while some members of the Kinesin-13 class have their catalytic domain in the middle of the sequence and are “+” end motors [74]. The structure of kinesins is varied further by their quaternary structures. For instance, conventional kinesin includes homodimers of the heavy chains (head, neck, and tail) as well as homodimers of the light chains. In contrast, the Kinesin-2 class members are heterotrimeric and the Kinesin-3 class members are monomeric. The double-headed conventional kinesin is thought to walk along microtubules in an ATPase-driven, asymmetrical hand-over-hand manner [75, 76]. This asymmetric walk is hypothesized to result from the over-twist of the coiled-coil region in the neck after the trailing head makes its stride. This makes the leading head take a shorter

34

THE LIVER: MYOSIN MOTOR PROTEINS

stride, giving conventional kinesin and related family members a limp as they walk along the microtubule [76, 77]. However, this model remains controversial. KIF1A, in contrast, can function as a single-head monomer or can work synergistically as a functional double-headed dimer [78]. KIF1A is processive as both a monomer and a dimer, however the details of its walk along microtubules have not been completely elucidated [78].

MYOSIN MOTOR PROTEINS Non-muscle actin was discovered in slime mold, Physarum polycephalum, and later in Acanthamoeba [79, 80], leading to speculation that myosins might exist in tissues besides muscle tissue. In non-muscle cells, actin filaments appeared to be enriched in structures involved in cell motility called lamellipodia and pseudopodia [18, 19]. Lamellipodia are manifested by ruffling of the plasma membrane, a process by which motile cells create the mechanical force for propulsion [19]. When myosins were discovered in non-muscle cells, scientists hypothesized that myosin in non-muscle cells behaved the same as myosin in muscle, functioning with actin to generate mechanical force [81, 82]. In muscle, actin and myosin are the active components for contraction, and in non-muscle cells, actin and myosin were thought to be involved in propulsion [83]. This mechanistic view of actin and myosin function in non-muscle tissues began to appear more complicated when the first unconventional myosin was discovered; this myosin was later termed “myosin 1” [82]. The term “unconventional” was applied to distinguish myosin 1 from the conventional muscle and non-muscle myosin 2 [80, 84, 85]. Later, as more myosins were discovered, the term “unconventional” was applied to all myosins except for the filament-forming conventional myosin 2. In polarized epithelial enterocytes, myosin 1 is involved in the bundling of actin filaments that runs throughout the core of the microvilli [42]. More evidence began to emerge that actin and nonmuscle myosin might not only be involved with the maintenance of cellular structure and cellular propulsion, but also in vesicular traffic [82, 86–88]. Evidence for myosin’s role in vesicular traffic first emerged when it was observed that phagocytic cups in Acanthamoeba were enriched in myosin 1, and vesicle transport in vitro was possible with myosin 1 [82]. However it was not until the discovery of the dilute mouse that solid evidence emerged for a role for myosins in vesicular traffic [89, 90]. The phenotype for the dilute mouse is albinism and severe neurological impairment. It was discovered that the dilute mouse has a loss-of-function mutation in another previously uncharacterized unconventional myosin called myosin 5a [89]. This phenotype is attributed to the inability of myosin 5a to transport melanosomes (pigment granules) along actin filaments to the plasma membrane, leading to accumulation in the perinuclear region of the melanocyte,

and resulting in lightening of the coat color [91]. Similarly, the neurological impairment is attributed to the inability of myosin 5a to move synaptic vesicles toward the dendritic membrane, fuse them to the membrane and discharge their neurotransmitter load [90, 92].

Myosins To date, 20 myosin classes have been discovered since the discovery of the first unconventional myosin, myosin 1 [93, 94]. Most myosins have a similar conserved tripartite structure consisting of a semiconserved N-terminal motor domain that associates with actin filaments and can generate force utilizing ATP hydrolysis; a neck region that can regulate head domain/actin filament interaction; and a C-terminal highly-divergent tail domain that mediates diverse functions such as intracellular membrane trafficking, tethering, and polarization [93–96]. The amino acid sequence of the tail domain confers function, identity, and membership into one of the 19 classes of unconventional or 1 class of conventional myosins. While the head region is semiconserved, the total structure of myosins can vary. For instance, the class V, VI, and II myosins exist as homodimers with two heads, while myosin 1 exists as a monomer with just one head. The myosin family is a diverse one, labeled as a superfamily, with a similar structural and functional diversity exemplified by the kinesin superfamily. While the functions of different classes of myosins are just as diverse as their structures, the focus of this chapter is on their known functions in vesicle and organelle transport in epithelial cells. Myosins, particularly class V, VI, II, and I myosins, are involved in vesicle transport by binding to vesicle cargo through their tail domains [93, 94]. ATPase activity, as well as actin binding within the head domain, confers the ability of myosins to walk along the tracks provided by actin filaments [93, 97]. In the case of the double-headed myosins like class V, VI, and II, the ATPase activity is staggered so each head is in a different state of ATP hydrolysis [93,97–99]. These conformational changes give myosins their ability to walk vesicles along actin filaments. For the double-headed myosins, this walk is akin to a “hand-over-hand” motion, while the single-headed myosins propel vesicles by using a ratchet-like movement of the head [39, 100]. Furthermore, myosins have an inherent directionality to their transport [93, 100]. Class V, I, and II move vesicle to the “+” ends and class VI myosins move vesicles to the “−” end of the actin filament [93]. This inherent directionality preference has implications for the direction of vesicle motility, because of filament orientation at the pericanalicular/subapical region of polarized epithelial cells. For instance, myosin 5a and myosin 5c have been implicated in the movement of secretory vesicles to the APM to participate in exocytosis in pancreatic beta MIN6 cells and lacrimal acinar cells, while myosin 6 has been

3: MICROTUBULES, ACTIN FILAMENTS AND MOTOR-MEDIATED VESICULAR TRANSPORT

implicated in clathrin-mediated apical endocytosis in Caco-2 cells [101–103].

CYTOSKELETON AND INTRACELLULAR TRAFFICKING IN EPITHELIAL CELLS Cytoskeleton and motor proteins are key participants in essential membrane trafficking events in polarized epithelial cells such as endocytosis, transcytosis, and exocytosis. Furthermore, cytoskeleton and motor proteins have a role in the spatial organization of organelles and the maintenance of distinct apical and basolateral membrane domains. Thus, cytoskeleton and motor proteins are essential to the basic functions of epithelial cells. Exocytosis is the constitutive or regulated release of contents from vesicular stores at either apical or basolateral plasma membranes (BLMs) to the external environment. The earliest indication of the importance of the cytoskeleton to this function in hepatocytes was observed when rats injected with phalloidin (actin filament stabilizer) or colchicine (microtubule polymerization inhibitor) showed decreased bile acid secretion from the canalicular membrane [104–106]. Similar experiments using cytoskeleton-disrupting or -stabilizing agents in exocrine epithelial cells such as lacrimal and pancreatic acini exhibited various disruptions in exocytosis [48, 107, 108]. Endocytosis occurs with the uptake of membraneassociated or soluble macromolecules from the extracellular space into the cell by plasma membrane invagination and budding. Endocytosis was also shown to be affected when cultured hepatocytes were incubated with the microtubule stabilizer taxol [33]. Transcytosis is the movement of vesicles endocytosed from the BLMs toward the APM followed by the release of vesicular contents at the APM. Transcytosis can occur in reverse, from the APM to the BLMs. This mechanism is principally used by cells to move essential macromolecular cargo across a cell layer [109], since movement between epithelial cells (paracellular transport) is impeded by junctional connections between cells. Inhibitor studies show that microtubules and actin filaments are likewise involved in transcytosis [110]. Incubation of polarized Madin–Darby canine kidney (MDCK) cells with the tubulin-sequestering agent nocodazole reduced transcytosis by ∼60% [110, 111]. When these same cells were treated with the F-actin depolymerization agent cytochalasin D, transcytosis was inhibited by ∼45%. When MDCK cells were treated with both nocodazole and cytochalasin D, transcytosis was inhibited by 95%. Incubation of the hepatocytic cell line HepG2 at 4 ◦ C (microtubule depolymerization) caused a decrease in the transport of phospholipids to the canaliculus, suggesting a decrease in transcytosis [112]. Hepatocytes and hepatocyte-derived cell lines are known to use the

35

transcytotic pathway from the BLMs to the APM for several proteins such as polymeric IgA receptor (pIgAR), dipeptidyl peptidase IV and GPI-anchored 5′ nucleotidase [113–115]. These studies using cytoskeleton-disrupting agents collectively provide evidence that actin filaments and microtubules are important to the normal vesicular transport functions of exocytosis, endocytosis, and transcytosis in hepatocytes and other epithelial cells. The importance of cytoskeleton in many of these instances has subsequently been shown to be due to impairment of motor protein function, which inevitably follows from the disruption of cytoskeleton dynamics.

MYOSIN FUNCTION IN EPITHELIAL CELLS Polarized epithelial cells express several classes of myosin. The best characterized of the myosins expressed in polarized epithelial cells are the class I, II, V, and VI myosins.

Myosin 1 Vertebrates express six class I myosins: myosin 1a, myosin 1b, myosin 1c, myosin 1d, myosin 1e, and myosin 1f [116]. While cells can express several members of the class I myosins concurrently, some expression of class I isoforms is cell-specific. Myosin 1b and 1c have been characterized in hepatocytes [117, 118]. Myosin 1a is almost exclusively expressed in the brush border membrane of intestinal enterocytes. In intestinal enterocytes, myosin 1a (also known as Brush Border Myosin 1) is responsible for the binding of the core actin bundles to the plasma membrane in the microvilli of enterocytes. Myosin 1b is ubiquitously expressed in most cell types, including hepatocytes [117]. Myosin 1b purified from liver shows an ability to bundle or crosslink actin. This may function to maintain actin-rich regions of the cell such as the cortical actin. In a liver tumor-derived non-polarized hepatoma cell line, myosin 1b was shown to be involved with the traffic between endosomes and lysosomes [119]. Myosin 1b is localized in multivesicular endosomes in non-polarized melanosomes [120]. In addition, intracellular localization studies revealed significant myosin 1b enrichment in the plasma membrane of hepatocytes [99, 118]. Since myosin 1b has been found to be enriched in cortical actin in hepatocytes, is implicated in the endosomal pathway, and has actin bundling capability, myosin 1b may be involved in actin reorganization during endocytosis in hepatocytes. Myosin 1c is also expressed in polarized epithelial cells and has been detected in hepatocytes [118]. Localization studies of myosin 1c in hepatocytes reveal that myosin 1c is enriched in two membranes, the Golgi apparatus and the APM [99, 118]. A pleckstrin homology (PH)

36

THE LIVER: MYOSIN FUNCTION IN EPITHELIAL CELLS

domain is present in myosin 1c, which enables myosin 1c to bind to membrane domains rich in phosphoinositides [99]. Furthermore, myosin 1c has been implicated in the transport of lipid rafts from the trans-Golgi network (TGN) to the APM in non-polar cells. Since myosin 1c is enriched in both the TGN and APM of hepatocytes, myosin 1c may have a role in the transport of lipid rafts to the APM from the TGN in this cell type.

Non-muscle Myosin 2 Non-muscle myosin 2 has been implicated in exocytosis in polarized epithelial cells of many cell types [108]. In pancreatic acinar cells, non-muscle myosin 2 is involved in the transport of zymogen granules to the APM for subsequent fusion during stimulated exocytosis [121]. In lacrimal gland acinar cells, non-muscle myosin 2 has been proposed to play a role in a type of exocytosis called multivesicular exocytosis, in which actin filaments reorganize around clusters of adjacent secretory vesicles (Plate 3.3). Non-muscle myosin 2, with actin, then facilitates the intracellular compression of these clusters of secretory vesicles before content extrusion through fusion pores at the APM. In MDCK cells, bile salt export pump (BSEP) is exported and internalized to and from the APM in a non-muscle myosin 2-dependent manner [122]. Furthermore, myosin 2 light chain kinase co-immunoprecipitates with BSEP from liver. This implies that non-muscle myosin 2 is involved in the transport by BSEP to the APM through the recycling endosomes. Similar studies show that non-muscle myosin 2 proteins are implicated in the direct transport of mdr-1 (hydrophobic cation exporter) and mdr-2 (phosphotidycholine flippase) directly to the APM from the TGN in liver-derived cell lines [122, 123].

Myosin 5 Vertebrates express three class V myosins: myosin 5a, myosin 5b, and myosin 5c [89, 95]. In the dilute mouse, which has a loss-of-function mutation to the myosin 5a gene, melanosomes (pigment granules) accumulate in a perinuclear region of the melanocyte, resulting in a lightening of the coat color [91]. The coat color phenotype is the result of a failure of myosin 5a to capture, tether, and maintain the melanosomes in the actin-rich periphery of the melanocyte [124]. Animals lacking myosin 5a function also suffer from a lethal neurological disorder and fail to properly localize an endoplasmic reticulum-like structure in dendritic spines of Purkinje neurons of the brain [125]. These defects and melanosome accumulations can be emulated by expressing a dominant negative tail, thus confirming the importance of motor activity for proper targeting of these organelles [124, 126]. Myosin 5a is also expressed in polarized epithelial cells including acinar epithelial cells [103].

Myosin 5b, the second of the class V myosins expressed in vertebrates, is associated with a plasma membrane recycling compartment in several cell types [127–133]. Based on studies with a dominant negative tail, myosin 5b inhibits proper recycling of several cell surface receptors including transferrin receptor, CXCR2, and the M4 muscarinic receptor [129, 130, 134]. Interestingly, myosin 5b has been implicated in the establishment of apical or canalicular polarity in hepatic cell lines. Inhibition of myosin 5b function with dominant negative myosin 5b expression caused accumulation of apical resident proteins trafficking from the TGN and the basolateral membrane in hepatic cell lines. Myosin 5b functioning in the trafficking of proteins and lipid components to the plasma membrane may therefore be critical to the establishment of canalicular polarity [115, 128]. Myosin 5c, the third member of the class V myosin family in vertebrates is expressed most abundantly in exocrine secretory tissues, where it localizes in the apical domain of epithelial cells [89]. In HeLa cells, the expression of a dominant negative myosin 5c tail led to an accumulation of transferrin receptors in large cytoplasmic puncta and inhibited transferrin recycling. Recently, myosin 5c was identified in a proteomics study as a component on secretory granules of pancreatic acinar cells [135]. Moreover, in lacrimal gland acini, myosin 5c has been characterized as a major contributor to the reorganization of actin around clusters of secretory vesicles and the ultimate exocytosis of the contents within these clusters at the APM [103] (Plate 3.3).

Myosin 6 Myosin 6 has been implicated in inward movement of endocytic vesicles from the APM of polarized epithelial cells [46, 102, 136]. Since actin filaments are oriented with their “+” ends to the APM, the obvious motor to be involved in actin-dependent events in endocytosis or movement away from the APM is the “−” end-directed myosin 6. This was confirmed through co-localization studies with clathrin as well as with functional analysis [137]. In addition, one of the four isoforms of myosin 6 has a role in basolateral membrane sorting and secretion in polarized cells by functioning in tubular extension and fission from the recycling endosome and TGN, respectively [138]. Loss-of-function mutants of the myosin 6 allele in mice, Snell’s waltzer, result in deafness [139, 140]. Snell’s waltzer phenotype results from the inability of myosin 6 to maintain the structure of stereocilia in epithelial cells of the cochlea [141]. Myosin 6 has also been implicated in the anchoring and maintenance of the Golgi [102]. Myosin 6 may have a role in actin reorganization during lamellipodia and filopodia formation in motile non-polarized cells. There is controversy about whether myosin 6 can exist in vivo as a dimer, monomer or both, depending on its function [102, 142]. No in-depth analysis of myosin 6 function in hepatocytes has been done, but one can presume that

3: MICROTUBULES, ACTIN FILAMENTS AND MOTOR-MEDIATED VESICULAR TRANSPORT

37

Table 3.1 Actin- and microtubule-dependent motor proteins implicated in specific trafficking events in epithelial cells Motor protein

Function

Reference

Myosin 1a (Brush Border Myosin)

Actin-based “+” end motor. Exclusively expressed in enterocytes. Mediates actin bundle binding to the plasma membrane in the microvilli. Responsible for the structural integrity of the microvilli. Actin-based “+” end motor. Involved in the endocytotic pathway. Localized to endosomes and lysosomes in unpolarized hepatoma cell lines. Localized in multivesicular endosomes in unpolarized melanoma cell lines. Crosslinks actin in hepatocytes. Detected on the plasma membrane of hepatocytes. Actin-based “+” end motor. Trafficking of lipid rafts to the canalicular membrane. May be involved in the last stages of exocytosis. Implicated in membrane ruffling and regulation of neuronal growth cone extension. Localized in stereocillia tips. Also, associated with the Golgi in hepatocytes. Actin-based “+” end motor. Role in transferrin trafficking from the early endosomes to the recycling endosomes in MDCK cells. Actin-based “+” end motor. Associates with actin bundles and provides actin-dependent movement associated with compression of actin bundles. Facilitates homotypic fusion of secretory vesicles during multivesicular or compound exocytosis. Also implicated in actin reorganization in endocytosis. Controls phagocytosis. Actin-based “+” end motor. Exocytosis of synaptic vesicles, secretory vesicles and melanosomes. Maintenance of ER. Tethering of organelles. Negative regulator of endocytosis. Actin reorganization in growth cone filopodia in neurons. Present in acinar cells. Actin-based “+” end motor. Enriched in recycling endosomes. Recycling of transferrin receptor and other receptors to the APM. Recycling trafficking of BSEP to the APM in WIF-B9 hepatic cell lines. Actin-based “+” end motor. Associated with secretory vesicles in acinar cells. Involved in the trafficking of transferrin receptor. Implicated in actin reorganization during exocytosis in secretory epithelial cells. Actin based “–” end motor. Implicated in clathrin-dependent endocytosis in epithelial cells. Actin reorganization during cell migration. Structural maintenance of stereocilia. Movement of ligand-bound receptors down the microvilli. Movement of vesicles through the terminal web/cortical actin to early endosomes. Microtubule-based “+” end motor. Involved in early endocytosis and fission of endocytic vesicles in hepatocytes.

[88, 94, 143]

Myosin 1b

Myosin 1c

Myosin 1d Non-muscle Myosin 2

Myosin 5a

Myosin 5b

Myosin 5c

Myosin 6

KIF5B (Conventional Kinesin) KIFC1 KIFC2 Cytoplasmic Dynein 1

Microtubule-based “–” end motor. Involved in early endocytosis and fission of endocytic vesicles in hepatocytes. Microtubule-based “–” end motor. Involved in early endocytosis and fission of endocytic vesicles in hepatocytes. Microtubule-based “–” end motor. Associates with late endocytotic vesicles in hepatocytes. Associates with immature secretory vesicles in acinar cells that are targeted to the apical plasma membrane.

myosin 6 may have the same function in hepatocytes as it does in other polarized epithelia. Table 3.1 summarizes the myosin motors so far identified in epithelial cells as well as some of their possible functions.

KINESINS AND CYTOPLASMIC DYNEIN IN EPITHELIAL CELLS In primary hepatocytes, the predominant location of the MTOCs is the perinuclear region, and in the

[117–120]

[94, 99, 118]

[144] [94, 108, 145, 146]

[94, 103, 124, 145–148]

[94, 115, 128, 130, 134, 149] [89, 103]

[50, 94, 102, 137, 145, 146, 150, 151]

[152]

[152] [153] [50, 63, 71, 72]

hepatocyte-derived cell line WIF-B9 the predominant location is immediately below the APM [33–35]. The orientation of microtubules, in particular their polar ends, has a major impact on the molecular machinery required for events such as endocytosis, exocytosis, and transcytosis. In hepatocytes, microtubule-based movement to the APM may require “+” end-directed motors, while in WIF-B9 cells and other polarized epithelial cells like acinar cells, movement to the APM requires “−” end-directed motors.

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THE LIVER: REFERENCES

In primary hepatocytes, early endocytic vesicles are internalized and trafficked toward the “−” end of the microtubules [50]. Isolated early endocytic vesicles from rat liver associate with both the “+” end kinesin KIF5B and the “−” end kinesin KIFC2 [153]. It is hypothesized that the two opposing motors enable early endocytic fission during receptor recycling once the vesicle is internalized; the receptor moving along the “+” end directed back to the plasma membrane, and the ligand moving to the “−” end toward the perinuclear region of the cell [153, 154]. In mouse, kinesin-mediated fission through KIF5B and KIFC2 may sort heterogeneous endosomal contents into different vesicles [153, 154]. However, this observation was not seen in KIFC2 knockout mouse liver; rather a population of mouse early endocytic vesicles was enriched in KIF5B and another “−” end kinesin, KIFC1, which appears to function in endocytic vesicle fission in the same manner as KIFC2 in rat hepatocytes [152]. Cytoplasmic dynein is involved in the endocytotic pathway in hepatocytes [71], specifically by participating in the late endocytic traffic of ligand-containing vesicles to the lysosomes after the segregation from early endocytic vesicles [152, 155]. In lacrimal gland acinar cells, cytoplasmic dynein is responsible for transport of cargo destined for the exocytotic pathway to the APM, while kinesin is involved in membrane retrieval from this domain [72, 156]. Beyond hepatocytes and lacrimal acini, microtubule orientation and the role of microtubule-based motors have not been comprehensively studied in other primary polarized epithelial cells and often conflicting reports are published in the literature. Table 3.1 summarizes the function of several microtubule-based motors in epithelial cells as well as their proposed functions. The integrated functioning of microtubules, actin filaments and their associated motor proteins contributes greatly to the unique trafficking patterns and transport of diverse cargo in epithelial cells. Plate 3.3 depicts the action of microtubule-based and myosin motors in secretagogue-stimulated exocytosis of tear proteins from the lacrimal gland acinar cells at the APM. Plate 3.4 depicts the action of microtubule-based and myosin motors in facilitating endocytosis at the sinusoidal membrane, as well as exocytosis and recycling at the canalicular membrane in hepatocytes.

CONCLUSION Cytoskeleton and motor proteins mediate many essential events in polarized epithelial cells. In polarized epithelial cells, actin filaments and microtubules are enriched in discrete regions in the cell. Disruption of either one or both can lead to the inability of the cell to perform vital functions such as endocytosis, exocytosis, and transcytosis. Furthermore, these functions are reliant on the proper function of motor proteins that utilize the cytoskeleton to promote key steps in membrane

traffic. Evidence is emerging that motor proteins and cytoskeleton interact in an increasingly complex way. For instance, myosin 5 has been observed to associate and walk on microtubules [157, 158]. In addition, kinesins have been observed to directly bind with myosin 5 and the cytoplasmic dynein-associated protein, dynein light chain kinase, has been shown to bind to myosin 5a [159, 160]. While cytoskeleton and motor proteins have been shown to be integral to the proper function of polarized epithelial cells, elucidation of the complexity, and regulation of their interaction remains a work in progress.

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N.A. and Meisler, M.H. (2000) The mouse neurological mutant flailer expresses a novel hybrid gene derived by exon shuffling between Gnb5 and Myo5a. Hum Mol Genet , 9, 821–28. Lise, M.F., Wong, T.P., Trinh, A., Hines, R.M., Liu, L., Kang, R., Hines, D.J., Lu, J., Goldenring, J.R., Wang, Y.T. and El-Husseini, A. (2006) Involvement of myosin Vb in glutamate receptor trafficking. J Biol Chem, 281, 3669–78. Wakabayashi, Y., Dutt, P., Lippincott-Schwartz, J. and Arias, I.M. (2005) Rab11a and myosin Vb are required for bile canalicular formation in WIF-B9 cells. Proc Natl Acad Sci USA, 102, 15087–92. Fan, G.H., Lapierre, L.A., Goldenring, J.R., Sai, J. and Richmond, A. (2004) Rab11-family interacting protein 2 and myosin Vb are required for CXCR2 recycling and receptor-mediated chemotaxis. Mol Biol Cell , 15, 2456–69. Lapierre, L.A., Kumar, R., Hales, C.M., Navarre, J., Bhartur, S.G., Burnette, J.O., Provance, D.W. Jr., Mercer, J.A., Bahler, M. and Goldenring, J.R. (2001) Myosin Vb is associated with plasma membrane recycling systems. Mol Biol Cell , 12, 1843–57. Zhao, L.P., Koslovsky, J.S., Reinhard, J., Bahler, M., Witt, A.E., Provance, D.W. Jr. and Mercer, J.A. (1996) Cloning and characterization of myr 6, an unconventional myosin of the dilute/myosin-V family. Proc Natl Acad Sci USA, 93, 10826–31. Nedvetsky, P.I., Stefan, E., Frische, S., Santamaria, K., Wiesner, B., Valenti, G., Hammer, J.A. III, Nielsen, S., Goldenring, J.R., Rosenthal, W. and Klussmann, E. (2007) A role of myosin Vb and Rab11-FIP2 in the aquaporin-2 shuttle. Traffic, 8, 110–23. Swiatecka-Urban, A., Talebian, L., Kanno, E., MoreauMarquis, S., Coutermarsh, B., Hansen, K., Karlson, K.H., Barnaby, R., Cheney, R.E., Langford, G.M., Fukuda, M. and Stanton, B.A. (2007) Myosin Vb is required for trafficking of CFTR in RAB11A-specific apical recycling endosomes in polarized human airway epithelial cells. J Biol Chem, 282, 23725–36. Volpicelli, L.A., Lah, J.J., Fang, G., Goldenring, J.R. and Levey, A.I. (2002) Rab11a and myosin Vb regulate recycling of the M4 muscarinic acetylcholine receptor. J Neurosci , 22, 9776–84. Chen, X., Walker, A.K. et al. (2006) Organellar proteomics: analysis of pancreatic zymogen granule membranes. Mol Cell Proteomics, 5, 306–312. Buss, F. and Kendrick-Jones, J. (2008) How are the cellular functions of myosin VI regulated within the cell? Biochem Biophys Res Commun, 369, 165–75. Hasson, T. (2003) Myosin VI: two distinct roles in endocytosis. J Cell Sci , 116, 3453–61. Au, J.S., Puri, C., Ihrke, G., Kendrick-Jones, J. and Buss, F. (2007) Myosin VI is required for sorting of AP-1B-dependent cargo to the basolateral domain in polarized MDCK cells. J Cell Biol , 177, 103–114. Avraham, K.B., Hasson, T., Steel, K.P., Kingsley, D.M., Russell, L.B., Mooseker, M.S., Copeland, N.G. and Jenkins, N.A. (1995) The mouse Snell’s waltzer deafness gene encodes an unconventional myosin required

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for structural integrity of inner ear hair cells. Nat Genet , 11, 369–75. Avraham, K.B., Hasson, T., Sobe, T., Balsara, B., Testa, J.R., Skvorak, A.B., Morton, C.C., Copeland, N.G. and Jenkins, N.A. (1997) Characterization of unconventional MYO6, the human homologue of the gene responsible for deafness in Snell’s waltzer mice. Hum Mol Genet , 6, 1225–31. Seiler, C., Ben-David, O., Sidi, S., Hendrich, O., Rusch, A., Burnside, B., Avraham, K.B. and Nicolson, T. (2004) Myosin VI is required for structural integrity of the apical surface of sensory hair cells in zebrafish. Dev Biol , 272, 328–38. Lister, I., Roberts, R., Schmitz, S., Walker, M., Trinick, J., Veigel, C., Buss, F. and Kendrick-Jones, J. (2004) Myosin VI: a multifunctional motor. Biochem Soc Trans, 32, 685–88. Wolenski, J.S., Hayden, S.M., Forscher, P. and Mooseker, M.S. (1993) Calcium-calmodulin and regulation of brush border myosin-I MgATPase and mechanochemistry. J Cell Biol , 122, 613–21. Huber, L.A., Fialka, I., Paiha, K., Hunziker, W., Sacks, D.B., Bahler, M., Way, M., Gagescu, R. and Gruenberg, J. (2000) Both calmodulin and the unconventional myosin Myr4 regulate membrane trafficking along the recycling pathway of MDCK cells. Traffic, 1, 494–503. Holt, J.P., Bottomly, K. and Mooseker, M.S. (2007) Assessment of myosin II, Va, VI and VIIa loss of function on endocytosis and endocytic vesicle motility in bone marrow-derived dendritic cells. Cell Motil Cytoskeleton, 64, 756–66. Mermall, V., Post, P.L. and Mooseker, M.S. (1998) Unconventional myosins in cell movement, membrane traffic, and signal transduction. Science, 279, 527–33. Wu, X., Rao, K., Bowers, M.B., Copeland, N.G., Jenkins, N.A. and Hammer, J.A. III (2001) Rab27a enables myosin Va-dependent melanosome capture by recruiting the myosin to the organelle. J Cell Sci , 114, 1091–100. Rudolf, R., Kogel, T., Kuznetsov, S.A., Salm, T., Schlicker, O., Hellwig, A., Hammer, J.A. III and Gerdes, H.H. (2003) Myosin Va facilitates the distribution of secretory granules in the F-actin rich cortex of PC12 cells. J Cell Sci , 116, 1339–48. Lapierre, L.A. and Goldenring, J.R. (2005) Interactions of myosin Vb with RAB11 family members and cargoes traversing the plasma membrane recycling system. Methods Enzymol , 403, 715–23.

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150. Dance, A.L., Miller, M., Seragaki, S., Aryal, P., White, B., Aschenbrenner, L. and Hasson, T. (2004) Regulation of myosin-VI targeting to endocytic compartments. Traffic, 5, 798–813. 151. Sweeney, H.L. and Houdusse, A. (2007) What can myosin VI do in cells? Curr Opin Cell Biol , 19, 57–66. 152. Nath, S., Bananis, E., Sarkar, S., Stockert, R.J., Sperry, A.O., Murray, J.W. and Wolkoff, A.W. (2007) Kif5B and Kifc1 interact and are required for motility and fission of early endocytic vesicles in mouse liver. Mol Biol Cell , 18, 1839–49. 153. Bananis, E., Murray, J.W., Stockert, R.J., Satir, P. and Wolkoff, A.W. (2003) Regulation of early endocytic vesicle motility and fission in a reconstituted system. J Cell Sci , 116, 2749–61. 154. Bananis, E., Murray, J.W., Stockert, R.J., Satir, P. and Wolkoff, A.W. (2000) Microtubule and motor-dependent endocytic vesicle sorting in vitro. J Cell Biol , 151, 179–86. 155. Bananis, E., Nath, S., Gordon, K., Satir, P., Stockert, R.J., Murray, J.W. and Wolkoff, A.W. (2004) Microtubule-dependent movement of late endocytic vesicles in vitro: requirements for Dynein and Kinesin. Mol Biol Cell , 15, 3688–97. 156. Hamm-Alvarez, S.F., Da Costa, S., Yang, T., Wei, X., Gierow, J.P. and Mircheff, A.K. (1997) Cholinergic stimulation of lacrimal acinar cells promotes redistribution of membrane-associated kinesin and the secretory protein, beta-hexosaminidase, and increases kinesin motor activity. Exp Eye Res, 64, 141–56. 157. Cao, T.T., Chang, W., Masters, S.E. and Mooseker, M.S. (2004) Myosin-Va binds to and mechanochemically couples microtubules to actin filaments. Mol Biol Cell , 15, 151–61. 158. Ali, M.Y., Krementsova, E.B., Kennedy, G.G., Mahaffy, R., Pollard, T.D., Trybus, K.M. and Warshaw, D.M. (2007) Myosin Va maneuvers through actin intersections and diffuses along microtubules. Proc Natl Acad Sci USA, 104, 4332–36. 159. Langford, G.M. (2002) Myosin-V, a versatile motor for short-range vesicle transport. Traffic, 3, 859–65. 160. Espindola, F.S., Suter, D.M., Partata, L.B., Cao, T., Wolenski, J.S., Cheney, R.E., King, S.M. and Mooseker, M.S. (2000) The light chain composition of chicken brain myosin-Va: calmodulin, myosin-II essential light chains, and 8-kDa dynein light chain/PIN. Cell Motil Cytoskeleton, 47, 269–81.

4

Molecular Motors Peter Satir Department of Anatomy and Structural Biology, Albert Einstein College of Medicine, Bronx, New York, NY, USA

INTRODUCTION Almost all cells of the liver, including hepatocytes, intrahepatic bile duct epithelial cells (cholangiocytes), liver stem cells, capillary endothelium, and sinusoidal lining cells, excluding the mature erythrocyte, have trafficking that operates via molecular motors of the cytoskeleton. In previous editions of this volume, I have reviewed molecular motors of the hepatocyte [1]; here I broaden the topic to include molecular motors operating in the primary cilium of cholangiocytes, fibroblasts, and perhaps other liver cell types. This aspect of the subject has received considerable recent attention, and is related to the development of several pathological conditions, including cystic liver disease and hepatic fibrosis. Molecular motors of the cytoskeleton fall into three major superfamilies: the myosins, the kinesins, and the dyneins. Using ATP hydrolysis as a source of energy, the motors operate by moving specified cargo, both vesicular and non-vesicular, through the cytoplasm along tracks defined by actin microfilaments (MFs), in the case of myosin, or tubulin-based microtubules (MTs) for kinesins and dynein. In addition, the motors can move MFs or MTs themselves, together with intermediate filaments, to change cell shape or to produce cell movement. In some instances, molecular motors either by themselves or via accessory proteins may be able to cross-react with both MFs and MTs or to influence both systems. Myosins and kinesins, which are ATPases related to the small G-proteins in structure, are found in large superfamilies with a variety of molecular configurations, distributions, polarities of movement, and functions. In contrast, while there are perhaps as many as 25 different dynein isoforms, only one of these, cytoplasmic dynein 1

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

(DYNC1), functions specifically in the cell cytoplasm; the rest, including cytoplasmic dynein 2 (DYNC2), are associated with cilia. Cytoplasmic dynein is homodimeric, comprised of two identical ATPase isoforms attached by their cargo-bearing tails. Unlike kinesin or myosin, cytoplasmic dynein, like other dyneins, is an AAA type protein, whose ATPase activity depends on a conformational switch in the six-membered ring of each heavy chain, and whose MT attachment site is a rigid coiled coil. In the hepatocyte, Novikoff et al. [2] described the distribution of MF and MT cytoskeletons in some detail. The actin cytoskeleton mainly lies immediately subjacent to the cell membrane. It is also the cytoskeleton of the microvillus core of the bile canalicular surface and of the adherens junction of the hepatocyte apical surface, encircling the canaliculus. Within the cytoplasm proper, there are few actin MFs; instead the cytoplasm is filled with intermediate filaments (cytokeratin) and MTs that focus on the centrosome, the amorphous organelle surrounding a pair of centrioles, normally located in the cell center, just adjacent to the nucleus. In the cholangiocyte [3] and in quiescent fibroblasts, the centrosome is found at the apical surface of the cell, where a primary cilium grows from the older of the two centrioles. In general, this cytoskeletal distribution implies that interactions between the cell surface and the underlying cytoplasm are first mediated through the actin cytoskeleton and cargos moved by myosins, while traffic through the bulk of the cytoplasm toward or away from the cell center involves the MT cytoskeleton and cargos moved by kinesins or by cytoplasmic dynein. In addition, MT-based molecular motors are probably responsible for the overall spatial distribution of cell organelles such as the Golgi apparatus, which flies

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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THE LIVER: THE MYOSINS

apart when the MT cytoskeleton or dynein-based movement are disrupted. MT molecular motors are primarily responsible for building or resorption of the primary cilium, where this occurs. In cells of the liver which divide, the distribution of the cytoskeleton and the types and functions of molecular motors of the cell change as the cell enters its division cycle. Kinesins and cytoplasmic dynein are important for spindle assembly and MT capture by chromosomal kinetochores, for producing a balance of forces that assists chromosome alignment at the metaphase plate, and for chromosome movement at anaphase [4]. Myosin 2 acting on the contractile ring of actin MFs is essential for cytokinesis [5]. Both actin MFs and tubulin-based MTs are built of polarized subunits, such that the two ends of the polymer are not equivalent. Within the cell cytoplasm, MFs are usually arranged with their fast polymerizing “+” barbed ends attached to the cell membrane and their pointed “−” ends extending into the cytoplasm. MTs have their fast growing “+” ends near the membrane and their slow growing “−” ends usually embedded in the centrosome. In the absence of cell movement, although there is some dynamic behavior in both the MF and MT cytoskeletons, the overall arrangement is largely stable. Generally, each molecular motor is also polarized to move in one direction along the polymer, either moving material into the cell center or out from the edge of the nucleus to the cell surface. In a few motors, the direction of movement can potentially be regulated and reversed by signaling molecules or perhaps by load. Structural biology techniques including X-ray diffraction, NMR, and cryoelectron microscopy have been employed to define the arrangement of subunits in MFs and MTs at atomic resolution. Similarly, the detailed tertiary structure of individual isolated motor molecules has been studied. Combined images indicate how motors dock on the cytoskeletal elements, and biophysical studies visualizing single motors dynamically walking along MFs or MTs have confirmed step size, direction of movement, and processivity of selected motors. While the relationship of some molecular motors to the cytoskeleton has been extensively studied, much less is known about the interactions of motors and cargos. As detailed below, physiologically, cargos such as an endosome interact successively with several different motors during their travel through the hepatocyte. The mechanisms by which an orderly and useful succession is achieved are largely unknown, but there is considerable specificity and cellular control, for example by interactions of cargo and motors with various small cytoplasmic G proteins. In general, the cargo-binding ends of the motors lie at the opposite ends from the ATPase sites; often the heavy, higher Mr, ATPase chains of the motor are bound to light chains, some of which modulate heavy chain activity, some of which bind cargo.

Table 4.1 Molecular motors of the hepatocyte Molecular motor Myosins I II V VI VII IX Kinesins 1 2 3 14

Myo1b (130 kDa) Myo1c (110 kDa) Myo1e Myo2a Myo2b Myo5b Myo6 Myo7a Myo9b KIF5B KIF3A KIF1B KIFC1 KIFC2

Dyneins DYNC1 DYNC2 Original literature references to these motors in hepatocytes under older designations are given in [1], except that Myo 2b is described in [6], Myo 6 is described in [7], KIF3A and KIF5B are shown in [8], KIFC1 is shown in [9].

Previously, this review of molecular motors [1] detailed the motors that were present in the hepatocyte—and by extension in other cells of the liver. Table 4.1 updates this list. As can be seen, there are many different motors in the cytoplasm of a single cell; each has a specific role in the physiology of the hepatocyte. Some properties of key motors are detailed in Table 4.2. The motors are matched to their cargos and they sometimes can be shown to act in sequence, but sequential targeting and release and activation/inactivation mechanisms for each motor are generally not well understood. Some relevant aspects of specific motors are discussed in the next sections. For additional descriptions, see the dynein, kinesin and myosin home pages http://www.dynein.leeds.ac.uk/, http://www.proweb.org/kinesin/ and http://www.mrc-lmb. cam.ac.uk/myosin/myosin.html.

THE MYOSINS There are now some 20 different classes of myosin, each with distinct structures and functions, of which at least 12 classes are present in humans, encoded by about 40 myosin genes. The functional diversity of the myosins is reviewed in Krendel and Mooseker [6]; the terminology for the classes used in that review (Myo1, 2, etc.) is adopted here. See also Berg et al. [10] and Coluccio [11].

Myo2 Conventional myosin, the myosin of striated muscle, is Myo2, a two-headed molecule. Each myosin head

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47

Table 4.2 General properties of some typical molecular motors Duty phase ratio Force per ATP Cytoskeletal partner Step Size Speed in vitro Polarity

Myo2b

KIF5B

DYNC1

0.05 5–8pN Actin microfilament 4–5 nm 0.25–8 µm s−1 “+” end

0.85 5–8pN MT 8 nm 0.8 µm s−1 “+” end

0.6 ∼7pN MT 8 nm 1.25 µm s−1 “−” end

Duty phase ratio is time of strong attachment and force-generating phase (ts) divided by total time of one mechanochemical cycle (tc). Speed in vitro taken from Howard [27]. ATP: adenosine triphosphate; MT: microtubule.

contains a motor domain that binds to actin and hydrolyzes ATP to walk along the actin MFs toward the “+” (barbed) end. The coiled-coil tails of the Myo2 molecule permit polymerization and self-assembly of Myo2 into a thick bipolar filament. Within cells, actin filaments of opposite polarity slide toward each other by the action of the oppositely directed heads at the two ends of the thick filament. In non-muscle cells, such as the hepatocyte, Myo2 forms short bipolar filaments that are responsible for sliding of actin filaments of the contractile ring at cytokinesis. Activity is regulated by light chain phosphorylation. In hepatocytes, cytokinesis is often incomplete during post-natal liver growth because recruitment of Myo2 to the cortex is impaired [12]. In non-dividing hepatocytes, Myo2 is localized to the actin filaments supporting the bile canaliculus, equivalent to the adherens junction MF belt of other epithelial cells. There are at least two human isoforms of Myo2: Myo2a is a low duty ratio form associated with rapid contraction, while Myo2b is a high duty ratio form associated with maintenance of cortical tension. Both forms may be expressed in a single cell and both are expressed in liver.

Myo1 Myo1 is a single-headed molecule with a short neck and basic tail. The neck binds light chains including calmodulin and function is Ca2+ regulated. The Myo1 family illustrates the expansion of myosin function to the control of organelle position and movement with the regulation of activity via multiple cell signaling cascades. There are now at least five and probably seven different MYO1 genes (MYO1 A–H) in humans [13], at least three of which are expressed in liver. Myo1a is a 110 kDa brush border myosin, well characterized in rat liver, mainly associated with Golgi and plasma membranes [14]. Myo1b is a 130 kDa protein involved in the distribution and movement of endosomes and lysosomes in the cell [15, 16]. It is capable of cross-linking actin filaments, presumably by virtue of a second actin-binding site. Like other myosins involved in vesicular trafficking, it can act as an intracellular mechanosensor, sensing load of vesicular cargo or cortical tension by changing its duty cycle

time of attachment to actin [17] Myo1e contains an SH3 domain and may stabilize adherens junctions and operate in “purse string” contractions at the bile canaliculus [18].

Myo5 and Myo6 These are two-headed myosins important in organelle transport in the cell that act similarly but in opposite directions. Myo6 is a processive “−” end-directed motor, whose processivity is Ca2+ regulated, which would move vesicles along actin MFs from the cell surface toward the cell center. It localizes to clathrin-coated vesicles and vesicles that lose their clathrin coat and is probably important in receptor-mediated endocytosis [7], possibly including uptake of transferrin in the liver. Myo6 may allow endosomes to transverse peripheral actin meshworks at the cell cortex. Similarly, Myo5 is a processive “+” end-directed motor with several isoforms whose role in vesicle trafficking and organelle transport is well established [19]. Myo5-associated organelles often interact with MT motors as well, such that Myo5 activity may counteract or regulate faster MT-based motility. Myo5 is targeted to specific organelles by tail interaction with specific Rabs. Myo5b is associated with recycling endosomes, including recycling of the transferrin receptor [20]. Rab11a and Myo5b are required for bile canalicular formation in WIF-B9 cells, a polarized epithelial cell line [21]. More generally, Myo5b seems responsible for microvillar formation in cells of the digestive tract, especially enterocytes, where mutations in the protein lead to microvillus inclusion disease (MVID), characterized by lack of microvilli on the apical cell surface [22].

Myo7a and Myo9 Myo7a, a two-headed myosin, has been found in embryonic liver and other epithelial cells. The Myo7a tail contains a FERM domain (a specific amino acid sequence related to proteins such as ezrin), which presumably mediates binding to the cytoplasmic side of transmembrane proteins and may help organize adherens junctions. Myo9 is a single-headed myosin that contains a tail domain with homology to the GTPase-activating proteins (GAPs) of

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THE LIVER: CYTOPLASMIC DYNEIN

the Rho family of small G-proteins. It would seem that Myo9 is involved in attaching to actin filaments to organize them in relation to membranes under specific signal transduction conditions, mimicking the effects of Rho.

530-kD cytoplasmic dynein heavy chain MT

CYTOPLASMIC DYNEIN 74-kD intermediate chain

Cytoplasmic dynein is found in two distinct forms, both of which are present in liver: cytoplasmic dynein 1, a ubiquitous molecule of all eukaryotic cells, designated DYNC1, and cytoplasmic dynein 2, DYNC2, a specialized form that functions in ciliogenesis (of which more below) [23].

55-kD light intermediate chain Glued

Dynamitin

p150 p135 Glued p62

Capping protein

Spectrin Ankyrin

DYNC1

Arp1, Actin Vesicle

DYNC1 is a complex molecule composed of two homodimeric ∼530 kDa heavy chains each consisting of an AAA subunit head with a protruding stalk and tail, together with two intermediate chains (about 74 kDa), two light-intermediate chains (53–59 kDa), and light chains (10–14 kDa) of at least three different protein families: TcTex 1 (T), Roadblock (RB), and LC8, also found in several axonemal dyneins [24]. The TcTex 1 family is a group of proteins first identified in mice, where they are encoded by the t locus. RB is a Drosophila protein that affects MT-based transport in that organism. LC8 is a common stoichiometric constituent of DYNC1 and Myo5. It has a direct association with a form of nitric acid synthetase (NOS). One unidentified light chain of cytoplasmic dynein in hepatocytes is probably orthologous to a regulatory light chain of 22S axonemal dynein [25]. Other light chains of interest, present on certain axonemal dyneins, which could also be present in some isoforms of DYNC1, are a Ca2+ binding protein such as centrin, and thioredoxin, which could regulate dynein in relation to redox state and disulfide bond formation. Combinatorial possibilities of the standard heavy chains with different intermediate, light-intermediate, and light chains may give rise to molecular diversity and functional specificity. The coiled-coil stalks on the AAA heads attach to cytoplasmic MTs on which the dynein walks. Cryo electron microscopy of attachment using an isolated single-headed DYNC1 to bind to MTs shows the stalk extending perpendicular to the MT with the head in various configurations [26]. ATP hydrolysis produces a conformational change, swinging the head and tail in relation to the stalk to produce a “−” end-directed movement [28, 29]). Cytoplasmic dynein is usually depicted as a two-headed bouquet with the motor domains protruding from a base containing the heavy chain tail together with intermediate and light chain components (Figure 4.1), which themselves bind to specific targets that are transported as cargo [22, 30], but the in vivo picture may be more compact. DYNC1 transports vesicles such as endosomes

Figure 4.1 Proposed organization of DYNC1 and dynactin in relation to vesicle trafficking. In this model, the actin base of dynactin is integrated into the membrane skeleton of the vesicle. Dynein binds to the dynamitin subunits of dynactin and both dynein and dynactin interact with the MT. The substructure of the DYNC1 heavy chain is indicated. Modified from Hirokawa [30]

along the MT cytoskeleton. DYNC1 is also the motor for ribonucleoprotein particle (RNP) transport along MTs, which is important for embryonic development and cell differentiation [31]. Based on single molecule studies, two-headed cytoplasmic dynein is a processive motor [32] whose duty ratio is not as high as in Kin5B but considerably higher than in Myo2b or 22S axonemal dynein. The mechanochemical cycle of DYNC1 has been studied in detail [33]. DYNC1 primarily advances by 8 nm steps, although other step sizes are seen and there are occasional backwards steps. Continuous motion at speeds measured for MT gliding probably requires the action and cooperativity of multiple molecules. Only a few motors can be bound to a vesicle or RNP particle being transported at any one time, and there is always the possibility that these could detach. For DYNC1 this possibility is minimized in the cell by the presence of dynactin [34, 35]. Dynactin is a 20S multisubunit protein complex that binds to and co-localizes with DYNC1. It is essential for the function of cytoplasmic dynein in organelle transport. It increases the processivity of the dynein, which allows the molecule to move along an MT for long distances. Its subunit composition and ultrastructure are well characterized (Figure 4.1). The backbone of dynactin is an extended subunit p150Glued attached to a short filament of actin-related subunits (Arp1) and possibly authentic actin, and capped at both ends, the barbed end by the capping protein CapZ, the pointed end by p62. The Arp1 filament interacts with or perhaps forms part of an ankyrin-spectrin

4: MOLECULAR MOTORS

membrane skeleton around vesicular cargo [36, 37]. At its projecting head end, p150Glued binds to MTs, while the coiled-coil region of the protein interacts with LC74 of DYNC1. A protein of Mr 50 kDa (p50), called dynamitin, connects the p150Glued arm to the Arp1 filament. Overexpression of dynamitin disrupts the binding of dynactin-dynein to its cargo [38–40], leading to dispersion of Golgi, lysosome, and late endosomal membranes to the cell periphery [41, 42], and disruption of endoplasmic reticulum (ER)-to-Golgi transport [43]. This suggests that DYNC1 is the effective “−” end motor of these membranes, working against a “+” end motor to achieve appropriate positioning and stability of the Golgi–lysosome complex in the cell center. Early endosomes and receptor recycling are unaffected [42]. As I discuss below, an in vitro approach specifically validates these findings for the hepatocyte.

49

family (Kinesin13) of M kinesins with the motor domain in the middle of the molecule. There are many different kinesins present in the cytoplasm of epithelial and connective tissue cells. Although only a few of these have been specifically reported to be present in various liver cells, there is reason to believe that many are present and functional, at least in small amounts at certain times. A functional analysis of human kinesins using RNA interference procedures to identify function revealed that, in addition to vesicular trafficking, at least 12 different kinesins, members of the Kinesin3, 4, 6, 7, 8, 10, 13, and 14 families, are involved in mitosis and cytokinesis [4]. Kinesins are also involved in the transport of intermediate filaments and their organization in the cytoplasm [48], which supports a role for kinesin in overall organization of cytoskeleton and membrane positioning in the cell.

KIF5 (Kinesin1 family) DYNC2 DYNC2 is assembled together with about 20 other cytoplasmic proteins, but not dynactin or dynamitin, into complexes at the base of primary cilia, where such are present on many types of liver cell—for example, cholangiocytes and so on, as mentioned above—although not normally on hepatocytes. DYNC2 is then transported passively along the axonemal MTs of the cilium by active kinesin motors toward the ciliary tip in a complex with these protein carriers and cargos such as membrane receptors and proteins that build and maintain axonemal structure in a process known as “intraflagellar transport” (IFT) [44]. Kinesins, along with receptors, proteins of the IFT complexes, and cargos, are brought back to the cell body by active DYNC2. This process as it applies to cholangiocytes is described in detail below.

KINESIN The kinesin superfamily consists of at least 14 families of kinesins, now labeled Kinesin1, Kinesin2, and so on, replacing the previous older mixed nomenclature [45]. There are about 45 kinesin family (KIF) genes in the human genome [46]. However, alternative splicing can produce several mRNAs for each gene, with different tail domains that bind to different cargos, giving rise to perhaps 90 or more KIF proteins. The role of kinesins in intracellular transport has been reviewed recently by Hirokawa and Noda [47]; molecular notation here follows this review. The globular motor domains of all members of the kinesin superfamily are similar, but their positions in the molecule vary. Most of the kinesins possess an N-terminal motor domain and are anterograde transporters, moving cargo toward the “+” ends of MTs, but there is one family (Kinesin14) of C-terminal motor domains which function as “−” end transporters and one

The original kinesin (conventional kinesin) is Kinesin1. It is a dimeric molecule with an N-terminal motor domain, identified as the major “+” end anterograde transporter of vesicles along axonal MTs. There are three conventional kinesin1 family (KIF) genes in mammals, one of which—KIF5B—is found in most cells, including hepatocytes, while the others, like KIFs of other kinesin families, are mainly expressed in the brain. KIF5 heavy chain binds light chains (KLC) through domains in its stalk and tail, and in turn the stalk or the KLCs bind specific cargos. Targeting of KIF5 to particular vesicles is usually via the KLCs, generally with the aid of adaptor proteins. In epithelial cells, kinesin is often localized to the region of the Golgi apparatus. KIF5 functions in Golgi-to-ER recycling, lysosomal and endosomal movement (as will be described below), perhaps in other exocytic events, and in mitochondrial movement. Other kinesins, mainly of the monomeric Kinesin3 and heterotrimeric Kinesin2 families, complement KIF5 as the primary anterograde motors in intracellular trafficking. KIF5 is also a transporter of mRNA. Conventional kinesin (KIF5), like cytoplasmic dynein, is an MT-activated ATPase. However, KIF5 differs from DYNC1, not only in the direction of force generation along the MT, but also in its duty ratio (Table 4.2) and increased processivity. The mechanochemical cycle of kinesin is such that the molecule remains attached to the MT for over 80% of a cycle [27]. Effectively, a dimeric kinesin like KIF5 never detaches completely from the tubulin lattice, and there is no need for a dynactin-like component. A kinesin like KIF5 walks along an MT with a head-over-head action in a straight line along an MT protofilament in 8 nm steps for about 100 steps at a time.

KIF3A (Kinesin2) The Kinesin2 family consists of homodimeric (KIF17) and heterotrimeric kinesins (KIF3A, 3B). The two

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THE LIVER: EXAMPLES OF MOTOR MOLECULE FUNCTION IN LIVER CELLS

different kinesins of the heterotrimeric complex are capped by a non-motor globular subunit (KAP) at their tail ends. These are “+” end motors involved in vesicular trafficking and/or in movement of non-vesicular membrane protein rafts or axonemal components along the ciliary axoneme in IFT (see below). Members of the Kinesin2 family are also candidates for motors that transport mRNAs through the cytoplasm.

KIF1B (Kinesin3) The Kinesin3 family consists of monomeric and dimeric kinesins, including KIF1B, a ubiquitously expressed monomeric kinesin found in liver, associated with “+” end-directed transport of mitochondria and KIF1C, a dimeric kinesin associated with ER-to-Golgi transport. Although monomeric KIF5 is not an effective motor since it is freely released by ATP addition, KIF1B is capable of moving along MTs and transporting cargo, probably by utilizing an electrostatic interaction to maintain continuous contact with the MT [49].

KIFC1 and KIFC2 (Kinesin14) C-type kinesins such as KIFC1 and KIFC2 are “−” end-directed motors that move endosomes or multivesicular bodies in various cells in place of DYNC1. KIFC1 and KIFC2 are associated with and move early endosomes in liver hepatocytes of various species (see below).

EXAMPLES OF MOTOR MOLECULE FUNCTION IN LIVER CELLS Endocytic trafficking in the hepatocyte The cytoskeleton of the hepatocyte is arranged so that endosomes on their way to lysosomal fusion must penetrate a cortical actin-based network before arriving in the vicinity of a microtubular umbrella that funnels toward the centrosome around which the lysosomes and Golgi membranes are distributed [2]. They reach the center of the cell by motor molecule-directed movement, certainly moving along MTs for most of the distance, but also probably originally along actin filaments using myosins (Figure 4.2), as do clathrin-coated endosomes in lymphocytes [50]. However, to traverse the maximum ∼20 µm distance from surface to nucleus along an MT takes perhaps 30 minutes, while motor-based transport along MTs is about 1 µm s−1 . Consider now what would happen if KIF5 and DYNC1, or more generally a “+” end-directed motor and a “−”

Figure 4.2 A proposed model of motor involvement in receptor-mediated endocytosis in hepatocytes. Movement of the clathrin-coated vesicle through the actin belt might require Myo6 (MVI). The uncoated acidified vesicle would reach the MT cytoskeleton, attach and begin to undergo fission upon hydrolysis of Rab4-GTP, activating KIFC2 (or KIFC1) and producing a ligand-rich daughter and a receptor-rich daughter pulled by KIF5B (K) to begin recycling. As maturation proceeded after repeated fissions, the late endosome would replace the “−” end kinesin with dynactin (Dy) and DYNC1 (D) for the journey to the lysosome, and the “+” end kinesin with KIF3A. The recycling vesicle would pass through the actin belt aided by Myo5 (MV). Several types of motor are present on a single vesicle, and differential activity can be controlled by cell signal pathways, including phosphorylation of the motor subunits. Based on work in [8, 9, 51, 52]

end-directed motor, both worked on the same MT. Vale et al. [53] showed that in vitro MTs will translocate first one way, then the other. When movement occurs with the “−” end of the MT leading, the kinesins are active, the dyneins inactive; when the “+” end is leading, the reverse is true. Switching between the directions occurs randomly, but for equal timing there needs to be about an order of magnitude more dynein on the substratum than kinesin. For vesicular trafficking, this difference might be largely overcome by dynactin. In vitro and in the hepatocyte, vesicles move back and forth in this manner, contributing to the increased time of transit to the cell center. With a “−“ end kinesin substituting for dynein, the numbers of molecules involved and the switching will depend on the specific properties of the motors. Kinesin is largely inhibited by the non-hydrolyzable ATP analog AMP-PNP and little affected by 5 µM vanadate, while movement of cytoplasmic dynein is largely unaffected by AMP-PNP but stopped by vanadate [54]. This difference provides a convenient initial assay to determine whether a given movement along MTs is kinesin- or dynein-based.

4: MOLECULAR MOTORS

An in vitro approach has proven very useful in studying the MT-based molecular motors of hepatocytes in vesicular trafficking in receptor-mediated endocytosis, specifically of fluorescently-labeled asialo-orosomucoid (ASOR), and the details have yielded some insights that are probably usefully generalized to other cell types and other ligand-receptor pairs [8, 9, 51, 52]. Endosomes are normally prepared from rat liver. About a quarter of the labeled endosomes attached to MTs move in the presence of ATP at average rates of about 0.7 µm s−1 , with about half moving in a “+” end direction, half in a “−” end direction. Motility is unaffected by addition of exogenous motor molecules, suggesting that the purified endosomes are saturated with motor molecules and that only a small number of the motors are required for motility. If the endosomes are prepared five minutes after IV injection of labeled ASOR, the preparation is enriched in early pre-segregation endosomes and virtually all of the labeled endosomes that bind to MTs also contain receptor [51]. About 10–15% of the moving endosomes undergo fission, which results in segregation of ligand and receptor into two daughter vesicles, one enriched in ligand relative to receptor, and the other containing about 80% of the original receptor. Movement is unbiased, with about 50% of vesicles moving to the “+” end of the MT and about 50% to the “−” end. Movement in both directions and all fission events are abolished by AMP-PNP, even in the presence of ATP, but are not affected by vanadate, suggesting that these processes are independent of DYNC1 but dependent on kinesins. In rat hepatocytes, the “+” end motor was identified as Kinesin 1 (later as KIF5B), while the “−” end motor was KIFC2. In a further study using in vitro observations and cultured mouse hepatocytes, Nath et al. [9] showed that over 90% of early endosomes of mouse hepatocytes were associated with both “+” and “−” end kinesins. Although the “+” end motor again was KIF5B, surprisingly the “−” end motor was KIFC1, not KIFC2, as previously shown for rat liver. The two motors on a single vesicle interact with one another. Fission was reduced when either motor was inhibited, indicating that opposing forces from both motors produce a “tug of war” that promotes fission. An initial analysis revealed that dynamin and the small G-protein Rab4 were also present on a large percentage of early endosomes [52]. These proteins are part of the membrane scaffold that together with the kinesins probably act on the early endosome to regulate and produce fission. In particular, when exogenous Rab4 is added together with GTPγS, a non-hydrolyzable analog of GTP, or GTPγS is added to endogenous endosomal Rab4, motility and fission are suppressed, while addition of GDP enhances both motility and fission. A careful analysis reveals that while GDP has little effect on “+” end motility, “−” end motility of the early endosomes is doubled. This suggests that when the endosome forms, Rab4-GTP incorporated into the scaffold near or directly

51

interacting with the kinesins initially inhibits motor activity. Hydrolysis to Rab4-GDP (with the aid of a GAP?) increases the activity of the “−” end kinesin, perhaps stretching the vesicle to facilitate fission (by the action of dynamin?). Action of Rab-GDI (guanine nucleotide dissociation inhibitor) then releases the Rab4 from the scaffold, whereby both “+” and “−” end kinesins become equally active. Presumably, in the hepatocyte, cytosolic Rab4-GDP would interact with a guanine nucleotide exchange factor (GEF) to reform Rab4-GTP, which perhaps would rebind to an unfissioned endosome, for another cycle of the sorting process. Bananis et al. [8] used fluorescent activated cell sorting to purify the early endosomes and also to study a corresponding fraction of late endocytic vesicles prepared from rat livers 15 minutes after injection of labeled ASOR. Both preparations were subject to in vitro motility and immunoblot analysis and the similarities and differences are instructive in terms of molecular motor activity. Both vesicle populations move equally well on MTs, but many more of the early endosomes undergo fission. Consistent with the early endosomes being pre-segregation endosomes and the late endosomes being post-sorting endosomes, the early endosome fraction is enriched in asialoglycoprotein receptor (ASGPR) and Rab4, unlike the late endosome fraction. While movement of early endosomes is blocked only by AMP-PNP and not by vanadate, movement of late endosomes is sensitive to both vanadate and AMP-PNP. In particular, vanadate blocks “−” end movement of late endosomes, supporting the hypothesis that as the endosome matures, KIFC is replaced by DYNC1 as the “−” end directed motor. This is confirmed by immunoblot, which shows that in addition to DYNC1 heavy chain and DYNC1 intermediate chain, the p150 subunit of dynactin is found on late but not early endosomes. In addition, the binding of late but not early endosomes to MTs is affected by dynamitin. The “+” end motor is also different between early and late endosomes. In early endosomes, the predominant “+” end kinesin is KIF5B, which is virtually absent from the late endosome preparation, where the “+” end motor is KIF3A, of the Kinesin2 family. Rab7 is also present on late but not early endosomes. These results indicate that as the endosome matures, its entire scaffold is reorganized so that motors and control molecules are replaced, functioning in specific sequence, as shown in Figure 4.2. Clearly, in hepatocytes, multiple motors are present on a single vesicle, and they are assembled and disassociated at various times for interactions with other scaffold molecules and with the cytoskeleton. The motor toolbox is utilized to carry out defined physiological tasks, presumably under the control of cellular signal transduction systems, placing cellular components in a localized functional space for further molecular interactions. It is reasonable to speculate that the cellular signals and controls that affect DYNC1 are different from those acting on KIFC and that therefore different motors

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THE LIVER: EXAMPLES OF MOTOR MOLECULE FUNCTION IN LIVER CELLS

moving vesicles in the same direction might be useful at different points in endosome maturation. Nevertheless, the molecular tools, for example “−” end kinesins, are redundant, so that the nature of the toolbox is evolutionarily imprecise and different for even closely related organisms. The precise cellular rationale for different motor substitutions during a process such as receptor-mediated endocytosis remains largely unknown. However, as the studies cited indicate, much can be learned about which of various molecular motors acts in the hepatocyte and when and where they act, which deepens our understanding of the role of these motors in liver physiology.

Molecular motors of the cholangiocyte primary cilium Many cells of the liver, but not hepatocytes (except under pathological conditions—for example [55]), grow primary cilia. In contrast to the well known 9 + 2 motile cilia of, for instance, the respiratory tract, primary cilia are single, non-motile cilia consisting of an axoneme of nine doublet MTs without a central pair, hence 9 + 0, and missing the axonemal dyneins responsible for motility [56]. Initially from studies of ciliogenesis in the single-celled organism Chlamydomonas [44], it has become clear that cells such as cholangiocytes, fibroblasts, kidney epithelium, human embryonic stem cells, and neurons utilize the primary cilium as a sensory antenna to detect environmental signals including mechanical distortion, various growth factors and hormones, and osmolarity. Important receptors for these processes become localized to the cilium. Maysuk, LaRusso, and their collaborators provide particular insights into cholangiocyte primary cilia. In normal cell lines of rodent cholangiocytes, primary cilia appear at day 3 post-confluence and grow to 7–10 µm length. By day 10, the majority of cells possess 9 + 0 cilia. Cholangiocyte primary cilia project from the cells into the lumen of the bile duct and are thus positioned to detect changes in bile flow, composition, or osmolality. Masyuk et al. [3] suggest that receptor proteins localized to the cilia function for each of these modalities, such that cholangiocyte cilia are mechanosensors, chemosensors, and osmosensors. However, the cilia are heterogenous in length along the biliary tree and may also be functionally heterogenous in small and large bile ducts. Mechanosensing depends on localization of polycystins PC-1 and PC-2 to the cilia. The polycystin Ca2+ channel opens in response to bending of the cilium by flow. The mechanism of mechanosensation is identical to that in the kidney, except a different fluid flow is being detected, and Tg737 mice that are missing functional primary cilia develop polycystic liver disease, as well as polycystic kidney disease. For osmosensing, a transient receptor potential channel, TRPV4, becomes localized to the cilia [57]. Activation of this channel by hypotonicity leads to an increase in intracellular Ca2+ , which in turn affects ductal

bile secretion. Chemosensory function in cholangiocyte cilia involves the purinergic membrane receptor P2Y12 and adenylyl cyclase localized in the ciliary membrane. Biliary ATP and ADP act on the receptor to cause changes in ciliary cAMP mediated through an A-kinase anchoring protein (AKAP) complex. Potential signaling pathways are diagrammed in Figure 4.3a. The primary cilium forms during G0 phase of the cell cycle, when the cell is quiescent and non-dividing and there are two centrioles present. The MTs of the axoneme extend from a basal body, which is a mother centriole that becomes attached to the cell membrane by a ciliary necklace. They are polarized so that their “+” ends are nearest the growing tip of the cilium. Molecular motors are necessary to build the axoneme and to place and position the appropriate receptors in the ciliary membrane. As mentioned previously, the motors are members of the Kinesin2 family; “+” end motors which move cargo from the basal body to the growing tip in anterograde transport; and DYNC2, a “−” end cytoplasmic dynein responsible for retrograde transport, moving material out of the cilium for signaling, recycling, or degradation. The kinesin transporters include both heterotrimeric and homodimeric Kinesin2, the former KIF3A, 3B, and KAP, the latter KIF17, which move different cargos in IFT. In some cases it has been shown that the heterotrimeric molecule transports cargos that build the axoneme, while the homodimeric kinesin moves cargos to the tip of the cilium. There may be other kinesins and perhaps myosins that transport specific cargos, particularly membrane receptors or channels, in certain cilia. The motors are attached to two protein complexes, A and B, consisting of at least 16 proteins. Complex B includes the well-studied IFT protein, IFT88 (Polaris); this complex with kinesin is probably responsible for active anterograde transport, while complex A with dynein is responsible for retrograde transport. The loading zone for IFT complex, motor, and cargo assembly is the ciliary necklace region. The loading and motor activation processes are not well understood. It is important to realize that the anterograde complex, upon loading, has activated kinesin moving along the axonemal MTs but is passively transporting dynein, while at the cilium tip, dynein with complex A becomes activated while kinesin and complex B are turned off and carried passively back to the cell, as shown in Figure 4.3b. The complex A and B proteins, the motors and transport mechanisms, are highly conserved throughout evolution; and human as well as mouse orthologs of cloned Chlamydomonas proteins for the IFT components have been defined. Knockout of kinesin or IFT88 (as in the Tg737 mouse) results in defective ciliogenesis, leading to very short or absent cilia; knockout of dynein or complex A proteins leads mainly to shortened cilia with bulbous tips containing complex B proteins. A more complete review of IFT proteins and motors in the assembly of cilia is found in Pedersen et al. [58]. Because receptors and specific channel proteins must be localized

4: MOLECULAR MOTORS

53

3. P2Y12

4.

ATP/ADP

AC6 cAMP

5. Ca2+

TRPV4 FC Ca2+

retrograde IFT

Bile tonicity

ATP

anterograde IFT

Bile flow

PC-1 PC-2

cytoplasmic dynein 1b/2 2.

FCCT

kinesin-II

Ca2+

complex A BA

nucleus

AB

complex B cilia precursor

ER 1.

(a)

6.

cilia turn over product

(b)

Figure 4.3 Molecular motors of the cholangiocyte primary cilium. (a) A working model of sensory functions of the cholangiocyte primary cilium. Receptors (e.g. P2Yα) and channels (e.g. PC2, TRPV4) are placed in the ciliary membrane by IFT. They respond to bile flow and tonicity and extracellular ligands by changing second messenger concentrations and signaling molecules within the cilium. Signal transduction pathways lead to secretion and to gene activation within the nucleus. Reproduced from [3] with permission. Courtesy of A. Masyuk and Develop. Dynamics (Wiley-Liss). (b) A model of IFT, applicable to cholangiocyte cilia. Anterograde IFT (Steps 1–3) relies on active Kinesin2. Step 1: Loading at the ciliary necklace. Step 2: Movement along the axonemal MTs. Step 3: Dissociation. Retrograde IFT (Steps 4–6) relies on active DYNC2. Step 4: Reassembly at the cilium tip. Step 5: Movement along axonemal MTs. Step 6: Dissolution. Reproduced from [58] with permission. Courtesy of L. Petersen and Develop. Dynamics (Wiley-Liss).

to the cilium to signal properly in control of bile secretion and resorption by cholangiocytes, but also more generally to control cell division, cell migration, apoptosis, and ultimately tissue development and function, defects in the cilium or in receptor localization lead to important developmental defects and to ciliopathies [59], including polycystic liver disease [60].

CONCLUSION Multiple molecular motors functioning in the cytoplasm of various liver cells, including hepatocytes and cholangiocytes, and indeed almost every cell type in the body, are necessary for membrane trafficking, organelle placement, general cell organization, and signal transduction processes that occur in normal developmental and homeostatic events. The motors become part of larger protein complexes that bind to and transport molecules, particularly receptors and channels into specific membranes. The compositions of such complexes and scaffolds, and of the cargos transported, is highly dynamic. Motors on the same membrane can act antagonistically to one another, and there is evolutionary and perhaps some functional flexibility, but the net result is that molecular motors are the powerhouses of the biological nanomachines that operate

cell physiology, including major aspects in the control of liver biology in health and disease.

ACKNOWLEDGMENTS This work was supported in part by grants DK41918 and DK41296. I thank Shailesh Shenoy for help with the plates and tables.

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43. Presley, J.F., Zaal, K.J.M., Schroer, T.A. et al. (1997) ER to Golgi transport visualized in living cells. Nature, 389, 81–85. 44. Rosenbaum, J.L. and Witman, G.B. (2002) Intraflagellar transport. Nat Rev Mol Cell Biol , 3, 813–25. 45. Lawrence, C.J., Dawe, R.K., Christie, K.R. et al. (2004) A standardized kinesin nomenclature. J Cell Biol , 167, 19–22. 46. Miki, H., Setou, M., Kaneshiro, K. and Hirokawa, N. (2001) All kinesin superfamily protein, KIF, genes in mouse and human. Proc Natl Acad Sci USA, 98, 7004–11. 47. Hirokawa, N. and Noda, Y. (2008) Intracellular transport and kinesin superfamily proteins, KIFs: structure, function, and dynamics. Physiol Rev , 88, 1089– 118. 48. Prahlad, V., Yoon, M., Moir, R.D. et al. (1998) Rapid movements of vimentin on microtubule tracks: kinesin-dependent assembly of intermediate filament networks. J Cell Biol , 143, 159–70. 49. Okada, Y. and Hirokawa, N. (2000) Mechanism of the single headed processivity: diffusional anchoring between the K-loop of kinesin and the C-terminus of tubulin. Proc Natl Acad Sci USA, 97, 640–45. 50. Salisbury, J.L., Condeelis, J.S. and Satir, P. (1983) Receptor-mediated endocytosis: machinery and regulation of the clathrin-coated vesicle pathway. Int Rev Exp Pathol , 24, 1–62. 51. Bananis, E., Murray, J.W., Stockert, R.J. et al. (2000) Microtubules and motor dependent endocytic vesicle sorting in vitro. J Cell Biol , 151, 179–86.

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52. Bananis, E., Murray, J.W., Stockert, R.J. et al. (2003) Regulation of early endocytic vesicle and fission in a reconstituted system. J Cell Sci , 116, 2749–61. 53. Vale, R.D., Malik, F. and Brown, D. (1992) Directional instability of microtubule transport in the presence of kinesin and dynein, two opposite polarity motors. J Cell Biol , 119, 1589–96. 54. Murray, J.W., Bananis, E. and Wolkoff, A.W. (2000) Reconstitution of ATP-dependent movement of endocytic vesicles along microtubules in vitro: an oscillatory bidirectional process. Mol Biol Cell , 11, 419–33. 55. Sobel, H.J. and Marguet, E. (1983) Zipper-like structure and centrioles in hepatocytes. Ultrastruct Pathol , 4, 115–19. 56. Satir, P. and Christensen, S.T. (2007) Overview of structure and function of mammalian cilia. Annu Rev Physiol, 69, 14.1–14.24. 57. Gradilone, S.A., Maysuk, A.I., Splinter, P.L. et al. (2007) Cholangiocyte cilia express TRPV4 and detect changes in luminal tonicity inducing bicarbonate secretion. Proc Natl Acad Sci USA, 104, 19138–43. 58. Pedersen, L.B., Veland, I.R., Schrøder, J. and Christensen, S.T. (2008) Assembly of primary cilia develop. Dynamics, 237, 1993–2006. 59. Bardano, J.L., Mitsuma, N., Beales, P.L. and Katsanis, N. (2006) The ciliopathies: an emerging class of human genetic disorders. Annu Rev Genomics Hum Genet , 7, 125–48. 60. Masyuk, T. and LaRusso, N. (2006) Polycystic liver disease: new insights into disease pathogenesis. Hepatology, 43, 906–8.

5

Ion Pumps and Molecular Motors: P-, F-, and V-type ATPases Sarah Bond, Daniel J. Cipriano and Michael Forgac Department of Physiology, Tufts University School of Medicine, Boston, MA, USA

P-TYPE ATPASES The P-type ATPase family transports a variety of substrates across biological membranes. Its name derives from a phosphorylated intermediate that is generated during the reaction cycle. In addition to creating Na+ , K+ , and Ca2+ gradients in cells, P-type ATPases are important for heavy-metal homeostasis and the maintenance of lipid asymmetry in membranes.

P-type ATPase family members and their functions The P-type ATPase family can be classified by sequence homology into five branches, types I–V, and further divided into 10 sub-types based on substrate specificity (Table 5.1) [1]. Type I P-type ATPases include the type IA bacterial K+ -ATPases and type IB soft-metal-ion-transporting ATPases [1]. The Cu2+ -transporting ATPases that are defective in the hereditary and lethal Menkes and Wilson diseases, ATP7A and ATP7B, belong to this class [2, 3]. These pumps play important roles in Cu2+ homeostasis. Following uptake of copper from the intestinal lumen into intestinal epithelial cells via the copper transporter CTR1, ATP7A functions to transfer Cu2+ across the basolateral membrane into the blood. Mutations in ATP7A give rise The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

to Menkes disease, characterized by copper deficiency and problems that result from an inability to provide essential cuproenzymes with copper. In hepatocytes, ATP7B mediates the excretion of excess copper across the apical membrane into the bile, and mutations in this pump lead to Wilson disease. Clinical features of Wilson disease result primarily from toxic accumulation of copper in the liver and brain. Both pumps normally reside on Golgi membranes but undergo translocation to the cell surface to aid in elimination of copper from the cell in response to elevated cytoplasmic copper levels [4]. Type II P-type ATPases are the best-studied class, and include the Na+ /K+ -ATPase as well as the calcium pumps [1]. Calcium pumps lower resting cytosolic calcium levels to the nanometer range [5]. Opening of calcium channels elevates cytosolic concentrations and initiates signaling events that are then terminated by calcium pumps. In muscle cells, cytosolic calcium initiates muscle contraction, and removal of calcium causes relaxation [6, 7]. The sarco(endo)plasmic reticulum Ca2+ -ATPase (SERCA) belongs to the type IIA P-type ATPase subfamily [1]. This pump resides on the sarcoplasmic reticulum (SR) of muscle cells and pumps two Ca2+ ions into the SR lumen per ATP in exchange for two or three protons [8]. Several different SERCA isoforms have been identified, which have various functions [6]. SERCA2a is found in cardiac muscle, and in humans determines the rate of removal of more than 70% of

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

58

THE LIVER: P-TYPE ATPASES

Table 5.1 Class

P-type ATPase family

Sub-type

Substrate specificities

I

A B

Bacterial Kdp-like K+ -ATPases Soft-metal-ion-transporting ATPases

II

A B C D

Sarco(endo)plasmic reticulum (SR) Ca2+ -ATPases Plasma membrane Ca2+ -ATPases Na+ /K+ -ATPases and H+ /K+ -ATPases Fungal Na+ -ATPases

A B

Plant/fungal H+ -ATPases Bacterial Mg2+ -ATPases

III IV

“Lipid flippases”

V

Unknown substrate specificity

cytosolic calcium [9]. SERCA2a activity thus determines the rate of relaxation of the heart as well as the size of the calcium store available for release in the next beat, influencing contractility. SERCA1a is found on the SR of skeletal muscle cells, where it terminates contraction events [10]. A major breakthrough in the field came with the publication of the X-ray crystal structure of SERCA1a, which defined the major structural domains common to all P-type ATPases, and revealed the location of the cation binding sites within the membrane [11]. Mutations in SERCA1a cause Brody disease, an inherited disorder characterized by an exercise-induced delay in muscle-cell relaxation [12]. SERCA2b functions as a “housekeeping” isoform, and mutations in this isoform lead to the skin disorder Darier disease [13]. Type IIB P-type ATPases include the plasma membrane Ca2+ -ATPases (PMCAs), which reside on the plasma membrane of cells and pump calcium from the cytosol into the extracellular space [1]. These pumps are structurally similar to SERCA, except that they contain a longer C-terminal tail that mediates regulation by calmodulin [6, 14, 15]. PMCA is encoded by four different genes, PMCA1–4, which give rise to at least 20 alternatively spliced isoforms. These isoforms have different tissue expression patterns, activities, and calmodulin affinities. PMCA1 has been described as the “housekeeping” pump and, along with PMCA4, may be present in all cells. PMCA4 has the lowest basal activity but shows the greatest stimulation by calmodulin, and a role in shaping Ca2+ signals has been suggested for this isoform. PMCA2 has been implicated in hearing loss, and PMCA3 is present only in neuronal tissues. The Na+ /K+ -ATPase and gastric H+ /K+ -ATPase comprise the type IIC P-type ATPases [1]. The Na+ /K+ ATPase was the first P-type ATPase identified [16]. Located on the plasma membrane, the Na+ /K+ -ATPase pumps three Na+ ions out of the cell in exchange for two K+ ions [17]. This activity is essential for generation of the membrane potential in animal cells. The membrane potential is required for processes such as: nerve conduction; cellular uptake of ions, nutrients, and

neurotransmitters; and regulation of pH and cell volume. For example, the Na+ gradient drives removal of H+ ions from the cytoplasm via the Na+ /H+ exchanger to neutralize cytosolic pH. The Na+ gradient also energizes the uptake of sugars and amino acids across the plasma membrane of intestinal epithelial cells. The Na+ /K+ -ATPase indirectly influences removal of calcium from cells by powering the Na+ /Ca2+ exchanger, an antiporter located on the surface of most cells that exchanges one cytosolic Ca2+ ion for three extracellular Na+ ions [6]. The H+ /K+ -ATPase resides on the surface of cells lining the stomach, where it pumps protons into the extracellular space in exchange for K+ ions. Both type IIC members are important drug targets in humans [18]. The anti-ulcer drug omeprazole, for example, targets the H+ /K+ -ATPase. The Na+ /K+ -ATPase is the target of cardiac glycosides such as Digoxin that are used in the treatment of congestive heart failure [19]. Inhibition of Na+ /K+ -ATPase activity reduces the driving force for the Na+ /Ca2+ exchanger, leading to higher levels of intracellular calcium and a stronger contraction. Endogenous cardiac glycosides have also been identified in mammals. The Na+ /K+ - and H+ /K+ -ATPases exist as heterooligomers, consisting of α- and β-subunits that are often co-expressed with a third regulatory subunit [17, 20]. The α-subunit is homologous to other single-subunit P-type ATPases, contains the ATP and ion binding sites, and mediates Na+ and K+ translocation. The β-subunit is a single transmembrane-spanning glycoprotein that is unique to K+ -counter-transporting P-type ATPases [21, 22]. It is required for routing of the α-subunit to the plasma membrane and occlusion of the K+ ions during the pumping cycle. The α- and β-subunits are often found with a third regulatory subunit that is a member of the FXYD family of single-span transmembrane proteins [20, 23]. In mammals, four α-subunit isoforms of the Na+ /K+ -ATPase have been identified, which are expressed in a tissue-specific manner and perform different functions [24]. The α1-isoform is uniformly distributed in the plasma membrane, is found in every tissue type, and is thought to act as a “housekeeping” isoform. The α2-isoform colocalizes with the Na+ /Ca2+ exchanger on discrete areas of the plasma membrane overlying the SR/ER (endoplasmic reticulum). It has a higher affinity for cardiac glycosides than the α1-isoform and may play a role in muscle contraction by regulating transient Ca2+ levels. Mice expressing half the normal amount of α2-isoform exhibit increased force of contraction in skeletal and cardiac muscle. The α4-isoform is exclusively found in testes and is required for sperm motility. While Na+ and K+ gradients produce the membrane potential in animal cells, plants and fungi utilize a proton gradient, which is established and maintained by members of the type III class of P-type ATPases [1, 18]. Representative members of this class include the Arabidopsis thaliana auto-inhibited H+ -ATPase 2 (AHA2) and the Pma1 H+ -ATPase of Saccharomyces cerevisiae.

5: ION PUMPS AND MOLECULAR MOTORS: P-, F-, AND V-TYPE ATPASES

These pumps transport protons out of the cell and can maintain an intracellular pH of ∼6.6 against an extracellular pH of 3.5, which corresponds to a membrane potential of −180 mV. H+ -ATPases contain an autoinhibitory C-terminal regulatory domain and do not utilize a counter-ion for transport. Type IV P-type ATPases are thought to act as “lipid flippases,” transferring lipids from the outer to the inner leaflet of the plasma membrane to maintain lipid asymmetry [1, 25, 26]. They are the only members of the family reported to transport substrates other than cations. P-type ATPases of class V have been identified in eukaryotic genomes, but their substrate specificities are unknown [1].

P-type ATPase structure and mechanism Transport of ions by P-type ATPases is usually described in terms of the Post–Albers scheme for the Na+ /K+ -ATPase [27, 28], in which the pump adopts two main conformations, E1 and E2 (Plate 5.1a). The cycle features alternating steps of ion binding and phosphoryl transfer and hydrolysis, with temporary occlusion of ions within the membrane during transition states to ensure vectorial transport [8, 18, 29]. In the E1 state, a cytoplasmic access channel allows binding of ion 1 from the cytoplasm. Ion 1 binding promotes phosphorylation of the pump by bound ATP. ADP is then released and the E2 state forms, characterized by a dramatic decrease in affinity for ion 1 and increase in affinity for ion 2 at the cation binding sites as well as an extracellular/luminal access channel that allows ion 1 to dissociate and ion 2 to enter. Binding of ion 2 triggers dephosphorylation of the pump. The enzyme then reverts back to the E1 state, allowing ion 2 to dissociate into the cytosol and another cycle to begin. Completion of one cycle yields 1–3 ions of type 1 expelled and 1–3 ions of type 2 imported per ATP hydrolyzed. The first X-ray crystal structure of SERCA1a showed the enzyme in the E1 state with two bound and occluded Ca2+ ions [11]. Additional structures of SERCA1a mimicking various transition states followed, elucidating the structural changes that occur during the cycle to couple ATP hydrolysis with the uphill transport of Ca2+ ions [30–36]. Recent crystal structures of the Na+ /K+ -ATPase and the plant H+ -ATPase reveal that the overall structure and transport mechanisms are conserved among the members of the P-type ATPase family [37, 38]. Plate 5.1b shows crystal structures of SERCA1a, representing the E1 state with bound ATP and Ca2+ ions immediately before phosphoryl transfer [30], and the E2 state after ADP dissociation but before phosphate hydrolysis, revealing a luminal exit channel [31]. Four structural and functional domains can be identified in P-type ATPases: the membrane (M) domain,

59

phosphorylation (P) domain, nucleotide-binding (N) domain, and actuator/anchor (A) domain [8, 11, 18]. The M-domain of SERCA1a is composed of ten membrane-spanning helices, M1–10, four of which extend into the cytoplasm. The first six transmembrane helices, M1–6, are the canonical transmembrane helices common to all family members, while M7–10 are absent in some P-type ATPases [1]. The structure of the E1-2Ca2+ state of SERCA1a shows that specific residues within the transmembrane helices coordinate the precise geometry of bound Ca2+ ions and explains the micromolar binding affinity for Ca2+ in this state [8, 11]. These sites are located at the interface of amphipathic helices (M4, M5, M6, and M8) that contribute specific side chains to the Ca2+ binding sites. Remarkably, part of these sites is also formed by partial unwinding of one of the transmembrane helices, thus providing backbone carbonyl oxygen atoms. The structure confirms the results of extensive mutagenesis studies that identified polar and charged residues essential for Ca2+ binding [39]. Changes that occur in the M-domain during the E1–E2 transition disrupt these binding sites, resulting in a 1000-fold change in affinity and promoting dissociation of the bound Ca2+ ions into the SR lumen [8, 35]. These changes are accompanied by the opening of a funnel-shaped, polar luminal exit pathway, shown in Plate 5.1b [31]. The location of the cytoplasmic pathway in the E1 state has not yet been visualized in a crystal structure. Interestingly, the crystal structure of the Na+ /K+ ATPase reveals a similar cation binding pocket to SERCA1a, suggesting that substrate specificity lies within subtle differences in the positions of side chains and water molecules at that site rather than in gross changes in M-domain structure [37]. This structure also reveals the position of the Na+ /K+ -ATPase β-subunit, covering the extracellular αM5–αM6 and αM7–αM8 loops and relating to its role in K+ ion occlusion. The structure of the M-domain of the plant H+ -ATPase suggested a mechanism for proton translocation in the absence of a counter-ion [38]. A buried aspartic acid residue that is reversibly protonated during the pumping cycle has been shown to be located near a positively charged arginine residue. This arginine is positioned to act as a built-in counter-ion in the E2 state, promoting dissociation of the proton into the extracellular space, stabilizing the aspartate in its negatively charged deprotonated state, and preventing backflow, or re-protonation of the aspartate. The highly conserved cytoplasmic domains of P-type ATPases are derived mainly from two large cytoplasmic loops between M2/M3 and M4/M5 [8, 18]. These domains undergo dramatic, rigid-body movements during the catalytic cycle. Due to the physical connections between the cytoplasmic domains and transmembrane helices, the large movements that take place in the cytoplasm are transduced into the changes in the transmembrane helices described above. The P-domain is the

60

THE LIVER: ATP SYNTHASE

catalytic core of P-type ATPases because it contains the invariant DKTGTLT sequence with the aspartate residue (D) that is reversibly phosphorylated during the reaction cycle. The P-domain is connected to the transmembrane domain through cytoplasmic extensions of M4 and M5. The N-domain is an insert within the P-domain that contains the ATP binding site. Only the adenosine base of ATP sits in the N-domain binding pocket, with the triphosphate group protruding out toward the P-domain. A flexible hinge between the N- and P-domains allows the N-domain to move toward the P-domain so that the γ-phosphate can reach its target aspartate in the P-domain. A Mg2+ ion in the phosphorylation site is thought to facilitate phosphoryl transfer from ATP to the aspartate by reducing electrostatic repulsion and stabilizing the pentavalent transition state. The A-domain is located near the N-terminal region of the enzyme and is tethered to M2 and M3 by two flexible loops [8, 18]. This domain contains a highly conserved TGES motif. Movement of the A-domain during the reaction cycle has significant consequences for the arrangement of helices in the M-domain. Analysis of SERCA1a transition state crystal structures shows that binding of ATP and Mg2+ to the Ca2+ -bound E1 state leads to a 30◦ rotation of the A-domain, which pulls on transmembrane helices (primarily M1) to close the cytoplasmic gate and occlude bound Ca2+ ions [30]. Following phosphoryl transfer, the A-domain rotates 120◦ to bring the TGES loop to the phosphorylation site, allowing the TGES loop to form a tight interaction and replace ADP and the N-domain at this site [31]. This movement pulls on M1–M2 and M3–M4 and forces a rearrangement of the transmembrane helices, ultimately creating a luminal exit pathway and distorting the high-affinity Ca2+ -binding sites. Following counter-ion binding, the TGES loop also stimulates dephosphorylation, leading to reversion of the pump to the E1 state [18]. Thus, the connections between the cytoplasmic domains and transmembrane helices provide the basis for coupling the events at the phosphorylation sites to those at the cation binding sites.

contractility on subsequent beats due to faster accumulation of Ca2+ in the SR. Mutations in PLN cause dilated cardiomyopathy in humans. SERCA1a is regulated by sarcolipin, a membrane protein expressed in fast-twitch skeletal muscle that is homologous to PLN [12, 40]. As mentioned above, Na+ /K+ -ATPases are regulated by members of the FXYD family of proteins [23]. This family contains seven members that are expressed in a tissue-specific manner, including phospholemman, which is expressed in heart and skeletal muscle, and the γ-subunit found exclusively in kidney. FXYD proteins are not required for Na+ /K+ -ATPase function, but may play a role in fine-tuning the activity of pump according to the needs of the cell. In addition to interacting proteins, Na+ /K+ -ATPase activity can be controlled by cardiac glycosides and membrane potential [19, 41, 42]. The crystal structure of the Na+ /K+ -ATPase has identified a cluster of arginines near the α-subunit C-terminus at the membrane edge that contributes a positive charge to that region of the protein [37]. Due to its location, this cluster has been speculated to act as a voltage sensor in regulating activity of the pump in response to membrane potential. Certain P-type ATPase subtypes contain regulatory domains that are fused to the main chain of the enzyme. PMCA has an autoinhibitory C-terminal tail that is displaced upon binding of Ca2+ -calmodulin, leading to increased activity [15]. Plant and fungal H+ -ATPases also have C-terminal extensions that function as autoinhibitory regulatory domains [18]. Soft-metal-ion-transporting ATPases have long N-terminal extensions containing 2–6 repeats of a heavy-metal-binding motif, plus two extra membrane-spanning helices. Interaction of metal binding sites with their corresponding ions is hypothesized to alter the activity of the pump. The Cu2+ -transporting ATPases affected in Menkes and Wilson diseases are controlled by trafficking [4]. In response to elevated cellular copper levels, ATP7A relocalizes to the basolateral membrane of polarized epithelial cells, while ATP7B translocates to vesicular compartments near the apical surface of hepatocytes. Mutations that affect this regulatory mechanism lead to disease.

Regulation of P-type ATPases

ATP SYNTHASE

The activity of P-type ATPases can be controlled through interactions with peptides, inhibitors, regulatory subunits, or N- or C-terminal domains that are fused to the main chain of the enzyme. SERCA2a activity is regulated by phospholamban (PLN), a small transmembrane protein found in cardiac SR [9]. Under resting conditions, PLN binds and inhibits SERCA2a. Activation of β-adrenergic receptors in the myocyte plasma membrane leads to PKA-dependent phosphorylation of PLN, disrupting the PLN/SERCA2a interaction and relieving inhibition of the pump. This results in an increased rate of relaxation and enhanced

ATP synthase (also referred to as the F-type ATPase) is the enzyme responsible for coupling the energy stored in a transmembrane electrochemical proton gradient to the synthesis of ATP. ATP synthases are found in both prokaryotic and eukaryotic cells (for recent reviews see [43–47]). Although this article will focus primarily on the ATP synthase from mitochondria, information on the bacterial enzyme will also be included. While the function of the mitochondrial and chloroplast enzymes is to synthesize ATP, the bacterial enzyme can also act as an ATP-driven proton pump to energize the inner membrane.

5: ION PUMPS AND MOLECULAR MOTORS: P-, F-, AND V-TYPE ATPASES

Structure of the ATP synthase complex ATP synthase is a large protein complex containing 28 protein subunits in the mammalian enzyme. A model of the bovine mitochondrial ATP synthase is shown in Figure 5.1a. A high-resolution structure for the entire ATP synthase has yet to be solved; however, the current model is based on the known high-resolution

structures of isolated subunits or subcomplexes, as well as biochemical data. The ATP synthase complex is composed of two opposing molecular rotary motors, a membrane-peripheral F1 motor and the membrane-integral F0 motor. Energy-dependent rotation of one motor thus drives the other motor in reverse. The F1 motor is composed of five different polypeptides with a stoichiometry of α3 β3 γδε, and an additional inhibitor protein, IF1 . Crystal structures of the bovine F1

F-type

V-type

β

A

B

β

α

B

A

OSCP

F1

61

F6

V1

G

B

A E

E

G

d H g −

d

b

C D

a

F

e d

a F0

C10

+e,f,

V0

g,A6L

e

C

C’

C”

+ (a) ADP +Pi

ADP +Pi

L

L

O

T

1

O

T

ATP

ATP

C C Energy

C

C C

C

C C

C

C

2 ADP +Pi

ATP

L

a

ATP

T

T O

3 H2O

(b)

L

O ATP

(c)

Figure 5.1 Structure and mechanism of the F-type and V-type ATPases. (a) Schematic representation comparing the subunit organization of the F-type and V-type ATPases. The models are based on a combination of high-resolution structures of individual subunits and subcomplexes, electron microscopy of the entire complexes, and a plethora of genetic and biochemical studies. (b) The binding change mechanism. The binding change mechanism for ATP synthesis is shown; however, the scheme can work in reverse for ATP hydrolysis. Energy input from proton translocation causes the sequential conformation changes in the β-subunits that catalyze the formation of ATP. ADP and Pi bind with high affinity to the “loose” conformation (L). The newly bound ADP and Pi are spontaneously turned into ATP when the loose conformation converts into the “tight” conformation (T). The tight conformation is converted to an “open” conformation (O) that then releases ATP. The sequential conformational changes from O→L→T→O promote the formation of ATP. (c) Proton translocation through F0 . Protons (solid black circles) enter the a-subunit through the hemi-channel located on the intermembrane space side (bottom in the figure) of the mitochondrial inner membrane and protonate cAsp-61. As the c-oligomer rotates, the protonated cAsp-61 travels through the lipid milieu in a neutral state, eventually returning to a and aligning with a second hemi-channel on the matrix side of the membrane (top in the figure). De-protonation of cAsp-61 releases the proton into the channel and into the mitochondrial matrix. A similar mechanism occurs in V0 ; however, there are only six proteolipid subunits that become protonated on an essential glutamate residue, and the direction of rotation (and thus proton movements) is reversed

62

THE LIVER: ATP SYNTHASE

subcomplex solved in the presence of different nucleotides have helped elucidate the catalytic mechanism [48, 49]. The three α- and β-subunits form an alternating hexameric ring that surrounds a central γ-subunit. The three catalytic ATP binding sites are situated in the clefts between the αand β-subunits, with most of the contacts contributed by residues on β. Hydrolysis of ATP causes conformational changes in the catalytic β-subunit that drives rotation of the central γ-rotor. The minor δ- and ε-subunits are required for the binding of F1 onto F0 . In addition, δ has been shown to play a role in energy coupling between F1 and F0 [50, 51]. Limited structural information is available for the F0 motor. It is composed of seven different types of membrane-integral proteins with a stoichiometry of a, b, c 10 , e, f , g, and A6L, and one copy each of the membrane-associated proteins d , F6 , and OSCP. Subunit c contains a critical aspartic acid residue (Asp-61) that is essential to proton movement through F0 . NMR analysis of c shows that it folds as a hairpin of two transmembrane helices. It oligomerizes into a decameric ring [52, 53], and through interactions with the a-subunit forms a proton channel through the membrane. Movement of protons through this channel drives the rotation of the c-ring with respect to the remainder of F0 . Hydropathy, residue accessibility, and cysteine-mediated crosslinking studies have led to a topology of subunit a containing five transmembrane helices [54]. Subunit a forms two channels that allow protons access to cAsp-61 [55]. Residue Arg-210 is critical to proton transport [56] and is thought to modulate protonation of cAsp-61 [57]. Subunits e and g have been shown to be involved in ATP synthase dimerization (see below). In assembled ATP synthase, the rotary motors are connected by two stalks. The central stalk, or rotor, composed of the c-oligomer of F0 and the γ-, δ-, and ε-subunits of F1 , forms a physical connection between F1 and F0 and transmits mechanical energy between the two sectors via rotation. An obligatory consequence of the rotary mechanism is that the α3 β3 hexamer must be anchored to the stationary parts of F0 . The peripheral stalk (or “stator”) is on the periphery of the complex and accomplishes this anchoring function. The stator is the part of the enzyme that has undergone the most evolutionary divergence. In the bovine enzyme it is composed of b, d , F6 , and OSCP. The b-subunit spans the membrane twice and reaches from the membrane up the side of F1 and makes contact with OSCP near the apex of the α3 β3 hexamer. Recently a high-resolution structure of a b, d , and F6 subcomplex has been solved showing a long extended protein of mostly α-helix, with a slight ˚ structure obtained by curve, that fits nicely into a 32-A cryoelectron microscopy [58]. In the bacterial enzyme the peripheral stalk is composed of a b 2 δ complex, with two copies of the b-subunit that spans the membrane once, and one copy of δ, the homolog of OSCP. The b-subunits are mostly α-helical [59] and

dimerize into a novel right-handed coiled-coil [60] that extends from the membrane, up the side of F1 and contacts δ at the apex of the α3 β3 hexamer. It is proposed that the right-handed twist and elastic deformation of the helical interactions in the b-dimer may store energy and correct for the symmetry mismatch of having a 3-stepping F1 and a 10-stepping F0 motor [60].

The binding change mechanism of ATP synthesis A schematic representation of the binding change mechanism during ATP synthesis proposed by Boyer [61] is shown in Figure 5.1b. The three catalytic sites sequentially and cooperatively cycle between the “open,” “loose,” and “tight” conformations. ADP and Pi bind with high affinity to a catalytic site in a loose conformation. The loose conformation is converted to a tight conformation which causes the spontaneous formation of ATP. The tight conformation then changes to an open conformation which has a low affinity for nucleotides and releases the newly formed ATP. Each of the three catalytic sites is always in a different conformation and thus each of the steps is happening concurrently. Energy from proton translocation drives the conformational changes in the catalytic sites, and thus that energy is actually used to promote the binding of substrates and release of product.

Rotary catalytic mechanism Rotational catalysis (as postulated in [62]) provides a mechanism to transduce energy from F0 to F1 . Rotation of a central part of the enzyme drives the repetitive conformational changes that are integral to the binding change mechanism. Movement of protons down an electrochemical gradient drives the rotation of the c-oligomer, which in turns drives rotation of the δ-, ε-, and γ-subunits with respect to the remainder of the enzyme. The bovine heart mitochondrial F1 structure [48] provided validation for Boyer’s theory, showing the γ-subunit forms an asymmetrical coiled-coil that protrudes through the center of the α3 β3 hexamer, and that each of the three catalytic β-subunits is in a different conformation with different nucleotide occupancies. Rotation of γ would cause the three catalytic β-subunits to cycle between the three different conformations. The mechanism of proton-driven rotation through F0 [63] is shown in Figure 5.1c. Protons from the mitochondrial intermembrane space enter the a-subunit through a hemi-channel, gaining access to and protonating cAsp-61. When the c-ring rotates due to Brownian motion, the protonated Asp-61 cannot go back into position with a and thus acts as a Brownian ratchet. The protonated Asp-61 travels through the lipid milieu in an

5: ION PUMPS AND MOLECULAR MOTORS: P-, F-, AND V-TYPE ATPASES

uncharged state as the c-oligomer rotates in a counterclockwise direction (when looking from the matrix side of the membrane), eventually reaching the other side of the a-subunit. The proton is released from cAsp-61 into a second hemi-channel which leads to the mitochondrial matrix. aArg-210 stimulates deprotonation by stabilizing Asp-61 in the charged state. Subunit rotation in ATP synthase has been shown unequivocally. Duncan et al. [64] have used disulfide crosslinks between the γ-subunit and the β-subunit in the E. coli enzyme. Consistent with γ-rotation, this drastically inhibits ATPase activity. In an elegant series of experiments, Noji et al. [65] attached a fluorescently labeled actin filament to the γ of the thermophilic Bacillus PS3 F1 and directly observed ATP-dependent rotation using single-molecule fluorescence microscopy. The same experiment was repeated showing rotation of ε (mitochondrial δ) in F1 [66], and rotation of the c-oligomer in the complete ATP synthase [67]. Single-molecule fluorescence resonance energy transfer has also been used to show the stepwise rotation of both γ and ε in lipid-embedded ATP synthase [68].

Assembly and regulation of ATP synthase Of the 16 different gene products that make up the ATP synthase and its inhibitor, all but 2 (subunit a and A6L) are nuclear encoded, and thus must be imported into the mitochondria. In yeast, an ordered assembly pathway has been demonstrated (reviewed in [69]). F1 assembly occurs independent of F0 ; however, in the absence of F1 , no F0 can be detected. Subunit c appears to be the starting point of F0 assembly, with the a-subunit and A6L assembling onto F0 last. Once in the mitochondria, ATPase subunits require a number of chaperones to assist in folding and assembly. In yeast, Atp11p and Atp12p assist in the assembly of the F1 domain by acting as chaperones assisting the folding of the β- and α-subunits, respectively [70]. F0 assembly is dependent on two chaperones, Atp10p [71] and Atp22p [72], with Atp10p interacting directly with subunit a. In mammals, homologs have been identified for Atp11p and Atp12p, however no Atp10p or Atp22p homologs have been identified. In the bacterial system the product of the uncI gene acts as a chaperone in assembling the c-ring [73]. It has recently been discovered that mitochondrial ATP synthase dimerization affects mitochondrial morphology [74]. Subunits e and g are required to maintain dimeric ATP synthase [75], and deletion of either subunit results in altered mitochondrial cristae morphology [76]. EM images of dimeric ATP synthase reveal that the two complexes are arranged at a 40◦ angle and it has been proposed that this defines the mitochondrial cristae curvature [77]. Control of ATP synthase biogenesis involves both transcriptional and post-transcriptional regulation and

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shares common regulatory cascades with other mitochondrial complexes (reviewed in [69]). In most tissues the ATP synthase content relative to other respiratory chain enzymes is constant; however, in brown fat decreased transcription of subunit c is correlated with a 10-fold reduction in ATP synthase. Overexpression of c results in a normal level of ATP synthase in brown fat, suggesting that the amount of ATP synthase present in a cell may be controlled by varying the amount of subunit c. In bacteria, the catalytic activity of ATP synthase is partially controlled by the ε-subunit (δ- in mitochondria) (reviewed in [44]). During assembly, ε is a potent inhibitor of F1 -ATPase, preventing the unwanted hydrolysis of cellular ATP. Recently it has been suggested that in the holoenzyme ε modulates both the activity and the coupling of the enzyme in an ATP, ADP, Pi , and proton-motive force-dependent manner [45]. A slightly different hypothesis has been proposed for the E. coli enzyme, where ε senses and integrates the phosphorylation potential and proton-motive force, acting like a switch, and adapting a conformation that allows for rotation only in the direction of ATP hydrolysis when the enzyme is undergoing ATP-driven proton pumping, and only in the direction of synthesis when the enzyme is synthesizing ATP [44, 50, 51]. Thus ε is a coupling factor, undergoing direction-dependent conformational changes that keep the enzyme efficient. The mitochondrial ATP synthase is regulated by an additional protein subunit, IF1 , that inhibits ATP hydrolysis, but not the ATP synthesis activity of ATP synthase (reviewed in [78]). The active inhibitor is a dimer that oligomerizes in a pH-dependent manner. At pH values over 7, IF1 oligomerizes and does not associate with the ATPase. During ischemia, glycolysis is the only source of ATP production and hence cellular and mitochondrial pH drop. When the pH is low, IF1 forms a dimer that binds to and inhibits hydrolysis of two ATPase complexes.

V-TYPE ATPASES V-type ATPase function The vacuolar (H+ )-ATPases (or V-type ATPases) are ATP-driven proton pumps present in both intracellular compartments, such as endosomes, lysosomes, and secretory vesicles, and in the plasma membrane of certain animal cells [79]. V-type ATPases couple the energy derived from ATP hydrolysis to the active transport of protons across the membrane. V-type ATPases play an important role in both endocytic and intracellular membrane traffic (Figure 5.2). In endocytosis, acidification of early endosomes triggers the release of internalized ligands from their receptors, thus facilitating receptor recycling to the cell surface [80]. Acidification of early endosomes is also required for budding of endosomal carrier vesicles that

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THE LIVER: V-TYPE ATPASES

(a)

(b)

Figure 5.2 (a) Function of intracellular V-type ATPases. V-type ATPases in the endosome activate ligand-receptor dissociation and receptor recycling, playing a similar role in the trafficking of newly synthesized lysosomal enzymes via the Man-6-P receptor. V-type ATPases in lysosomes activate protein degradation, and in secretory vesicles they drive coupled transport of neurotransmitters. (b) Function of plasma membrane V-type ATPases. Plasma membrane V-type ATPases serve important roles in renal acidification in alpha-intercalated cells, in bone resorption by osteoclasts, and in the invasiveness of certain tumor cells. They are thus potential drug targets in treating osteoporosis and cancer

carry released ligands from early to late endsosomes. This process requires beta-COP proteins, which bind to early endosomes in a pH-dependent manner [81]. The V-type ATPase itself may serve as a pH sensor in this process by binding the G protein Arf6 and its cognate guanine nucleotide exchange factor (ARNO) [82]. Low pH serves an analogous role in the trafficking of lysosomal enzymes, which bind to the mannose-6-phosphate receptor in the trans-Golgi network (TGN) and dissociate in response to low pH in late endosomes, thus allowing the unoccupied receptors to recycle to the Golgi [80]. Certain pathogens also utilize the acidic pH of endosomal compartments to gain access to the cytoplasm of

infected cells [83]. Following internalization, influenza virus undergoes fusion with the endosomal membrane, thus releasing the viral RNA. This fusion is mediated by the viral hemagglutinin (HA) molecule, which acts as a pH-dependent fusogen. Various bacterial toxins, including diphtheria toxin and anthrax toxin, also gain entry via acidic endosomes. For diphtheria toxin, the low pH causes the B chain to form a pore in the endosomal membrane through which the cytotoxic A chain passes. A similar process occurs for anthrax toxin, but in late endosomes. V-type ATPases in lysosomes and other digestive compartments create an acidic environment that both activates proteases and drives the coupled transport of degradation

5: ION PUMPS AND MOLECULAR MOTORS: P-, F-, AND V-TYPE ATPASES

products and other small molecules and ions [80]. In secretory vesicles, such as synaptic vesicles, V-type ATPases create the proton gradient and membrane potential that drives the coupled transport of neurotransmitters, such as glutamate and norepinephrine. The acidic environment also activates processing enzymes that convert precursor forms of hormones and growth factors to their mature forms. V-type ATPases also play an important role in the plasma membrane of various cell types. V-type ATPases in the apical membrane of cells of the renal proximal tubule assist in bicarbonate reabsorption by secreting protons into the renal fluid, thus promoting conversion of bicarbonate to carbon dioxide [84]. In the distal tubule and collecting duct, alpha intercalated cells use V-type ATPases at the apical membrane to secrete acid into the urine. Defects in specific V-type ATPase isoforms expressed in these cells lead to renal tubule acidosis [84]. Plasma membrane V-type ATPases are essential for bone resorption by osteoclasts, since they create the acidic extracellular environment that degrades bone [85]. Defects in the osteoclast V-type ATPase cause the disease osteopetrosis, characterized by inadequate bone resorption [80]. V-type ATPases in the plasma membrane of clear cells in the epididymus and vas deferens create the acidic lumenal environment necessary for sperm maturation and stability [86]. Recently, plasma membrane V-type ATPases have been identified in certain tumor cells, where they appear to assist in invasion [87]. Thus, highly invasive MB231 human breast tumor cells possess active V-type ATPase at the cell surface, and the in vitro invasiveness of these cells is significantly reduced by the specific V-type ATPase inhibitor bafilomycin. Because of these diverse functions, coupled with a comparable isoform diversity (see below), V-type ATPases are being investigated as potential therapeutic targets in treating such diseases as osteoporosis and cancer.

V-type ATPase structure and mechanism V-type ATPases resemble in overall structure the archaebacterial ATPases and F1 F0 ATP synthases (see above) and, like these enzymes, operate by a rotary mechanism (Figure 5.1a) [79]. The V-type ATPase complex is composed of a peripheral ATP hydrolyzing domain (V1 ) and a membrane-integral proton translocation domain (V0 ) [79, 88]. The V1 domain contains eight different subunits (A–H) in a stoichiometry of A3 B3 CDE2 FG2 H [79]. The nucleotide binding sites are located at the interface of the A- and B-subunits, which are arranged in an alternating fashion in a hexameric ring. The catalytic sites are located mainly on the A-subunits, whereas “non-catalytic” sites (of unknown function) are mainly on the B-subunits [80]. The remaining subunits in V1 serve to attach the V1

65

domain to V0 and are arranged in one of two types of stalks. Subunits have been assigned to these two stalks based on cysteine-mediated crosslinking using the photoactivated crosslinking reagent maleimidobenzophenone (MBP) [89–91] as well as electron microscopic (EM) analysis [91–93] and co-immunoprecipitation studies [94]. The central (or rotary) stalk contains the V1 subunits D and F, whereas the peripheral (or stator) stalk contains the V1 subunits C, E, G, and H. The V0 domain (in yeast) contains six different subunits in a stoichiometry adc 4 c’c”e [80, 95]. Mammalian V-type ATPase lacks subunit c’ but contains an additional accessory subunit (Ac45) [80]. The three proteolipid subunits (c, c’, and c”) are highly hydrophobic proteins that form a six-membered ring. Subunits c and c’ each contain four transmembrane helices (TMs) while subunit c” contains five TMs. Each proteolipid subunit contains a buried glutamic acid residue (present in TM4 of subunit c and c’ and TM3 of subunit c”) that is essential for proton transport [96]. The proteolipid subunits adopt a unique arrangement in the ring, with subunits c’ and c” adjacent to each other [97]. Subunit d is a hydrophilic protein which sits on the cytoplasmic side of the proteolipid ring [98] and connects it to the central rotary subunits (D and F) of V1 . Subunit e is a small hydrophobic protein of unknown function. The last subunit of V0 is subunit a, which is a 100 kDa transmembrane protein containing a 50 kDa hydrophilic N-terminal domain located on the cytoplasmic side of the membrane and a 50 kDa hydrophobic C-terminal domain containing eight to nine TMs [99]. The N-terminal domain forms part of the peripheral stalk by interacting with subunits A, C, and H [90, 94]. A buried Arg residue in TM7 of subunit a is essential for proton transport [100] and has been shown using disulfide-mediated crosslinking to come into close proximity to the carboxyl groups on the proteolipid ring [101]. As with the F-type and A-type ATPases, the V-type ATPases utilize a rotary mechanism to couple ATP hydrolysis to proton translocation [102, 103]. ATP hydrolysis at the catalytic sites of V1 drives rotation of the central rotary complex containing subunits D and F attached to subunit d and the proteolipid ring of V0 . During this process, subunit a is held fixed relative to the A3 B3 hexamer of V1 by the peripheral stalk. Unlike the F-type ATPases, the V-type ATPases contain two peripheral stalks [91], one of which appears to contain an EG heterodimer plus subunit C, whereas the other contains an EG heterodimer and subunit H. As described above for the F-type ATPases, the hydrophobic domain of subunit a plays two important functions in proton transport. It allows protons to reach and leave the buried carboxyl groups on the proteolipid ring via cytoplasmic and luminal hemi-channels and it stabilizes these carboxyl groups in their charged, deprotonated form through interaction with the buried Arg residue in TM7 [100]. Protons enter via the cytoplasmic hemi-channel on subunit a, protonate the glutamic acid residues on the proteolipid ring, rotate

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THE LIVER: V-TYPE ATPASES

through the hydrophobic phase of the membrane, and are displaced into the luminal hemi-channel by interaction between the glutmate residues and the subunit a Arg residue. Evidence for swiveling of the critical helices in subunit a and the proteolipid subunits has emerged from disulfide-mediated crosslinking studies [101], and characterization of the binding site for the specific inhibitors bafilomycin and concanamycin [104] suggests that helical swiveling may be required for activity.

V-type ATPase assembly and targeting V-type ATPases are large, multi-subunit complexes which require dedicated chaperones resident to the ER for assembly. In yeast, these chaperones are Vma12p, Vma21p, Vma22p, and Pkr1p [105]. Vma21p interacts with subunit c’ and promotes its assembly with subunits c, c”, and d in the ER. Vma12p and Vma22p form a complex that transiently binds to subunit a and promotes its assembly with the remainder of the V0 complex. Vma21p then accompanies the completed V0 domain to the Golgi, where it dissociates and recycles back to the ER. The V1 domain assembles in the cytosol and attaches to V0 in the Golgi. A second pathway in which assembly of V1 and V0 occurs in a more coordinated fashion has also been described [88]. Targeting of V-type ATPases to different cellular membranes is controlled by isoforms of subunit a [79]. In yeast there are two isoforms, with Vph1p targeting the V-type ATPase to the vacuole and Stv1p targeting to the Golgi [106]. In mammalian cells there are four a-subunit isoforms (a1–a4). The a1 isoform is present in synaptic vesicles and the presynaptic membrane in neurons [107], a2 is present in Golgi and apical endosomes in renal cells [82, 85], a3 is present in the plasma membrane of osteoclasts, the lysosomes of osteoclast precursors, and insulin-containing secretory vesicles in pancreatic islet cells [85, 108], and a4 is present in the apical membrane of renal intercalated cells and epididymal clear cells [86]. Studies of chimeric a-subunit constructs in yeast have demonstrated that the targeting information is located in the cytoplasmic N-terminal domain of subunit a [109].

Regulation of V-type ATPase activity Different intracellular compartments are maintained at different pH values, with lysosomes more acidic than late endosomes, which are in turn more acidic than early endosomes and the Golgi [80]. The amount of proton transport across the plasma membrane of cells is also typically tightly controlled. Because V-type ATPases serve important functions in both of these environments, regulation of

V-type ATPase activity is of crucial importance to cells and organisms. Several mechanisms have been shown to function in the regulation of V-type ATPase activity in vivo. Among the most important regulatory mechanisms is reversible dissociation of the V1 and V0 domains [79, 88]. This process has been demonstrated in a variety of cell types, including yeast, goblet cells of the insect midgut, dendritic cells involved in antigen processing, and epithelial cells of the renal proximal tubule. In yeast and insect cells, in vivo dissociation serves to conserve cellular energy stores, since it occurs in response to glucose depletion and molting, respectively [88, 110]. Activation of antigen processing in dendritic cells causes increased assembly of the V-type ATPase on the membrane of the antigen processing compartment, thus providing the requisite increase in acidity [111]. V-type ATPase assembly in renal cells also increases in response to elevated glucose levels [112], but the physiological function of this response is not certain. Reversible dissociation of the V-type ATPase has been best characterized in yeast. Dissociation occurs rapidly and without the need for new protein synthesis [88]. Dissociation and reassembly appear to be independently controlled processes. Thus dissociation (but not reassembly) requires intact microtubules [113], whereas reassembly (but not dissociation) requires a complex known as RAVE (regulator of vacuolar and endosome acidification) [114]. The RAVE complex contains the Rav1p and Rav2p proteins as well as the ubiquitin ligase subunit Skp1p. RAVE is required for both biosynthetic and glucose-dependent assembly and appears to bind to the E and G subunits of V1 , and to stabilize V1 in a form competent for assembly with V0 [115]. Another protein required for glucose-dependent reassembly is the glycolytic enzyme aldolase. Aldolase binds to the intact V1 V0 complex in a glucose-dependent manner and mutations in aldolase that interfere with binding to the V-type ATPase complex prevent V-type ATPase assembly [116]. Aldolase may thus be serving as a glucose sensor. A second candidate for the glucose sensor is the “non-homologous domain” of the V-type ATPase A-subunit. The non-homologous domain is a 90 amino acid insert present in all A-subunit sequences but absent from the corresponding F-type ATPase β-subunit (hence the name). Mutations in this region have been shown to block in vivo dissociation without affecting catalytic activity [117]. Moreover, the non-homologous domain alone, expressed in the absence of the remainder of the A-subunit, is able to bind to the V0 domain without other V1 subunits in a glucose-dependent manner [118]. The dependence of in vivo dissociation on subunit a isoforms and cellular environment has been studied in yeast. Vph1p-containing complexes normally target to the vacuole, while Stv1p-containing complexes are retained in the Golgi. If Stv1p is overexpressed, most Stv1p-containing complexes also appear in the vacuole [106]. Both Vph1p and overexpressed Stv1p utilize the

5: ION PUMPS AND MOLECULAR MOTORS: P-, F-, AND V-TYPE ATPASES

carboxypeptidase Y (CPY) pathway for vacuole delivery, with vacuolar targeting interrupted by disruption of the vps (vacuolar protein sorting) genes. Disruption of Vps21p results in accumulation of proteins in a post-Golgi compartment, whereas disruption of Vps27p causes proteins to accumulate in the prevacuolar compartment [106]. Recent studies have shown that in vivo dissociation of the V-type ATPase complex is primarily determined by the cellular environment rather that the a-subunit isoform present. Thus complexes localized to the same compartment but containing different a-subunit isoforms show very similar in vivo dissociation [119]. Although the precise environmental factors affecting dissociation are uncertain, the luminal pH appears to be one determining factor [118]. The signaling pathways controlling reversible dissociation in yeast have also not been identified. However, PI-3 kinase and protein kinase A (PKA) have been implicated in controlling dissociation in renal tubule cells and insect salivary gland, respectively [112, 120]. An important property of the separated V1 and V0 domains is that they do not possess ATP hydrolytic activity nor passive proton conductance. This avoids non-productive ATP hydrolysis and dissipation of intracellular pH gradients. Subunit H appears critical to silencing ATPase activity of free V1 [121]. Subunit H accomplishes this function by bridging the peripheral and rotor domains and physically preventing rotation in free V1 [122]. A second important regulatory mechanism appears to be the changes in coupling efficiency between proton transport and ATP hydrolysis [80]. V-type ATPase complexes containing Vph1p show much tighter coupling of proton transport and ATP hydrolysis than complexes containing Stv1p [106]. This finding, together with the much lower assembly of Stv1p-containing V0 complexes with V1 relative to Vph1p-containing V0 complexes, may explain the more acidic pH of the vacuole compared to the Golgi. V-type ATPase complexes also appear to be poised to either increase or decrease their coupling efficiency, as mutagenesis studies indicate that the wild-type V-type ATPase is not optimally coupled [117]. The in vivo factors controlling coupling efficiency have not yet been elucidated. A number of other regulatory mechanisms have also been identified. A particularly important one in controlling proton transport across the plasma membrane of epithelial cells is control of pump number through reversible fusion of V-type ATPase containing vesicles with the plasma membrane. This mechanism is used in both renal intercalated cells and epididymal clear cells [84, 123]. In the case of clear cells, fusion is under the control of a bicarbonate-sensitive adenylyl cyclase, which increases cellular cAMP in response to elevated bicarbonate levels. Covalent modification of the catalytic site of the V-type ATPase has also been shown to regulate activity. Reversible disulfide bond formation between conserved cysteine residues at the catalytic site restricts the ability of substrate to bind, thus inhibiting activity

67

[80]. A final regulatory mechanism that does not involve direct changes in the V-type ATPase is modification of proton and other ion conductances. Because the V-type ATPase is electrogenic it requires movement of another charged species to allow net acidification of intracellular compartments. This appears to primarily involve intracellular chloride channels of the Clc family, whose activity is in turn regulated by various mechanisms, including PKA [80]. Because the pH of intracellular compartments is a balance between active transport and passive leakage, regulation can also be accomplished at the level of passive proton channels, of which Zn2+ -sensitive channels are one example [124]. The diversity of processes in which the V-type ATPases participate may help to explain the multiple mechanisms that have evolved to regulate V-type ATPase activity in vivo.

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96. Hirata, R., Graham, L.A., Takatsuki, A. et al. (1997) VMA11 and VMA16 encode second and third proteolipid subunits of the Saccharomyces cerevisiae vacuolar membrane H+-ATPase. J Biol Chem, 272, 4795–803. 97. Wang, Y., Cipriano, D.J. and Forgac, M. (2007) Arrangement of subunits in the proteolipid ring of the V-ATPase. J Biol Chem, 282, 34058–65. 98. Iwata, M., Imamura, H., Stambouli, E. et al. (2004) Crystal structure of a central stalk subunit C and reversible association/dissociation of vacuole-type ATPase. Proc Natl Acad Sci U S A, 101, 59–64. 99. Leng, X.H., Nishi, T. and Forgac, M. (1999) Transmembrane topography of the 100-kDa a subunit (Vph1p) of the yeast vacuolar proton-translocating ATPase. J Biol Chem, 274, 14655–61. 100. Kawasaki-Nishi, S., Nishi, T. and Forgac, M. (2001) Arg-735 of the 100-kDa subunit a of the yeast V-ATPase is essential for proton translocation. Proc Natl Acad Sci U S A, 98, 12397–402. 101. Wang, Y., Inoue, T. and Forgac, M. (2004) TM2 but not TM4 of subunit c” interacts with TM7 of subunit a of the yeast V-ATPase as defined by disulfide-mediated cross-linking. J Biol Chem, 279, 44628–38. 102. Hirata, T., Iwamoto-Kihara, A., Sun-Wada, G.H. et al. (2003) Subunit rotation of vacuolar-type proton pumping ATPase: relative rotation of the G and C subunits. J Biol Chem, 278, 23714–19. 103. Imamura, H., Nakano, M., Noji, H. et al. (2003) Evidence for rotation of V1-ATPase. Proc Natl Acad Sci U S A, 100, 2312–15. 104. Bowman, B.J., McCall, M.E., Baertsch, R. et al. (2006) A model for the proteolipid ring and bafilomycin/concanamycin-binding site in the vacuolar ATPase of Neurospora crassa. J Biol Chem, 281, 31885–93. 105. Malkus, P., Graham, L.A., Stevens, T.H. et al. (2004) Role of Vma21p in assembly and transport of the yeast vacuolar ATPase. Mol Biol Cell , 15, 5075–91. 106. Kawasaki-Nishi, S., Nishi, T. and Forgac, M. (2001) Yeast V-ATPase complexes containing different isoforms of the 100-kDa a-subunit differ in coupling efficiency and in vivo dissociation. J Biol Chem, 276, 17941–48. 107. Morel, N., Dedieu, J.C. and Philippe, J.M. (2003) Specific sorting of the a1 isoform of the V-H+ATPase a subunit to nerve terminals where it associates with both synaptic vesicles and the presynaptic plasma membrane. J Cell Sci , 116, 4751–62. 108. Sun-Wada, G.H., Toyomura, T., Murata, Y. et al. (2006) The a3 isoform of V-ATPase regulates insulin secretion from pancreatic beta-cells. J Cell Sci , 119, 4531–40. 109. Kawasaki-Nishi, S., Bowers, K., Nishi, T. et al. (2001) The amino-terminal domain of the vacuolar proton-translocating ATPase a subunit controls targeting and in vivo dissociation, and the carboxyl-terminal domain affects coupling of proton transport and ATP hydrolysis. J Biol Chem, 276, 47411–20. 110. Beyenbach, K.W. and Wieczorek, H. (2006) The V-type H+ ATPase: molecular structure and function,

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6

Hepatocyte Surface Polarity: Its Dynamic Maintenance and Establishment Lelita T. Braiterman and Ann L. Hubbard Department of Cell Biology, Johns Hopkins University School of Medicine, Baltimore, MD, USA

INTRODUCTION The hepatocyte, the major epithelial cell of the liver, performs many crucial functions that stem largely from its strategic position between two different environments, the blood plasma and the bile. The functions carried out at the two fronts are distinct, which means that the hepatocyte surface is asymmetric or polarized. Polarity is a fundamental characteristic of most eukaryotic cells, either as a transient phenomenon (e.g. in a moving fibroblast) or a permanent feature (e.g. of an epithelial layer) [1]. In epithelial cells, polarity is evident at many levels. At the cell surface, the basolateral and apical membrane domains face different environments (internal and external, respectively) and each membrane contains a distinct set of proteins and lipids [2]. The stereotypical locations of different organelles within a particular epithelial cell type are additional indicators of polarity. For example, the Golgi is positioned between the nucleus and apical surface in most epithelial cells; this relationship reflects the polarized organization of microtubules. Acquisition of the fully polarized phenotype requires coordination of multiple processes: cell–substrate and cell–cell adhesion; activation of multiple signal transduction pathways; reorganization of cytoskeletal elements; assembly of tight The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

junctions (TJs) and adherens junctions (AJs); and polarized vesicle traffic [3]. Furthermore, though the epithelial phenotype is temporally and spatially stable, its maintenance requires all the above processes to be continuously “on,” prompting the addition of “dynamic” to the title. In this chapter, we discuss the establishment and maintenance of hepatocyte surface polarity. After presenting selected features of cellular organization, we first focus on the maintenance of hepatic surface polarity. Plasma membrane (PM) biogenesis and turnover involve organelles of the cell’s secretory and endocytic pathways, where signals, sorting, membrane carrier formation, and transport between successive compartments are central features (covered in other chapters). Our emphasis in this edition is on PM protein biogenesis. From research over the past three decades, it is quite clear that similarities and differences in the pathways and mechanisms of PM biogenesis exist between hepatocytes and simpler, bipolar epithelial cells. Because the latter cells (e.g. kidney and intestine) have been used more extensively than hepatic cells, of necessity, we will discuss advances from studies of these simpler cells and consider how hepatocytes are similar and different. In the second half of this chapter, we discuss the establishment of hepatocyte polarity. We summarize

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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work from genetically tractable organisms that opened the way to studies of mammalian epithelial polarity, then discuss TJ/AJ proteins and an assortment of proteins we designate “polarity modulators” that may play roles in hepatic polarity. Throughout, we raise unresolved issues to stimulate interest in solving them. Our apologies to those whose work we do not cite; space was limited. Instead, we cite reviews that we found particularly helpful and/or whose bibliographies were extensive. The reader is directed to our earlier chapters [4].

HEPATOCYTE ORGANIZATION Structural aspects Liver architecture, in which hepatocytes are arranged in cords, is unique among epithelial organs. An individ-

ual cell is polygonal and faces at least two blood sinusoids (the basal domain). A branched network of grooves between adjacent cells forms the bile canaliculus (apical domain), which in a single optical section can appear variously round or tubular (Figure 6.1a,b). Such a polygonal shape means that hepatocytes do not have a single basolateral-to-apical axis as do the simple bipolar epithelial cells lining kidney tubules or the intestinal lumen. Moreover, the hepatocyte organelles of the secretory and endocytic pathways must simultaneously serve these “fragmented” PM domains. The low-magnification view of hepatocytes in situ (Figure 6.1c) doesn’t reveal many clues as to how this is accomplished. In fact, aside from the rough endoplasmic reticulum (ER), the major organelles making up the two pathways (Golgi, secretory vesicles, endosomes, and lysosomes) occupy a small fraction of the hepatocyte cytoplasm (shaded areas of Figure 6.1d). Furthermore, the Golgi and lysosomes have

Sinusoidal Front BC

Golgi complex

BC

High concentrations of endo/exocytotic organelles

BC BC

Figure 6.1 The architecture of the liver is unique. (a) A scanning electron micrograph of a portion of a liver lobule is shown. A continuous network of bile canaliculi runs along the exposed cell surfaces of the liver plate. (b) The two distinct PM domains are visualized by immunofluorescence detection of the basolateral PM protein, HA 321/BEN and the apical PM protein, HA4/cell-CAM105/ectoATPase. An electron micrograph of a hepatocyte (c) and a corresponding schematic drawing (d) highlight the “active zones” in vesicular trafficking. The major sorting organelles (TGN and endosomes) and transport vesicles are concentrated in small “clear” zones that are probably the most active in vesicle traffic. These zones are located between the Golgi and the apical PM and near the basolateral PM (the shaded regions in (d))

6: HEPATOCYTE SURFACE POLARITY: ITS DYNAMIC MAINTENANCE AND ESTABLISHMENT

seemingly paradoxical locations: they are concentrated around the apical pole, despite the fact that they serve predominantly the sinusoidal pole. Most newly synthesized membrane and secretory proteins exiting the trans-Golgi network (TGN) are directed to the basolateral front, with less TGN-derived traffic directed to the apical front. Similarly, macromolecular cargo (including PM components) internalized at the sinusoidal front is delivered to lysosomes, the apical PM, or bile. Because we know that multiple vesicular steps in the biosynthetic and endocytic pathways rely on microtubules, this cytoskeletal system (along with its associated motors) is strongly implicated as the link between the apical and basolateral regions of polarized epithelial cells.

Molecular aspects The hepatocyte PM has long been studied as a biochemical entity, and the differences in the protein and lipid compositions among the domains were appreciated early. With few exceptions, disagreements about assignments for certain PM proteins continue; domain-specificity seems to be the general rule in hepatocytes and other epithelia. That is, in a particular cell type an integral PM protein is restricted to a single PM domain (Figure 6.1b). However, there is great plasticity in PM protein locations among epithelial cells in different organs; a basolateral PM protein in liver hepatocytes (e.g. NaK-ATPase) might be apical in another organ (e.g. retinal pigment epithelial cells [5]). The increasing number of such examples of cell-specific localization—“protein location following function”—is important to remember when investigating implications for cell type-specific sorting and targeting mechanisms. Paradoxically, an important family of integral PM proteins involved in vesicle targeting, the t-SNAREs, does not conform to the general rule of domain-specificity in epithelial tissue (more below). Although the PM lipids of epithelial cells do not show the same absolute domain-specificity as most classes of PM proteins, differences between the domains exist. In hepatocytes, as in most epithelial cells, sphingomyelin and cholesterol are relatively enriched in apical versus basolateral PM subcellular fractions. Additionally, 75% of the major hepatocyte gangliosides (GM1 and GM2) have been found in the PM, but individual domains have not been analyzed [6]. Phospho-inositides (PI) are increasingly being recognized as key molecules in both signaling pathways and membrane traffic. There is differential enrichment of specific phospho-forms of PI in the cytoplasmic leaflet of the Golgi, endosomal, and PM domains. The hepatic organelles and PM domains need to be examined. Despite the apparent similarities with other epithelial cells, the lipid dynamics, if not the lipids themselves, of the hepatocyte apical PM must be unique since this PM functions as the site of bile secretion, which includes detergent-like bile salts, phosphatidyl choline (PC),

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and cholesterol. The fact that the PC present in the apical PM contains longer and more unsaturated fatty acyl chains than that in bile, which is predominantly sn1-16 : 0/sn2-18 : 1 or 2 [8], points to exquisite selectivity in the mechanisms for removal/transport of PC from/through the apical PM bilayer. MDR3 (ABCB4) carries out this important function. More detailed aspects of lipids are covered in Chapter 7. However, relevant to our discussion is the rate of PC release into bile, reported to be equivalent to ∼10% of the apical PM outer (lumenal) leaflet per minute in vivo [8]. This extraordinary flux of lipid through the apical PM bilayer must place constraints on its physical structure.

Dynamic aspects The amount of traffic through the secretory and endocytic/transcytotic pathways of the hepatocyte is truly remarkable. In the secretory pathway, the quantity of plasma proteins made and shipped out each day by a single hepatocyte nearly equals the total cell content of protein [9]; ∼120 × 103 albumin molecules are synthesized per minute per cell. Vesicles ∼200–400 nm in diameter deliver this and other secretory cargo to the basolateral PM, which increases its surface area by ∼0.5% min cell−1 . Fluid-phase endocytosis at the basolateral surface of hepatocytes is even more robust, with an estimated 8% of the PM domain surface area internalized per minute per cell [8]. Despite this flux of membrane into and out of the basolateral PM, there is homeostasis in membrane surface area and identity. The dynamic nature of the apical PM is just as impressive. From studies of fluid phase transcytosis, ∼600–850 vesicles (100 nm in diameter) are estimated to fuse with the apical surface every minute [8]. This means that the apical membrane surface area would double or triple every 20 minutes if there was no mechanism for removal of molecular components. Clearly, maintenance of the polarized phenotype is a continuous process.

CELL SYSTEMS FOR STUDYING THE ITINERARIES OF PM PROTEINS Intact tissue There are advantages to studying PM dynamics in whole animals or the perfused liver: cell polarity and tissue architecture are maintained; and, since the liver is composed of ∼70% hepatocytes, organelles from a single cell type can be isolated in abundance and relative purity. Among the disadvantages are: physiological variation among animals; dilution of precursors, inhibitors, and so on, throughout the whole body; and difficulties in quickly modulating experimental conditions (e.g. temperatures, inhibitors, etc.).

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Nonetheless, results from work in the whole organ have established the basic parameters of hepatocyte PM dynamics. This is important, since the fidelity and rates of many processes are much higher in vivo than they are in the in vitro cell systems so far examined [10]. Recent progress in understanding the molecular aspects of PM traffic in epithelial cells has come from the use of in vitro cell systems.

Simple epithelial cell systems Development of polarized cell lines has been a goal for decades and several have been popular due to the relative ease with which they can be polarized. These include cells from normal (?) kidney (MDCK, Madin–Darby canine kidney and LLCPK1, LLC pig kidney, with characteristics of the distal and the proximal tubular nephron, respectively); a human colon carcinoma (Caco-2) with characteristics of the small intestine; and rat thyroid (FRT, Fisher rat thyroid). In Table 6.1, we list several of their molecular features (and the two hepatic cell lines described below), because these are important for interpreting/understanding the differences among the cells with regard to PM protein sorting and targeting.

Primary hepatocytes Dissociation of rat liver tissue into isolated cells that are suitable for culture was first described over 25–30 years ago [4]. However, the loss of structural polarity and mixing of membrane domains made them poor models for studies of polarity. The development of isolated rat hepatocyte couplets was a significant advance for short-term studies of polarized hepatocyte functions, such as transcytosis, and bile formation and Ca regulation of bile flow [39, 40]. As the importance the extracellular matrix (ECM) plays in maintaining gene expression and promoting cell repolarization became apparent, investigators began culturing hepatocytes on different matrix components in various geometries and physical states. The most successful reconstitution of polarity has come from use of sandwiches of collagen gels or Matrigel (see [41–44]). Despite these advances, the limited life span of primary hepatocytes and the difficulties in reproducibly obtaining such polarized cultures prompted many investigators to use secondary hepatocyte cell lines.

HepG2 cells The human hepatoma HepG2 (Table 6.1) is currently being used for studies of polarized membrane traffic. This line, which expresses many liver-specific genes, was

generated in the late 1970s from liver tumor biopsies in which the histology presented as “well-differentiated hepatocellular carcinoma with a trabecular pattern” [34]. The existence of bile canalicular-like cysts within and between cultured HepG2 cells was first reported in 1990 [45]; limited characterization has been reported despite extensive use of these cells, whose polarity has been estimated to be ∼10–40 [46]. Such a low polarity index necessarily limits one to morphological approaches. Hoekstra, van Izjendoorn, and colleagues have studied sphingolipid and PM protein traffic in these cells (reviewed in [46–49]), as have others (e.g. ATP7B [50–52], pIgA receptor [53, 54]; SR-B1 [55], MRP2 [56]). The field needs development of a HepG2 line with reproducibly high polarity.

WIF-B cells D. Cassio and colleagues have generated many somatic cell hybrid lines in their studies of the genetic basis of liver-specific gene expression [57, 58]. One line (WIF) was derived from fusion of the differentiated (i.e. expressing liver-specific genes) but unpolarized rat hepatoma cells (Fao) with human skin fibroblasts (WI-38 cells). Subsequent clonal selections yielded many polarized lines, which have been carefully characterized for hepatic functions (reviewed in [59]). We have used the WIF-B cell line (Table 6.1) in our studies of PM biogenesis, after first characterizing the mature phenotype with regard to its polarity [37] (which can reach 80% or more under the appropriate culture conditions), microtubule organization [60], and the structural and functional properties of the TJ boundary [37]. This systematic characterization convinced us that the WIF-B cells were a suitable model for the study of membrane trafficking and targeting in hepatocytes. We and others have used these cells, or the WIF-B9 clonal derivative [61], for studies of many PM proteins (see [59]). In vitro models always have limitations that it is important to be aware of, and the polarized WIF-B cells are no exception. To achieve maximal polarity, the cells require strict culture conditions. Their “bile canaliculi” are spherical and closed, making difficult the types of experimentation performed in simpler, columnar epithelia where there is easy access to the apical PM. The extent to which this bile canalicular morphology reflects a cholestatic state is not known. A new polarized hepatic line, Can-10, derived from rat Fao cells [58], has an extensive, interconnected bile canaliculus, which is similar to that in vivo [62]. More details on WIF-B cells are presented at www.bs.jhmi.edu/wifb. Other hepatic lines, whose characteristics are reviewed elsewhere [59], have not been shown to be polarized.

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Table 6.1 Polarized epithelial cell lines used to study aspects of PM traffic MDCK Madin–Darby canine kidney

LLC-PK1 Lilly Laboratories Culture pig kidney

Caco-2 Human colorectal carcinoma

FRT Fisher rat thyroid

HepG2 Human hepatoma from lung exudate

WIF-B Human fibroblast–rat hepatoma hybrid

Renal proximal tubule N—[13]

Small intestine

Thyroid epithelium Y—[12, 14]

Hepatocyte

Hepatocyte

MAL?

Renal distal tubule Y—[11, 12]

N—[15, 16]

N—[15]

MAL2?

Y—[17, 18]

Unknown

Y—[19]

Y—[17, 20]

AP-1A? AP-1B? Early refs

Y—[21] Y—[21–23] Y—[21, 23] Y—[21, 25, 26] N—[21, 27, 28] Y—[21, 23] [29, 30] [31] [32]

Unknown Unknown [7, 33]

Unknown Unknown [34–36]

Y—P. Tuma, personal communication Y—[24] N—[24] [37, 38]

Name/origin In vitro model

N—P. Tuma, personal communication Y—[17]

The presence of selected molecular features of cell lines are designated, Y-yes, N-no

PM BIOGENESIS: SIGNALS, SORTING MECHANISMS, DELIVERY SYSTEMS Early work The pathways and mechanisms of post-TGN membrane protein trafficking and lipid transport to the PM have been studied intensely (reviewed in [63–67]). Early work in simple epithelial cells, predominantly the MDCK cells, demonstrated that most newly-synthesized PM proteins were sorted in the TGN into vesicles that delivered them directly to their appropriate PM destination without first appearing at the opposite or both PM domains. This was called the “direct” route, which predicted that at least two PM-destined vesicles containing either basolaterallyor apically-directed cargo budded from the TGN in the same cell. The prediction was confirmed using two ectopically expressed viral proteins in non-polarized cells and in MDCK cells [2, 68]. Extensive dissection of each “direct” pathway in MDCK cells using both natural and reporter PM proteins subsequently revealed the existence of multiple routes, which included endosomal compartments. It is nonetheless important to remember that this is one cell type with distal tubular cell features. Other epithelial cells, including the LLC-PK1, Caco-2, and FRT lines, show important and useful differences, which are increasingly appreciated and being exploited. In contrast to the results in MDCK, in studies conducted on hepatocytes in vivo we reported that newly-synthesized single-transmembrane domain (TMD) apical PM pro-

teins took an “indirect” route via the basolateral PM before transcytosing across the cell [69]. Subsequently, we showed that the glycosyl-phosphatidyl-inositol (GPI)-anchored apical protein, 5′ nucleotidase, followed the same indirect route [70], which is generally called “transcytotic.” As in many simple epithelia, the basolateral PM proteins we examined took a “direct” route to the basolateral PM in vivo. At the same time, both direct and indirect routes to the apical surface were found to operate in the delivery of several single-TMD apical proteins in intestinal epithelial cells in culture and in vivo [71, 72]. As more epithelial cell types and classes of PM proteins were studied, it became quite clear that steady-state PM protein localizations are accomplished in a variety of ways. In Figures 6.2–6.4, we distill this large body of work into three diagrams, which depict the multiple pathways shown to be taken by basolateral and apical PM proteins in four epithelial cell lines. Below, we highlight new work, which has advanced understanding of PM recognition and targeting mechanisms.

Signals How do integral PM proteins delivered to a single PM domain find their way into the right intracellular path? Structural signals, such as amino acid sequences, conformations, or modifications are on the protein itself. These specify a correct destination that is decoded by a recognition mechanism, which directs the protein into the appropriate membrane carrier (vesicle/tubule) for correct delivery. Multiple PM signals may be present on a single PM protein, but one will prevail in a particular location.

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THE LIVER: PM BIOGENESIS: SIGNALS, SORTING MECHANISMS, DELIVERY SYSTEMS

APICAL 1

2

BSEP -myo2LC -myoVb -rab11a

MRP2 MDR1 ABST

3

4

GPI-Prots Influenz. HA -PI4P -FAPP2 -glu-cer/chol -rafts?

5

? Suc-Iso -annexin2

p75-NTR -LIM-K -Cofilin cortactin -dynamin -actin

6 ARE Lactase

? p75-NTR

CEACAM -galectin 3

TGN

TGF-a -Naked2

7

NaK-ATPase -clathrin-indep.

8

VSV-G, ASGP-R H1 (Y+) Tf-R1, LDL-R (non-Y) E-cad, CD147 (L+) NCAM (40aa), EGF-R (L+,P+) -clathrin-dependent

VSV-G -rab8,10,13 -optineurin AP-1B-indep. -myoVI -AP1 -clathrin

AP-1B-dep.

9

10

plgA-R (non-Y) -AP-2 -dlgA -clathrin -EE

11

BASOLATERAL

Figure 6.2 Biosynthetic pathways used to deliver PM proteins from the TGN (trans-Golgi network) to the basolateral and apical surfaces of MDCK Cells. The boxes associated with each pathway list the PM proteins (bold) documented to take that pathway. Also listed in the box are proteins and/or lipids documented to regulate the PM protein’s sorting and/or trafficking [73]. Abbreviations and references specific to each pathway are given with that pathway’s description below. Path 1—TGN-“indirect”–apical PM delivery of multi-TMD (BSEP, bile salt export protein, ABCB11, 1: [74], involving the apical recycling endosome, which uses rab11a and myoVb [75, 76]. Myo2LC (myosin 2 (regulatory) light chain) is required for BSEP’s apical delivery [77]. Path 2—direct TGN–apical PM delivery of three multi-TMD proteins, MDR1 (multi-drug resistance-1 (ABCB1)), MRP2 (MDR-related protein 2 (ABCC2)), and ABST (Na-dependent bile salt transporter), perhaps using a route and machinery similar to that of Path 1 [56, 78–80, 88, 89]. Path 3—direct TGN–apical delivery of a chimeric GPI-protein and influenza hemagglutinin (HA), both of which associate with detergent-resistant microdomains (DRM), requires a cytoplasmic protein (four phosphate adaptor protein 2) [81], PI4P (phosphatidylinositol-4-phosphate), perhaps glu-cer (glucosyl ceramide), and chol (cholesterol) in the donor TGN membrane [82]. Path 4—direct apical PM delivery of a single-TMD protein, Suc-Iso (sucrase-isomaltase), which associates with detergent-resistance microdomains and requires annexin 2, a cytoplasmic protein that binds cholesterol [83–85]. Path 5—direct TGN–apical delivery of p75-NTR (protein of 75 kDa/neurotrophin receptor), which requires LIM-K (LIM-kinase) and at least four other cytoplasmic proteins to be sorted and released from the TGN [86]. Path 6—direct TGN–apical delivery of three single-TMD proteins, lactase, p75-NTR (protein of 75kDa/neurotrophin receptor), and CEACAM (carcinoembryonic antigen-related cell adhesion molecule, sialo-glycoprotein of 114 kDa), which require a secreted protein, galectin 3 (for clustering?) [85]. Path 7—direct TGN–BL delivery of a single-TMD TGF-alpha precursor protein, which requires Naked 2, a cytoplasmic cargo recognition and targeting protein [87]. Path 8—direct TGN–BL delivery of a multi-TMD resident protein, NaK-ATPase, which does not require clathrin for delivery but does require non-erythroid ankyrin and spectrin to be retained at the BL surface [5]. Path 9—clathrin-dependent sorting of many single-TMD BL proteins, including four proteins that bind adaptor protein-1B, AP-1B (VSV-G (vesicular stomatitis virus) glycoprotein and ASGP-R (asialo-glycoprotein receptor, H1 subunit), through their cytoplasmic tail Y+ (tyrosine-containing) motifs; TfR1 (transferrin receptor 1) and LDL-R (low density lipoprotein receptor), through a non-Y (tyrosine) motif), and four proteins that require clathrin but do not interact with the known clathrin adaptors (E-cad (E-cadherin) and CD147 (also EMMPRIN or CE9), which have L+ (leucine-containing) motifs; NCAM (neural cell adhesion molecule), with a 40 aa sorting signal; and EGF-R, whose cytoplasmic tail contains at least one bipartite BL sorting signal (L+P+, leucine and proline-containing)) [90–92]. Path 10—indirect TGN–endosome–BL delivery of the single-TMD model protein, VSV-G, which requires several Rabs for high-fidelity sorting and/or targeting, the protein optineurin, which binds myoVI and rab8 [93]. Path 11—TGN–BL–endosomes–apical route taken by the pIgA-R (polymeric IgA-receptor). A 14 aa signal in tail (no Y or L) is necessary and sufficient for BL targeting [94]; additional machinery is not known (clathrin?). AP2-mediated endocytosis from the BL PM is followed by sorting in endosomes and delivery to the apical PM, where the ectodomain is cleaved and released with dIgA as secretory IgA. The fate of the 30 kDa stalk + TMD + tail is not known (reviewed in [95])

6: HEPATOCYTE SURFACE POLARITY: ITS DYNAMIC MAINTENANCE AND ESTABLISHMENT

APICAL

Caco-2

APICAL

FRT

1

2

3

4

1

GPI-prot -chot -GSLs -VAMP7

Lac -gal 3

Sl -anx2

ABST NHE3

Ap-GPI-chol -GSLs -oligomers -VAMP 7

TGN DPP4 APN -VAMP 8

79

2

DPP4 APN -VAMP 7

TGN ATP7A Cu1+

BL-GPI-chol -GSLs

6 Endo

3 5 BASOLATERAL

BASOLATERAL

Figure 6.3 Known biosynthetic pathways taken by select PM proteins in Caco-2 and FRT cells. (a) Pathways reported to deliver newly-synthesized PM proteins from the TGN (trans-Golgi network) to basolateral and apical surfaces of Caco-2 cells. The boxes associated with each pathway list PM proteins (bold) documented to take that pathway. Also listed are proteins and/or lipids documented to regulate the PM protein’s sorting and/or trafficking. Abbreviations and references specific to each pathway are given with that pathway’s description below. Path 1—direct TGN–apical delivery of GPI-proteins, which reside in detergent-resistant microdomains enriched in cholesterol and glycosphingolipids (chol + GSLs) and require VAMP7 (a v-SNARE) activity [96, 97]. Path 2—direct TGN–apical delivery of lactase, which requires clustering by a secreted protein, galectin 3 (gal 3) [83, 85]. Path 3—direct TGN delivery of sucrase-isomaltase (SI), which requires annexin 2 (anx 2, [98]), a cytoplasmic protein that binds cholesterol [85, 97]. Path 4—direct (presumed) TGN–apical delivery of two multi-TMD proteins, ABST (ileal bile salt transporter) and NHE3 (Na-H exchanger 3). ABST is presumed to use the same sorting signal and pathway as shown in Figure 6.2, Path 2. NHE3’s biosynthetic sorting signal has not studied, but apical retention is differentially regulated by three PDZ proteins, NHERF1, 2, and 3 [99, 100]. Path 5—TGN–BL–apical (transcytotic) delivery of two single-TMD glycoproteins, DPP4 (dipeptidyl peptidase 4) and APN (aminopeptidase N), which requires microtubules and VAMP8 (a v-SNARE) activity [96, 97]. Path 6—reversible (microtubule-independent) TGN–vesicle–BL delivery of ATP7A (Cu-ATPase) is activated by increased copper levels and requires a PDZ protein, AIPP1 (ATPase-Interacting PDZ Protein 1), to retain (presumed) protein in the BL region (small sub-BL vesicles?) [101–103]. Perhaps a di-leucine motif near the C-terminus constitutes a BL sorting signal [104, 105]. (b) Pathways reported to deliver select PM proteins from the TGN to basolateral and apical surface of FRT cells. See legend above (Caco-2 cells) for description of boxes. Path 1—direct TGN–apical delivery of (Ap)-GPI proteins, which reside in DRMs enriched in cholesterol and glycosphingolipids (chol+GSLs), oligomerize into SDS-resistant complexes, and require VAMP7 (a v-SNARE) activity [96]. Path 2—direct TGN–apical delivery of two single-TMD proteins, DPP4 and APN, requiring VAMP7 activity [96]. Path 3—direct TGN–BL delivery of BL-GPI proteins, whose GPI-attachment signals differ from those going apically; they reside in DRMs but do not further oligomerize like those in Path 3 [96]

The “Necessary and Sufficiency Rule” for a sorting signal

under study. Targeting (not a signal) is the delivery of carrier and cargo to the correct destination.

For a sequence, “patch,” or domain to constitute a sorting signal, it must be “necessary” for the intact protein to be included in the appropriate transport carrier. Mutation or deletion of the putative signal renders the mutant protein unrecognizable by the appropriate recognition machinery. Thus, the mutant is packaged incorrectly and sent to the wrong destination. In addition, when transferred onto a neutral reporter protein, which is normally sorted either randomly or to the opposite PM domain, the sorting signal is “sufficient” to redirect the reporter into the carrier going to the same PM domain as that of the wild-type protein

Known sorting BL sorting signals The overwhelming majority of BL proteins whose signals have been studied belong to the single-TMD protein class, and their signals reside in the cytoplasmic tails (reviewed in [116]). Most identified BL signals contain either a tyrosine in the context of a short degenerate sequence or a di-leucine motif. Proteins whose tyrosine signal overlaps with that used at the PM in receptor-mediated endocytosis are now thought to be a subclass, to which the reporter VSVG and the natural protein ASGP-R

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BASOLATERAL

2

1

Multi-TMD: AQPS (g’on) -cAMP BSEP (TC) -Pl 3-K -cAMP ATP7B (Cu 1+)

Alb HA321 CE9

7

8

Multi-TMD: NPLC 1

MTdep plus chol

BL-GPI hemojuvelin

minus chol

3 ARE TGN

TGN

APICAL

4

MDR 1

L

5

NTCP -cAMP -MTs

SA C LDL-R -PCSK 9

DPP4, APN, HA4, plgAR Ap-GPI 5’Nuc -chol+GSLs

Mutli-TMD SR-B1, OATP1 -PDZK1

EE

6

9

10

BASOLATERAL Figure 6.4 Known and hypothetical trafficking pathways of secretory and PM proteins in hepatocytes. The boxes associated with Paths 1 and 3–10 list PM proteins (bold) documented to take the pathway. Also listed are proteins and/or lipids documented to regulate the PM protein’s sorting and/or trafficking. Abbreviations and references specific to each pathway are given with each pathway’s description. Left: Pathways documented to deliver secretory and PM proteins from the TGN (trans-Golgi network) to the basolateral and apical surfaces. Path 1—biosynthetic delivery of single-TMD BL proteins HA321 (also BEN, DM-GRASP) and CE9 (also EMMPRIN, CD147, see Figure 6.2, Path 8). Path 2—biosynthetic delivery of secretory proteins from the TGN to the space of Disse. Alb (albumin), circles (very low-density lipoproteins). Refs [9] from last edition. Path 3—regulated (and reversible), microtubule-dependent (MT-dep.) recruitment of three multi-TMD apical proteins from TGN and/or an ARE (apical recycling endosome) to apical PM. AQP8 (aquaporin 8), g’on (glucagon; agonist acting via cAMP [106]; BSEP (bile salt export protein (ABCB11)); TC (taurocholate)) is agonist and PI3K activity (phosphatidyl-inositide-3 kinase) is required. MyosinVb and Rab11a operating at the ARE (apical recycling endosome) are also required [75], as are ATP7B (Cu-ATPase) and Cu1+ (both stimulator and substrate [107]). Path 4—direct apical delivery of MDR1, multi-resistance protein 1. Path 5—cAMP-stimulated BL delivery of NTCP (Na-dependent taurocholate co-transporting protein) from (presumed) EEs (early endosomes). Microtubules are required. NTCP [108]. Path 6—TGN–BL–apical (transcytotic) delivery of single-TMD apical proteins APN (aminopeptidase N), HA4 (cell–cell adhesion molecule of 105 kDa), pIgAR (polymeric immunoglobulin A receptor), and an Ap-GPI, apical glycosyl phosphoinositide protein 5’Nuc (5’nucleotidase). Cholesterol and glycosphingolipids (chol + GSLs) are needed for delivery from early endosomes (EE) through a sub-apical compartment (SAC) to the apical PM [76]. Right: Pathways likely taken by select hepatic PM proteins of cholesterol and iron metabolism. The boxes associated with each pathway list the classes and names of PM proteins (bold), which are reported to reside in the indicated PM domain, as well as accessory/regulatory proteins. Path 7—cholesterol-responsive, reversible trafficking of NPC1-L1, Niemann-Pick-C1-like 1, a cholesterol importer reported to retrieve biliary cholesterol [109]. Path 8—proposed biosynthetic route for a BL-GPI protein, hemojuvelin, involved in iron homeostasis [110–112]. Path 9—BL delivery of newly-synthesized LDL-R (low-density lipoprotein receptor) followed by its downregulation from the BL PM by an hepatic secretory protein, PCSK9 (proprotein convertase subtilisin kexin 9), that binds and directs LDL-R to lysosomes (L) for degradation [113]. The hepatic biosynthetic pathway/mechanism for BL delivery of LDL-R does not involve AP-1B, as shown in Figure 6.2, Path 9 for MDCK cells [24]. Path 10—presumed delivery and BL retention of two multi-TMD proteins, SR-B1 (scavenger receptor-B1, [114]) and OATP1 (organic anion transporter protein-1, [115]) by the multi-PDZ protein, PDZK1

6: HEPATOCYTE SURFACE POLARITY: ITS DYNAMIC MAINTENANCE AND ESTABLISHMENT

belong. Another BL protein subclass is defined by separate BL sorting and PM endocytic signals; a third has a single or di-leucine motif (which includes E-cadherin); and a fourth group has unique signals. Interestingly, the first three types of BL sorting signal are specifically recognized by the µ subunit of the Golgi tetrameric adaptor protein-1A (AP-1A) (Chapter 8). We will return to these subdivisions in the next section, when recognition mechanisms are discussed. A GPI anchor, long recognized as the clearest apical sorting signal, has recently been shown conclusively to constitute a BL sorting signal in FRT cells (Figure 6.3, Path 3). The GPI-attachment signals in these BL proteins apparently differ sufficiently from those in apical GPI protein to be recognized and sorted differently [96, 117, 118]. Hemojuvelin (Figure 6.4, Path 8) is an hepatic BL-GPI protein that plays a central role in iron homeostasis [110, 111]. Its GPI-attachment signal and final GPI anchor need to be identified. Finally, a localization signal in basolateral and apical PM proteins consists of short degenerate sequences, often at the C-terminus, that are recognized by PDZ proteins. Members of the fast-growing PDZ protein family, named after the three founding members (PSD-95, Discs Large, and ZO-1, reviewed in [119]), all contain one or more globular PDZ motifs that are 80–100 amino acids in length and are proposed to function as scaffolds in multiprotein complexes. The function of PDZ proteins that bind to PM proteins may be to retain the molecule once it has reached the basolateral PM rather than to sort it into basolaterally-destined vesicles in the TGN [120]. However, a targeting function has been ascribed to several artificial PDZ modules [121]. PDZ proteins that bind apical PM proteins have also been identified (e.g. [122]), but whether they serve a sorting or a retention function is not yet clear [123].

Apical sorting signals As mentioned above, a GPI anchor is the clearest apical sorting signal, since its transfer onto a randomly secreted protein is sufficient to redirect the protein into the apical pathway of most cells [124]. The transmembrane region of some single-TMD proteins (e.g. influenza hemagglutinin (HA); Figure 6.2, Path 3) may share similarities with the GPI anchor and be sorted by a similar mechanism (see next section). Yet another class of sorting apical signal has gained prominence with the identification of a possible recognition mechanism. O- or N-glycan modifications were postulated to be recognized and sorted by luminal lectins, one of which may be galectin 3 (Figure 6.2, Path 6).

81

Recognition mechanisms Basolateral mechanisms The existence of two AP-1 isoforms and their differential localization in MDCK cells, together with early trafficking studies reporting differences among several newly-synthesized BL PM proteins in the pathways taken in LLC-PK1 versus MDCK cells, spawned numerous subsequent studies which have revealed a fascinating complexity to the AP-1 recognition mechanisms, their functional locations, and the role of clathrin in apical sorting and delivery. There is differential expression of the two AP-1 isoforms, with LLC-PK1 cells lacking the B isoform (Table 6.1). Paths 9 and 10 in Figure 6.2 list the BL proteins taking an AP1-clathrin-dependent pathway as well as several cytoplasmic targeting and motor proteins that have been implicated [125]. The essential role of clathrin in BL targeting appears firmly established with the elegant study of Rodriguez-Boulan and colleagues [90]. The finding that the BL PM protein NCAM (which has a unique sorting signal containing neither Y nor L) requires clathrin indicates that yet another adaptor must be involved in BL recognition. Several other BL targeting mechanisms have been identified. The most unusual is that of Naked 2, a cytoplasmic protein that specifically binds the tail of the TGF-alpha precursor, which is synthesized as a single-TMD precursor and released after cleavage at the BL PM (Figure 6.2, Path 7) [87, 126]. The second most unusual is the PDZ protein PDZK1 (NHERF3), which is required to localize two BL TMD proteins in hepatocytes (Figure 6.4, Path 10).

Apical mechanisms A tetra-spanning TMD protein of ∼17 kDa, first identified in myelin and lymphocytes (hence the name MAL), has been implicated in the apical sorting of a single-TMD apical protein surrogate, the influenza HA, by Puertollano et al. [11]. MAL is expressed in many epithelial cells, where it is concentrated in the TGN region [127]. Using an anti-sense approach to study the apical delivery of HA in MDCK cells lacking MAL, these investigators found HA’s transport to be less efficient and less specific in the absence of MAL; the ectopic expression of human MAL rescued the defect. Therefore, it is intriguing that liver does not express MAL, a finding consistent with the absence of a direct apical delivery mechanism for the single-TMD class of apical PM proteins in hepatocytes. When exogenous MAL was expressed in WIF-B cells, which lack it, a GPI and several single-TMD apical PM

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THE LIVER: PM BIOGENESIS: SIGNALS, SORTING MECHANISMS, DELIVERY SYSTEMS

proteins were redirected from the transcytotic to the direct pathway [15]. Membrane domains, termed “rafts,” that are enriched in glycoplipids and cholesterol have been proposed as “platforms” upon which apical PM protein sorting occurs in MDCK cells (reviewed in [128–130]). The rafts are defined operationally as detergent-insoluble, lipid-containing complexes that float in sucrose density gradients [133]. As some newly-synthesized single-TMD and GPI-anchored PM proteins transit through the Golgi, they appear in such complexes. This behavior correlates with the efficiency of subsequent delivery to the apical PM. Lowering cholesterol levels through use of metabolic inhibitors and/or reagents that extract cholesterol acutely from living cells disrupts the rafts (i.e. decreases the detergent insolubility of the proteins) and reduces the specificity of apical delivery of some proteins ([134], but see [135]). Although there is now information that detergent-resistant microdomains exist in the hepatocyte PM and Golgi [136, 137], their possible role as a recognition system is in doubt. Cholesterol and glycosphingolipids are required for transcytotic efflux from early endosomes of WIF-B cells [137], but DRMS seem not to be involved.

PM targeting machinery How do cargo-bearing vesicles deliver their contents to the correct target domain? Morphological, biochemical, and genetic approaches have successfully identified many molecules involved in “vesicle targeting,” a complex series of molecular events that minimally includes docking and fusion. These approaches indicate that the basic mechanisms are conserved among the membrane compartments throughout the biosynthetic and endocytic pathways and in organisms ranging from yeast to humans. The players so far identified fall into two broad categories: some are used repeatedly throughout the pathways, others belong to discrete protein families, where one or a few members act at one or a few transport sites. Members of three protein families are central players in vesicle targeting and fusion (for recent reviews, see [125, 138]). They are: (i) the SNAREs, which, in general, are a group of cytoplasmically oriented integral membrane proteins that are present on vesicles (v-SNAREs) or target membranes (t-SNAREs); (ii) the Sec1/Munc18 proteins; and (iii) small-molecular-weight GTP-binding proteins, the Rabs. Recent reviews cover the current status of SNARE hypothesis and components [131, 132]. We focus briefly on the Rab proteins, since many have now been implicated in PM protein trafficking. The Rab proteins belong to the largest family of small-molecular-weight (20–30 kDa) GTP-binding proteins, with >60 at last count (for recent reviews, see [139–141]). Examination of transfected cells overexpressing either wild-type or dominant-negative mutant (usually

the GDP-bound conformer) forms of various Rabs shows they either stimulate or inhibit protein transport and in some cases alter organelle morphology. They have been shown to function in one of three ways: (i) by facilitating vectorial traffic via associations with the cytoskeleton; (ii) by regulating vesicle docking by recruiting effector molecules, thereby promoting the formation of “molecular tethers”; or (iii) by “priming” docking and fusion by activating SNARE molecules [142, 143].

What about the Hepatocyte? Compared to the impressive recent advances in our understanding of sorting signals, recognition machinery, and pathways in simple epithelial cells, PM protein trafficking in hepatocytes remains enigmatic. Below, we raise three questions that we would like to see answered. • Is (are) the apical sorting signal(s) in each multi-TMD protein unique?: The formation of bile is a central function of hepatocytes and a process requiring many multi-TMD apical PM proteins. Additional export processes are carried out at the canalicular PM by other multi-TMD proteins (Figure 6.4, Paths 3 and 4). The apical transporters studied to date appear to have unique and complex sorting signals, and the recognition systems have yet to be identified. More study is needed, particularly of patient mutations whose protein phenotypes (in hepatic cells) may give clues about sorting signals (e.g. [107]). • Are several apical PM protein recognition systems absent from hepatocytes?: The clear difference in the biosynthetic pathways taken by all tested GPI-anchored and single-TMD apical PM proteins in hepatic cells (transcytotic) versus those in MDCK cells (direct) suggests that the former cells lack certain key recognition mechanisms operating in the latter. We know that apical sorting signals are present on the hepatic PM proteins, because several have been expressed in MDCK cells and shown to be trafficked directly to the apical PM. The two best-characterized systems from work in MDCK cells are those recognizing lipid raft-associated and O- or N-glycans on apical PM proteins (Figure 6.2, Paths 3 + 4 and 5 + 6, respectively). This suggests to us that components of the FAPP2/PI-4P and galectin 3/LIM-kinase recognition systems (or their homologs) are either missing, inactive, or located outside of the TGN in hepatic cells. The relevant proteins should be studied. • How do hepatic BL protein sorting, recognition, and targeting work?: The recent results showing that clathrin plays a central role in the sorting of many single BL PM proteins are exciting (Figure 6.2, Path 9) [90]. Multiple clathrin adaptors appear to be involved, in addition to the two AP-1 isoforms, AP-1A

6: HEPATOCYTE SURFACE POLARITY: ITS DYNAMIC MAINTENANCE AND ESTABLISHMENT

and AP-1B. Since the latter AP-1 isoform is absent from hepatic cells, the question is how do hepatic BL PM proteins needing this adaptor get properly sorted? Study of the biosynthetic pathway taken by a disease-causing mutant low-density lipoprotein (LDL) receptor in polarized WIF-B cells has given some clues [24]. The mutation, G823D, is part of the BL sorting signal, and the mutant BL protein took a transcytotic route to the apical PM, including a stop at the BL PM. These results, together with those of an additional mutant LDL receptor lacking both BL and PM endocytic sorting signals, suggested that hepatic AP-1A must be acting, at least, in the hepatic BL endosomal compartment in order to recognize an internalized LDL receptor and redirect it back to the BL PM. Such a scenario is different from that operating in MDCK cells, where AP-1A operates in the TGN and AP-1B in recycling endosomes [22, 23, 25, 63]. Thus, we hypothesize that normal sorting of single-TMD BL and apical PM proteins in hepatic cells occurs predominantly or exclusively in basolateral endosomes, not the TGN. “Why?” is a difficult question to answer, but it may be the (evolutionary?) consequence of a central homeostatic function performed by hepatocytes—that of lipid and cholesterol regulation and trafficking of a major secretory (TGN) pathway process. Testing this hypothesis will be a challenge.

ESTABLISHING EPITHELIAL CELL POLARITY Development of hepatic polarity In Vivo Though hepatocyte polarity is well-described morphologically, we know very little about the molecules required for polarity formation or their temporal expression, because there has been no ideal system for study. Since the liver is a hematopoietic organ early in development, genes expressed during this time represent at least four programs: hematopoietic, mitogenic, liver-specific (functional differentiation), and morphogenetic (polarity). Nonetheless, there is good information on liver development (see Chapter 2 and [144]). During mouse development, a liver bud composed of non-polarized hepatoblasts is detected at embryonic day 9.5, when it is already a hematopoietic organ. Our own studies have demonstrated that expression of hepatocyte polarity occurs very early in development (day 15, [145]). From embryonic days 17 to 21, the hepatoblasts form clusters resembling acini, which exhibit a simple polar phenotype, with their apical surfaces facing a central lumen (see [145, 146]). The multi-polar hepatic phenotype gradually

Simple epithelial cell

83

Hepatocyte

apical PM +

+ − −

− −

Golgi

−

nuc + +

−

nuc

Golgi

+ + basolateral PM

+

− Golgi apcial PM

nuc −

+

Polarized Phenotype

Feature

Polarized Phenotype

Minus ends-apical

Microtubules

Minus ends-apical

Exposed

Apical PM

sequestered

Direct (TGN→Apical) Apical PM protein route Indirect (transcytotic)

Figure 6.5 Simple and complex polarity. Features of simple epithelial cells (a) and hepatocytes (b). In simple epithelial cells, a single exposed apical and basal surface oppose each other and lateral surfaces participate in cell–cell associations forming a sheet or tube structure. Hepatocytes are polygonal and multipolar, with at least two basal surfaces facing the circulation and a branched network of grooves that forms between adjacent cells and constitutes the apical, or bile canalicular, surface. Black rectangle designates TJs; nuc, nucleus; +/–, microtubule orientation

manifests itself during the postnatal period, although cell clusters whose apical poles face a small lumen are still evident 12 days after birth [146]. In fact, characterization of the hepatic cell line WIF-B9 demonstrates that they recapitulate this biphasic polarity phenotype in culture [61]. The molecular mechanisms directing conversion from simple to polygonal hepatic polarity and cord organization of the adult liver are unknown. A working model for the development of epithelial cell polarity postulates that extracellular signals generated from cell–cell and cell–substrate interactions initiate the multiple cellular processes that must occur to form a polarized epithelium [3]. These processes include: (i) an organized and stabilized cytoskeleton; (ii) sorted apical and basolateral components; (iii) arrangement of intracellular organelles [147]; and (iv) formation of AJs and TJs to act as a diffusion barrier. Most epithelial cells exhibit a simple polarity, consisting of single apical and basal surfaces that are opposite each other and lateral surfaces that participate in cell–cell associations to form a sheet or tube structure. In contrast, hepatocytes are polygonal and multi-polar, with at least two basal surfaces facing the circulation and a branched network of grooves that forms between adjacent cells and constitutes the apical, or bile canalicular, surface (Figure 6.5). Recent studies have increasingly pointed to the TJ and AJ as important macromolecular complexes that establish the apical and basolateral poles and/or serve as landmarks for vesicle targeting in epithelial cells (e.g. exocyst, [148]); however, this model is being challenged by several recent findings, as discussed below.

84

THE LIVER: ESTABLISHING EPITHELIAL CELL POLARITY

Polarity complexes play key roles in establishing cell polarity Advances in understanding the development of epithelial cell polarity have come from studies of model genetic systems such as Drosophila and C. elegans. Three membrane-associated protein complexes required for this process have been identified (Table 6.2a, [149–152]). The components have been localized to the basolateral membrane and to junctional elements at the apical-lateral margin (ALM) of various epithelia exhibiting a simple polarized phenotype [149, 153]. Many proteins have one to multiple PDZ domains, which are proposed to cluster and coordinate other molecules at the cell cortex, thereby promoting development of apical/basal polarity [154, 155]. The PAR–aPKC complex was identified in C. elegans by its requirement in asymmetric cell division of the one-cell embryo [155]. Studies of this complex in flies demonstrate its essential role in establishing apical/basal polarity [156], yet neither the membrane anchor nor the mechanism of membrane association is known for either model system. Interestingly, mammalian studies reveal that junction adhesion molecules (JAMs) bind PAR3 [159] and anchor this complex at the TJ, a structure maintaining the boundary between the apical/basal surfaces and regulating paracellular permeability in an epithelial sheet [157, 158]. Two other complexes were identified genetically in Drosophila. One complex is composed of Scribble, Discs Large (both PDZ proteins), and Lethal Giant Larvae (LGL). The other complex includes the transmembrane protein Crumbs, which is required for the organization of epithelia [186]. Crumbs localizes to the most apical aspect of the ALM and recruits the PDZ proteins Discs Lost [365] and Stardust [366, 367]. Disruption of any one complex member leads to general loss of epithelial organization and overproliferation [149]. Genetic studies in Drosophila demonstrate that it is the integrated activity of these polarity-specifying complexes that regulates epithelial polarity [186, 368]. Study of these complexes in mammalian systems demonstrated that conserved mechanisms regulate apical/ basal polarity as well as AJ and TJ formation (see citations in Table 6.2(a) and [64]). In fact, loss of the single Polarity complex protein, mouse PAR3, altered the organization of an epithelium [161]. From Table 6.2a, it is obvious that study of these polarity complexes in hepatic cell polarity is needed.

The junction proteins have key roles in establishing polarity Animal models and cell culture systems are being used to identify and determine the roles of junction proteins in establishing/maintaining apical/basal polarity. While

most junction proteins listed in Table 6.2 are expressed in hepatocytes, their roles in establishing hepatic polarity are largely unknown. There is a growing collection of liver-specific knockout (KO) animals with deletion of genes shown to be important for establishing polarity. Surprisingly, although many animals develop hepatocellular carcinoma (HCC), some KO animals exhibit no reported liver pathology. Because HCC in humans is often triggered by viral infections or cirrhosis, we propose that study of these KO animals (e.g. occludin, ZO-3, PAR1, see Table 6.2) following liver insult may provide clues regarding the role of the missing protein in hepatic polarity. Additionally, a number of the polarity proteins and mediators listed are misregulated in human HCC (Table 6.2). Thus, there is a strong correlation between the development of HCC and altered polarity protein expression. Although no pattern has emerged, Table 6.2b,c lists many of the polarity proteins found at mammalian AJs and TJs, their role in establishing epithelial simple polarity, and any documented role in establishing hepatic polarity. Current reviews and key references are also provided. While this list is extensive, a recent proteomics study uncovered additional TJ proteins that deserve further attention [369].

Adherens junctions The cadherin family of Ca2+ -dependent cell–cell adhesion proteins is recognized as essential for the formation of a properly organized epithelium [197] (Table 6.2b). They are multifunctional since they are involved in differentiation, cell proliferation, signaling, and cell fate. The E-cadherin/β-catenin complex was long thought to bind the underlying actin cytoskeleton directly and act as an anchor in order to establish AJs. Recent studies now indicate that the cadherin/catenin complex associations with actin are dynamic [370]. How do these dynamics impact AJ formation and epithelial polarity? Several groups used KO studies to address the role of the cadherin/catenin complex in establishing cell polarity. Thus it was surprising that upon knockdown of E-cadherin, AJs appeared to be intact in HepG2 and MDCK cells [198, 371]. The finding that E-cadherin knockdown in HepG2 cells did not alter TJ formation or polarity is also puzzling [198], given that its loss in MDCK cells prevented TJ assembly after Ca2+ switch but not its maintenance [371]. In contrast, hepatocyte-specific knockdown of β-catenin inhibited development of hepatic cell polarity, hepatoblast expansion, and survival [201]. A delay in hepatocyte proliferation following partial hepatectomy was also observed [372]. Taken together, these results indicate that regulation of E-cadherin and β-catenin levels is important for hepatocyte AJ and possibly TJ assembly. Animal KO of afadin, which binds nectin, results in embryonic lethality [213]. To study proteins whose deletion is embryonic-lethal, KO embryonic stem (ES) cells

Functional domains

S/T kinase

aPKC

MAGUK, PDZ, SH3

Tumor suppressor; Lgl2 phosphorylated by Par6/aPKC [165, 166]

Tumor suppressor; likely a scaffold [180]; binds oncoviral protein PBM [181]

Tumor suppressor; targeted degradation via oncoviral protein PBM [175]; Scrib mutant mice (circletail) exhibit no polarity defects [176]

Scaffold; binds aPKC [160]; JAM [159], required for establishing apical cortical domains of epicardial progenitor cells [161] Links Cdc42/aPKC to the downstream effectors PAR3 [150, 151] and Lgl2 [165] [166]; negatively regulates TJ assembly [167] Plays critical role in establishing TJ structures [169] through regulation of actin [170]; phosphorylates Lgl [165], PAR3 [171], PAR1b [172], and Drosophila Crumbs [173]

Role in polarity (reports)

E-cadherin

Cell adhesion glycoprotein, Cadherin repeat domain, 1 TM

Ca2+ -dependent cell–cell adhesion, phosphorylation-dependent binding of β-catenin and p120 catenin (see review)

Binds adaptors PAR6 [185]; Stardust [186]; PALS [187]; required for TJ formation in mammary MCF10A cells [188] PALS (protein MAGUK, PDZ , SH3 Scaffold; binds Crumbs, PATJ [187], and PAR6 [185]; associated with reduced expression results in multiple polarity lin seven 1, aka defects [190]; its loss reduces E-cadherin trafficking Stardust) [191] PATJ (aka hINDL) Multiple PDZ domains Scaffold; binds PALS1 [192]; reduced expression delays TJ formation [193]; mislocalization by oncoviral protein via PBM disrupts TJ [194] (b) Proteins associated with adherens junctions

Lgl (Lethal Giant WD40; C-term Larva) 1 and 2 Lgl-specific domain Crumbs/PALS/PATJ Crumbs (crumbs3 1 TM, FERM binding in mammals) motif, PBM

DLG (Discs Large)

Scribble/DLG/LGL Scribble LRR, 4 PDZ

PB1, CRIB, 1 PDZ

PAR6

(a) Polarity complexes PAR–aPKC complex PAR3 (partitioning 3 PDZ domains defective)

Protein

ND

[195, 196]

Y—E-cadherin/β-catenin-based adherens junctions are dispensable for TJ formation and apical surface formation in HepG2 cells [198] (continued overleaf )

ND

[182, 189]

[197]

ND

Y—Present in WIF-B [179] and liver primordium; at E10.5 liver is overgrown in circletail mutant mice [176] Y—Binds two GPCRs found on endothelial cell membranes in embryonic liver [183] ND

Y—TJ and apical surface WIFB and liver [163]

Y—TJ in hepatocytes and WIFB [163]

Y—TJ in hepatocytes and WIFB [163]; downregulated in HCC [164]

Present in liver? Y/N—finding and reference

[122, 162, 189]

[184]

[182]

[177, 178]

[174]

See above and [168]

[162]

Reviews

Table 6.2 Catalog of proteins associated with (a) Polarity complexes, (b) adherens junction (c) tight junctions (d) promoting polarity (polarity mediators) (e) the hepatic-like phenotype in MDCK cells and (f) polarity and vesicle trafficking 6: HEPATOCYTE SURFACE POLARITY: ITS DYNAMIC MAINTENANCE AND ESTABLISHMENT 85

[209]

Ca2+ -independent cell–cell adhesion; bind afadin (see below, [208])

Cell adhesion molecule of Ig super family, 1 TM, C-term PBM N-term RAS binding domain, PDZ

4 TMs, PBM 4 TMs, PBM

2 Ig extracellular loops, 1 TM, PBM 2 Ig extracellular loops, 1 TM, and PBM 2 Ig extracellular loops, 1 TM, PBM

Claudin 2

Claudin multigene family

JAMA (junction adhesion molecule)

JAMC

CAR (coxsackie and adenovirus receptor) and other JAM family members

4 TMs, PBM

Claudin 1

(c) Proteins associated with tight junctions Occludin 4 TMs

Afadin (aka AF-6)

Roles unknown; ZO-1 co-precipitates with CAR [236], JAM-B KO mouse viable [237]

Selective TJ “gate” function; associated with TJ strands [230]; binds PAR3[159, 230]; ZO-1/ZO-2, afadin [212] Selective TJ “gate” function [234]; required for spermatid differentiation [235]

Role unclear; mouse KO is viable with complex abnormalities in various organs [215]; loss of occludin alters tricellulin localization [216] Selective TJ “gate” function [219]; binds ZO-1, ZO-2, ZO-3 [220]; KO demonstrates essential role in epidermis [221] Selective TJ “gate” function [219]; binds ZO-1, ZO-2, ZO-3 [220] Selective TJ “gate” function [227]; binds ZO-1, ZO-2, ZO-3 [220]

Adaptor, binds actin, many scaffolding and F-actin binding proteins. Also binds nectins (AJ) and JAM (TJ, [212]). KO is embryonic-lethal [213]

[205, 206]

Gamma-catenin (plakoglobin) KO mice develop hair coat abnormalities [204]

Armadillo repeat motifs

Other catenin isoforms (i.e. p120, gamma-catenin, aka plakoglobin) Nectin family

See above and [158, 238]

See above

[157, 231, 232]

[228]

[225]

[222]

[158, 217]

[214]

[197]

When phosphorylated, binds cadherins and alpha-catenin; mouse KO embryonic-lethal [199]

Armadillo repeat motifs

β-catenin

Reviews

Role in polarity (reports)

Protein

Functional domains

Table 6.2 (continued)

Y—[238, 239]

Y—Present in hepatocytes by RT-PCR [163]

Y—Loss correlates with HCC [223]; TGF-beta induces EMT and loss of claudin 1 expression [224] Y—KD in WIF-B9 loss of apical surface [226] Y—Claudin 1, 2, 3, 5 present in liver [59]; increased Claudin 10 in HCC [229] Y—Essential and inhibitory to hepatic polarity [163], also see [233]

Y—KO cells apoptotic and claudin 2 expression increased [218]

ND

Y—Critical for postnatal liver growth [200]; liver-specific KO disrupts hepatocyte morphogenesis and survival [201]; role in rat HCC [202]; mutations in exon-3 found in HCC [203] Y—Expression of three catenins (gamma) is upregulated in HCC, with least differentiated having greatest expression [207] Y—Northern blot shows abundant levels of Nectin-2 and -3 [210, 211]

Present in liver? Y/N—finding and reference

86 THE LIVER: ESTABLISHING EPITHELIAL CELL POLARITY

Globular head, central coiled-coiled domain Coiled-coiled domain similarity to cingulin

Cingulin

ABC signature motif, SMC family

MAGUK inverted orientation, WW, multiple PDZ domains

7H6 (aka barmotin)

MAGI-1 (aka BAP 1) and MAGI-3

NLS

3 PDZ, MAGUK, SH3

ZO-3

JACOP (junction associated coiled-coil protein), aka Paracingulin Symplekin

3 PDZ, MAGUK, SH3

3 PDZ, MAGUK, SH3

1 TM (type IV) with coiled-coil

ZO-2

VAP-33 (VAMP (vesicle-associated membrane protein)-associated protein, 33 kDa) ZO-1 (Zonula occludins 1)

CLMP 2 extracellular Ig (coxsackie-adenovirus loops, 1 TM receptor-like membrane protein) Tricellulin 4 TMs

Role unknown; presence at TJ is not dependent upon GUK or WW domains [266]; binds JAM-4 [267], ESAM [268], and oncoviral proteins in vitro and in fibroblasts [269]

Role unknown, also localized in nucleus in a wide range of cell types [261] Regulatory role in TJ barrier function [263]

Scaffold; binds claudins [220], occludin [243], and actin [244]; also in non-epithelial cells; KO embryonic-lethal [245] Scaffold; binds claudins [220], occludin [247], and actin [248]; also in non-epithelial cells, mislocalization by oncoviral protein PBM disrupts TJ [194]; KO embryonic-lethal [249] Scaffold; binds claudin [220], occludin [251], actin [248], and PATJ [252]; expression epithelial-specific, KO mice viable without TJ abnormalities [253] Scaffold [255]; binds ZO-1, ZO-2, ZO-3, and actin [256]; deletion of head domain does not alter TJ structure [257]. Scaffold; involved in anchoring the apical junctional complex to actin-based cytoskeleton [259, 260]

[158]

Important for barrier function, concentrated at tricellular contacts, C-term similar to occludin [241] Role unknown, implicated in vesicle docking/fusion [242]

[258]

[264]

[258, 262]

[258]

[258]

See above

See above

[182, 222]

—

—

Role unknown; widely expressed, mediates cell adhesion when exogenously expressed [240]

Y—Monoclonal antibody recognizing a novel hepatocyte TJ protein [265]; HCC severity in rat correlates with its loss at hepatocyte TJ [263] Y—Upregulated with increased TJ formation in connexin32 KO immortalized hepatocytes following connexin32 transfection; these cells exhibit a simple polar phenotype [270] (continued overleaf )

ND

Y—Localized to hepatocyte TJ by immunoEM [259].

Y—Localized to hepatocyte TJ with JACOP [259].

Y—Abundant transcripts and localized to hepatocyte TJ; [254]

Y—Localized to hepatocyte TJ [250]

Y—First isolated from liver [246]

Y—Liver cDNA library used to identify occluding-binding proteins [242]

Y—Present in liver by Northern blot [241]

Y—Low abundance message and protein [240]

6: HEPATOCYTE SURFACE POLARITY: ITS DYNAMIC MAINTENANCE AND ESTABLISHMENT 87

[300]

[301]

Energy-dependent regulation of TJ assembly in MDCK cells [306] Regulator of translation; phosphorylates 4E-BP1 and p70 Sk kinase [307]; involved in neuronal polarity [308]

S/T kinase heterotrimeric complex S/T kinase

AMPK (AMP-activated protein kinase) mTOR (mammalian target of rapamycin)

[299–302]

Tumor suppressor; master kinase [295]; mutated in PJS [296, 297]; cell polarity of a single intestinal cell occurs upon activation [298]

S/T kinase; farnesylated at C-terminus

[290]

LKB1 (aka PAR4)

Required for gastrulation and regulated gene expression in the visceral endoderm [289]

Transcription factor

[285, 286]

[279–281]

[273]

[194, 258]

Reviews

HNF4α (hepatocyte nuclear factor)

SMADs (small mothers against deca-pentaplegic)

Cell–substrate adhesion; apical distribution required for MDCK cyst formation [277]; works with Rac1 and laminin to orient polarity [278] Intracellular mediators of TGF-beta signaling [283, 284]

Coiled-coiled proline-rich domain

1 TM; forms heterodimers with α-integrins Transcription factors

Role unknown; isolated using GUK region of DLG in a 2-H screen; does not overlap with DLG by IF but rather with ZO-1 at TJ in mouse small intestine [276]

Coiled-coil, PBM

AMOT (angiomotin)/JEAP Family PILT (protein incorporated later into TJs)

(d) Polarity mediators β1 integrins

Role unknown; a likely scaffold; binds JAM-A, claudins [271], CAR, and oncoviral proteins in fibroblasts [272] Role unknown [274]; binds MUPP1 and PATJ via PBM [273]; regulates endothelial cell–cell junctions [275]

Role in polarity (reports)

Multiple PDZ domains

Functional domains

MUPP1 (PATJ paralog)

Protein

Table 6.2 (continued)

Y—mTOR pathway activated in HCC; sustained activation impairs differentiation of hepatocyte polarity (HepaFR cells, [309]); MDCK cells grown in collagen gels with rapamycin (inhibits mTOR) exhibited the hepatic phenotype [310]

Y—β1 integrin KO ES cells used to establish chimeric embryos; integrin KO cells did not colonize the liver [282] Y—Smad2, Smad3 heterozygotes died at midgestation with liver hypoplasia, levels of β1 integrin decreased, and E-cadherin mislocalized [287]; TGF-beta/SMAD signaling in injured liver reviewed in [288] Y—Hepatocyte-specific HNF4 KO mice develop HCC [291]; HNF4 null liver does not show normal polarity or epithelialization [292–294] Y—LKB1 (+/–) mice develop HCC [303]; accelerated onset of hepatic adenomas/carcinomas in LKB (+/–) p53 (–/–) mice [304]; LKB1 activates AMPK in liver [305] Y—LKB1 activates AMPK in liver [305]

Y—Enriched in AJ and TJ fraction with DLG using subcellular distribution of rat liver [276]

Y—Family members AMOT and JEAP localized to TJ of hepatocytes [273].

Y—Present in bile canaliculi fraction isolated from mouse liver [273]

Present in liver? Y/N—finding and reference

88 THE LIVER: ESTABLISHING EPITHELIAL CELL POLARITY

lipid and protein phosphatase

Tumor suppressor; mediates apical distribution of phosphoinositides, which in turn control epithelial polarity through Cdc42 [311] Multiple, armadillo Tumor suppressor; binds microtubules [314]; regulates repeats, β-catenin levels [315]; involved in cell migration oligomerizes [316] Cdc42 (rho family of Small GTPase Binds PAR6 to generate polarity [319–321]; GTPases) knockdown in Caco-2 cysts alters spindle orientation, which in turn alters the position of apical surface [322] TIAM1 (T-lymphoma GEF Rac activator [325]; required for cadherin-based invasion and adhesion [326]; PAR3 controls TJ assembly via metastasis) TIAM1 [327, 328] ROCK Rho kinase Critical for TJ assembly and actin cytoskeleton organization [330], and polarity orientation [331] ERM proteins (ezrin, F(4.1) ERM domain, Members of FERM superfamily (see review); provide radixin, moesin) C-terminal F-actin regulated linkage between membrane proteins and binding site F-actin to interface signaling and the cytoskeleton; ROCK substrate [334]; moesin KO mice are normal [335]; ezrin KO mice have defective terminal web of intestinal cells [336] EBP50 (ERM-binding 2 PDZs, C-terminal Binds ezrin and localizes apically [340]; KO mice phosphoprotein, aka ERM binding exhibit mild phosphatemia, decreased levels of ERM NHERF1) domain proteins, aberrant intestinal microvilli with a thickened actin-rich terminal web [341] (e) Proteins that induce the hepatic-like phenotype in MDCK cells PAR1b (aka STK11 or S/T kinase MARK family of S/T kinases; phosphorylates PAR3, EMK1) MAPs and Rab11-FIP2; substrate for aPKC [172], KO mouse viable [343] Myosin II Motor head domain, Important for polarity orientation [331] tail domain Rho family Small GTPase RhoA frequently hyperactivated in epithelial cancers; works with Rock I and myosin II to orient polarity [331] Rac Small GTPase Responsible for trafficking of apical proteins in intestinal epithelial cell in mice (f) Vesicle trafficking proteins Rab8 Small GTPase Mouse KO studies show mislocalization of apical proteins [350]

PTEN (phosphatase and tensin homology) APC (adenomatous polyposis coli) Y—Liver-specific Cdc42 KO mice develop HCC [324]

Y—Overexpressed in HCC [329]

Y—ROCK2 frequently overexpressed in HCC [333] Y—Radixin is found at bile canaliculi; KO mice have hyperbilirubinemia with decreased MRP2 at bile canaliculi [339]

Y—Upregulated in HCC [342]

Y—First to show the hepatic phenotype develops in MDCK cells following overexpression of a polarity protein [345–347]

[323]

[327]

[332] [337, 338]

[99]

[344]

[140]

See above

[323]

ND (continued overleaf )

RhoA levels increased in HCC tissue [400]; hepatic phenotype develops in MDCK cells following overexpression of DN RhoA [349] Hepatic phenotype develops in MDCK cells following overexpression of active Rac [349]

Y—Liver-specific APC KO mice develop HCC, β-catenin signaling activated [318]

[317]

[348]

Y—PTEN hepatocyte-specific KO mice develop HCC [313]

[312]

6: HEPATOCYTE SURFACE POLARITY: ITS DYNAMIC MAINTENANCE AND ESTABLISHMENT 89

Small GTPase

Unconventional myosin Unconventional myosin

Rab13

Myosin Vc

C2 domain, C-terminal amphipathic α-helix

Rab11-FIP2 (family of interacting protein)

Membrane traffic regulator; direct and functional link with protein kinase A signaling during junction assembly in epithelial cells [351] Novel class expressed in epithelial tissues, likely involved in membrane trafficking [354] Role in vesicle trafficking via binding Rab11a [356]; mutations cause microvillus inclusion disease; defective trafficking, apically localized TfR propose a role in maintenance of cell polarity [357] Regulation of apical vesicle trafficking/transcytosis [359, 360] Binds Rab11a and dominant active form (Rab11aS20V), Rab11b, and Rab25 [362]; substrate for PAR1b and is involved in an alternate pathway modulating establishment of polarity [363, 364]

Role in polarity (reports)

[139]

[361]

[358]

[355]

[352]

Reviews

Y—Required for canalicular formation in WIF-B9 [75] ND

Y—Required for canalicular formation in WIF-B9 [75]

ND

Y—Localized to hepatocyte TJ [353]

Present in liver? Y/N—finding and reference

Key: ABC, ATP binding cassette; aPKC, atypical protein kinase C; CRIB, Cdc42/Rac-interactive binding; DN, dominant negative; ES, embryonic stem; EMT, epithelial mesenchymal transition; ESAM, endothelial cell-selective adhesion molecule; FERM, 4.1, Ezrin, radixin, moesin; GEF, Guanine nucleotide-exchange factor; GPCR, G-protein coupled receptor; HCC, hepatocellular carcinoma; IF, immunofluorescence; Ig, immunoglobulin; KO, knockout; LRR, leucine rich repeat; MAGUK, membrane associated guanylate kinase; MAP, microtubule associated proteins; ND, not determined; NLS, nuclear localization signal; p70 Sk, p70 S6 kinase; PB1, Phox and Bem1p; PBM, PDZ binding motif; PJS, Puetz-Jergers Syndrome; SH, src homology; SMC, structural Maintenance of Chromosome; S/T, serine/threonine; TGF, transforming growth factor; TM, transmembrane span; WD40, ∼40 aa domain with conserved trp and asp often found as repeats that form a propeller structure; WW, ∼40 aa domain with 2 conserved trp ∼20 aa apart; Y/N- yes/no; 2-H, two-hybrid; 4E-BP1, eukaryotic initiation factor 4E (eIF4E) binding protein 1.

Small GTPase

Rab11a

Myosin Vb

Functional domains

Protein

Table 6.2 (continued)

90 THE LIVER: ESTABLISHING EPITHELIAL CELL POLARITY

6: HEPATOCYTE SURFACE POLARITY: ITS DYNAMIC MAINTENANCE AND ESTABLISHMENT

can be induced to differentiate in culture to produce embryoid bodies (EBs). Examination of afadin KO EBs revealed AJs and TJs disrupted and mislocalized PAR3 and PAR6, and reduced activation of Cdc42 and aPKC. In addition, levels of integrin alpha-6 and integrin beta-1 were reduced, indicating cross-talk between integrins and junction components. These findings were consistent with previous studies where afadin and its association with nectin were shown to organize cell–cell junctions (reviewed in [373]). While afadin plays a key role is establishing junctions of cells with simple polarity, little is known about it or nectin in hepatocytes; studies are needed. A spontaneous mutation in the polarity/tumor suppressor gene, Scrib (circletail mouse), caused defective neural tube closure as well as liver overgrowth at ∼E 10.5 [176]. While no polarity defects were found in neuroepithelium, the planar cell polarity (PCP) organization of the inner ear hairs was defective (for review see [374]). Genomic analysis of the circletail mouse revealed that a single base insertion resulted in a protein truncated after PDZ 2, demonstrating an essential role for Scrib’s C-terminus, which has two additional PDZ domains and ∼440 aa lacking any recognizable motifs. Could the lack of polarity defects in the circletail mouse be explained by Scrib’s documented interaction with two polarity proteins, Vangl2 and Lgl2 [179]? Interestingly, Scrib knockdown in MDCK cells exhibited aberrant AJs at steady-state and delayed TJ formation following Ca switch [375]. This study showed that effects of Scrib knockdown may be due to loss of E-cadherin-mediated adhesiveness at AJs. Could PCP genes play a role in organizing hepatocyte cords in liver? Hepatocyte-specific KO studies of PCP genes could be used to test this hypothesis.

Tight junctions A large body of work has revealed that many proteins form a complex network at the TJ. The proteins are listed in Table 6.2c (also see, [195, 258, 352]). Surprisingly, KO of occludin and claudin 1 did not result in any overt pathology; however, the animals exhibit some tissue-specific defects [215, 221]. Studies of two other TJ proteins revealed their important role in hepatic polarity; loss of the hepatic apical surface was observed with knockdown of claudin 2 in WIF-B9 cells [226] and JAM-A in either HepG2 [233] or WIF-B cells [163]. Interestingly, only limited tissue-specific effects were reported following single disruption of JAM-A, B, or C in animals [235, 237, 376], suggesting the need for double or triple KO given gene redundancy. Multiple transcriptional changes were documented following JAM-A depletion in HepG2 cells, including fivefold E-cadherin upregulation. Restoration of apical surfaces was accomplished by re-expressing JAM-A, but only when E-cadherin expression was downregulated. This finding suggests that

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cross-talk between TJ and AJ components may occur [233]. In fact, molecular cross-talk at cell contacts may mediate assembly of both junctions (reviewed in [377, 378]), a poorly understood process at present. Thus, hepatocyte-specific junction factors and the molecular mechanisms regarding the development of hepatic polarity are still poorly understood. A combination of animal and cell-culture KO studies of the TJ proteins ZO-1, ZO-2, and ZO-3, highlights their important role at the TJ and beyond. Animal studies demonstrated that the TJ proteins ZO-1 and ZO-2 [245, 249] are essential but ZO-3 is not [249]. ZO-1 and ZO-2 functions are both likely required prior to TJ formation. Consistent with this finding, TJs were intact following ZO-1 deletion in a mouse mammary epithelial cell line, EPH4 [379]. However, simultaneous KO and knockdown of ZO-1 and ZO-2, respectively, in EPH4 cells resulted in no TJ formation and severely compromised TJ barrier function [379]. Reconstitution studies showed that either ZO-1 or ZO-2 can independently recruit claudins to form TJ strands. Unexpectedly, the double KO/knockdown cells were well-polarized, as demonstrated by distribution of several markers [379]. Thus, this study demonstrated that cells can polarize without TJs, a key structural aspect to normal epithelial tissue physiology and a likely signaling center for proper vesicle trafficking [152, 258, 351]. Can hepatic cells polarize without TJs? What are the cues that allowed these simple polarized cells to orient their apical surfaces?

Signaling molecules, effectors, and modulators Many studies have uncovered a wide variety of proteins that are not exclusively found at junctions yet impact apical/basal polarity. Since this is such a broad topic, we tabulated only those proteins where reports in the literature indicate that they might play key roles in hepatic polarity, and limited the following discussion (Table 6.2d). Hepatocyte nuclear factors (HNFs) have a wellrecognized role in liver development [292, 293]. In fact, HNF4α plays an important role in hepatic cell polarity, since embryonic (e)18.5 livers from HNF4α liver-specific KO animals lack normal hepatic architecture [380]. These results suggest that this transcription program drives the morphological changes that occur during hepatocyte differentiation. In fact, including the present one, several studies have demonstrated that forced expression of HNF4α in cultured cells leads to a polar phenotype [380, 381]. A role for HNFs in the development of HCC has also been investigated [291]. An animal model was developed whereby cultured HCC cells were injected subcutaneously into recipient mice for subsequent study. Hepatocytes lost polarity and exhibited reduced cell–cell and cell–ECM adhesion as well as reduced expression of

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liver transcription factors including HNF4. Interestingly, cells exhibited an epithelial phenotype and formed cell–substrate contacts upon forced re-expression of HNF4α alone either in vitro or in vivo [291]. Does HNF4α drive the ECM transcription program? The cell–ECM adhesion also has a well-recognized role in liver differentiation [382, 383]. Although beta-1-integrin functions are extensive, we only discuss two studies that indicate beta-1-integrin outside/in signaling is important for early liver development. Beta-1-integrin forms heterodimers with 13 of the 18 alpha-integrin family members (reviewed in [279, 280]), has widespread tissue expression during development, and is embryonic-lethal on deletion [282, 384]. First, study of beta-1-integrin KO ES cells revealed a role for this molecule in liver development through generation of chimeric mice that lived to adulthood [282]. The adult livers were devoid of KO cells but all other analyzed tissues (except spleen) contained substantial numbers. Second, study of mice heterozygous for deletion of Smad2/3 (activated by TGF-beta signaling) indicated that beta-1-integrin levels were reduced and E-cadherin was misprocessed and mislocalized [287]. The animals died in midgestation with liver hypoplasia and anemia, suggesting that the levels of activated Smad2/3 were inadequate for proper development [287]. Upon Smad2/3 activation by phosphorylation, these intracellular mediators associate with Smad4; then, in the nucleus, Smad2/4 and Smad3/4 complexes regulate transcription of TGF-beta-responsive genes, which in turn regulate many cellular responses. Misregulation of TGF-beta signaling can induce epithelial mesenchymal transition (EMT), which causes disruption of TJs and loss of cell polarity [285]. When cultured isolated hepatocytes were treated with TGF-beta, reduced claudin-1 levels were observed, which likely impacted the observed increase in paracellular permeability [224]. Thus, cross-talk between integrins, E-cadherin, and TGF-beta signaling plays a key role in liver development and likely hepatic polarity [385]. As discussed above, studies of ZO-1/ZO-2 depletion revealed that cell polarity can be uncoupled from TJ formation in cultured cells [379]. Consistent with this finding is the demonstration that a single autonomous cell can polarize without the aid of cell–cell contact [298]. This study demonstrated that LKB1 (also known as the serine/threonine kinase 11 (STK11) and PAR4 in C. elegans) activation by STRAD (STe20 Related ADaptor, [386]) resulted in a polarized distribution of microvilli and actin in a single Caco-2 cell [298]. In fact, PAR4/dLKB1 plays a key role in cell polarity of C. elegans and D. melanogaster (reviewed in [299]). LKB1 is also a tumor suppressor since inactivating mutations are often found in patients with Peutz–Jeghers cancer syndrome [387]. Among the LKB1 substrates is AMP kinase (AMPK), which activates many downstream cellular pathways including TGF-beta (reviewed in [300]) and mammalian target of rapamycin

(mTOR) [388]. Recent studies indicate that both AMPK and mTOR also have roles in cell polarity (Table 6.2d). These results suggest that LKB1 indeed functions as a master kinase that regulates diverse cellular functions. Interestingly, two other well-studied tumor suppressors, the lipid phosphatase PTEN [389] and adenomatous polyposis coli (APC, [390]), have important roles in cell polarity. Among the many roles of PTEN, it mediates apical distribution of phosphoinositides, which in turn control epithelial polarity through Cdc42 [311]. APC is also multifunctional; it binds microtubules [314], regulates β-catenin levels [315, 391], and is involved in cell migration [316]. Liver-specific KO studies of each tumor suppressor resulted in development of HCC (see Table 6.2d). Three Rho GTPases, Rho, Rac, and Cdc42, impact cell polarity in a variety of ways (reviewed in [392]). Studies indicate that the PAR–aPKC complex likely participates in balancing their GTPase activity (reviewed in [393]). In fact, the apical distribution of the actin cytoskeleton is a hallmark of apical/basal polarity, and signals driving this localization, an area of active research, are just being uncovered (see [394–397]). Recently, a liver-specific Cre-recombinant Cdc42 KO animal was generated, which exhibited severe liver morphological defects by two months and developed HCC by eight months [324]. At two months, when low levels of Cdc42 were still present, the hepatocyte plate architecture was irregular and bile canaliculi were enlarged. While an abnormal actin cytoskeleton was observed, which may be related to the reduced levels of E-cadherin, ultrastructural analysis showed that TJs were intact, as were levels of TJ proteins. Components of the PAR/aPKC complex were also properly localized, a surprising finding given that Cdc42—which binds PAR6—is implicated in Drosophila and MDCK cell junction assembly (Table 6.2a, [311]). Regenerating liver studies in Cdc42 KO mice showed that cord structure was disorganized and bile canaliculi were dilated [398]. The hepatectomized livers were slow to regenerate and multiple pathways involved in cell proliferation were delayed [398]. In contrast, study of Cdc42 KO EBs revealed defective AJs and TJs [399]. Could another RhoGTPase be acting in liver? Perhaps RhoA, since its increased expression correlates with progression of HCC [400].

MDCK cells can adopt a hepatic-like phenotype Unexpectedly, the study of hepatic polarity has been aided by overexpressing and studying select proteins implicated in polarity in MDCK cells grown in collagen overlays. This technique has been used extensively to elucidate the requirements for the development of tubulogenesis, a process whereby MDCK cells reorganize to form cysts or tubes with cells expressing the simple polarized phenotype ([401] and reviewed in [65]). Study of selected

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overexpressed proteins in cultured MDCK cells grown on collagen gels results in the appearance of a hepatic-like phenotype (i.e. lateral lumen formation) [345]. This finding led us to propose this culture system as a tool to examine molecular mechanisms that may initiate the development of hepatic polarity. By studying overexpression of various polarity modulators/effectors (Table 6.2e), mechanistic clues have emerged regarding the establishment of hepatic polarity. The first clue came when overexpression of the STK PAR1b did not result in a culture with tubules or with cells exhibiting simple polarity; rather, an altered polarity phenotype was observed such that the apical membrane formed a lumen between two cells, giving rise to what is now termed a “hepatic-like phenotype” [346]. Characterization of these cells revealed that MTs were reorientated [346], apical proteins trafficked via transcytosis [346], and myosin II and E-cadherin-dependent signaling were required [347]. In support of a role for PAR1b in the development of hepatic polarity is the finding that overexpression of a mutant PAR1b, KNPAR1b, in WIF-B9 cells blocked formation of their apical lumens [346]. Par1 (also known as MARK or EMK (for ELKL motif kinase)) is a member of a small family of serine/threonine protein kinases involved in the control of cell polarity, microtubule stability, and cancer (reviewed in [344]). PAR1 was first identified as one of four maternal genes important for early development whose absence inhibited asymmetric cell division of the C. elegans zygote [402]. In mammals, Par1b phosphorylates microtubule-associated proteins and triggers microtubule disruption [403]. Surprisingly, two Par1 KO mouse models do not display polarity defects, but rather slight growth retardation, late onset immunological defects, and changes in glucose uptake ([343, 404], reviewed in [344]). Again, gene redundancy likely obscured the role of Par1b in polarity, since Par1a/MARK3 and Par1b/MARK2 appear to be ubiquitously expressed (including liver), with MARK1 expression highest in brain tissue. Each is associated with the PM, while another family member, MARK4, appears to be microtubule-associated. There is a link between PAR1 and LKB1; however, studies in mammalian cells and D. melanogaster differ in the activation sequence. Mammalian studies demonstrate that LKB1 phosphorylates PAR1 homologs on their regulatory loops [405], while studies in Drosophila indicate that PAR1 acts on dLKB1 [406]. How is Par1b working to promote reorientation of the apical surface? What signaling pathways are activated? Are cell–substrate cues, outside/in signaling via integrins, Wnts, or growth receptors required? This MDCK system should be exploited as a first step toward answering these questions. The hepatic-like phenotype (lateral lumens) in MDCK cells was also observed following overexpression of dominant negative RhoA (RhoN19) and constitutively active Rac1 (RacV12) [349]. Rho and Rac have antagonistic roles that have been well characterized in actin dynamics

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regulating myosin II and are now thought to have a role in AJ and TJ assembly and function [348]. MDCK cells also exhibited the hepatic phenotype when grown in collagen gels with rapamycin, which inhibits mTOR (Table 6.2d and [310]). Consistent with this finding is a study which shows that sustained activation of mTOR impairs the development of hepatic polarity in HepaFR cells [309]. Microtubule orientation and transcytotic studies are needed to validate the extent of hepatic-like phenotype in these systems. In summary, through studies that were directed at perturbing tubulogenesis in MDCK cells, a well-defined system has evolved that may be used to study molecular mechanisms involved in the development of hepatic polarity. In fact, these studies point to the idea that cells undergoing architectural remodeling are faced with a branch point whereby levels of certain polarity proteins, protein phosphorylation status, activation state, and apical retention mechanisms determine polarity phenotype.

Apical/basal polarity requires vesicle trafficking Myosin Vb (myoVb) is an unconventional myosin with a role in vesicle trafficking via binding Rab11a ([356], reviewed in [358]). Recently, mutations in myoVb were found to cause microvillus inclusion disease, which is characterized by a lack of apical microvilli and intracellular structures containing microvilli [357]. The transferrin receptor was found at the apical surface instead of the basolateral surface. These results indicate that loss of myoVb causes defective vesicle trafficking of apical and basolateral proteins, linking myoVb to the maintenance of cell polarity. Vesicle trafficking has also been linked to the development of hepatic polarity. In knockdown studies carried out in cultured WIF-B9s, a loss of either myoVb or Rab11a prevented formation of apical surfaces [75]. MyoVb binds Rab11a, a small GTPase involved in regulated apical vesicle trafficking/transcytosis ([359, 360] and reviewed in [361]). These results indicate that hepatic polarity may require recruitment of Rab11a and myoVb to intracellular vesicles containing apical ABC transporters and transcytotic markers. The Rab11 binding protein, Rab11-FIP2 (family of interacting protein with a C2 domain and a C-terminal amphipathic α-helix), is a Par1b substrate and has been shown to be involved in an alternate pathway modulating establishment of polarity in MDCK cells ([363, 364], reviewed in [139]). Rab11-FIP2 also binds the dominant active form (Rab11aS20V), Rab11b, and Rab25 [362]. How do these proteins work together or in sequence to impact polarity? How does polarization occur upon delivery of Rab11a/myoVb vesicles to the PM? What induces differentiation of the PM into an apical domain? The hepatic phenotype was also absent upon loss of the TJ proteins JAM-A [163] and claudin-2 [226]; is Rab11a

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involved in trafficking of TJ proteins? Rab13 has been localized to the hepatocyte TJ [353], is documented to regulate membrane traffic, and has a direct and functional link with protein kinase A signaling during junction assembly in epithelial cells ([351], reviewed in [352]). Does Rab13 work with myoVb or perhaps myoVc, another unconventional myosin, which is also expressed in epithelial tissues and is likely involved in membrane trafficking ([354], reviewed in [355])?

ACKNOWLEDGEMENTS We thank A. Musch for helpful discussions. Research on hepatocyte polarity in the Hubbard Laboratory was supported by a grant awarded to ALH by the NIH (DK063096).

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Endocytosis as an Essential Process in Liver Function and Pathology Barbara Schroeder and Mark McNiven Department of Biochemistry & Molecular Biology, Miles and Shirley Fiterman Center for Digestive Diseases, Mayo Clinic & Foundation, Rochester, MN, USA

INTRODUCTION

VESICLE FORMATION IN THE ENDOCYTIC PATHWAY

A prominent function of the hepatocyte is the regulated endocytic uptake of various proteins and lipids from the blood sinusoid for subsequent processing and transport into bile (Figure 7.1). This function depends on elaborate vesicle trafficking machinery that shares substantial homology with that utilized in the secretory pathway. Coat proteins and adaptors that are linked to specific lipid-membrane subdomains and the cytoskeletal matrix provide a mechanism to sequester and internalize trophic receptor/ligand complexes such as epidermal growth factor, hepatocyte growth factor, and iron-bound transferrin and to help maintain normal lipid serum levels through the endocytosis of low-density lipoproteins (LDLs). Of equal importance is the fact that this highly evolved machinery can be “hijacked” by many pathogens including bacteria, viruses, and parasites to infect the liver, leading to inflammation and hepatitis. This review will provide a general overview of the essential components of the hepatocyte endocytic machinery and how it supports crucial hepatic functions in health and disease.

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

The endosomal apparatus is a heterogeneous population of membrane-bound structures that are involved in the uptake of exogenous substances into cells. Internalization of materials can occur in a non-selective manner, as in macropinocytosis where cells non-specifically take up small droplets of fluid, or selectively, as in receptormediated endocytosis (RME) where specific soluble molecules are internalized. The translocation of internalized cargo proteins from the cell surface to the early endosome (EE) and the subsequent delivery of lysosomally destined components to late endosomes (LEs) and lysosomes involves multiple vesicular trafficking events, in which endosomal compartments are thought to undergo maturation to temporally later compartments. In this model, coated vesicles derived from the plasma membrane (PM) lose their coat and fuse together to assemble an EE, which undergoes gradual remodeling and transformation to an LE and eventually a dense lysosome.

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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Figure 7.1 Endocytic membrane trafficking is an essential process in the hepatocyte. Thin-section electron micrographs of rat hepatocytes from original collections of Dr. Keith Porter. (a) High-magnification image of the apical, canalicular domain of a hepatocyte that shows a bile canaliculus (BC) surrounded by many electron-dense lysosomes (Lys). (b) A high-magnification electron micrograph of a basolateral hepatocyte sinusoidal space (Sin) flanked by endothelial cells (E). The subcortical region of the hepatocyte shows many different types of endocytic vesicle that are forming and internalizing. Magnification for (a) and (b): 9100 ×

CLATHRIN-COATED VESICLES Coated pits/vesicles and receptor-mediated endocytosis Clathrin-dependent endocytosis is one of the major and most intensely studied mechanisms by which surface receptors are internalized. The initial steps in the process of receptor internalization, during which itinerant membrane proteins are segregated from resident PM proteins and are internalized by the cell, are mediated by clathrin-coat assembly (Plate 7.1, Figure 7.2). Coated pits are relatively uniform in size (100–150 nm profile diameter) in all cell types (Plate 7.1d). They were first observed by Roth and Porter, who noted that the endocytosis of yolk protein into the oocyte of the mosquito was associated with a marked increase in invaginations of the oocyte cell membrane, which was coated on the cytoplasmic face with what they termed a bristle coat [1]. Soon thereafter, clathrin was identified as the major protein

component of the coat [2], and the concentration of a specific ligand in coated pits followed by its internalization was demonstrated in cultured fibroblasts using ferritin-LDL [3]. Coated pits occupy up to 2% of the cell surface area, as determined by calculations for human fibroblasts [3], rat hepatocytes [4], and other cells [5, 6]. Coated pits assemble and bud with a half-time of ∼5 minutes in broken A-431 cells [7] and in K+ -depleted fibroblasts that are incubated in the presence of KCl [8]. Morphometric quantitation of the number of coated pits at the PM in baby hamster kidney cells estimated the lifetime of coated pits as 1–2 minutes [5]. Earlier estimates from quantitative ligand (α2 -macroglobulin (MG)) uptake suggested that each coated pit might transfer ligands into endocytic vesicles every 20 seconds at 37 ◦ C [9]. However, biochemical analyses of receptor internalization kinetics in rat liver parenchyma demonstrated that following ligand stimulation, receptors known to utilize the coated pit pathway (i.e. epidermal growth factor receptor (EGFR) and insulin receptor) are rapidly lost from the PM (t1/2 ∼1 minute) [10, 11]. In addition to soluble ligands, many viruses enter cells by coated pits. These include adenovirus, vesicular stomatitis virus (VSV), Rous sarcoma virus, and Semliki forest virus. Viruses possessing a lipid membrane surrounding the nucleocapsid contain glycoproteins that are involved in receptor recognition and virus entry into cells [12]. Clathrin-coated vesicles are also found at the Golgi apparatus, where they mediate (i) the transport of lysosomal enzymes from the trans-Golgi network (TGN) to the endosomal apparatus, (ii) the formation of secretory granules in regulated secretory cells, and (iii) the maturation of immature zymogen granules into mature secretory granules.

Coated pit ultrastructure and coat proteins The purification of coated vesicles has allowed the major structural units of the coat, namely clathrin triskelions and adaptors, to be identified [13]. The characteristic honeycomb lattice of coats [14] as visualized by quick-freeze, deep-etch, rotary shadowing is made up of clathrin (Plate 7.1b), and the inner shell of coats consists of adaptor proteins (APs) (see below). Structurally, clathrin triskelions have a flexible design that allows them to pack together to form polyhedral lattices [15]. Each triskelion is a three-legged structure consisting of three heavy chains and three associated light chains. Whereas the heavy chains are capable of binding to adaptins and other accessory proteins that help mediate endocytosis, the light chains contribute to lattice assembly (see below). Clathrin is linked to the cargo by so-called adaptins or APs. So far, four structurally-related classes of APs are known, and all are expressed ubiquitously. In general, adaptins select and concentrate cargo proteins into

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Figure 7.2 The vesicle fission machinery and associated actin cytoskeleton act in concert to liberate nascent vesicles from the plasma membrane. (a) A complex actin cytoskeletal network together with the clathrin-based sorting and budding machinery function to complete the process of cargo sequestration, vesicle formation and membrane scission. (b,c) Fluorescence micrographs of cells stained for Dyn2 (b) and cortactin (c) as examples of components of the fission machinery and actin cytoskeleton that are present at the plasma membrane. This figure is modified from [148] with permission from Science

clathrin-coated pits, recruit endocytic regulatory proteins, and attach clathrin to the PM. Within this family, PM-residing AP-2 and Golgi-localized AP-1 are probably the best-characterized members [16, 17]. The other two members, AP-3 and AP-4, are localized to the Golgi region and endosomes [16, 18]. In addition, both AP-1 and AP-3 have tissue-specific isoforms designated AP-1B (epithelial cells) and AP-3B (neurons), respectively [18, 19]. Each AP consists of four subunits: two heavy chains (γ/α/δ/ε and β1/β2/β3/β4 of ∼90–140 kDa), one medium chain (µ1/µ2/µ3/µ4 of ∼50 kDa), and one light chain (σ1/σ2/σ3/σ4 of ∼20 kDa). The different adaptin subunits serve distinct functions. The β and µ chains are implicated in cargo selection, and different sorting signals on cargo proteins have been identified. These sorting signals include NPXY (four-amino-acid sequence consisting of asparagine, phenylalanine, any amino acid, tyrosine, one-letter code), as well as tyrosineand dileucine-based sorting motifs [20–22]. The α and β ear domains are responsible for recruiting accessory proteins to the PM, where they participate in vesicle budding and uncoating. The α and µ chains of AP-2 both contain a PdtIns(4,5)P2 -binding module that enables them to associate with the PM, which brings AP-2 in close proximity with the cargo proteins. In addition, the interaction of α and µ with PdtIns(4,5)P2 induces a conformational change that leads to the uncovering of a hidden Tyr-based cargo binding site [17]. Furthermore, the β chains of AP-1, AP-2, and AP-3 contain a so-called “clathrin box” that mediates interaction with the clathrin heavy chain (CHC) [23, 24]. This clathrin-binding motif is absent in AP-4; therefore, whether or not AP-4 works in concert with clathrin is still under debate. In contrast to the various functional roles of the heavy and medium chains, the role of the light chains (σ) appears to be strictly structural [25]. Whereas AP-2 is involved in endocytosis from the PM, AP-1 mediatesTGN-to-endosome transport, and AP-3

facilitates trafficking from the TGN to lysosomes (see Chapter 8). The precise role of AP-4 is still underexplored, but it might also have a role in post-Golgi transport processes [19, 26].

Coated pit/vesicle formation and associated factors The process of coated pit formation involves the recruitment and assembly of clathrin and associated coat proteins and other factors at the PM to form a pit, which subsequently invaginates and buds to produce a coated vesicle. Coat proteins assemble onto the cell surface as planar structures that later gain curvature to form invaginated pits [8, 14]. Although the clathrin light chains were initially believed to be non-essential for cage assembly [27], some evidence suggests that light chains participate in the regulation of clathrin assembly and disassembly in vivo by preventing cage assembly in the cytosol [28]. This hypothesis is based on the external position of the clathrin light chains on the coat [29], their possession of calcium-binding sites [30] and phosphorylation sites [31], their interactions with uncoating ATPase [32], and their negative regulation of polymerization in vitro. Clathrin triskelions can join together to form flat hexagonal lattices that gain curvature by the introduction of pentagons. However, this change in lattice arrangement and the energetics of clathrin polymerization do not appear to be sufficient to induce membrane curvature [33]. Therefore, additional factors are required for effective membrane bending. The uncoating ATPase Hsc70, its co-chaperone GAK/auxilin [34], and synaptojanin-1 [35]—all of which are recruited with clathrin to the coated pits [36]—likely play roles in this process [37, 38].

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Accessory proteins in clathrinmediated endocytosis and coated vesicle scission A battery of accessory proteins (e.g. epsin, Eps15, amphiphysin, and dynamin) involved in vesicle formation/scission and substrate recruitment was identified recently in GST–α-appendage pull-down binding experiments. These proteins are known to interact with each other and with other proteins via a number of specific protein-binding sequence motifs, including EH, coiled-coil (CC), proline-rich domain (PRD), and SH3 domains [39] (Plate 7.1f–h). A role for Eps15 (EGFR protein substrate 15, Figure 7.2h) and epsin in coated pit assembly and endocytosis has been suggested from experiments in which their functions were disrupted, resulting in inhibition of clathrin-mediated endocytosis (CME) [39–41]. Eps15 is complexed to AP-2 in the cell; binding to the α-appendage of AP-2 occurs via repeated DPF motifs (amino acid sequence consisting of aspartic acid, proline, phenylalanine in single letter code) in the C-terminus of Eps15. In the vesicle-forming process, Eps15 recruitment to the PM appears to necessary for subsequent AP-2 association [40]. Interestingly, detailed electron microscopy (EM) immunolocalization studies of Eps15 in fibroblasts has revealed that the AP-2 adaptor is distributed homogenously over the entire coated pit surface, whereas Eps15 is restricted to the rim, or the growing part, of the budding coated pit [42]. This observation suggests that Eps15 may dissociate from AP-2 during coat and vesicle formation. Clathrin appears to be a competing factor in the Eps15–AP-2 interaction and has been shown to release Eps15 from AP-2 during coated pit formation [43]. Moreover, Eps15 is associated constitutively with Ese1 (EH domain and SH3 domain regulator of endocytosis), a multifunctional endocytic AP that can link Eps15 to dynamin 2 (see below) via EH and SH3 domain interactions, respectively [44]. Eps homology domains, or EH domains, are protein– protein interaction domains that were originally identified as a repeated sequence at the N-termini of Eps15 and Eps15R (Eps15-related molecule). In addition to its interaction with Eps15, AP-2 has been shown to interact with the central region of epsin [45], another protein implicated in CME. In its C-terminal region, epsin harbors repeats of an EH domain-binding consensus motif, NPF, which has been shown to interact with Eps15 and intersectin, another recently identified EH domain-containing family member [39]. Intersectin also contains five SH3 domains [39], which are 50–70-amino-acid modules through which intersectin can bind to proline-rich ligands. As already described for Ese1, these protein–protein interaction domains allow intersectin to form various macromolecular complexes and bridges between proteins with binding regions for EH domains (e.g. NPF repeats in epsin) and

for SH3 domains (e.g. PRDs in dynamin (see below) and synaptojanin, an inositol-5-phosphatase implicated in intracellular signaling and vesicular trafficking events). The phosphorylation status of the accessory proteins (Figure 7.3) appears to be a means by which vesicle formation is regulated at the cell surface. Many accessory proteins (Eps15, epsin, dynamin, amphiphysin, and AP-2) are constitutively phosphorylated on serine and threonine residues. In neurons, these molecules are known to be dephosphorylated and thus activated upon a depolarization stimulus [39]. Activation allows the accessory proteins to be recruited to membranes from their predominantly cytosolic locations [46] and to associate with the AP-2 adaptor [47]. Late stages in coated vesicle formation, namely invagination and liberation of coated vesicles, appear to be mediated predominantly by the large GTPase dynamin. In mammals, the dynamin family comprises at least three members that are encoded by distinct genes. All members contain four conserved domains: an N-terminal, highly-conserved GTP-binding domain; followed by a pleckstrin homology (PH) domain; a CC domain; and a PRD, which is less conserved than the other domains [48–50]. The PH domain allows dynamin to bind membranes, whereas the PRD mediates association with multiple effector proteins [51–53]. Dynamin has been referred to as a “pinchase” because of its ability to generate discrete vesicles from invaginated coated pits. Dynamin, like clathrin, can self-assemble in the absence of nucleotide. Polymeric dynamin complexes have been visualized along lipid vesicles [54] and membrane tubules in rat brain synaptosomal membranes under specified conditions [55]. In vitro studies by Sweitzer and Hinschaw revealed that purified dynamin in the absence of any nucleotide can tubulate the originally spherical liposome; in the presence of GTP, the elongated tubules are constricted and fragmented into small vesicles [56]. In contrast, the use of a kinase-deficient dynamin K44A-mutant or of GTPγS as a non-hydrolyzable analogue of GTP resulted in tubulation but not constriction and vesiculation of liposomes, which indicates that GTP hydrolysis is the driving force for membrane fission [57, 58]. These and other recent studies have led to various models for dynamin function in vesicle formation [17]. Several studies have indicated that endocytosis via clathrin-coated pits requires a functional dynamin GTPase, although CME is not the only endocytic process dependent on dynamin function (see below).

Regulators of clathrin-mediated endocytosis and vesicle uncoating The budding of coated vesicles from the PM may also be mediated by the actin cytoskeleton beneath the cell membrane. Although most studies have been carried out in yeast, evidence of a role for actin in endocytosis

7: ENDOCYTOSIS AS AN ESSENTIAL PROCESS IN LIVER FUNCTION AND PATHOLOGY

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Figure 7.3 Membrane-tubulating proteins at the PM. (a) Along with conventional clathrin coats and adaptors, a variety of BAR and F-BAR/EFC domain-containing proteins bind and deform membranes into tubules in concert with the large GTPase dynamin and the actin cytoskeleton. (b,c) Electron micrograph of negative-stained liposomes tubulated in vitro by the addition of purified amphiphysin I (b) and endophillin B (c). The accompanying fluorescence micrographs (d,e) demonstrate the tubulating action of these proteins upon overexpression in living cells. Insets show higher magnifications of the boxed regions, emphasizing the massive membrane tubulation induced by these proteins. The figure is modified from [148] with permission from Science

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in mammalian cells has accumulated over recent years. Mutations in a yeast actin-binding protein, Sla2p, were shown to lead to altered endocytosis and actin cytoskeletal organization [59]. A mammalian homologue of Sla2p, huntingtin-interacting protein 1 (Hip1) colocalizes with clathrin, AP-2, and endocytosed transferrin. As a component of clathrin-coated pits, Hip1 could link the actin cytoskeleton to endocytosis [60, 61]. In addition, recent studies investigating the role of actin motors in endocytic events indicated an involvement of the “−” end motor myosin VI and the “+” end motor myosin 1e in CME. Myosin IV can interact with the adaptor Dab2 and PdtIns(4,5)P2 , whereas myosin 1e is linked to clathrin via SH3-mediated interactions with dynamin and synaptojanin 1 [62, 63]. Other actin links to the endocytic machinery include the actin-binding proteins profilin, synapsin, syndapin, and cortactin. These proteins are known binding partners of dynamin and have been shown to colocalize with the molecular pinchase ([64, 65] and McNiven, M, Kim, L, Krueger, EW, Cao, H, Wong, TW, personal communication). Morphological studies have shown that once formed, coated vesicles uncoat rapidly [66] and fuse with each other or with peripheral endosomes [67]. The half-life of a coated vesicle was reported to be less than 1 minute [67], and photobleaching studies following rhodamine-clathrin microinjection estimated the half-life of polymerized clathrin to be ∼10–15 seconds [68]. Studies using green fluorescent protein-tagged clathrin light chain suggested that coated vesicles release their coat very close to the coated pit [69]. Both heavy and light chains of clathrin are released by uncoating ATPase Hsc70 and its co-chaperone GAK/auxilin [34, 70]. GAK is recruited to the nascent coated pit before actin-mediated scission begins [71]. GAK recruitment can be enhanced by the activity of synaptojanin, which converts PdtIns(4,5)P2 into the PdtIns(4)P that is necessary for GAK binding [36, 71]. After auxilin/GAK recruitment, the clathrin coat is released from the vesicle, as visualized by decreasing clathrin fluorescence. It is worth noting that auxilin/GAK activity is not restricted to clathrin uncoating but is also required for the dynamin-dependent constriction of coated pits [72]. A model for clathrin coat assembly/disassembly is shown in Plate 7.1e.

UBIQUITINYLATION AS A RECEPTOR DEGRADATION SIGNAL Ligand-induced internalization, downregulation, and degradation of receptor tyrosine kinases are very important for maintenance versus attenuation of growth signaling cascades. Over the years it has became apparent that the attachment of a ubiquitin (Ub) moiety to proteins

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plays a major role in lysosomal targeting and degradation. However, this post-translational modification is restricted to higher eukaryotes [73, 74] and is independent of Ub-induced proteasomal degradation. Ub is an 8-kDa, highly-conserved protein that can be covalently linked to lysine residues in target proteins. The attachment of Ub can occur in different ways: as a chain (poly-ubiquitylation) or as a single molecule attached to one (mono-ubiquitylation) or multiple (multi-ubiquitylation) lysine residues. Poly-ubiquitinated proteins are targeted to the proteasome, where the protein is degraded and the Ub moiety is recycled [75, 76]. It has become increasingly obvious that poly-ubiquitylation serves additional roles because different linkages can be used between the individual Ub molecules [77, 78]. Mono- and multi-ubiquitylation are required for the passage of certain cargo molecules into/out of vesicles of the endocytic pathway, as well as for signaling processes [79–81]. Ub-mediated downregulation of receptor tyrosine kinases has been observed for a number of different receptors including cMet (the receptor for hepatocyte growth factor/scatter factor (HGF/SF) [82, 83]), plated-derived growth factor receptor (PDGFR) [84, 85], and Flk-1 [86], with the best-studied example being the EGFR. Various groups have shown that EGFR is ubiquitylated upon ligand addition, and this ubiquitylation targets EGFR to the lysosome for degradation [87, 88]. Studies using a modified EGFR with mutated lysine residues to prevent Ub modification revealed that this receptor could be internalized properly but was recycled back to the PM to avoid degradation [89, 90]. Further studies demonstrated that attachment of the Ub moiety, although dispensable for the initial internalization, is required for the exit of EGFR from the EE and thus for its degradation [89, 90]. Not surprisingly, epithelial cells utilize a number of different Ub-binding proteins as adaptors to direct modified cargo to the lysosome. Ub-binding domains are found in various endocytic proteins such as Eps15, GGAs (Golgi-localized, gamma-ear-containing, ADPribosylation-factor-binding proteins), and proteins associated with the ESCRT (endosomal sorting complex required for transport, see below) machinery such as Hrs (hepatocyte growth factor-regulated tyrosine kinase substrate) and STAM (signal-transducing adaptor molecule), as well as some of the ESCRT subunits themselves (Tsg101 and Vps 36 [91, 92]). The adaptors Hrs and STAM are of particular interest because they localize specifically to EEs [93, 94] and are essential for the selection of ubiquitinated cargo and thus the formation of multivesicular bodies (MVBs). Knockout (KO) of either Hrs or STAM in mice is embryonic-lethal, and fibroblasts obtained from these KO mice display enlarged vacuolar structures [95, 96]. However, ligand-induced downregulation of growth factor receptors such as EGFR and cMet is only partially impaired in Hrs-depleted cells [97, 98]. Together, Hrs and STAM are often referred to as ESCRT-0 because they are

responsible for the initial recognition and sequestration of Ub-cargo at the EE. In addition, Hrs recruits downstream ESCRT proteins through direct interaction with Tsg101, an ESCRT-I component [99]. In general, three different ESCRT complexes named ESCRT-I, -II, and -III [100–102] can be distinguished. These heteromeric protein complexes comprise up to six subunits and are recruited from the cytoplasm to the endosomal membrane, where they sequentially sort transmembrane proteins into MVBs to target them for degradation. As mentioned above, some of the ESCRT subunits contain Ub-binding motifs, suggesting that the ubiquitinated cargo is handed over from one complex to the other, although there is no direct evidence for this mechanism so far [103, 104]. Although most of the players in the MVB pathway appear to have been identified, some key questions remain unanswered. For example, it is unclear whether the lipid composition of the endosomal membrane or the subunits of the ESCRT complexes contribute to inward budding (away from the cytosol). An interesting parallel to this process is the outward budding of HIV, in which some of the ESCRT subunits are recruited to budding sites and are incorporated into the virions [105, 106]. As the ESCRT complex mediates budding of cytoplasmic vesicles into a lysosomal void, a modified complex is believed to support a similar process, namely the fusion and release of membrane-encased virions into the extracellular space.

CAVEOLAE IN THE HEPATOCYTE Endocytosis of membrane and fluid also occurs through a pathway that is independent of clathrin. This clathrinindependent internalization is believed to be responsible for the uptake of molecules that do not utilize coated pits, such as glycosyl-phosphatidyl-inositol (GPI)-anchored proteins [107] and cholera and tetanus toxins [108], and may play a role in the turnover of membrane proteins. Here we focus on caveolae, which constitute the best-studied clathrin-independent endocytic process. Caveolae are flask-shaped, specialized, sphingolipidand cholesterol-rich microdomains with a diameter of 60–80 nm. Caveolae have been reported on the surfaces of various cells including fibroblasts [108, 109] (Plate 7.2a–c, f–h), different liver-derived cell lines [108, 109], and whole rat liver [110]. Notably, smooth membrane microinvaginations ∼60–70 nm in diameter in a cultured liver cell line were described in classic studies by Lelio Orci’s group as being involved in the initial binding of gold-labeled cholera and tetanus toxins, two known ligands of caveolar membrane glycolipids [108]. Caveolae have a unique lipid makeup and are often associated with a spiral coat on their cytoplasmic side that consists in part of the integral membrane protein caveolin (Cav) [111]. Cavs are the main membrane proteins in caveolae and are critical for caveolar function (see below). Three members

7: ENDOCYTOSIS AS AN ESSENTIAL PROCESS IN LIVER FUNCTION AND PATHOLOGY

of the Cav family have been identified: Cav-1 and Cav-2 are expressed in most cell types (Plate 7.2d, e), whereas Cav-3 is found only in skeletal and cardiac muscle and some smooth muscle cells [112, 113]. Cav-1 and Cav-3 can form invaginated caveolae, but Cav-2 appears unable to do so without Cav-1 [114, 115]. All Cavs contain a long hairpin domain between their N- and C-termini that allows insertion into the PM. Further membrane association is achieved by palmytoylation at the C-terminus [119] and an ability to associate with cholesterol [120]. Functionally, caveolae have been implicated in a number of cellular processes, including endocytosis and signal transduction events, as well as transcytosis of various proteins such as toxins and viruses [121, 122]. Cholera toxin, for example, takes advantage of the high content of GM1 gangliosides in caveolae by binding to these lipid constituents and is subsequently internalized in a pathway involving the endosomal and Golgi compartments, followed by retrograde transport to the endoplasmic reticulum (ER). Furthermore, caveolae are involved in cell adhesion, sterol and fatty acid regulation, and mechanosensing [122].

Lipid composition and caveolar ultrastructural integrity Caveolae have been characterized as unique lipid microdomains at the cell surface. Cholesterol and sphingolipids are enriched in caveolae, whereas the rest of the PM consists mainly of phospholipids. Local high concentrations of cholesterol and sphingolipids, which may or may not be associated with caveolae, have been referred to as detergent-insoluble glycolipid (DIG)-enriched domains due to their insolubility in Triton X-100 [123, 124]. Whether these “lipid rafts” truly exist in vivo, however, is still a matter of debate [125]. The lipid composition of caveolae appears to be important for maintenance of both their structure and their function. Given that cholesterol is a major component of caveolae, the perturbation of cholesterol trafficking or depletion of free cholesterol has been shown to strongly decrease the number of caveolae in cells [126, 127]. At the EM level, a complete loss of morphologically identifiable caveolae was reported in two epithelial cell lines (MDCK [128] and HEp-2 cells [127]) depleted of cholesterol with methyl-β-cyclodextrin. Levels of Cav protein in MDCK cells were decreased 50% in the presence of methyl-β-cyclodextrin [128], which suggests that a necessary steady-state balance exists between Cav protein levels and cholesterol content in caveolae. Indeed, Cav-1 (see below) can bind to cholesterol [120] and its expression is upregulated when levels of intracellular cholesterol are increased [128]. Moreover, disruption of the cholesterol composition of the PM with cholesterol oxidase leads to the rapid

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redistribution of Cav from the PM to the Golgi region [129]. In addition, treatment of fibroblasts with progesterone induces the loss of Cav from caveolae and its accumulation in internal membranes. Furthermore, the movement of newly synthesized cholesterol to caveolae was inhibited when cells were incubated with progesterone [130–132]. In support of these observations, caveolae are known to influence cholesterol trafficking [130, 131] because Cav can bind directly to cholesterol [133–135]. Thus, cholesterol is likely critical for both the structure and the function of caveolae [111].

Biogenesis and endocytosis of caveolae Cav is known to be synthesized as an integral membrane protein in the ER, where it can also oligomerize [136, 137]. In addition, there is a Golgi-associated Cav pool that is distinct from the lipid-raft-associated, detergentresistant pool [138, 139], which indicates that Cav characteristics change after exit from the Golgi. The transition from the Golgi to the PM is influenced positively by cholesterol addition and negatively by glycosphingolipid depletion [138, 139]. Studies using GFP-Cav-1 indicate that exit from the Golgi might occur as a fully assembled caveolae-like structure that is transported to the PM [140]. However, this is just a model, and what drives the budding, how this event is regulated, and whether other, non-caveolar membranes also contribute to the assembly of exocytic caveolae-like structures remain to be determined [122]. In addition to its localization to the ER and TGN [132], Cav-1 has been visualized within the lumen of secretory vesicles in pancreatic acinar cells [141], suggesting a mechanism by which Cav might recycle between caveolae and the ER/Golgi apparatus. At the PM, caveolae form an immobile, stable functional unit that maintains its structure even upon endocytosis [140, 142]. Although caveolae can remain at the PM for a long period of time, their internalization can be stimulated by cargo such as SV40 virus [142, 143] or cholera toxin [144], as well as by glycosphingolipids or sterols [145]. This liberation of caveolae from the cell surface is also believed to be mediated by the large GTPase dynamin Dyn2 [146]. The fission of caveolae in bovine lung endothelial cells was shown to require wild-type cytosolic dynamin and GTP hydrolysis; overexpression of GTPase-deficient mutant dynamin abrogated caveolar budding and inhibited caveolae-mediated cholera toxin internalization [147]. Furthermore, microinjection of inhibitory Dyn2 antibodies in cultured mouse hepatocytes led to [1] the generation of long tubular invaginations of the plasmalemma with visible clathrin coats at the ends and [2] the accumulation of distinct, non-clathrin-coated, flask-shaped invaginations that were morphologically similar to caveolae at the PM (Figure 7.2c). Cholera toxin internalization was also inhibited in these antibody-injected

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cells. Moreover, immunofluorescence and biochemical experiments have demonstrated that dynamin and caveolae colocalize within intracellular vesicular structures in mouse hepatocytes [109]. Finally, EM immunogold labeling of lung endothelial cells revealed that dynamin was localized at the necks of caveolae, where it is believed to mediate their scission from the PM. Thus, the release of clathrin-coated pits and caveolae from the PM utilizes many similar proteins. For example, phosphorylation of Cav-1-Tyr14 by Src, Fyn, or c-abl contributes to an increased internalization rate and an increased cytoplasmic pool of Cav-1 [149, 150]. Phosphorylation of this critical Tyr residue is also required for the entry of SV-40 or coxsackievirus [143, 151]. In addition, Shajahan and colleagues showed that Src-mediated phosphorylation of dynamin is required for Cav-mediated endocytosis [152]. Furthermore, binding of Cav-1 to filamin [153] links caveolae to the actin cytoskeleton, which might contribute to caveolar movement, although that is still a matter of debate [154]. In addition, several reports showed that PKCα activity is necessary for caveolar endocytosis [155, 156].

LIPIDS AND PHOSPHOINOSITIDES AS REGULATORS OF ENDOCYTOSIS Considering the vast number of studies on the role of proteins in vesicle formation and trafficking, it has often been assumed that the membrane lipids of a vesicle do not play an active role. More and more, however, it appears that lipids of the membrane bilayer are important regulators of membrane dynamics, not only in endocytic events but also in endosomal trafficking and signaling from the PM [157, 158]. For example, studies in model membrane systems have revealed that vesicle budding can be facilitated if the lipid composition is modified to provide favorable membrane bending energy. Cone-shaped lipids influence negative curvature (if found in the inner membrane leaflet of a bud) or positive curvature (if found in the outer leaflet) of a membrane, depending on whether they are right-side-up or inverted, respectively [159]. Also, drugs involved in changing the membrane lipid content or organization, such as the cholesterol chelator methyl β-cyclodextrin, abrogate both clathrin- and caveolae-mediated endocytosis at the vesicle-budding stage [127]. In addition, PtdIns, in particular PtdIns(4,5)P2 , play important roles in endocytic events [157]. PtdIns(4,5)P2 and PtdIns(3,4,5)P3 interact with many of the clathrin adaptors including AP-2, epsin, and dynamin and therefore can function as co-receptors to selectively recruit these endocytic proteins to the PM [160, 161]. Moreover, PtdIns(4,5)P2 is also linked to the actin cytoskeleton, which has been implicated in the regulation of endocytic events (see above) [157, 162].

With regard to trafficking events, PtdIns are crucial for the biology of endosomes, and different PtdIns are distributed differentially. PtdIns(4,5)P2 and PtdIns(3,4,5)P3 are enriched at the PM. PtdIns(3,4)P2 is found mainly at EEs and PtdIns(3,5)P2 at LEs, where they bind to different modules found in lipid-binding proteins such as epsin, EEA1, and Ezrin [157] (Figure 7.4). In this way, the cell is able to achieve both diversity and specificity in the targeting of its endosomal traffic.

Rab PROTEINS IN ENDOCYTIC TRAFFIC Endocytic and post-endocytic processes in hepatocytes are tightly controlled in both spatial and temporal manners. A family of proteins that contributes to this regulation is the RabGTPases, which belong to the Ras superfamily of small GTPases that function as molecular switches by cycling between an inactive, cytosolic GDP-bound form and an active, membrane-associated GTP-bound form. Like all members of the Ras superfamily, RabGTPases are regulated by a subset of proteins that control their activation and membrane association [163, 164]. In their active, GTP-bound form, Rabs bind to various effectors and regulate different trafficking steps including cargo selection and vesicle budding, targeting, and fusion [165, 166]. Different Rab proteins localize specifically to a subset of membranes and vesicles and therefore can be used as marker proteins for these organelles. Five Rab proteins are of particular interest in the endocytic pathway: Rab4, Rab5, Rab7, Rab9, and Rab11 (Figure 7.4). Rab4 and Rab5 are localized to the EE, where Rab4 mediates rapid recycling to the PM [167, 168] and Rab5 promotes heterotypic fusion between clathrin-coated vesicles (CCVs) and the EE, as well as EE-to-EE homotypic fusion [169, 170]. Rab7 is found at LEs and is required for EE-to-LE and LE-to-lysosome transport [171, 172]. Rab9 is involved in retrograde trafficking from the endosome to the TGN [173, 174], whereas Rab11 resides at the recycling endosome and mediates slow recycling processes back to the PM [175, 176].

HEPATOCYTE-SPECIFIC ENDOCYTIC MEMBRANE TRAFFIC Iron homeostasis by specific trafficking of the transferrin receptors 1 and 2 Maintenance of iron homeostasis is an important function of the liver. This is achieved in part by dual receptor cascades (TfR (transferrin receptor) 1 and 2) that act to both

7: ENDOCYTOSIS AS AN ESSENTIAL PROCESS IN LIVER FUNCTION AND PATHOLOGY

Basolateral Membrane

Pdtins(4,5)P2

Pdtins(3,4)P2

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Pdtins(3,4,5)P3

CCP Rab4 ARE

Rab5 Rab4 Rab5 BEE Rab4

Pdtins(3)P Pdtins(3,4)P2

AEE

Rab5

BC

Rab7

Rab7

LYS

Pdtins(3,5)P2

Apical Membrane

LE/MVB Rab7

Figure 7.4 Distribution of RabGTPases and phosphatidylinositides at the various membranous compartments within the cell. RabGTPases mediate trafficking from coated vesicles to early endosomes (Rab5) or late endosomes (Rab7) as well as recycling back to the plasma membrane (Rab4). In addition, the plasma membrane and the endosomal pathway are enriched in a subset of phosphatidylinositides that contribute to Rab targeting and additional specificity of the different compartments. See text for details

internalize iron-coupled transferrin and stimulate peptide signals to iron transporters in other cells and tissues. TfR2 is expressed almost exclusively in the liver. TfR1, which is the “classic” receptor generally referred to as “the” TfR, is modestly expressed in the liver [177]. Importantly, these related receptors appear to be regulated differently, to stimulate distinct downstream effectors, and to be internalized by distinct mechanisms. Surprisingly, TfR1 is internalized by hepatocytes via a clathrin-dependent mechanism, whereas TfR2 localizes to lipid rafts and interacts with Cav-1 [178]. Furthermore, TfR1 is recycled back to the surface, but TfR2 is delivered to MVBs and subsequently trafficked to the lysosome for degradation [179]. Additional studies are needed to elucidate the molecular machinery that regulates the different endocytic pathways used by the two TfR types. For example, it is important to define how the related receptors are sequestered into distinct endocytic containers and whether TfR2, similar to EGFR, is ubiquitinated, thereby targeting TfR2 for entry into the degradative pathway.

HEPATITIS VIRUSES: MASTERS OF HEPATOCELLULAR ENDOCYTOSIS Human pathogens such as hepatitis B virus (HBV) and hepatitis C virus (HCV) are masters in the hijacking of cellular processes, in particular the endocytic pathway

that internalizes these virions and mediates their transport to the hepatocyte interior.

Hepatitis B (HBV) HBV belongs to the family of human Hepadnaviridae, which comprises encapsulated DNA viruses with a circular, partially double-stranded DNA genome [180]. The infectious virion, also called the “Dane particle,” consists of the inner core (or nucleocapsid) plus the outer surface coat. The inner core of the virion contains the viral DNA surrounded by the E-antigen (HBeAg) and the viral polymerase, encapsulated by a polymer of HBV core proteins (HBcAgs). Several hepatocyte receptors have been found to interact with HBV including TfR, IgA receptor, IL-6 receptor, and asialoglycoprotein receptor (ASGPR) [181–184], which suggests that endocytosis might be the initial uptake mechanism, instead of direct fusion with the PM. Indeed, it was demonstrated recently that virus entry into the target cell is dependent on CME because expression of the dominant-negative clathrin AP Eps15 or of dominant-negative Rab5 or cytosolic acidification abolished the uptake of core particles. In contrast, disruption of lipid rafts by nystatin had no effect [185]. The concept of clathrin-mediated entry of HBV is further supported by the work of Toh and colleagues, who showed an interaction of HBsAgs with AP-2, a PM-associated clathrin AP [186]. Once internalized, the endocytosed particles reach the EEs and are subsequently targeted to the lysosome [185],

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where the core antigen is cleaved by cathepsin B. This cleavage releases the nucleocapsid for transport to the nucleus in a microtubule-dependent manner [187]. For successful replication, the HBV is transported into the nucleus. Once released to the karyoplasm, the viral polymerase can initiate a new replication cycle for HBV, employing a variety of control mechanisms to ensure low toxicity, and a long cell lifetime to allow continuous virus production [188, 189].

Hepatitis C (HCV) In contrast to HBV, HCV belongs to the Flaviviridae family and is a small, enveloped, positive-stranded RNA virus. Like HBV, the HCV particle consists of a nucleocapsid surrounded by a lipid bilayer, which harbors the two viral envelope proteins E1 and E2. E1 and E2 can form heterodimers and play an important role in HCV entry [190, 191]. As is the case for HBV, the study of HCV entry into the target cell has been very difficult because of the lack of a suitable cell culture system that is efficient enough to produce infectious particles. The entry mechanism of HCV is similar to that of HBV, although the receptor proteins involved in HCV entry differ from the HBV receptors. Among the cellular factors that bind to HCV and help mediate internalization are glycosaminoglycans (GAGs) such as chondroitin sulfate, dermatan sulfate, keratin sulfate, and heparin sulfate, which seem to be the first attachment sites for the virus [192–194]. Furthermore, low-density lipoprotein receptor (LDL-R) appears to associate with HCV [192, 195], and antibodies against LDL-R or purified LDL inhibit cell surface adsorption of HCV isolated from patients [196, 197]. In addition, CD81, a member of the tetraspanin family [198], and the human scavenger receptor class B type I (SR-B1) [199] have been identified as binding partners for the viral E2 protein. The identification of these interaction partners suggested that CME might be responsible for HCV entry. Indeed, the use of siRNA against CHC and of chemical inhibitors provided evidence that HCV entry into the target cell is supported by CME [199, 200] and is pH-dependent, indicating that the fusion occurs after endocytosis [201, 202]. The entry of HCV into the target cell is considered to be a slow multistep process that involves a number of virus–cell interactions. In the current model, the first contact of HCV with the target cell is facilitated by GAGs and LDL-R, followed by interactions of HCV with SR-B1 and CD81 [191]. Then HCV is internalized by CME and fusion occurs, probably at the EE [200]. The details of the fusion process are still unclear, but some data indicate that E2 [203] and/or E1 [203, 204] might be involved due to their sequence homologies with other fusion proteins. Our understanding of how HBV and HCV exploit the endocytic machinery of the hepatocyte is still at an embryonic stage. Substantially more study is needed to

elucidate how these viral-based trafficking processes are regulated, organized, and targeted.

FUTURE DIRECTIONS New imaging and biochemical techniques have provided detailed insights into the mechanisms of vesicle formation and post-endocytic trafficking. Nevertheless, the list of protein and lipid components of the vesicle-forming machinery is still expanding and the functions of the components in specific transport processes need to be established. It is important to understand how the hepatocyte manages to coordinate and control a massive protein and lipid network that supports vesicular trafficking events. In this context, challenges for the future will be to determine how secretory and endocytic cargo are sequestered, how different subdomains in one compartment are maintained, and which signaling cascades control sequential steps in endocytic trafficking events. Insights into these processes will provide a better understanding of hepatocyte function.

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175. Ullrich, O., Reinsch, S., Urbe, S., Zerial, M. and Parton, R.G. (1996) Rab11 regulates recycling through the pericentriolar recycling endosome. J Cell Biol , 135 (4), 913–24. 176. Wang, X., Kumar, R., Navarre, J., Casanova, J.E. and Goldenring, J.R. (2000) Regulation of vesicle trafficking in madin-darby canine kidney cells by Rab11a and Rab25. J Biol Chem, 275 (37), 29138–46. 177. Kawabata, H., Germain, R.S., Vuong, P.T., Nakamaki, T., Said, J.W. and Koeffler, H.P. (2000) Transferrin receptor 2-alpha supports cell growth both in iron-chelated cultured cells and in vivo. J Biol Chem, 275 (22), 16618–25. 178. Calzolari, A., Raggi, C., Deaglio, S., Sposi, N.M., Stafsnes, M., Fecchi, K., Parolini, I., Malavasi, F., Peschle, C., Sargiacomo, M. and Testa, U. (2006) TfR2 localizes in lipid raft domains and is released in exosomes to activate signal transduction along the MAPK pathway. J Cell Sci , 119 (Pt 21), 4486–98. 179. Robb, A.D., Ericsson, M. and Wessling-Resnick, M. (2004) Transferrin receptor 2 mediates a biphasic pattern of transferrin uptake associated with ligand delivery to multivesicular bodies. Am J Physiol Cell Physiol , 287 (6), C1769–75. 180. Herzer, K., Sprinzl, M.F. and Galle, P.R. (2007) Hepatitis viruses: live and let die. Liver Int , 27 (3), 293–301. 181. Dash, S., Rao, K.V. and Panda, S.K. (1992) Receptor for pre-S1(21-47) component of hepatitis B virus on the liver cell: role in virus cell interaction. J Med Virol , 37 (2), 116–21. 182. Franco, A., Paroli, M., Testa, U., Benvenuto, R., Peschle, C., Balsano, F. and Barnaba, V. (1992) Transferrin receptor mediates uptake and presentation of hepatitis B envelope antigen by T lymphocytes. J Exp Med , 175 (5), 1195–205. 183. Neurath, A.R., Strick, N. and Li, Y.Y. (1992) Cells transfected with human interleukin 6 cDNA acquire binding sites for the hepatitis B virus envelope protein. J Exp Med , 176 (6), 1561–69. 184. Treichel, U., Meyer zum Buschenfelde, K.H., Dienes, H.P. and Gerken, G. (1997) Receptor-mediated entry of hepatitis B virus particles into liver cells. Arch Virol , 142 (3), 493–98. 185. Cooper, A. and Shaul, Y. (2006) Clathrin-mediated endocytosis and lysosomal cleavage of hepatitis B virus capsid-like core particles. J Biol Chem, 281 (24), 16 563–69. 186. Toh, Q.C., Tan, T.L., Teo, W.Q., Ho, C.Y., Parida, S. and Chen, W.N. (2005) Identification of cellular membrane proteins interacting with hepatitis B surface antigen using yeast split-ubiquitin system. Int J Med Sci , 2 (3), 114–17. 187. Sodeik, B. (2000) Mechanisms of viral transport in the cytoplasm. Trends Microbiol , 8 (10), 465–72. 188. Kann, M., Schmitz, A. and Rabe, B. (2007) Intracellular transport of hepatitis B virus. World J Gastroenterol , 13 (1), 39–47. 189. Beck, J. and Nassal, M. (2007) Hepatitis B virus replication. World J Gastroenterol , 13 (1), 48–64.

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190. Deleersnyder, V., Pillez, A., Wychowski, C., Blight, K., Xu, J., Hahn, Y.S., Rice, C.M. and Dubuisson, J. (1997) Formation of native hepatitis C virus glycoprotein complexes. J Virol , 71 (1), 697–704. 191. Helle, F. and Dubuisson, J. (2008) Hepatitis C virus entry into host cells. Cell Mol Life Sci , 65 (1), 100–12. 192. Andre, P., Komurian-Pradel, F., Deforges, S., Perret, M., Berland, J.L., Sodoyer, M., Pol, S., Brechot, C., Paranhos-Baccala, G. and Lotteau, V. (2002) Characterization of low- and very-low-density hepatitis C virus RNA-containing particles. J Virol , 76 (14), 6919–28. 193. Germi, R., Crance, J.M., Garin, D., Guimet, J., Lortat-Jacob, H., Ruigrok, R.W., Zarski, J.P. and Drouet, E. (2002) Heparan sulfate-mediated binding of infectious dengue virus type 2 and yellow fever virus. Virology, 292 (1), 162–68. 194. Hilgard, P. and Stockert, R. (2000) Heparan sulfate proteoglycans initiate dengue virus infection of hepatocytes. Hepatology, 32 (5), 1069–77. 195. Thomssen, R., Bonk, S., Propfe, C., Heermann, K.H., Kochel, H.G. and Uy, A. (1992) Association of hepatitis C virus in human sera with beta-lipoprotein. Med Microbiol Immunol , 181 (5), 293–300. 196. Agnello, V., Abel, G., Elfahal, M., Knight, G.B. and Zhang, Q.X. (1999) Hepatitis C virus and other flaviviridae viruses enter cells via low density lipoprotein receptor. Proc Natl Acad Sci USA, 96 (22), 12766–71. 197. Wunschmann, S., Medh, J.D., Klinzmann, D., Schmidt, W.N. and Stapleton, J.T. (2000) Characterization of hepatitis C virus (HCV) and HCV E2 interactions with CD81 and the low-density lipoprotein receptor. J Virol , 74 (21), 10055–62.

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198. Pileri, P., Uematsu, Y., Campagnoli, S., Galli, G., Falugi, F., Petracca, R., Weiner, A.J., Houghton, M., Rosa, D., Grandi, G. and Abrignani, S. (1998) Binding of hepatitis C virus to CD81. Science, 282 (5390), 938–41. 199. Blanchard, E., Belouzard, S., Goueslain, L., Wakita, T., Dubuisson, J., Wychowski, C. and Rouille, Y. (2006) Hepatitis C virus entry depends on clathrin-mediated endocytosis. J Virol , 80 (14), 6964–72. 200. Meertens, L., Bertaux, C. and Dragic, T. (2006) Hepatitis C virus entry requires a critical postinternalization step and delivery to early endosomes via clathrin-coated vesicles. J Virol , 80 (23), 11571–78. 201. Hsu, M., Zhang, J., Flint, M., Logvinoff, C., Cheng-Mayer, C., Rice, C.M. and McKeating, J.A. (2003) Hepatitis C virus glycoproteins mediate pH-dependent cell entry of pseudotyped retroviral particles. Proc Natl Acad Sci USA, 100 (12), 7271–76. 202. Tscherne, D.M., Jones, C.T., Evans, M.J., Lindenbach, B.D., McKeating, J.A. and Rice, C.M. (2006) Time-and temperature-dependent activation of hepatitis C virus for low-pH-triggered entry. J Virol , 80 (4), 1734–41. 203. Drummer, H.E., Boo, I. and Poumbourios, P. (2007) Mutagenesis of a conserved fusion peptide-like motif and membrane-proximal heptad-repeat region of hepatitis C virus glycoprotein E1. J Gen Virol , 88 (Pt 4), 1144–48. 204. Garry, R.F. and Dash, S. (2003) Proteomics computational analyses suggest that hepatitis C virus E1 and pestivirus E2 envelope glycoproteins are truncated class II fusion proteins. Virology, 307 (2), 255–65. 205. Heuser, J. (1989) Effects of cytoplasmic acidification on clathrin lattice morphology. J Cell Biol , 108, 401–11.

8

Membrane Transport in Hepatocellular Secretion Susan Chi and Mark McNiven Department of Biochemistry and Molecular Biology, Miles and Shirley Fiterman Center for Digestive Diseases, Mayo Clinic & Foundation, Rochester, MN, USA

INTRODUCTION The major functions of the hepatocyte are the synthesis, packaging, and directed transport into the sinusoidal space of multiple plasma proteins. These processes depend on a complex, highly-organized vesicular trafficking machinery that requires the coordinated synergistic action of scores of enzymes, cytoskeletal proteins, molecular motors, and coat proteins. Through conventional biochemical and molecular methods, the use of genetic model organisms, and recent advancements in live cell imaging, much has been learned about these trafficking pathways in hepatocytes and other epithelial cells. This review will focus on the molecular mechanisms by which nascent proteins are sequestered, packaged into vesicle carriers, and targeted to specific hepatocellular destinations during the secretory process.

VESICLE FORMATION IN THE SECRETORY PATHWAY The basic structural framework of the secretory apparatus includes the endoplasmic reticulum (ER) and the Golgi apparatus (Figure 8.1). Usually located at the periphery of the cell, the ER is characterized by a network of interconnected, highly-convoluted membranous cisternae. The Golgi apparatus, in contrast, is localized to the perinuclear region of the cell and consists of organized stacks of flattened saccules and associated secretory

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

vesicles. The cis-Golgi network (CGN) and trans-Golgi network (TGN) are membranous tubular reticula at the entry and exit sides of the Golgi apparatus, respectively. As the early hypotheses of protein secretion were being evaluated, the ER and Golgi apparatus were analyzed extensively in exocrine pancreas and rat liver due to the ease of secretory product visualization. These organelles were postulated to act sequentially in the synthesis, transport, and packaging of proteins destined for extracellular discharge by exocytosis. The mechanisms by which proteins are transported anterogradely from the ER to the Golgi apparatus and vectorially through the Golgi apparatus itself are thought to involve vesicular carriers. At the ER, constituents of the COP (coat promoter) II coat are recruited to form buds and generate vesicles known as vesicular tubular clusters (VTCs), which have been proposed to coalesce to form a complex network of tubules called the endoplasmic reticulum/Golgi intermediate compartment (ERGIC) [1]. From the ERGIC, COPI coat components are recruited and induce the formation of vesicles that mediate ERGIC-to-Golgi protein/membrane transport. COPI-coated vesicles are also thought to mediate the retrograde transport of resident ER proteins and Golgi enzymes from the Golgi apparatus to the ERGIC and ER or to the CGN, respectively. In addition to secretory proteins, membrane proteins and lysosomal enzymes pass through the secretory apparatus to reach their final destinations, which reflects the important sorting role of the Golgi apparatus. This sorting function has been attributed

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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COPI-COATED VESICLES COPI-coated vesicles and protein trafficking through the ER and Golgi apparatus

Figure 8.1 Elaborate vesicular processes in the hepatocyte. Thin-section electron micrograph of a rat liver hepatocyte from the original collections of Dr. Keith Porter, about 1962. Deep in the cytoplasm, parallel cisternae of the endoplasmic reticulum (ER) appear to lead into the flattened, stacked saccules of the Golgi apparatus (G). Scores of small vesicular profiles can be seen in the Golgi region, as well as larger lipoprotein-filled secretory vesicles budding from the trans side of the Golgi. Specific coats, accessory proteins and address tags characterize many of these vesicles. G: Golgi; ER: endoplasmic reticulum; Lys: lysosome

predominantly to, and is best characterized for, the TGN. At this site, several distinct vesicle types containing different cargo proteins and harboring particular coat proteins have been reported to arise. Non-coated secretory vesicles from the TGN are thought to transport constitutively secreted proteins to the cell surface. Notably, the formation of COPI- and COPII-coated vesicles and TGN-derived vesicles requires similar molecular components. In these vesicle populations, the priming of vesicle budding is initiated by the binding of specific small GTPases (ADP ribosylation factor (ARF) for both COPI-coated vesicles and clathrin-coated adaptor protein (AP)-1-positive vesicles; Sar1 for COPII-coated vesicles) at the site of vesicle budding. Moreover, unique GTP-GDP exchange factors (GEFs) and GTP-activating proteins (GAPs) for each vesicle type activate the membrane-binding and GTP-hydrolysis activities of the GTPases, respectively. Once bound to the membrane, the GTPases induce the recruitment of COPI coat components, COPII, or the AP-1 complex in clathrin-coated vesicles (CCVs). The coat proteins, as well as accessory factors, are necessary components for vesicle biogenesis from various compartments along the secretory pathway.

COPI-coated vesicles have been studied primarily in the retrieval of ER resident proteins from the Golgi. Selected COPI subunits are known to associate with dilysine amino acid motifs in the cytoplasmic domains of ER-resident enzymes [2]. Furthermore, several COPI temperature-sensitive mutants in yeast have revealed defects in Golgi-to-ER protein transport but not in ER-to-Golgi transport at the restrictive temperature [3]. Further, COPI-coated vesicles have been shown to mediate retrograde transport of resident ER proteins by using the KDEL (single-letter code for amino acids) receptor [4], confirming a retrieval role for COPI coat proteins. Whether COPI sorts cargoes in opposing anterograde and retrograde directions at the ERGIC was studied recently [5]. Interestingly, different isoforms of the small GTPase ARF were reported to be involved in the support of this opposing traffic. Knockdown of Arf1 and Arf3 impaired β-COP-positive ERGIC-to-cis-Golgi transport, whereas knockdown of Arf4+Arf5 or Arf1+Arf5 arrested the KDEL receptor in the cis-Golgi or ERGIC, respectively [6]. Rab-dependent sorting also provides another mechanism of specificity. Overexpression of an inactive form of Rab1b failed to recruit COPI [7], and Rab2 was shown to promote the release of retrograde-directed COPI vesicles [8]. Thus, directional transport from the ERGIC could be controlled by distinct ARF- or Rab-dependent sorting by COPI. The COPI coat consists of seven coat proteins (α, β, β′ , γ, δ, ε, ζ COP) that form a 680-kDa complex and associate with the small GTP-binding protein Arf1. After initiation by GTP-ARF, COPI bud formation is thought to be mediated by a cytoplasmic pool of coats recruited en bloc as a preassembled coat complex to budding regions on membranes [9] (Figure 8.2a,b). GTP binding to ARF, and the resultant activation of the protein, is catalyzed by a membrane-associated GTP-GEF such as GBF1, which is sensitive to the fungal agent brefeldin A [10]. The transient action of GBF1 on Arf1 at the membrane has been demonstrated by fluorescence recovery after photobleaching. After photobleaching of the YFP-GBF1 pool at the Golgi, YFP-GBF1 rapidly translocates from the cytoplasm to the Golgi, which demonstrates substantial dynamics of GBF1 at the membrane [11]. Recruitment of COPI coat proteins to target sites is thought to shape the membrane into buds (reviewed in [12, 13]). The shape of the COPI-coated bud might be

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(a)

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(b)

Figure 8.2 COPI-coated vesicles in protein trafficking through the secretory pathway. (a) A Golgi fraction isolated from rat liver shows Golgi stacks of two to three flattened cisternae (G) and scores of vesicular and budding profiles. The bristle coat of clathrin on some of these profiles (arrowheads) is visibly different from the thin electron-dense coat of COPI buds and vesicles (arrowheads). Images from Dr. John Bergeron, McGill University, Montreal, Canada. Inset: purified yeast COPI vesicles. Reprinted with permission from [13]. Copyright 1986 American Association for the Advancement of Science. (b) COPI-coated vesicle formation involves the COPI coatomer complex, which consists of seven subunit proteins. Two of these, β-COP and γ-COP, mediate important interactions with the small GTP-binding protein Arf1 and with the cytosolic tails (specifically dilysine, or KK, motifs) of transmembrane proteins (such as p24 family members; see text). ARF-GDP/GTP exchange is one of the initial steps in the recruitment of the coatomer complex for COPI vesicle biogenesis

determined by the uniform organization of coatomer and ARF in their polymerized form. p24 family members with roles as putative cargo receptors induce polymerization of the coatomer complex. Thus, a trimeric complex of coatomer, ARF, and a tetramer of p24 cytoplasmic tails is postulated to provide the vesicle-budding machinery in the biogenesis of COPI vesicles (reviewed in [14]). Hydrolysis of ARF-bound GTP initiates the uncoating of coated vesicles, which is critical for their fusion to the acceptor membrane. Once in a GDP-bound form, ARF dissociates from the membrane and causes coatomer to detach, yielding uncoated vesicle. Notably, ARF-GAP, which is an activator of ARF GTP-hydrolysis activity, is recruited to membranes by interaction with the KDEL receptor, which suggests that this receptor tail might regulate COPI coat assembly at the Golgi [14].

COPII-COATED VESICLES COPII-coated vesicles and ER-to-Golgi protein trafficking Protein transport from the ER to the Golgi complex is believed to be mediated by vesicular carriers that originate from morphologically defined, specialized ribosome-free

regions of the ER called transitional elements (tER) [15]. ER buds and an abundance of small vesicular profiles at these sites, which are distinct from smooth ER, could be clearly visualized in the classic electron micrographs of pancreatic acinar cells by Jamieson and Palade [16]. Recently, studies with live cell imaging demonstrated that these transitional areas are formed transiently and randomly from the ER [17]. The cargo carriers that mediate ER-to-Golgi protein transport have been subjects of intense study. As for many of the protein trafficking steps along the secretory pathway, the answers arose from studies in yeast Saccharomyces cerevisiae conditional sec (secretion) mutants. By using yeast membranes for ER vesicle budding in vitro, cell-free assays could be reconstituted. These systems enabled the isolation of ER-derived transport vesicles (Plate 8.1a, bottom inset) that were 60–65 nm in diameter and exhibited a distinct electron-dense coat on their surface, termed COPII [18]. COPII coat components were identified from the purified vesicles and revealed to be distinct from those of COPI coats [18]. Immunoreactivity of antibodies to yeast COPII coat components could be detected throughout groups of vesicles in ER cisternae of pancreatic β-cells and acinar cells [19, 20]. These COPII-positive vesicular profiles were postulated to be anterograde transport carriers because of their ER-proximal location. Indeed, two independent

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studies in normal rat kidney (NRK) cells [21] and in human hepatoma HepG2 cells [22] reported that cargo protein was present at bud sites upon ER exit and within small 40–80 nm carrier vesicles and tubules referred to as vesicular-tubular clusters (Plate 8.1a). These results confirmed the forward-transport-carrier role of these groups of vesicles. The ER cisternae near VTCs often exhibited budding profiles that were decorated with an electron-dense, honeycomb-patterned coat (Plate 8.1a, top inset), which was identified as COPII by immunogold labeling of NRK cell sections with antibodies against COPII coat components. However, more recent data generated by electron microscopy and tomography have suggested that large tubular protrusions are not always coated with COPII [23]. Thus, a current model for the formation of VTCs suggests that vesicles are formed by a COPII complex at the tER, resulting in membrane curvature. Following rapid disassembly of the coat, uncoated vesicles are formed. Alternatively, membrane protrusion from sites adjacent to the tER could lead to the formation of a tubular structure that is not coated with COPII proteins. Either vesicles or tubules are likely to be precursors of VTCs (reviewed in [24]).

COPII-coated vesicle protein components and vesicle formation The COPII coat components, which are distinct from the COPI coatomer, were postulated to induce vesicle budding of ER-derived transport vesicles because only the GTPase Sar1, Sec13/31, and Sec23/24 were found to be necessary for the generation of COPII-coated vesicles from washed microsomes [18]. Sec12, a GTP-GEF for Sar1 that is localized to the ER, initiates COPII vesicle biogenesis. After the activation of Sar1, the adaptor coat components Sec23/24 are recruited to bind cargo. Subsequently, the structural Sec13/31 coats are recruited to form the coat and to contribute to membrane deformation (reviewed in [25]) (Plate 8.1b). The selection of cargo including transmembrane proteins and v-SNAREs (soluble NSF attachment receptors) by Sec23/24 has been reported [26, 27]. Sec24 has three different sites for recognizing ER-to-Golgi v-SNAREs such as Sed5, Bet1, and Sec22. The interaction is regulated by the assembly state of the SNAREs, which suggests that COPII proteins can be involved in vesicle fusion specificity in the ER-to-Golgi step [27]. Although the minimal requirements for in vitro vesicle biogenesis are the five molecules listed above, the essential gene product of Sec16 appears to be required for vesicle formation in vivo [13]. Sec16 on the vesicle interacts with Sec23/Sec24 [13], Sec31 [28], and Sed4, an ER membrane protein that is homologous to Sec12. Sec16 likely plays a regulatory role, which can be bypassed in in vitro experiments [13]. In mammalian cells, GST–hSec23-binding columns allowed the

identification of a 250 kDa protein that could be detected by Western blotting with an antibody against yeast Sec16, showing that Sec16 orthologs are also present in mammalian cells [29]. p250 is recruited to ER membranes in a Sar1-dependent manner, and depletion of p250 partially impaired VSV-G transport from the ER, which suggests a role for p250 in membrane traffic from the ER [30]. Another protein isolated from the GST–hSec23-binding columns, p125, is likely expressed in liver, as demonstrated by Northern blots [29], and is localized to ER exit sites that participate in the organization of this compartment [31]. Amino acid analysis showed sequence homology between p125 and phosphatidic acid (PA)-preferring phospholipase A1, and seemed to indicate the importance of phospholipid metabolism in vesicle formation [29]. In the final stages of COPII vesicle biogenesis, the GTP used during vesicle formation is hydrolyzed by Sec23p, a Sar1p-specific GAP. The fission of a COPII vesicle is believed to require the N-terminal helix of Sar1. Upon GTP binding by Sar1, membrane insertion of the N-terminal α helix deforms synthetic liposomes into narrow tubules. Mutation of the helix led to a defect in membrane curvature and the formation of vesicles from native ER, although the recruitment of coat proteins appeared to be normal [32]. Moreover, inhibition of GTP hydrolysis by Sar1 resulted in COPII-coated vesicles that failed to detach from the ER, suggesting that regulation of the N-terminus of Sar1 by GTP binding and hydrolysis controls COPII vesicle fission [33]. In addition to Sar1, other small GTPases called Rabs support this transport process. Rab1 is known to function in vesicles budding from the ER. Another Rab GTPase, Rab11, is required in the late Golgi (reviewed in [34]) (Figure 8.3).

TGN-DERIVED VESICLES Protein sorting and trafficking events at the TGN The TGN is a tubulovesicular network that acts primarily in sorting proteins to their final destinations (Plate 8.2a). Within this membranous reticulum, cargo proteins are segregated efficiently into distinct vesicles that will be targeted to various compartments, including the endosome/lysosome system, the apical or basolateral domains in polarized cells, and the secretory granule pool. In regulated secretory cells, this granule pool awaits the appropriate extracellular stimulus to exocytose its contents. The TGN is known to interact intimately and dynamically with the endosomal apparatus, especially during events related to receptor recycling, endosomal maturation, and the delivery of lysosomal enzymes (reviewed in [35]). Exocytic secretion can be classified roughly into two categories: clathrin-coated and non-clathrin-coated

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Figure 8.3 Specific small Rab GTPases and phosphoinositides are localized to distinct membrane compartments along the hepatocyte secretory pathway. Rab1 and Rab2 are believed to mediate the traffic of nascent proteins from the ER to the cis-Golgi. Rab6 is a kinesin-associated Rab that mediates transport within the Golgi stacks, Rab11 regulates exit from the TGN, and Rab8 functions in the targeting of secretory vesicles from the Golgi to the plasma membrane. Rab9 has a role in late endosome-to-TGN transport. The precise functions of these rab proteins are currently unclear. PtdIns(4)P is significantly enriched in the Golgi and is believed to mediate the recruitment of multiple proteins that support vesicle formation at the TGN. PtdIns(4,5)P2 at the Golgi functions with Arf1 and phospholipase D (PLD) and is also involved in vesicle formation. LE/MVB: late endosome/multivesicular body; ER: endoplasmic reticulum; BC: bile canalicular membrane

vesicles. The former are responsible for concentrating (i) newly synthesized lysosomal hydrolases for delivery to the endosomal apparatus and then to the lysosomes and (ii) enzymes and hormones in the secretory granules of cells that have regulated secretory pathways [35, 36]. In contrast, non-CCVs are believed to package mainly cargo proteins that are secreted constitutively. Specific sets of proteins and lipids are associated with both types of vesicle, as discussed below [37, 38].

Clathrin-coated TGN-derived vesicles Clathrin was the first coat to be visualized in the Golgi region of mammalian cells. An electron-dense, spiked coat on TGN membranes in liver hepatocytes could be seen clearly in the original micrographs of Novikoff and Yam [39] (Plate 8.2a). Similar coats were visible on buds and vesicles of the TGN compartment in the principal epithelial cells of the epididymis [40], and clathrin patches have been reported along the delimiting membrane of secretory granules in regulated secretory cells [41]. TGN-derived CCVs are now known to have a coat composed of clathrin triskelia and the AP complexes AP-1 (made up of γ-, β1-, µ1-, and σ1-adaptin subunits) and AP-3 (δ-, β3-, µ3-, and σ3-adaptin subunits) [42]. AP-1 and AP-3 are found predominantly in vesicular structures in the Golgi region;

however, an endosomal localization has also been reported [43]. The subunit µ1 of AP-1 is responsible for binding to YXX
motifs within cargo proteins, whereas β1 interacts with dileucine-based signals in cargo, in addition to clathrin. Recently, it was shown that the hinge domain of γ-adaptin can also bind to clathrin. The γ-adaptin subunit recruits accessory proteins to the site of vesicle formation (reviewed in [44]). Several groups have identified a family of monomeric adaptors, the Golgi-localized, γ-ear-containing, ARFbinding proteins (GGAs) [45]. There are three known GGA proteins: GGA1, GGA2, and GGA3 [46–48]. Each GGA protein contains three domains: VHS (Vps27, Hrs, Stam), GAT (GGA and TOM), and gamma-adaptin ear (GAE). Through the VHS domain, the GGAs recognize DXXLL motifs in specific cargo such as the mannose 6-phosphate receptor (MPR) tail and thus provide a sorting function between the TGN and endosomes [46]. The GAT domain interacts with ARF; mutations in this domain abrogated recruitment of GGAs to the TGN as well as binding to ARF [47, 48]. The GAE domain is homologous to the GAE domain of the AP-1 complex-recruiting accessory proteins [49]. The GGAs are essential for packaging MPRs into CCVs and for transporting them from the TGN to endosomes. Overexpression of a truncated GGA containing only VHS and GAT domains resulted in accumulation of MPRs in the TGN [50]. GGAs also have a ubiquitin-binding

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domain [51], which suggests that they might mediate the sorting of ubiquitinated cargo at the TGN. Another adaptor complex at the TGN, AP-3, is expressed ubiquitously, is localized to buds/vesicles associated with the TGN [42], and can interact with clathrin [52] and with sorting signals in the cytoplasmic tails of lysosomal membrane proteins [53]. Correct targeting of lysosomal-associated membrane protein-1 (LAMP-1) and lysosomal integral membrane protein-2 (LIMP-2) appears to be mediated by the AP-3 adaptor complex [54]. CCVs generated from the TGN are involved in the transport of newly synthesized lysosomal enzymes. Newly synthesized, soluble lysosomal enzymes acquire mannose 6-phosphate residues at the CGN, and these residues are recognized by the MPRs at the TGN [35]. At this site, the AP-1 adaptor complex has long been thought to bind to the MPRs and then to recruit clathrin during bud and vesicle formation. These CCVs subsequently mediate the transport of MPR-lysosomal enzyme complexes to endosomes [55]. However, the recently identified GGA proteins are now known to be essential for the transport of MPRs from the TGN to the endosome [45]. Thus, it is unclear whether AP-1 and GGA proteins cooperate or function separately in mediating the transport of MPRs. How are CCVs formed at the TGN? Part of the answer lies in what is already known about clathrin coats at the plasma membrane. Adaptors at both the cell surface (AP-2) and at the Golgi apparatus (AP-1) are thought to initiate coat assembly by recruiting clathrin triskelia that polymerize to form the coat [43]. As with the AP-2 adaptor at the plasma membrane, the AP-1 β1 subunit promotes clathrin cage assembly in vitro [56], and the µ1 subunit associates with specific sorting motifs in the cytosolic tails of MPR and TGN38 (a membrane protein that is localized predominantly to the TGN) [53]. Similar to the COPI coatomer, AP-1 and GGA recruitment to TGN membranes is sensitive to BFA and thus is regulated by ARF (see Section “COPI-coated Vesicles”) [57, 58] (Plate 8.2d). Recently, new clathrin adaptors and accessory proteins that support CCV formation at the TGN have been identified. Since the adaptor γ-synergin was first identified as a binding partner for the γ-adaptin subunit of AP-1 [59], several accessory proteins such as Rabaptin-5 [60], EpsinR [61], and γ-BAR [62] have been reported to interact with γ-adaptin of AP-1 and are therefore implicated in CCV formation at the TGN. Although the in vivo order of association of ARF, adaptors (AP-1 and GGAs), and MPR (transmembrane cargo receptor) has not been established, a model for the formation of CCVs at the TGN has been proposed. This model suggests that membrane-tethered, activated ARF recruits AP-1 and/or GGAs to the membrane, and those adaptors bind to cargo, clathrin, and accessory proteins with their various domains.

Non-clathrin-coated TGN-derived vesicles In a purified rat liver stacked Golgi fraction that was immobilized on magnetic beads, vesicle budding was shown to be ATP-, cytosol-, and temperature-dependent [63]. Vesicles purified from the budding fraction were 50–200 nm in diameter and devoid of any visible coat. An 85 kDa cytosolic complex (p62 and a 25 kDa GTPase) and TGN38 were found to be essential for vesicle formation from the TGN [64, 65]. The 85 kDa cytosolic complex was later identified as a regulatory subunit of phosphatidylinositol 3-kinase (PI3-kinase) and shown to play a role in PI3-kinase activity in TGN-derived exocytic vesicle formation [66]. Another cytosolic protein, p200, was found to bind selectively to Golgi membranes and to be involved in vesicle budding from the TGN upon activation of heterotrimeric G proteins with GTPγS (a non-hydrolyzable analog of GTP), AlF4 − , or mastoparan [67]. The p200 protein is the heavy chain of non-muscle myosin 2, a protein that is involved in the generation of constitutive transport vesicles containing vesicular stomatitis virus G protein from the TGN of polarized semi-intact MDCK cells [68]. Golgins (p230/golgin-245, golgin-97, GCC88, and GCC185), structural proteins that maintain Golgi structure, are localized to the TGN by their C-terminal GRIP domain [69]. Of the four TGN golgins, p230 was reported to be involved in non-CCV formation, because p230 was found to recycle between the cytosol and TGN membrane buds and vesicles in a G protein-dependent fashion [70]. In contrast, GCC88 and GCC185 were required for retrograde transport from the early endosomes to the TGN [69, 71, 72]. The Rab8 GTPase has been implicated in vesicle transport from the Golgi to the plasma membrane. Interaction of Rab8 with one of the exocyst subunits prepares the exocytic vesicle for fusion to the plasma membrane (reviewed in [34]) (Figure 8.3).

Scission of TGN-derived vesicles Since the localization of the large GTPase dynamin to the Golgi apparatus, in addition to its known cell surface distribution, was reported, it has been suggested that vesicle fission at the Golgi apparatus is mediated by this molecular pinchase. Dynamin’s association with Golgi membranes was demonstrated biochemically with specific antibodies that detected dynamin in immunoblots of purified rat liver Golgi fractions and by the immunoisolation of Golgi membranes on magnetic beads coated with those dynamin antibodies [73]. Immunogold

8: MEMBRANE TRANSPORT IN HEPATOCELLULAR SECRETION

localization of Dyn2 on the TGN of HeLa cells [74] and the Golgi distribution of various GFP-tagged dynamin isoforms (especially Dyn2(aa)) [75] confirmed these findings. Cortactin, an actin-binding protein, in a complex with dynamin, functions in post-Golgi transport in liver cells. By using in vitro or intact cell experiments, it was shown that activation of Arf1 recruits actin, cortactin, and dynamin to Golgi membranes. Disruption of cortactin–dynamin interactions reduced Dyn2 recruitment to the Golgi and blocked the transit of nascent proteins from the TGN, which indicates an essential role of the cortactin–Dyn2 complex in TGN function [76]. The dynamins are believed to form large helical polymers from which many interactive proline-rich tail domains extend. These domains bind to a variety of SH3domain-containing proteins, many of which appear to be actin-binding proteins such as Abp1 and syndapin. These findings suggest that the dynamin family acts as a “polymeric contractile scaffold” at the interface between biological membranes and filamentous actin [77] (Plate 8.3).

LIPID REGULATORS INVOLVED IN VESICLE BIOGENESIS Several lipids and membrane lipid-modifying proteins appear to be involved in the formation of different types of vesicle that bud from the Golgi apparatus. PtdIns(4,5)P2 is required for the interaction of Arf1 with phospholipase D (PLD) [78, 79], a protein with abundant activity in Golgi-enriched membranes [80] (Figure 8.3). Together with activated ARF, PtdIns(4,5)P2 activates PLD1 to generate PA from phosphatidylcholine (PC). PA is required for the in vitro formation of coated vesicles from Golgi membranes in Chinese hamster ovary cells [81] and has been suggested to be an important factor in the mediation of a shape change in the lipid bilayer to facilitate invagination. Another specific phosphoinositide (PI) component at the Golgi is phosphatidylinositol 4-phosphate (PtdIns(4)P) (Figure 8.3). Experiments using RNAi to reduce the amount of the resident PI4KIIα, phosphatidylinositol 4-kinase, showed that decreased Golgi PtdIns(4)P blocks the targeting of the clathrin AP-1 to the Golgi [82]. EpsinR, an accessory protein at the Golgi that is recruited by AP-1, is also selective for PtdIns(4)P through its Epsin N-terminal homology (ENTH) domain [61]. Four-phosphate-adaptor protein (FAPP) is also known to bind to PtdIns(4)P and ARF through its pleckstrin homology (PH) domain. Knockdown of FAPP blocked Golgi-to-cell surface membrane trafficking, and over-expression of the FAPP PH domain impaired fission,

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which indicates that FAPP binding to PtdIns(4)P and ARF is essential to control post-Golgi trafficking [83]. More recently, protein kinase D (PKD) was shown to be required for membrane fission at the TGN. When a kinase-inactive form of PKD was over-expressed, cargo transport from the TGN to the cell surface was blocked, which caused the formation of many membrane tubules that could not detach from the TGN [84, 85]. Diacylglycerol (DAG) at the TGN recruits PKD and activates PKCη. In turn, the activated PKCη stimulates PKD. Inactivation of PKCη inhibits protein transport from the TGN to the cell surface [86, 87]. These data suggest that PKD acts downstream of DAG and PKCη to regulate membrane fission from the TGN (reviewed in [37]). The formation of both constitutive secretory vesicles and immature secretory granules from the TGN was stimulated by the concerted action of phosphatidylinositol transfer protein (PITP) and PLD in a cell-free system with PC12 membrane fractions [88]. This stimulatory effect was inhibited by geneticin, an antibiotic that binds PIs, which demonstrates a role for PIs in promoting secretory vesicle formation downstream of PITP and PLD. PITP and PI3-kinase are also essential components in the cell-free formation of polymeric immunoglobulin A receptor-containing exocytic vesicles in Golgi fractions isolated from rat liver [89]. Together, PITP and PI3-kinase generate a pool of PtdIns(3)P, which is thought to be essential for vesicle formation from the TGN.

FUTURE DIRECTIONS Membrane trafficking is essential for the secretion of nascent proteins into the blood space and for the formation of bile, which are major hepatocellular functions. This summary has described a small number of the adaptor, cytoskeletal, and enzyme components required to direct a nascent protein from the ER to the Golgi apparatus and then to the desired location. This machinery is conserved in all cells but is likely modified for the specific trafficking processes performed by the liver. Attaining a better understanding of how these processes are utilized by the liver under normal and pathophysiological conditions is a current and future challenge for liver cell biologists. Some areas of interest include the assembly, packaging, and transport of multiple infectious viruses (Figure 8.4a,b) [90]; the directed insertion of the ATP-binding cassette (ABC) transporters into the canalicular domain for bile formation (Figure 8.4c–e) [91]; and the transferrin-induced packaging and secretion into the blood of prohepcidin, which regulates iron homeostasis (Figure 8.4f–h) [92]. Many more examples of liver-specific secretory processes exist and require substantial cell biological investigation.

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(a)

(b)

(d)

(c) (e)

(f)

(g)

(h)

Figure 8.4 Membrane traffic in hepatocellular secretion. (a,b) Electron micrographs of HBV particles located in an undefined cytoplasmic compartment in HepG2.2.15 cells. Expression of a mutant dynamin 2 (K44A) protein induces substantial accumulation of the virus in this compartment (b, arrows) whereas expression of wild-type dynamin does not (a). a and b Reprinted from [90], Copyright  2003, with permission from Elsevier (c–e) Intracellular movement of the ABC transporter mdr1-GFP between the Golgi and the canalicular membrane in a polarized hepatocyte cell model, WIF-B9 cells. Smaller panels in (d) are time sequences of the boxed area from (c). The Golgi (G) and bile canalicular membranes (BC) are labeled. Arrows point to mdr1-GFP vesicles that move in a linear fashion from the Golgi to the BC. In (e), tubulovesicular protrusions of mdr1-GFP extend outward from the canalicular membrane. Reproduced from [91] with permission from Journal of Cell Science (f,h) Immunofluorescence images of a primary mouse hepatocyte stained with antibodies to prohepcidin (f), which appears to accumulate in the Golgi, as shown by co-staining for the peripheral Golgi protein p115 (g). N: nucleus. Reprinted from [92], Copyright  2005, with permission from Elsevier

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D mediates ADP ribosylation factor-dependent formation of Golgi coated vesicles. J Cell Biol , 134, 295–306. Wang, Y.J., Wang, J., Sun, H.Q., Martinez, M., Sun, Y.X., Macia, E., Kirchhausen, T., Albanesi, J.P., Roth, M.G. and Yin, H.L. (2003) Phosphatidylinositol 4 phosphate regulates targeting of clathrin adaptor AP-1 complexes to the Golgi. Cell , 114, 299–310. Godi, A., Di Campli, A., Konstantakopoulos, A., Di Tullio, G., Alessi, D.R., Kular, G.S., Daniele, T., Marra, P., Lucocq, J.M. and De Matteis, M.A. (2004) FAPPs control Golgi-to-cell-surface membrane traffic by binding to ARF and PtdIns(4)P. Nat Cell Biol , 6, 393–404. Liljedahl, M., Maeda, Y., Colanzi, A., Ayala, I., Van Lint, J. and Malhotra, V. (2001) Protein kinase D regulates the fission of cell surface destined transport carriers from the trans-Golgi network. Cell , 104, 409–20. Yeaman, C., Ayala, M.I., Wright, J.R., Bard, F., Bossard, C., Ang, A., Maeda, Y., Seufferlein, T., Mellman, I., Nelson, W.J. and Malhotra, V. (2004) Protein kinase D regulates basolateral membrane protein exit from trans-Golgi network. Nat Cell Biol , 6, 106–112. Diaz Anel, A.M. and Malhotra, V. (2005) PKCeta is required for β-1-γ-2/β-3-γ-2- and PKD-mediated transport to the cell surface and the organization of the Golgi apparatus. J Cell Biol , 169, 83–91. Maeda, Y., Beznoussenko, G.V., Van Lint, J., Mironov, A.A. and Malhotra, V. (2001) Recruitment of protein kinase D to the trans-Golgi network via the first cysteine-rich domain. EMBO J , 20, 5982–90. Tuscher, O., Lorra, C., Bouma, B., Wirtz, K.W. and Huttner, W.B. (1997) Cooperativity of phosphatidylinositol transfer protein and phospholipase D in secretory vesicle formation from the TGN–phosphoinositides as a common denominator? FEBS Lett , 419, 271–75. Jones, S.M., Howell, K.E., Henley, J.R., Cao, H. and McNiven, M.A. (1998) Role of dynamin in the formation of transport vesicles from the trans-Golgi network. Science, 279, 573–77. Abdulkarim, A.S., Cao, H., Huang, B. and McNiven, M.A. (2003) The large GTPase dynamin is required for hepatitis B virus protein secretion from hepatocytes. J Hepatol , 38, 76–83. Sai, Y., Nies, A.T. and Arias, I.M. (1999) Bile acid secretion and direct targeting of mdr1-green fluorescent protein from Golgi to the canalicular membrane in polarized WIF-B cells. J Cell Sci , 112 (Pt 24), 4535–45. Wallace, D.F., Summerville, L., Lusby, P.E. and Subramaniam, V.N. (2005) Prohepcidin localises to the Golgi compartment and secretory pathway in hepatocytes. J Hepatol , 43, 720–28. Dell’Angelica, E.C., Puertollano, R., Mullins, C., Aguilar, R.C., Vargas, J.D., Hartnell, L.M. and Bonifacino, J.S. (2000) GGAs: a family of ADP ribosylation factor-binding proteins related to adaptors and associated with the Golgi complex. J Cell Biol , 149, 81–94.

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Mitochondria Kasturi Mitra Cell Biology and Metabolism Program, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD, USA

INTRODUCTION The name “mitochondria” was given to cellular structures of varied morphology that were observed by nineteenth century morphologists; the nomenclature is a fusion of the Greek words “mito” meaning filaments and “chondro” meaning grains. These organelles are thought to have originated from aerobic bacteria that were engulfed by early eukaryotic cells and have maintained an endosymbiotic relationship with the host eukaryotic cells ever since. This idea, proposed as “the endosymbiotic theory” by Lynn Margulis [1], has been substantiated with various evidences and is widely accepted now. Mitochondria help the aerobic organisms by aiding the efficient conversion of energy trapped in food to its cellular form, ATP, in presence of oxygen. The abundance of mitochondria varies in different cell types, abundance being maximal in cells of aerobic tissues. Liver cells, being part of an aerobic tissue, have 20% of their cytoplasmic volume occupied by mitochondria. The mitochondrial story has been told by scientists from all arenas of biology. Biochemists in the early twentieth century directed their efforts in trying to understand the properties of these double-membrane-bound organelles and mitochondria emerged as the seat of energy production, hence the name “powerhouse of the cell.” Mitochondria were also identified as sites of intermediate metabolism, calcium regulation, and redox homeostasis in cells. Only after the middle of twentieth century did scientists begin to appreciate the importance of mitochondria in human diseases, the range of diseases being from metabolic to neurodegenerative to cancer, including hepatocellular carcinoma. Clinicians, along with geneticists, discovered and mapped a range of mitochondrial DNA mutations associated with diseases (Mitomap web The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

site, http://www.mitomap.org). Cell biologists and microscopists began taking interest in mitochondria with the advancement in both microscopy techniques and mitochondrial labeling techniques in live cells. Their contribution to mitochondrial biology helped us to appreciate the highly complex dynamic structure of mitochondria and that they are a regulatory part of the cell death machinery. Today, the mitochondrial story is taking different turns as we realize that there is much more to mitochondria than being a “powerhouse” of the cell. This chapter describes different aspects of mitochondrial biology that have been revealed to date. The role of mitochondria in energy metabolism, calcium regulation, apoptosis, aging, and cancer will be discussed. A special focus is provided on understanding the dynamic nature of mitochondria and the physiological importance of such dynamism, which is one of the most active current areas of mitochondrial research.

STRUCTURE Each cell harbors numerous mitochondria, which appear like bacteria and whose length can extend from 500 nm to 1 mm or more (in some specialized cells), the average diameter being 500 nm [2]. Mitochondria differ from other membrane-bound organelles in cells in that they are double-membrane bound and host circular DNA. The structure and composition of the outer membrane resembles a standard lipid bilayer, while that of the inner membrane is highly specialized. The inner membrane, which surrounds the mitochondrial matrix, is unique in protein and lipid composition. It is devoid of cholesterol and rich in a special lipid called cardiolipin, which is synthesized inside the mitochondria. Cardiolipin has been

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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found to be important for maintaining the organization and activity of the inner-membrane proteins [3]. The net protein-to-phospholipid ratio of the inner membrane is 3 : 1, which is several times higher than that of any other membrane bilayer in the cell. The ATP synthesis machinery, including the respiratory chain complexes and the uncoupling proteins (see next section), constitutes the major protein content of the inner mitochondrial membrane. Advanced electron microscopy on rat liver mitochondria [4] has revealed that the inner membrane that separates the matrix from the intermembrane space is organized in two distinct domains—the part that is juxtaposed to the outer membrane is called the inner boundary membrane, and the part that folds into the matrix is called the cristae (Figure 9.1, [5]). The cristae can be tubular or lamellar and the points of origin of the cristae from the inner boundary membrane are called cristae junctions. It is proposed that the cristae organization and the size of the cristae junctions can be controlled by certain inner-membrane proteins [6] and that their organization can also alter mobility of proteins in the intermembrane space [7]. The respiratory chain complexes (Oxidative Phosphorylation OXPHOS) and ATP synthase machinery have been found to exist in the inner membrane in the form of supercomplexes, presumably to enhance the efficiency of sequential transfer of electrons from one complex to the other, which is key for ATP generation [8]. Thirteen of the 90 proteins constituting the respiratory chain complexes are coded by the mitochondrial DNA that resides within the mitochon-

drial matrix. The ∼16 kb circular mitochondrial DNA also codes for 2 ribosomal and 22 transfer RNAs. Each somatic cell contains from 1000 to 10,000 copies of the mitochondrial DNA and about 2 to 10 mtDNA molecules, along with a set of ∼30 proteins to form organized structures called nucleoids. This set of proteins includes mitochondrial replication factor (polymerase gamma), transcription factor (TFAM), single-strand binding (SSB) protein, mitochondria DNA packaging factors (Abf2), and proteins of yet unknown functions. Mitochondrial DNA replication and transcription are coupled to one another and are highly regulated (see [9] for review). The transcribed mRNAs are translated by the ribosomes in the mitochondrial matrix and inserted into the inner membrane cotranslationally.

ENERGY PRODUCTION: ROS UNCOUPLING Mitochondria are capable of producing 30 molecules of ATP from a single molecule of glucose, which gets oxidized through glycolysis in cytosol and then through the Tricarboxylic Acid (TCA Cycle) in mitochondria. These pathways generate reducing equivalents in the form of NADH molecules, which undergo oxidative phosphorylation to ultimately generate ATP. This is brought about by the concerted action of the OXPHOS complexes in the

Crista junction diameter 28 nm Outer–inner membrane 20 nm

Crista width 27 nm

Contact site diameter 14 nm

Contact width 14 nm T/BS

Figure 9.1 Ultrastructure of mitochondria. 3D tomogram of mitochondria of chick cerebellum. Specific regions are shown in higher zoom. The dimensions are average values of mitochondria from neural tissue, brown adipose tissue, and Neurospora crassa. Reproduced with permission from [5], Copyright  2000, with permission from Elsevier

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inner membrane, pumping electrons derived from NADH from the matrix to the intermembrane space. This action creates a pH gradient and an electrical gradient across the membrane, the summation of these gradients being referred to as the transmembrane potential of mitochondria. The proton motive force generated by the downgradient flow of hydrogen ions from the intermembrane space to the matrix through the ATP synthase complex is utilized by this same protein complex to add the high-energy phosphate bond to ADP to form ATP (for detailed understanding, see review in [10]). An intact transmembrane potential is not only required for the driving of ATP synthesis but for myriad other mitochondrial functions such as calcium homeostasis, protein import, redox balance, and so on. The OXPHOS machinery constitutes several redox centers that might leak electrons from the mitochondria, which when they react with molecular oxygen generate superoxide radicals. About 2% of the molecular oxygen consumed during respiration is converted into superoxide radicals, which are precursors of other oxidative radicals generated inside cells that are collectively called reactive oxygen species (ROS). Manganese superoxide dismutase, which localizes in the mitochondria, is responsible for scavenging the ROS generated in mitochondria, lack of which causes neonatal lethality [11]. Any dysbalance in the ROS generation and scavenging system is potentially capable of causing oxidative stress. Alcoholic and non-alcoholic liver diseases have been linked to oxidative stress leading to lipid peroxidation and rendering hepatocytes susceptible to cell death. Depletion of reduced glutathione, which is also a part of the ROS scavenging system, or increased production of NADH could potentially be the cause of oxidative damage in chronic alcoholism [12]. Certain ion carrier proteins of the inner mitochondrial membrane, called uncoupling proteins, allow flow of proton from the intermembrane space to the matrix, thus reducing the protonmotive force to synthesize ATP. Several knockout and knockdown studies show that uncoupling proteins play an important role in controlling ROS production in several organs, including liver [13].

ORIGIN AND INHERITANCE OF MITOCHONDRIA The endosymbiotic theory on the origin of mitochondria from a prokaryote is supported by a number of pieces of evidence, mainly those which demonstrate the similarity of mitochondria to prokaryotes. Mitochondrial genome sequencing, which was the first successful endeavor [14] in the genomic era, revealed that mitochondrial DNA is closest to that of an alpha proteobacteria [15]. Signatures of mitochondrial genome have also been detected in nuclear DNA [16, 17], which led to the proposition that in the course of evolution the endosymbiont exchanged

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genetic material with the chromosomal DNA [18]. Unlike chromosomal DNA, mitochondrial DNA follows a non-Mendelian inheritance as the oocyte is the major contributor of mitochondria in the zygote. Mitochondrial genome has been found to be rapidly evolving and therefore has been successfully used as a tool for following the evolution of organisms. Variation in mitochondrial DNA sequences has led to the identification of different haplogroups in different ethnic populations, the most ancient and diverse being that of the Africans. Analysis of such ethnic haplogroups reveals migration of human mitochondrial DNA from Africa to the rest of the modern world ([19], http://www.mitomap.org). Another feature of mitochondrial genome is heteroplasmy—that is, all copies of the polyploid mitochondrial genome may not be identical. This concept is critical for the analysis of diseases related to mitochondrial DNA mutations and their inheritance. Liver failure is one of the common features in children affected with such diseases. In heteroplasmic mutation a threshold level of mutation is critical for biochemical defects and clinical expression of diseases. Mosaic expression of cytochrome c oxidase activity in patients harboring a mitochondrial tRNA point mutation exemplifies the heteroplasmic situation [20]. However, most patients with Leber hereditary optic neuropathy harbor homoplasmic mitochondrial DNA mutations [21]. Somatic mitochondrial DNA mutations have also been detected in a wide range of tumors [22] and in normal tissues during aging [23]. Although a causative connection has not been drawn between mitochondrial DNA mutations and diseases, serious efforts are being made to that end. For example, a particular mutation in NADH dehydrogenase subunit 6 has been shown to induce metastasis in certain tumors [24]. These kinds of study are accomplished using the “cybrid” or cytoplasmic-hybrid technology, which involves generating cells depleted of mitochondrial DNA (rho0 ) and repopulating them with a known mitochondrial DNA mutation in order to understand the physiological effects of that mutation. rho0 cells are generated in the laboratory by growing cells in the presence of agents that indirectly or directly inhibit mitochondrial DNA replication (such as ethidium bromide, rhodamine 6G, dominant negative mutant of polymerase gamma, or knocking down mitochondrial transcription). This leads to complete depletion of mitochondrial DNA, forming mitochondria ghosts without DNA called mitoids. Mitoids do not produce mitochondrial ATP but cells harboring them (rho0 ) survive and proliferate when supplemented with pyruvate and uridine [25]. The rho0 cells act as acceptors as they are fused with enucleated donor cells (cytoplasts) harboring mutated mitochondrial DNA, generating a cybrid wherein cytoplasmic mixing of both the cells will cause repopulation of the mitoids with mutated DNA. Although functional complementation takes place in the cybrid, mitochondrial nucleoids of the donor and acceptor may not physically mix with each

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other [26]. Cybrids thus generated have been used to assess the effects of mitochondrial DNA mutations on mitochondrial and cellular physiology.

APOPTOSIS Apoptosis or programed cell death is an energy-dependent regulated process of cellular demise characterized by cell shrinkage, nuclear DNA condensation, and generation of apoptotic bodies. This directed cell death is important for developmental processes as well as many diseases. Enhanced apoptosis with loss of regeneration is a characteristic feature of most liver disorders. More than two decades of research have unraveled the controlled signaling pathway involved in apoptosis, demonstrating mitochondria as a central regulatory component of the signaling cascade. However, certain apoptotic triggers (e.g. Fas ligand or tumor necrosis factor) can bypass the mitochondrial pathway. The key step toward apoptosis is to trigger the proteases called “caspases”, which brings about degradation of cellular machineries in a regulated fashion (see [27] for review). In this regulatory event, mitochondria contribute by sequestering factors responsible for activating the executionary caspases, in particular caspase 3. These factors, including cytochrome c, DIABLO (also SMAC or Second Mitochondria Derived Activator of Caspase, Apoptosis Inducing Factor (AIF), endonuclease G, and so on, normally reside within the intermembrane space of mitochondria and are released into the cytosol upon apoptotic trigger. The role of cytochrome C, which is a crucial component of the electron transfer chain within mitochondria, is the most well characterized of all the proteins in this category. Although it is well appreciated that release of cytochrome C into the cytosol from mitochondria is a highly regulated process, the exact molecular mechanism is shrouded with controversies. However, it is accepted that certain pores are formed in the outer mitochondrial membrane through which cytochrome C and other proteins escape into the cytosol. Therefore, pore formation is an essential step in apoptosis and is called mitochondrial outer membrane permeabilization [28]. The pores are thought to be formed by proteins such as BAX and BAK, which either reside on the mitochondrial membrane or are recruited to the mitochondrial membrane during apoptosis [29, 30]. These proteins belong to the Bcl2 family of proteins, which consists of both pro- and antiapototic proteins. BAX and BAK belong to the former class while Bcl2 and Bcl-XL belong to the latter. The Bcl2 protein was discovered as an oncogene and it was later realized that the antiapoptotic function of the protein [31] that might promote tumor growth. This protein has several Bcl2 homology domains (BH1–4) and proteins sharing one or more homologous domains are grouped in the Bcl2 family. A class of protein involved in apoptosis shares homology to Bcl2 only in the BH3 domain

and is called BH3-only protein. For a detailed list of the Bcl2 family of proteins and their functionalities, please refer to [32] . In brief, the proteins of the Bcl2 family interact with each other through these domains and can also influence each other’s activity. For example, the proapoptotic activity of the BH3-only proteins might be brought about when they interact with antiapoptotic proteins, thus liberating BAX and BAK to form pores in the outer mitochondrial membrane to cause permeabilization. Mitochondrial function, particularly transmembrane potential, decreases during apoptosis. Although it appears intuitive that release of cytochrome c, an integral part of the OXPHOS machinery, should be associated with loss of transmembrane potential, it has been demonstrated otherwise. A fraction of cytochrome c, presumably one that is not functionally involved in the OXPHOS machinery, is released from mitochondria maintaining transmembrane potential after an apoptotic trigger [33]. The trigger of cytochrome c release is all or nothing; that is, once cytochrome c leaks out from one mitochondrion, all other mitochondria quickly follow and complete release of cytochrome is accomplished within five minutes [34]. This release is followed by mitochondrial depolarization, an event possibly linked to caspase activation [35].

CROSSTALK WITH OTHER ORGANELLES Recently cell biologists have come to appreciate that mitochondria function by interacting with other organelles both physically and functionally. Interaction with nucleus appears intuitive because most of the genes coding for mitochondrial proteins reside in the nuclear genome. Alterations in mitochondrial physiology alter nuclear transcription of mitochondrial genes (and other genes involved in metabolic regulation), which are transcribed in the nucleus. The other organelle that is well studied with respect to mitochondrial interactions is the endoplasmic reticulum (ER). The driving force behind these studies was the realization that both mitochondria and ER act as calcium stores in cells. This nascent field of inter-organellar cross talk presents interesting clues that could explain the various complexities of cellular machineries.

Nucleus Of all the mitochondrial proteomes identified thus far, only 13 are coded by the mitochondrial genome. In contrast, 1500 mitochondrial proteins are coded by the nuclear genome, including proteins involved in mitochondrial DNA replication, transcription, and translation. It was first discovered in yeast that nuclear gene expression can be altered by changing the mitochondrial genotype [36]. Later studies in mammalian cells using rho0 cells

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and mitochondrial uncouplers confirmed that downregulation of mitochondrial function (i.e. mitochondrial stress) affects the expression profile of the nuclear genome to compensate for the loss of mitochondrial function [37]. This process is termed “retrograde regulation” in response to mitochondrial stress. The most obvious response is to increase the mitochondrial mass that is prompted by PGC1-alpha [38]. PGC1-alpha is a transcriptional coactivator which, when overexpressed, induces synthesis of molecules involved in oxidative phosphorylation, mitochondrial biogenesis, fatty acid oxidation, gluconeogenesis, and so on. Induction in oxidative phosphorylation and mitochondrial biogenesis is brought about by PGC1-alpha’s induction of transcription factors called nuclear respiratory factors (NRFs) 1 and 2. NRF1 and 2 induce mitochondrial DNA replication and transcription along with transcription of one or more OXPHOS subunits. An important sensor of mitochondrial bioenergetic status is AMP-kinase (AMPK). Loss of mitochondrial (and glycolytic) ability to convert ADP to ATP increases the conversion of ADP to AMP, which activates AMPK. AMPK activity is essential for inducing PGC1-alpha and mitochondrial biogenesis [39].

Endoplasmic reticulum Calcium is a second messenger in signaling cascades, and a stringent regulation of intracellular calcium is a prerequisite in intracellular signaling . In hepatocytes, epinephrine and non-epinephrine change calcium levels in cells [40]. The ER and mitochondria have a functional nexus through which the intracellular calcium pool is regulated. Cytosolic calcium can range from a very low 10 nM basal level to 2 µM in conditions that stimulate calcium release from the ER or the plasma membrane. Calcium from the ER is released through specialized channels (Inositol 3 Phosphate Receptors, IP3Rs) and then taken up by mitochondria to maintain calcium homeostasis. Upon stimulation, IP3R molecules, which are homogeneously present in the ER, redistribute into patches [41] that could be the zones of close vicinity with the mitochondria. The ER and mitochondria physically interact with each other to maintain calcium homeostasis. More than 20% of the mitochondrial surface is in very close vicinity to the ER and it has been postulated that these interacting zones are specialized in their molecular composition [42]. The two organelles are sometimes tethered to each other by 10 nm proteinaceous bridges, which may modulate functional interaction between the organelles [43]. The molecular chaperone GRP75 also participates in forming functional connections between the Voltage Dependent Anion Channel-1 (VDAC) channel on mitochondria and the IP3R channel on the ER [44]. Although it is not clear which set of molecules on each of the two organelles is directly involved in calcium exit and entry, evidence suggests the presence of uniporters on mitochondria

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that are specific for calcium. Biochemical fractionation of mitochondria also includes some ER membranes that can be further isolated from the mitochondrial fractions. This particular interacting zone between mitochondria and ER is called MAM (mitochondria-associated (ER) membrane). Detailed studies reveal that MAM is rich in lipid-modifying enzymes; phosphatidyl serine synthases in particular [45]. It has been hypothesized that phosphatidyl serine is synthesized in ER membranes and transferred through MAM to mitochondria, where phosphatidyl serine is converted to phosphatidyl ethanolamine. Sigma-1 receptors on MAM function in neuroprotection by modulating calcium exchange between the ER and mitochondria [46]. Another potential group of candidates for modulating ER–mitochondria interactions are the factors modulating mitochondrial dynamism (see below). Mitochondria exist in an equilibrium between various shapes and forms. Molecules affecting the dynamic equilibrium of mitochondrial shapes may affect calcium exchange with the ER, particularly in cell death.

DISTRIBUTION The presence of mitochondria in a certain part of the cell may reflect the energy demand for that part of the cell. Presence of mitochondria may also reflect other mitochondrial activities such as calcium regulation, redox homeostasis, lipid modification, and so on. Therefore, distribution of mitochondria to different cellular parts in a regulated fashion appears to be critical for the proper function of cells. Proper distribution of mitochondria can be achieved by two inherent properties of the organelles: (i) motility: mitochondria exhibit a wide range of motilities in cells; and (ii) dynamism: mitochondria change their shape and form in cells. Cytoskeletal elements, particularly microtubules, play a central role in maintaining proper mitochondrial distribution in mammalian cells. Alteration of microtubule structure affects mitochondrial motility, shape, and dynamism. Recent data suggest that microtubule depolymerization may alter mitochondrial oxidative capacity [47]. In neurons, movement of mitochondria is crucial as they need to travel down the axons to the synapses. In axons, mitochondrial motility may be anterograde or retrograde and could be up to 1 µm s−1 in velocity [48]. Kinesin and dynein motors on the microtubules are involved in moving mitochondria on these cytoskeletal structures. Syntabulin, which interacts with kinesin 1, mediates anterograde movement of mitochondria in neuronal processes [49]. Milton, a kinesin associated protein, and miro, a rhoGTPase on the outer mitochondrial membrane, serve as another important motor complex for anterograde mitochondrial movement in neurons [50]. Keeping mitochondria stationary in neuronal processes by tethering them to microtubules is important for neuronal activity, such as short-term facilitation [51].

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The dynamism of mitochondria is maintained by the activity of proteins, which aid mitochondrial fission and fusion. These molecules can either cause fission of one mitochondrion into two, or fuse two mitochondria residing next to each other. Fission and fusion of mitochondria happen incessantly inside cells. Most of the fission/fusion proteins, discovered thus far, have GTPase activity, inhibition of which blocks fission/fusion activity. Interestingly, addition of fusogenic mitochondrial outer-membrane fraction to permeabilized hepatocytes induces mitochondrial tubulation [52]. Inhibition of fission activity or downregulation of the fission proteins causes mitochondrial tabulation, whereas fusion proteins cause mitochondrial fragmentation [53]. Therefore, it has been postulated that net balance between the fission/fusion events dictates mitochondrial distribution/morphology in cells; normal mitochondrial distribution is intermediate between fully fused and fully fragmented forms (Figure 9.2). Many efforts are being made to understand how fission/fusion molecules are orchestrated on the mitochondrial surface. Because mitochondria are double-membrane-bound organelles, regulated fission/fusion events of both membranes requires a higher degree of control than the fusion of two membrane bilayers. Although some level of insight has been obtained into independent fusion and fission of the membranes, how the fission/fusion events of the two mitochondrial membranes are coupled is not completely understood. The most well characterized outer membrane fission element is DRP1, a protein recruited from cytosol to the outer mitochondrial membranes [54, 55]. The fission brought about by DRP1 is usually dependent on another outer-membrane protein, called Fis1, which might act as a receptor for recruiting DRP1 on the mitochondrial outer membrane [56].

Fission

Fusion

However, Fis1-independent recruitment of DRP1 to mitochondria has also been observed [57]. It is not fully understood how DRP1 brings about mitochondrial fission. Not all of the sites on mitochondria where DRP1 is recruited develop into fission sites. The GTPase activity of this protein has been associated with its property of forming multimeric ring structures around mitochondria [58]. The mechanochemical activity of these ring structures is thought to induce fission of the mitochondrial membranes. Interestingly, no protein has been found to be potentially a fission protein for the inner mitochondrial membrane. In contrast, fusion proteins have been found in both outer and inner mitochondrial membranes. Thus far, Mitofusin (MFN) 1 and 2 and OPA1 have been characterized as the fusion proteins for the outer and inner mitochondrial membrane, respectively. MFN1/2 are localized on the outer mitochondrial membrane, such that the major portion of each molecule hangs out of the mitochondrial membrane toward the cytosolic side. Data from MFN1/2’s structure analyses suggest that specific domains of MFN1/2 on two apposing surfaces of mitochondria interact with one another to form dimmers, which tether these apposing mitochondria together [59]. However, how actual fusion occurs between the tethered mitochondrial outer membranes is not known. For proper fusion of the mitochondrial inner membranes, an intact transmembrane potential influences the processing of OPA1 in the inner membrane [60, 61]. OPA1 is coded by a set of eight differentially spliced forms and each mRNA is further processed into a long and a short form. At least three mitochondrial proteases, namely paraplegin, Yme1L, and PARL, have been implicated in the complicated processing of OPA1. The functions of each

mito

chondro

Fission

Fusion

Fission

Fusion

Figure 9.2 Mitochondrial morphology is maintained by a balance of fission and fusion. The panel in the center shows normal mitochondrial morphology observed in live mammalian cells, which consists of some “mito” and some “chondro.” The panel on the right shows fragmented morphology, which is a result of predominant fission activity, while that on the left shows fused mitochondria resulting from predominant fusion

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splice variant remain to be investigated but the regulation of each isoform may be important for regulated fusion of the inner mitochondrial membrane [62]. The mechanism of mitochondrial fission/fusion is an active area of research and the list of proteins involved in mitochondrial fission/fusion is growing annually. Another area researchers are currently focusing on is unveiling the physiological relevance of mitochondrial dynamism. Two of the fission/fusion proteins, namely OPA1 and MFN1, were identified as mutated genes in specific neurodegenerative diseases. Mitochondrial fusion has been postulated to facilitate mitochondrial DNA exchange and complementation. Lack of this function could give rise to defects observed in neurodegenerative diseases caused by mutation of MFN1 [63]. Knockdown or knockout experimental models have been used to investigate the role of mitochondrial fission/fusion, and in each case lack of mitochondrial fission/fusion left a deleterious effect [2]. Although mitochondria undergo fragmentation during apoptosis, the causative role of fragmentation in apoptosis is debated. Apoptosis induced by certain drugs is delayed when the activity of DRP1 is inhibited [54], suggesting a role of mitochondrial fission in apoptosis. This finding has been confirmed by drug-induced inhibition of DRP1 activity that protects cells from apoptosis by inhibiting outer-membrane permeabilization [64]. However, the mitochondrial inner-membrane protein OPA1 protects cells from apoptosis by mitochondrial cristae reorganization [6]. Contrarily, induction of mitochondrial fission by overexpressing DRP1 protects against ceramide-induced apoptosis [65]. Fragmented mitochondria presumably may be weaker in responding to calcium signaling from the ER, which is key for ceramide-induced apoptosis. Interestingly, BAX and BAK also interact with mitofusins on the mitochondrial membrane, which may be important for regulating mitochondrial morphology during apoptosis [66, 67]. Further research needs to be concentrated on the association between mitochondrial fission/fusion apparatus and the cell death machinery. Inhibition of mitochondrial fission delays ROS production in hyperglycemia [68]. Mitochondria undergo fission during mitosis where phosphorylation at a specific site of DRP1 increases activity of the fission protein [69]. Phosphorylation at another site of the same protein however has been found to reduce the activity of the protein [70]. All these data suggest that the mitochondrial fission/fusion proteins have to be under tight regulation in different physiological conditions.

CANCER AND AGING Aging and cancer can be considered as opposite sides of the same coin. During aging, proliferating cells become senescent, whereas in cancer, certain cells acquire growth and proliferation advantage. Alteration of mitochondrial physiology has been implicated in both cancer and aging,

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and some common sets of molecules are involved in these two apparently diametrically opposite phenomena [71].

Aging The free-radical theory proposed by Denham Harman suggests that aging occurs because of cumulative damage caused by free radicals over a long period of time [23]. As mitochondria are the main source of free radicals in cells, this theory of aging implicates mitochondria’s causative role in aging. Reactive oxygen species generated by respiring mitochondria can also lead to damage of mitochondrial DNA, and indeed ROS-mediated oxidation products, like 8-hydroxy-2-deoxyguanosine, are prevalent in aged tissues [72]. Interestingly, targeted mutation in mitochondrial DNA polymerase increases mutations in mitochondrial DNA and such “mutator” mice have reduced life spans [73, 74]. Overexpression of catalase and super oxide dismutase, which will scavenge ROS, extends life span in flies [75]. Moreover, increased life span has been observed in mice engineered to overexpress human catalase in mitochondria [76]. However, direct involvement of ROS in mitochondrial DNA damage is still debated. Recent studies have implicated a component of the nutrient-sensing machinery, mTOR, in extending life span by altering mitochondrial respiration [77].

Cancer More than two decades ago, Otto Warburg proposed that cancer cells originate when mitochondrial respiration is downregulated and cells become dependent on glycolytic ATP [78]. Some of his predictions have been proven correct as researchers have unraveled the molecular basis of tumorigenesis. HIF1-alpha, a key molecule induced during hypoxia, upregulates glycolysis and downregulates mitochondrial respiration [79, 80]. Restoration of mitochondrial ATP production by activating key mitochondrial metabolic enzymes, such as PDH, may provide a target for anticancer therapeutic strategies in certain cases [81]. The classic tumor-suppressor p53 induces mitochondrial respiration through SCO2 [82] and reduces glycolysis through TIGAR [83]. Thus, the tumor-suppression action of p53 involves both upregulation of mitochondrial respiration and downregulation of glycolysis. Mitochondria in tumor cells are reprogramed to provide apoptotic resistance to the tumors. This alteration in metabolism in cancer cells may also occur in highly proliferating normal cells [84]. In this context, the emerging role of mitochondria is intriguing. A particular OXPHOS mutant that causes reduction in mitochondrial ATP is specifically blocked before DNA synthesis via downregulation of the relevant cyclin (cyclin E) [85]. This phase in the cell cycle holds the key to the controls

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that keep cells dependent on growth factors. Mutations in many regulatory molecules at this phase render cells independent of growth factor regulation. It is tempting to speculate that mitochondria might have a regulatory role in this critical phase of the cell cycle, which may be the root cause of the metabolic alteration observed in many cancers. Research in this direction might not only help our understanding of hepatocellular carcinoma but could also aid in our understanding of liver regeneration, which involves induction of cell proliferation and growth [86].

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10

Nuclear Pore Complex Joseph S. Glavy Department of Chemistry, Chemical Biology & Biomedical Engineering, Stevens Institute of Technology, Hoboken, NJ, USA

INTRODUCTION In eukaryotic cells, the separation of nuclear DNA from cytoplasmic organelles and cellular materials is governed by the nuclear envelope (NE). The NE is defined as a specialized endoplasmic reticulum membrane with a double bilayer containing an inner and outer membrane system. This partitioning is needed to organize nuclear entry and exit for basic cellular processes (Figure 10.1). Nuclear pore complexes (NPCs), the gateways for macromolecular traffic into and out of the nucleus, are set in circular openings of the double membrane of the NE [1, 2]. There are highly regulated pathways that control nuclear entry and exit of molecules such as transcription factors, RNAs, kinases, and viral particles. In broad spectrum, to be imported or exported from the nucleus, molecules (i) bind to transport receptors, (ii) are transported through NPCs present in the NE, and (iii) translocate from the NPCs to intranuclear or cytoplasmic target sites [1, 2]. NPCs are composed of proteins termed nucleoporins or Nups, which play a role in the structure of the NPC and in regulating the translocation of molecules through the NPC [1, 2]. Chromosomal translocations involving genes encoding Nups have been associated with many forms of leukemia [3]. In addition, Nups have been associated with primary biliary cirrhosis (PBC), viral infection, atrial fibrillation, and triple A syndrome [3].

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

THE STRUCTURE OF THE NUCLEAR PORE COMPLEX (NPC) The NPC is a large macromolecular assembly with an estimated size of 60 and 40 MDa in vertebrates and yeast, respectively (Figure 10.1). NPCs are composed of approximately 30 proteins in both yeast and higher eukaryotes [4, 5]. Cryoelectron tomography reconstructions have revealed the NPC as a cylindrical structure that consists of a central core with an eightfold rotational symmetry about a nucleocytoplasmic axis and twofold symmetry in the plane of the NE, with an outer diameter of 125 nm and an inner diameter of 60 nm (Figure 10.2) [6, 7]. Overall NPC architecture is conserved between yeast and higher eukaryotes. The framework of the NPC (Figure 10.2) can be divided into three subdomains: the nuclear basket (containing the nuclear ring (NR), nuclear filaments (NFs), and distal ring (DR)), the central core spoke ring (SR), and the cytoplasmic ring (CR). State-of-the-art cryoelectron tomography with three-dimensional reconstruction gives a resolution between 8.3 and 5.8 nm (Figure 10.2) [7]. The nuclear basket (NR, NF, DR) extends nearly 60 nm into the nucleus (Figure 10.2) [7]. The SR is estimated at less than 50 nm, while the envelope is approximately 50 nm [7]. The length of the CR is difficult to define given the

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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linked or scaffold Nups and phenylalanine-glycine flexible repeat unit (FG) Nups that are involved in transport [2]. The inner and outer rings help to facilitate membrane structure (Figure 10.2). The complete architecture of the NPC includes some non-Nup proteins: the nuclear membrane proteins, translocated promoter region protein (Tpr), and the nuclear lamina [11]. These non-Nup proteins make up the surrounding regions of the NPC and support its transport function. (a)

(b)

Figure 10.1 (a) Thin-section electron micrograph illustrating the NPC in isolated HeLa cell NE. (Courtesy of Dr. Samuel Dales.) (b) Artist’s rendition of an NPC. (Courtesy of Na-Young Kim.) 125 nm 60 nm 50 nm

CR SR

ONM LC INM NR

60 nm

NF

DR

Figure 10.2 Cross-section of the NPC from the cytoplasm (top) to the nucleoplasm (bottom); structure is based on cryoelectron tomography. The curvature of the NE is shown: outer nuclear membrane (ONM), luminal curve (LC), and inner nuclear membrane (INM). The NPC is labeled: cytoplasmic ring (CR), central core spoke ring (SR), and distal ring (DR). The structure includes the nuclear basket: nuclear ring (NR), nuclear filaments (NF), and distal ring (DR). (Courtesy of Martin Beck.)

flexibility of the overall domain. This structure shows that the cytoplasmic filaments adopt a highly “kinked or bent” structure [6–8]. The CR is believed to act as a docking site for protein transport [7]. In both yeast and higher eukaryotes, the nearly 30 proteins composing the NPC are called Nups. Names of Nups are designed mainly on molecular weight (Plate 10.1). Proteomic studies in yeast and metazoa have completely cataloged the Nups of the NPC [4, 5]. A recent novel approach using a combination of proteomic data and a computational platform has been applied to the architectures of the macromolecular assemblies of the NPC [9, 10]. The study concludes that the fundamental symmetry unit of the NPC is the spoke [9, 10]. A pronounced 16-fold repetition of Nup columns is seen; each spoke consists of nuclear and cytoplasmic halves. Furthermore, the NPC is divided into three rings: outer, inner, and membrane rings [9, 10]. Within the ringed structure are

NUCLEOPORINS (Nups) Nups are organized into macromolecular assemblies called subcomplexes (Plate 10.1). Nup subcomplexes release from the NPC during open mitosis and are the disassembly units of the NPC [12]. In the laboratory, interphase subcomplexes can be isolated through biochemical extraction of the NE with low-percentage non-ionic or zwitterionic detergent treatment (such as 2% Triton X-100 and/or 1% CHAPS) [13–15]. These modular units are present in multiple copies arranged around two- and eightfold axes of symmetry and are believed to generate discrete structures within the NPC [11]. Furthermore, Nups can be classified into two overall groups: scaffold Nups and FG Nups [2, 16], plus a mobile Nup (Nup98). Scaffold Nups have structural designs to interlock (coiled-coils, β-propellers, or disordered areas) while FG Nups contain flexible repeating units of phenylalanine-glycine (forming sea anemone-like fingers within the NPC that are implicated in active nuclear transport) [2, 11, 16]. Plate 10.1 shows the overall arrangement of subcomplexes and unique (non-subcomplex) Nups within the NPC.

Subclasses of nucleoporins: scaffold Nups, FG Nups, and mobile Nup Scaffold Nups Scaffold Nups are contained within the Nup107-160 and Nup93 subcomplexes. The Nup107-160 subcomplex has nine members: Nup160, Nup133, Nup107, Nup96, Nup75, Nup43, Nup37, Seh1, and Sec13 (Plate 10.1) [15, 17–19]. This subcomplex has been classified as a keystone of NPC assembly [20]. RNAi knockdown experiments of individual members within the Nup107-160 subcomplex affect select members plus some other FG Nups [14, 20]. These findings show the interdependence of subcomplex members and the overall NPC. Positioned at the curvatures of the membrane embedding the NPC, the Nup107-160 subcomplex acts to stabilize these bends in the membrane (Plate 10.1) [21]. In the “protocoatomer”

10: NUCLEAR PORE COMPLEX

hypothesis, components of the Nup107-160 subcomplex serve as membrane-curving modules similar to the members of the COPI, COPII, and clathrin complexes [22]. One member of the Nup107-160 subcomplex, Sec13, is also found as a key member of the COPII complex. Furthermore, ArfGAP1 lipid packing sensor (ALPS) domains are found within members of the subcomplex [9, 23]. ALPS domains are classified as membrane-binding amphipathic alpha-helix regions. Nup133 contains an ALPS domain shown to bind to isolated membrane [23]. Additional Nups including Nup107, Nup160, and Nup188 contain predicted ALPS domains [9]. Reconstitution experiments showed the Nup107-160 subcomplex assembled into a Y-shape visualized with negative staining electron microscopy [24]. The structural modules of the subcomplex are a collection of alpha-helical repeats and β-propellers. X-ray crystal structures have been reported for several members. Analysis of Nup133 revealed a seven-bladed β-propeller domain of the N-terminus [25]. The crystal structure of the alpha-helical interaction of Nup107-133 shows an elongated structure forming a compact interface in a tail-to-tail fashion [25]. This interaction is a critical attachment point for Nup133 [25]. Another crystal structure of Nup96-Sec13 forms a hetero-octamer [26]. The N-terminal of Nup96 invades the six-bladed β-propeller structure of Sec13, providing a seventh blade [26]. The remaining portion of Nup96 forms an anti-parallel alpha-helical domain. The potential interlocking modules of the subcomplex may be the meshwork of the overall macromolecule. The potential coating modules may form a cylinder layer that apposes the membrane [26]. Analysis of the Nup75-Seh1 X-ray structure shows that the scaffold Nups form a membrane-bordering lattice, providing attachment sites for additional Nups such as FG Nups, Nup98, and Nup155 [27]. The Nup93 subcomplex includes five members: Nup93, Nup205, Nup188, Nup155, and Nup35 [28]. Nup93 is a highly abundant protein with 32–48 or even more copies within the NPC [11]. The Nup93 subcomplex aids in inner-ring stabilization [29] and is needed for correct nuclear pore assembly and homeostasis of the NPC [29]. RNAi experiments suggest a functional link between NE transmembrane NDC1, Nup93, and Nup205. These results suggest an anchor point function for the Nup93 subcomplex [30]. Furthermore, Nup62 (FG Nup) has been shown to interact with Nup93, illustrating interdependence between inner-ring Nups and transport FG Nups [30]. The X-ray crystal structure of Nup93 reveals that the protein folds into an elongated, alpha-helical structure. This fold is evolutionarily conserved and therefore functionally maintained [29]. Members of the Nup93 subcomplex contain mainly alpha-helical domains. The Nup93 alpha-helical structure may be the centerpiece of the entire subcomplex [29].

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FG Nups FG Nups have stretches of FG (Phe-Gly) repetitive residues, which are separated by polar spacer regions of variable lengths [2, 16]. FG repeat domains form unstructured regions that produce weak interactions with transporting proteins called karyopherins (kaps) [2]. The CR is composed largely of FG Nups [6, 7]. Specifically, Nup358 and Nup214 (with its binding partner Nup88) are involved in the transport and selectivity of the NPC [2]. Another grouping of FG Nups is the Nup62 subcomplex, located in the inner pore region or SR [6, 7]. The Nup62 subcomplex includes Nup62, Nup58, Nup45, and Nup54 (Figure 10.2 and Plate 10.1) [11, 31]. This subcomplex is sometimes referred to as the central plug region of the NPC. While these transport Nups line the inner NPC, it is likely that they do not form a plug against transport, but rather a dynamic transport area of the complex (Figure 10.2). The FG repeat domains form a tentacle-like structure that emanates from and lines the channel of the pore [2, 32] (Figure 10.1b). Other FG Nups, Nup153 and Nup50 along with Tpr, make up the nuclear basket (NR, NF, DR) from Figure 10.2. Tpr is not classified as a Nup but rather as a filament protein that helps give support to the nuclear basket (Plate 10.1). Nup153 and Nup50 are involved in selective transport and release of substrates [2]. The FG regions remain a difficult domain for crystallization. Their floppy tentacle nature is not conducive to nucleation and therefore crystal formation. Thus far only non-FG regions of these Nups have been reported. The X-ray crystal structure of the non-FG repeat N-terminal of Nup214 reveals a seven-bladed β-propeller with a segment of its C-terminus bound to the propeller [33]. Furthermore, X-ray analysis of Nup58 and Nup45 revealed a possible circumferential sliding mechanism that adjusts the diameter of the central transport channel [31]. The alpha-helical region forms a distinct tetramer with a hydrophobic interface. The residues are laterally displaced in numerous tetramer conformations, giving the possibility of a sliding structure [31].

Mobile Nup: Nup98 Nup98 facilitates mRNA export from the nucleus [17, 34]. It is classified as a non-subcomplex Nup with multiple locations along the NPC (Plate 10.1) [17]. Nup98 arises from a Nup98-Nup96 precursor form that splits by a self-cleavage domain similar to those found in Drosophila Hedgehog and Flavobacterium glycosylasparaginase [17, 35]. The N-terminal of Nup98 contains FG repeats used as a docking site for kaps though it is not classified as FG but rather as a mobile Nup [17, 36]. Nup98 is found both at the NPC and within the nucleus [17]. One of Nup98’s interacting partners is Rae1; together they act as temporal

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regulators of the anaphase-promoting complex [37]. Also, a vesicular stomatitis virus (VSV) matrix protein binds Nup98 and inhibits nuclear export of mRNA [34].

NUCLEAR ENVELOPE AND MEMBRANE PROTEINS The NE is separated into three domains: the outer nuclear membrane (ONM), the pore membrane (POM), and the inner nuclear membrane (INM) [38]. NPCs are anchored to NE membrane proteins, NPC-anchoring membrane proteins including POM121, gp210, and NDC1, which are positioned at the curvatures of the NE (Plate 10.1, light brown) [38]. Elimination of the NPC membrane protein POM121 results in failure to assemble NPCs and formation of continuous nuclear membranes [39]. RNAi experiments suggest a functional link between NE membrane NDC1 and members of the subcomplex: Nup93 and Nup205 [40]. These results suggest an anchor point function for the Nup93 subcomplex [30]. The inner membrane of the NE contains a unique array of integral membrane proteins, including lamina-associated polypeptides 1 and 2 (LAP1 and LAP2), emerin, MAN1, nesprins-1 and -2, and lamin B receptor (LBR) [38]. These proteins contain an LEM domain, which is a 40-residue motif that interacts specifically with lamin A/C [41].

NUCLEAR LAMINA The shape of the NE is maintained by a meshwork of intermediate filaments of associated nuclear lamina [41–43]. This meshwork is 10–20 nm thick and composed of lamin A/C and lamin B [42, 43]. The lamina feature a coiled-coil structure flanked by non-helical head and tail domains. Interestingly, nuclear lamina are not found in lower eukaryotes like yeast [41]. Lamins have a scaffolding function for several nuclear processes such as transcription, chromatin organization, DNA replication, and maintenance of nuclear and cellular integrity. Lamins can bind directly to lamina-associated proteins (above LEM domain proteins) (LBR, LAP2, emerin, MAN1, nesprins-1 and -2) [38, 41]. Abnormalities within the structure or processing of the lamin A/C genes can lead to laminopathies that range from muscular disorders to premature aging (progeria) diseases [41].

NUCLEAR TRANSPORT CYCLE Molecules smaller than 40 kDa can passively diffuse across the NPC. With higher-molecular-weight proteins, an orchestrated receptor-mediated transport is needed to facilitate crossing into the nucleus (Figure 10.3) [2, 16, 44, 45]. Protein cargo recognition molecules, known as kaps,

bind to cargo in the cytosol and carry it into the nucleus, then in reverse they bind cargo in the nucleus and deliver it to the cytoplasm. Kaps recognize their cargo by binding short amino acid sequence segments of the protein [46]. For import, cargos have nuclear localization sequences (NLSs) that are classically rich in basic residues [2, 44, 46, 47]. For export, cargos have nuclear export sequences (NESs) that are classically leucine-rich [2, 44]. Masking or modifying an NLS or NES allows regulation of protein localization [47]. A majority of NPC traffic is achieved by coupling kap-cargo binding to a cycle of GTP-binding and hydrolysis conducted through the protein Ran [48]. Kaps deliver NLS-cargo and transport it through the NPC through weak transient interactions with FG Nups that extend into the NPC. As a small monomeric G-protein, Ran has a low inherent rate of GDP–GTP exchange and GTP hydrolysis [16, 44–46, 49]. It relies on interacting proteins to regulate its nucleotide state. Binding partners are separated according to their role in the cycle. Guanine nucleotide exchange factor (GEF) for Ran is found in the nucleus, while the GTPase-activating protein (GAP) for Ran is concentrated in the cytoplasm. As a result, nuclear Ran is in the GTP-bound form while the cytoplasm contains Ran predominantly in the GDP-bound form [44–46]. This established difference is then coupled to import through nucleotide-dependent binding of Ran to the kaps (Figure 10.3). For import, RanGTPs will dissociate their cargo, releasing it into the nucleus [49]. The kap–RanGTP complex can then travel back through the NPC to the cytoplasm, where RanGTP will be hydrolyzed and release the carry protein for another cycle with NLS-cargo binding and import [49]. Export occurs in the converse manner. Exportin β-kaps can bind when already bound to RanGTP [49]. Hydrolysis of the GTP dissociates the ternary complex in the cytoplasm and the kaps can travel back through the NPC for another cycle of export and can bind to an NLS-cargo for import. Figure 10.3 illustrates the process [44–46]. The key concept is the Ran gradient, with the concentration of RanGTP much higher in the nucleus. Therefore, once the complex has entered the nucleus, the high concentration allows for the disassembly and release of the components (Figure 10.3) [44]. INM proteins are thought to use a similar mechanism to explain their movement across the POM and eventual targeting [38, 50]. NLSs of the integral proteins signal their transports through the cargo-based transport system [38]. This mechanism implies an interaction between the integral protein and the FG Nups [50]. The kap plus its inner-membrane protein cargo would interact with FG Nups across or through scaffold Nup barriers. Fully understanding this inner-membrane transport represents a significant challenge. Furthermore, the targeted transport of POM membrane could occur through this cargo-based system. This may require a modification of the NLS or additional adaptor proteins to abort transport.

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151

6 α-kap

Importin β-kap

D D

T

1 T

2

5

NLS-Cargo

Cytoplasm

Nucleus T 4 T RanGTP

3

D RanGDP

T

T T Exportin β-kap

Figure 10.3 Karyopherin-dependent transport cycle. Key components for the cycle are RanGTP, NLS-cargo, kaps, and the NPC. Key concept is that the concentration of RanGTP is much higher in the nucleus. Steps: (1) Formation of the α-kap and importin β-kap transport complex. (2) α-Kap recognition of NLS-cargo with β-kap. (3) Selective transport through NPC. (4) Once inside, the high levels of nuclear RanGTP increase the likelihood of interaction and hence disassembly of the transport complex. (5) GTP bound β-kap is transported through the NPC to the cytosol, while α-kap forms a complex with GTP-bound exportin β-kap then exits by way of the near NPC. Left behind is the NLS-cargo. (6) Once back in the cytosol, GTP undergoes hydrolysis and kaps are recycled for the next round of transport. (Courtesy of Na-Young Kim.)

GENES, TRANSPORT, AND THE NPC Proposed by G¨unter Blobel, the “Gene Gating” hypothesis postulates the idea that transcribed genes are positioned in close vicinity to NPCs, thereby streamlining mRNA export [51]. The NPCs are envisioned to serve as gene-gating organelles capable of interacting specifically with expressing genes [51]. The non-random distribution of the NPC may mirror underlying genomic organization. Furthermore, it proposes that up to eight genes could be positioned due to the eightfold symmetry of the structure [51]. NPCs have been implicated in the sheltering of euchromatin regions from repressing factors [52–55]. Inside the nucleus, the nuclear basket interacts with proteins involved in trafficking of tRNA and mRNA [56, 57]. One example is the nuclear basket protein Nup2p, which has been shown to interact with the histone variant

H2AZ part of the euchromatin boundary formation and may position the euchromatin in closer proximity to the NPC [57–59]. Furthermore, studies combining peripheral positioning data with quantitative PCR of mRNA levels correlate with expression of some genes coupled to NPCs [60–62]. Also, studies have found DNA and Nups immunoprecipitated with NPC components in genome-wide analysis screening [63, 64]. However, it is not known if every gene is gated, so more extensive work is necessary in the future to fully prove this hypothesis [51, 54].

NUCLEAR ENVELOPE BREAKDOWN (NEBD) Cell division comprises two processes, namely mitosis and cytokinesis. Mitosis is the division of the genetic material into two equal halves while cytokinesis is the process by which the cell divides itself into two daughter cells. Mitosis is achieved by condensed chromosomes becoming

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attached in a bipolar fashion to the microtubule-based mitotic spindle and the subsequent pulling apart of the sister chromatids to opposite poles of the cell. Because the mitotic spindles form outside the NE in higher eukaryotes, the whole NE must disintegrate to progress through the cell cycle [12]. This process is termed nuclear envelope breakdown (NEBD) [12, 65]. In lower eukaryotes, the mitotic spindles embed in the inner NE and the process occurs within an intact envelope, yielding a “closed” mitosis, while higher eukaryotic NEBD gives an “open” mitosis [12]. The difference between the two forms has important implications in the events triggering the process in both. The NE is a complex dynamic barrier. NEBD occurs with the G2 /M phase transition at the very onset of open mitosis. NEBD is a phosphorylation-dependent process [12, 66]. Evidence indicates that mitotic disassembly of the NE and the NPC is driven by reversible phosphorylation of a subset of proteins [15, 30, 67, 68], Nups, lamina, and NE membrane proteins, which may disrupt structurally significant interactions. It has been shown that mitotic-activated cyclin-dependent kinase 1 (cdk1) activity is required for keeping NPCs dissociated during mitosis, while reassembly shows a phosphatase dependence [69]. In addition to kinase activity, a microtubule-based tearing process assists NE disassembly in cells [70, 71]. A key component in this process is dynein, which is recruited to the NE at the beginning of prophase and interacts with spindle microtubules [72]. This interaction and the resulting tension placed on the NE lead to its rupture [71, 72]. Microtubule-targeting drugs have been shown to interfere with the onset of this effect and thereby delay NEBD [71]. The combination of phosphorylation and microtubulebased tearing are believed to lead to the resulting NEBD. Although a general description of the dynamic process of NEBD is starting to emerge, very little is known about the molecular machinery behind it. Multiple signaling pathways may be involved in the final triggering. For example, it has been shown that high levels of RanGTP affect the dynamics of the late steps of NEBD and may play a pivotal role in mitotic entry [65]. So the full extent of the molecular components required is not fully understood.

THE NPC AND DISEASES Nups have emerged as factors associated with human diseases. It has been well documented that chromosomal translocations involving genes encoding Nups have been associated with many forms of leukemia, including acute myelogenous leukemia (AML), chronic myeloid leukemia (CML), myelodysplastic syndrome (MDS), and T-cell acute lymphoblastic leukemia (ALL) [3, 73–76]. Viruses such as adenovirus, VSV, and HIV have been shown to utilize the NPC, and in particular individual Nups, to gain entrance into the nucleus for integration [3]. In addition,

Nups have been associated with the autoimmune disease Primary Biliary Cirrhosis (PBC), which slowly destroys the bile ducts [3, 77, 78]. Even triple A syndrome, an adrenal insufficiency (achalasia-addisonianism-alacrima), is caused by mutations in the gene encoding the Nup designated as ALADIN [3]. Recently, atrial fibrillation, the most common form of sustained clinical arrhythmia, has been linked to Nup155 [79]. Loss of Nup155 function causes atrial fibrillation by altering mRNA and protein transport and links the NPC to cardiovascular disease [79]. PBC is an autoimmune disease of the liver [77]. Patients with PBC generate a panel of autoantibodies [80], primarily against mitochondrial antigens, but autoantibodies against FG Nup, Nup62, and NPC-anchoring membrane protein gp210 have been found in nearly a quarter of patients [78, 81]. Combined with mitochondrial antigen detection, α-gp210 and α-Nup62 antibodies may provide additional diagnostic and prognostic tools [80, 82]. But it is not yet known how these Nups are involved in the onset of the disease state [3, 82]. Several Nup genes are involved in chromosomal translocations that produce fusions of Nups and unrelated proteins. For example, Nup98, a mobile Nup, has been shown to have multiple fusions arising in and associated with leukemic transformations. In several cases, NUP98 translocations occur with genes of the homeobox family of transcription factors [76, 83]. These factors normally involve early embryonic development and regulation of hematopoiesis. In addition, Nup98 has been shown to participate in VSV replication through its interaction with VSV matrix protein [34]. Other Nups targeted in chromosomal rearrangements are FG Nups, namely Nup214 and Nup358 [84–86]. Also, several translocations of the TPR gene with specific protein kinases have been found in osteosarcoma, adenocarcinoma, and papillary thyroid carcinoma [87–89].

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11

Protein Maturation and Processing at the Endoplasmic Reticulum Ramanujan S. Hegde Cell Biology and Metabolism Program, National Institutes of Child Health and Human Development, National Institutes of Health, Bethesda, MD, USA

principles of secretory and membrane protein maturation and quality control in the ER.

INTRODUCTION The liver, and hepatocytes isolated from it, have long played a central role in our understanding of cellular organization, protein trafficking, and secretion. The abundance, accessibility, high secretory capacity, and wide range of functions made the liver an ideal choice for many of the earliest and seminal studies of cellular function. The first electron microscopic studies revealed a remarkable compartmentalization to hepatocytes, including the highly abundant membrane-bound organelles of the endoplasmic reticulum (ER), Golgi, mitochondria, peroxisomes, and others. Upon the development of subcellular fractionation methods, each of these organelles could be isolated and studied by classical biochemistry and enzymology [1, 2]. The isolation of liver and hepatocyte membrane fractions enriched in “rough” ER [3] was instrumental in the discovery that ER-bound ribosomes are synthesizing secretory and membrane proteins [4, 5]. These proteins were found to be selectively imported into the ER or inserted into the ER membrane. Indeed, the ER eventually proved to be the major site of secretory and membrane protein biosynthesis and maturation. Twenty years later, the ER was discovered to also be a major site of quality control, the process whereby immature, defective, or otherwise incompletely assembled proteins are triaged for selective degradation [6, 7]. This chapter will cover the basic The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

THE SEGREGATION OF SECRETORY AND MEMBRANE PROTEINS TO THE ER Essentially all secretory and membrane proteins destined for the cell surface or extracellular environment begin their biosynthesis at the ER. Secretory proteins are completely translocated across the ER membrane into the lumenal space, while membrane proteins are weaved into the ER membrane, with specific regions exposed to the cytosol and other portions exposed to the lumen (Figure 11.1). After achieving their correct topology (i.e. orientation relative to the membrane), these proteins undergo a series of maturation events in the ER to produce a functional protein. This involves proper folding, various modifications, and precise assembly with other proteins or co-factors. Only upon successful maturation are the secretory and membrane proteins allowed to exit the ER for their final destinations via vesicular trafficking pathways (see Chapter 7). Thus, the ER can be considered an intracellular factory for the biosynthesis and maturation of secretory and membrane proteins. The key steps in the assembly line

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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Figure 11.1 Segregation of secretory and membrane proteins to the ER. The key steps of co-translational translocation including: signal sequence recognition by SRP, targeting to the ER membrane via SRP-receptor (SR), and translocation through a channel. Membrane proteins utilize the same basic steps and machinery, but undergo integration into the membrane rather than complete translocation

of this factory are the selective targeting of secretory and membrane proteins to the ER, their translocation across or into the ER membrane, their folding into the correct conformation, and their assembly into a functional product.

Secretory and membrane proteins are recognized via signal sequences Cellular protein synthesis occurs on ribosomes located in the cytosol. Yet, over half of all proteins eventually reside in non-cytosolic locales within the highly compartmentalized eukaryotic cell. Thus, it is obvious that cells must effectively sort and segregate proteins among numerous intracellular organelles. One of the major conceptual advances in understanding this problem was the articulation of a cogent hypothesis for the basis of intracellular protein segregation. The “signal hypothesis” postulated that specific regions of a newly synthesized polypeptide (i.e. signal sequences) provide unique codes that specify the eventual destination for that protein [8–10]. This concept, originally developed to explain how secretory proteins are selectively segregated to the ER, eventually proved to be widely generalizable. Indeed, the signals for targeting to the ER, nucleus, mitochondria, peroxisomes, and other locations have been identified. In each case, the sorting signals are selectively recognized by dedicated machinery that mediates correct targeting. For secretory and membrane proteins, the distinguishing feature of their signal sequences essential for selective recognition is hydrophobicity (Figure 11.2; [11]). In many membrane proteins, the first transmembrane domain (TMD), which necessarily must be hydrophobic in

order to eventually reside stably in the lipid bilayer, serves as the signal sequence for selective recognition. However, secretory proteins are soluble in their mature form, and therefore do not typically contain long stretches of hydrophobic residues in their final primary sequence. Instead, they are usually made as precursors containing a cleavable signal sequence at the N-terminus. The cleavable signal is typically ∼15–40 amino acids long and contains a central hydrophobic region essential for its selective recognition and targeting to the ER. As implied by its name, this signal is removed at the ER, and is therefore not part of the final mature protein. Some membrane proteins also use a cleavable N-terminal signal for targeting to the ER. Thus, by either an N-terminal signal sequence or internal TMD, all secretory and membrane proteins are recognized via a hydrophobic motif that sets them apart from other non-ER targeted proteins.

Proteins are targeted to the ER membrane early in their synthesis N-terminal signals and TMDs are recognized cotranslational ly, as these domains first emerge from the ribosome. A large, ribosome-associated factor termed “signal recognition particle” (SRP) interacts directly with the hydrophobic signal sequence [18]. Co-translational SRP-mediated recognition has three important consequences. First, SRP shields the hydrophobic TMD or signal from the bulk aqueous environment to prevent inappropriate interactions or aggregation. In this sense, SRP can be viewed as a chaperone of sorts that protects nascent secretory and membrane proteins during the

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Figure 11.2 Signal sequences and transmembrane domains used in protein translocation. Note that they share no specific sequence, but rather general features such as hydrophobicity. N-terminal signal sequences are usually 15–40 residues long (although many exceptions exist), and contain a central hydrophobic region of at least 7 residues (underlined). Transmembrane domains are longer (16–25 residues) and also hydrophobic. These general biophysical properties (rather than a specific sequence motif) are thought to mediate recognition by the core targeting and translocation machinery. However, more nuanced sequence features may play regulatory roles in controlling substrate-specific translocation [12–15], or may be exploited for selective pharmacologic inhibition [16, 17]

earliest stages of their synthesis. Second, the SRP–signal interaction modulates ribosome function to slow the rate of translation. This translational arrest (which in reality is a transient slowing) provides time for targeting to the ER membrane. And third, SRP binding “marks” ribosomes that are synthesizing secretory and membrane proteins from other ribosomes. Thus, SRP is a key distinguishing feature that allows some ribosomes to be targeted to the ER, while others remain in the cytosol. Selective ER targeting of SRP-bound ribosomes is mediated by the presence in the ER membrane of a specific receptor for SRP. The interaction between SRP and SRP receptor mediates not only targeting, but also the transfer of the nascent polypeptide to a translocon, the next component in the assembly line. This unidirectional transfer of the nascent chain from SRP to the translocon is energy-dependent. Both SRP and SRP receptor are GTPases, and when they interact in the context of a ribosome, nascent chain, and translocon, they hydrolyze their GTPs to induce release of the signal sequence from SRP and transfer of the ribosome-nascent chain to the translocon. Although the details of this critical handoff remain to be worked out, the net effect is to deliver nascent secretory and membrane proteins to an ER-localized translocon while they are being synthesized .

This allows the translocon to transport proteins across the membrane in a linear fashion, before they acquire significant folding. Thus, a key feature of secretory and membrane protein biosynthesis is their co-translational targeting and translocation into the ER.

Translocons mediate protein translocation into the ER Quite remarkably, the same translocon machinery that mediates complete translocation of soluble secretory proteins also handles the insertion of membrane proteins. This is mandated because nearly all secretory and membrane proteins use the same SRP-based system to arrive at a translocon that cannot “know” ahead of time what will be synthesized next. Thus, the translocon, after receiving a nascent protein early in its synthesis, must interpret the subsequently synthesized sequence “on the fly” to mediate either translocation across the membrane (in the case of secretory proteins) or insertion into the membrane in a unique topology (in the case of membrane proteins). These requirements mean that the translocon must have several different functional properties, each of which is only partially understood at the molecular level.

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“Translocon” is a generic term used to describe the composite membrane-bound machinery involved in translocation [19]. The central component of the ER translocon is the Sec61 complex, composed of α, β, and γ subunits, which plays a key role in each of the translocons’ major functions [20]. Prime among these functions is the formation of a translocation channel through which the nascent secretory or membrane protein is transported. This channel is formed by Sec61α, although precisely how it is formed or regulated remains to be clearly delineated. Nonetheless, the channel is aqueous and shielded from the hydrophobic lipid bilayer, large enough to accommodate a nascent polypeptide (even in an alpha-helical configuration), and spans the entire bilayer when in use [21, 22]. Importantly, the opening and closing of this channel is regulated such that it is only opened when needed, and closed at other times. While the mechanisms of regulated opening and closing are not clear, it is generally assumed to be critical for preventing mixing of small molecules that are differentially concentrated in the ER lumen versus cytosol. The second function of the Sec61 complex is association with various partners that aid in translocation (Figure 11.3). The first obvious partner during co-translational translocation is the ribosome. By associating in a precise architecture, the ribosome–Sec61 complex is organized such that the tunnel inside the ribosome is aligned and continuous with the channel formed by Sec61 [22, 23]. Other partners include enzymes that mediate modifications of the translocating polypeptide (see Section “Other pathways of translocation and membrane protein insertion”), assist in signal sequence or TMD recognition by Sec61, stabilize the translocation channel, or recruit chaperones to the site of translocation to aid in polypeptide folding. Many of these associating partners

(a)

that compose the translocon are not necessarily absolutely required for translocation per se, but contribute to maximizing translocational efficiency, and aid in maturation of the translocating polypeptide. The third essential Sec61 function is to facilitate recognition and membrane insertion of TMDs as they emerge from the ribosome. The Sec61 complex has an intrinsic capacity to recognize hydrophobic domains such as signal sequences and TMDs [24, 25]. Indeed, upon initial SRP-mediated delivery, this recognition capacity of Sec61 is used to “proofread” nascent chains in case a cytosolic protein is inadvertently targeted [26]. It is thought that this same recognition function of Sec61 plays a role in TMD insertion, although the precise mechanics of this event remain to be elucidated. Upon complete emergence from the ribosome, the hydrophobic TMD is oriented perpendicular to the plane of the membrane in one of two ways. The sequences within and flanking the TMD are usually used to determine orientation [27]. Once positioned properly, the TMD is allowed to move from the aqueous channel within Sec61 to the lipid bilayer [28]. This lateral release is possible because the channel formed by the Sec61 complex can open sideways toward the lipid bilayer. TMD insertion may be facilitated in some cases by other factors that might “chaperone” it into the membrane. Thus, the translocon is a multifunctional channel that can open (in a regulated manner) in two directions: perpendicular to the membrane to allow translocation, and parallel to the membrane to allow lateral insertion of TMDs [20]. When combined with the ability to recognize and orient key sequence elements (e.g. signal sequences and TMDs), the ER translocon can handle a remarkable diversity of secretory and membrane proteins ranging widely in physical and topologic properties. Amazingly, all of this occurs simultaneously with ongoing

(b)

(c)

Figure 11.3 The translocon at the ER membrane. (a) The central translocon component, the Sec61 complex, forms the channel through which proteins are translocated. The crystal structure of a Sec61 homolog (from archaebacteria) in its inactive state is shown, although it remains unclear how this protein is configured during translocation. (b) Schematic diagram of an active translocon at the mammalian ER, showing the major associated factors: the ribosome, accessory factors in membrane (such as signal peptidase or oligosaccharyl transferase), and lumen (such as the chaperone BiP). (c) Top view of the translocon, illustrating its capacity to open toward the lipid bilayer to allow insertion of a transmembrane domain (TMD), perhaps with the assistance of a putative membrane protein chaperone (such as TRAM)

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protein synthesis occurring at a rate of ∼10 residues per second.

Other pathways of translocation and membrane protein insertion While the SRP- and Sec61-dependent pathway of co-translational translocation is the most broadly utilized and well-studied [29], other pathways are needed for certain specialized substrates. For example, some secretory proteins (including especially small proteins) may be translocated post-translationally by pathway(s) that are poorly understood [30]. Similarly, certain membrane proteins are also inserted post-translationally by an SRP-independent mechanism. One rather large family of post-translationally inserted membrane proteins are “tail-anchored” via a single TMD located close to the C-terminus [31]. Because the stop codon is reached before the TMD even emerges from the ribosome, tail-anchored proteins cannot be co-translationally recognized by SRP. Instead, they are recognized and targeted by another highly conserved and ubiquitous cytosolic factor termed “TRC” (for “TMD-recognition complex”) [32]. This pathway is poorly understood, but the central component (TRC40) is an ATPase that delivers substrates to a putative receptor at the ER and releases them upon ATP hydrolysis. The mechanisms of tail-anchored membrane insertion remain to be studied, but are unlikely to involve the known Sec61 machinery. And finally, there may also exist pathways of secretion that bypass the ER entirely [33]. Several secretory proteins including fibroblast growth factor 2 (FGF-2), interleukin 1-beta, and galectin appear to be transported directly across the plasma membrane by yet unknown mechanisms. The machinery and the physiological importance of such non-classical secretion pathways remain to be studied in detail.

FOLDING AND MATURATION OF SECRETORY AND MEMBRANE PROTEINS Correct ER targeting and translocation of a secretory or membrane protein are only the first steps of its eventual maturation into a functional product. As with any protein, the linear polypeptide must also be folded into a stable three-dimensional structure. Furthermore, many polypeptides also need to be processed, modified, and assembled with other proteins or co-factors before they leave the ER. Each protein has its own unique requirements; some are highly specialized in their maturation, while others use rather ubiquitous general machinery. While the mechanisms that coordinate the multi-step maturation events of

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complex proteins remain poorly understood, many of the individual steps have been better studied and are discussed here.

Chaperones assist in protein maturation Molecular chaperones are essential cellular factors that participate in the maturation of nearly every protein [34]. Chaperones assist in the folding and assembly of nascent polypeptides by direct binding, thereby preventing inappropriate interactions with other proteins in the highly crowded cellular environment. Of course, a nascent protein cannot fold when bound to a chaperone. Thus, chaperones periodically release the nascent protein and transiently afford it an opportunity to fold. If folding is unsuccessful, the chaperone quickly rebinds. By such repeated (and usually energy-dependent) cycles of binding and release, chaperones can facilitate protein folding before inappropriate interactions lead to non-productive aggregation. Essential to chaperone function is their ability to selectively bind immature, but not properly folded, conformations of a nascent protein. This selectivity is thought to be achieved in several ways, but typically involves the recognition of polypeptide segments that should normally be shielded in the final folded structure. For example, the buried core of a folded soluble protein is usually stabilized by hydrophobic parts of the polypeptide. Exposure of such hydrophobic patches, which necessarily means the protein is not properly folded, represents a common recognition motif for chaperone binding [35]. Similarly, hydrophilic residues within some TMDs become buried when multiple TMDs are assembled correctly. Exposure of such residues within the hydrophobic lipid bilayer can therefore signify a non-native conformation. In the same way, exposed cysteines that eventually should be disulfide bonded with other cysteines, can recruit certain types of chaperone. By recognizing these and other sequence elements, chaperones can distinguish between non-native and native folded states. It is therefore not surprising that almost immediately after a nascent secretory or membrane protein begins translocation, chaperones are recruited [36]. This is because proteins are translocated in a largely unfolded conformation, which presents an ideal substrate for chaperone binding. There are many chaperone systems in the ER (see below), and the choice of which chaperones are recruited probably depends on the specific sequence elements within the particular translocating protein [37]. The chaperones usually remain on the maturing polypeptide well after protein synthesis and translocation are completed, and different subsets of chaperones and maturation factors may associate with the same polypeptide at different stages of its maturation. This is especially likely if a protein’s maturation is complex, although such

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non-model proteins are relatively poorly studied. Because these chaperones are ER residents, their repeated binding to immature secretory and membrane proteins also serves the function of preventing the premature exit of non-functional products from the ER.

Multiple chaperone systems operate in the ER lumen Among the various ER chaperones, perhaps the most abundant and well-studied is a lumenal protein called BiP (IgG Binding Protein). BiP is a member of a very large and well-conserved family of chaperones (called the Hsp70 family) with homologs in all known organisms and essentially every cellular compartment [38]. These chaperones are ATPases that utilizes the energy of ATP hydrolysis to bind and release from exposed hydrophobic patches on substrates. In general, the ATP-bound state of the Hsp70s has high affinity for substrate; upon ATP hydrolysis, substrates are released so they can attempt to fold. The steps of ATP binding, ATP hydrolysis, and exchange of ADP for ATP are all tightly regulated by additional factors broadly termed co-chaperones. Thus, BiP and its various co-chaperones form a universal and general chaperone system in the ER lumen that recognizes and shields exposed hydrophobic patches of secretory and membrane proteins in the process of folding and assembly. Although far less studied, another very abundant ATPase in the ER lumen termed GRP94 also functions as a chaperone [39]. Like BiP, GRP94 has a cytosolic homolog (termed Hsp90), further illustrating the theme that similar conserved mechanisms of protein folding are utilized in multiple cellular compartments. Hsp90 interactions with its wide range of substrates are regulated by its ATPase cycle and co-chaperones [40]. It is likely that GRP94 may function analogously to Hsp90, although neither the substrates nor co-chaperones for GRP94 are well defined. While many mechanisms and machineries of protein folding in the ER are similar to those in the cytosol, there are several features unique to the ER folding environment that necessitate specialized folding machinery. One major ER-specific feature is asparagine-linked glycosylation (also called N-glycosylation, discussed below), a modification in which a 14-sugar carbohydrate chain is covalently attached to certain asparagine residues in the nascent protein [41]. Glycosylation is thought to have originally evolved in order to improve the stability of proteins that will eventually function in relatively harsh extracellular environments. Because most proteins transiting the ER become glycosylated, eukaryotic cells have evolved to exploit this modification for numerous other purposes including protein folding, quality control, trafficking, and protein–protein interactions. In the ER, a major use for N-linked glycans is the recruitment of a class of factors known as lectins (a general term which simply means “carbohydrate-binding”).

The two lectin-based chaperones in the ER are called calnexin (CNX, a membrane protein) and calreticulin (CRT, a soluble lumenal protein). They function by a very similar mechanism (see Figure 11.4) and are recruited to substrates via interaction with a specific isoform of the N-linked glycan [42]. As mentioned above, the initial glycan added to a protein consists of 14 sugars: two N-acetyl-glucosamines, nine mannoses, and three terminal glucoses. Shortly after addition of this core glycan, two of the terminal glucoses are removed by the action of glucosidases, resulting in a mono-glucosylated form of the N-glycan that is recognized specifically by CNX or CRT. Upon release from CNX/CRT, the remaining glucose is removed by a glucosidase and the protein has an opportunity to fold. If it fails to fold properly, an enzyme termed UDP-glucose:glycoprotein glucosyltransferase (UGGT) adds a single glucose back to the N-glycan, thereby making it a substrate for CNX/CRT binding and initiating another round of attempted folding. Thus, as with the general chaperones discussed above, repeated cycles of chaperone binding and release permit a substrate to attempt folding while being protected from inappropriate interactions. In this case, a reversible glucose tag is used to modulate substrate–chaperone interactions, with UGGT acting as a “folding sensor” to determine when the substrate has folded [43]. If a substrate passes UGGT inspection, it is not re-glucosylated and the glucose-free N-glycosylated protein is released from the CNX/CRT folding cycle. Another unique feature of the ER lumen is its oxidizing environment. This means that cysteines in nascent proteins will typically be oxidized into disulfide bonds with other cysteines (either within the same protein or in other proteins) [44]. Disulfide bonds can greatly stabilize the folded state of a protein or protein complex, thereby facilitating its function in the harsh extracellular environment. For example, antibodies (e.g. IgG) are composed of multiple polypeptide chains that are held together by a series of intra- and inter-molecular disulfide bonds, which lend them remarkable stability and longevity after secretion. The correct formation of disulfide bonds in a nascent protein is mediated by a class of chaperone enzymes termed oxidoreductases, the most well known of which is protein disulfide isomerase (PDI) [45]. These chaperones can interact directly with substrates (often via exposed hydrophobic patches). Upon substrate interaction, an internal disulfide bond within PDI can be reduced concomitantly with oxidation of cysteines in the substrate. Thus, a disulfide bond is effectively transferred from the PDI to a substrate with which it interacts. The PDI then becomes re-oxidized by other enzymes (e.g. the oxidase Ero1) for another round of substrate oxidation. PDI can also function in reverse, and has the capacity to reduce disulfide bonds within a substrate. This means that by reducing and re-oxidizing different pairs of cysteines, PDI can “shuffle” disulfide bonds until the correct protein fold is achieved (hence the name “disulfide isomerase”).

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Figure 11.4 Glycan-dependent protein folding by lectin chaperones in the ER. Many nascent proteins entering the ER are modified co-translationally with a 14-sugar core glycan containing two N-acetyl glucosamines (squares), nine mannoses (circles), and three glucoses (triangles). The glucoses are rapidly trimmed by glucosidases, allowing the mono-glucosylated glycan to recruit a lectin chaperone (calreticulin; CRT), along with associated factors (such as protein disulfide isomerase-like molecule ERP57). Protein folding is mediated by repeated cycles of binding and release from CRT, regulated by removal and re-addition of the terminal glucose. The enzyme UGGT only adds the glucose to non-native proteins, ensuring that CRT only rebinds if folding has not been achieved. If folding fails despite repeated cycles of CRT binding, the mannoses get further trimmed (by mannosidases such as EDEM), and the glycan now becomes a target for binding by another lectin that routes the protein for degradation. The same general scheme is used by other chaperones: they are recruited to substrates early in their biosynthesis, and undergo cycles of binding and release until the polypeptide either achieves its correct folded state or is transferred to the degradation pathway

Some oxidoreductases are recruited to substrates indirectly via interaction with other chaperones. For example, the oxidoreductase ERP57 interacts with CRT, thereby allowing it to operate on glycoproteins [46]. Thus, different members of the family of oxidoreductases in the ER lumen probably operate on different subsets of client substrates. Although the above chaperone systems are the most abundant and well-studied components of the protein maturation machinery in the ER, they are by no means the only ones. The sheer volume and diversity of substrates that transit the ER necessitate numerous other factors to ensure correct protein folding and maturation. For example, peptidyl-prolyl-isomerases catalyze the change in conformation of proline-containing peptide bonds between the cis and trans isomers [47]. Other chaperones, such as tapasin, help in the assembly of peptides onto MHC class I [48]. Yet other factors are likely to be specialized for the assembly of multimeric protein complexes or specifically involved in membrane protein folding and

assembly, although both of these complex processes are rather poorly understood. And finally, there are likely to exist numerous highly specialized chaperones dedicated to specific substrates with unique requirements. For example, collagen biosynthesis utilizes a chaperone called Hsp47 that is selectively expressed in fibroblasts [49]. In the liver, apolipoprotein B biosynthesis uniquely requires a factor termed microsomal triglyceride transfer protein (MTTP) that mediates non-covalent lipid–protein interactions needed for lipoprotein particle assembly [50].

Numerous Co- and posttranslational modifications accompany protein maturation In addition to the primarily non-covalent processes of folding and assembly, most secretory and membrane proteins are covalently modified during their

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maturation. As already mentioned, the most common modification is N-linked glycosylation. This usually occurs co-translationally, as a protein is entering the ER lumen through the translocon. In fact, the enzyme that mediates glycosylation, oligosaccharyl transferase (OST), is tightly associated with the translocon [51]. As certain sequences (Asn-Xxx-Ser or Asn-Xxx-Thr, where Xxx is any amino acid except proline) pass through the translocon into the ER lumen, OST transfers a fully assembled 14-sugar core glycan to the Asn residue. Not all consensus sites are necessarily glycosylated, and glycosylation is not uniformly efficient in all cell types or under all conditions. While the reasons for this heterogeneity are poorly understood, it is likely to influence the protein’s function and/or trafficking in subtle ways that are important in a tissue-specific manner. These subtleties notwithstanding, glycosylation as a whole is critical to the proper folding and maturation of a wide range of proteins, and failure or even partial deficiency of glycosylation can be disastrous for a cell (especially highly secretory cells such as hepatocytes). Another near-ubiquitous modification is signal sequence cleavage. As mentioned above, N-terminal signal sequences are temporary sequence elements that are used for correct targeting and translocation of secretory and membrane proteins, but are not part of the final mature product. They are therefore removed once they have completed their function; this removal usually happens co-translationally at the translocon. A proteolytic enzyme called signal peptidase interacts with the translocon in such a manner that its active site is positioned adjacent to the opening of the translocation channel in the ER lumen [52]. As a nascent polypeptide begins translocation, the boundary between a signal sequence and mature domain becomes exposed to signal peptidase, which efficiently cuts the polypeptide at a precise position. The third widely-used modification is attachment of a glycosyl-phosphatidylinositol (GPI) anchor to some proteins at their C-termini [53]. GPI anchors are essentially phospholipid molecules containing a glycan (i.e. a glycolipid). Attachment of the GPI anchor to an otherwise soluble protein therefore tethers it to the membrane. This allows GPI-anchored proteins to reside on the cell surface within certain microdomains that are favored by the lipids to which they are tethered. Furthermore, GPI anchors can be cleaved by extracellular phospholipases, allowing for regulated release (i.e. secretion) of certain proteins under specific conditions. GPI anchor addition occurs in the ER and is mediated by a GPI–transamidase complex. The choice of which proteins will be modified by the transamidase is determined by the presence at the very C-terminus of a hydrophobic “signal sequence” for GPI anchor addition. The transamidase recognizes this signal, and proteolytically removes it simultaneously with addition of the GPI anchor (in a transamidation reaction). Deficiencies in GPI anchor

biosynthesis or addition are lethal at the organismal level (although tolerated at the cellular level), illustrating the importance of this modification. Beyond these relatively universal ER-specific modifications, numerous substrate-specific and cell type-specific modifications have been described. For example, the prolines in collagen are hydroxylated, some secreted signaling molecules and morphogens are lipid-modified (e.g. with cholesterol), and other proteolytic processing events have been described in the ER and other compartments of the secretory pathway. In each case, the modifications are typically critical for the protein’s maturation, stability, and/or eventual function.

Many proteins are assembled into multimeric complexes Numerous proteins function as part of multi-protein complexes. Secretory and membrane protein complexes are often assembled in the ER by processes that are relatively poorly understood. In fact, some of the most important and highly expressed proteins transiting the secretory pathway are multi-protein complexes, including immunoglobulins, the T-cell receptor, ion channels, MHC complexes, and numerous others. The basic (and probably overly simplistic) concept that is typically considered in multi-protein assembly is that non-assembled subunits of a larger complex are chaperone-bound until they find and associate with the appropriate interacting partner(s). For example, BiP was originally discovered by virtue of its association with incompletely assembled IgG heavy chains. How assembly is coordinated is not known in most cases, and it is unclear whether it is simply a stochastic process of subunits finding each other by virtue of their mutual affinity, or if there are more regulated mechanisms that facilitate assembly. From this discussion, it should be obvious that the ER lumen is a remarkably complex protein-folding and -maturation factory. Some features of this factory are universal: it is highly rich in evolutionarily conserved chaperone systems that protect nascent proteins from inappropriate interactions and aggregation. Other aspects of the ER are unique to this organelle: N-linked glycosylation, lectin-based chaperones, an oxidizing environment favoring disulfide bond formation, and a very high flux of membrane proteins that need proper folding and maturation. Yet other ER-specific processes are highly specialized for unique substrates in only certain cell types. All of these pathways operate simultaneously; some are probably purely parallel processes, while most are partially overlapping and coordinated events. Thus, the loss or reduced capacity of some maturation pathways can likely be compensated for, at least temporarily for many substrates, by other maturation pathways. Indeed, the loss of a surprising number of the major chaperones (e.g. CRT, CNX,

11: PROTEIN MATURATION AND PROCESSING AT THE ENDOPLASMIC RETICULUM

GRP94, UGGT, and many co-chaperones) is compatible with cellular viability. Yet, in each case, the function of a complex organism or differentiated tissues is severely compromised (usually leading to embryonic lethality), illustrating the limits of redundancy. Although many basic principles have been delineated using model systems, our knowledge of protein maturation should be considered fragmentary and incomplete at best.

QUALITY CONTROL AND THE CULLING OF IMMATURE PROTEINS The relegation of secretory and membrane protein biosynthesis to an intracellular compartment (as opposed to the plasma membrane, as occurs in prokaryotes) provides many advantages to the eukaryotic cell. Perhaps the most obvious and single biggest benefit is the opportunity for considerably more control over what does and does not gain access to the cell surface. This control is manifested in many ways. For example, the cell can effectively regulate the quantity of a secretory or membrane protein

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released to the cell surface independently of the protein’s synthesis. Thus, key regulatory proteins like hormones, surface receptors, or ion channels can be made and stored intracellularly, then rapidly mobilized to the surface at a moment’s notice. This can provide cells with exquisite temporal control and responsiveness to changing environmental conditions. The other main regulatory benefit is quality control [54]. By making and inspecting all secretory and membrane proteins intracellularly, the cell can ensure that only properly folded and functional products have access to the surface. This is critical because partially functional or misfolded proteins can often be far worse than no protein at all. Thus, in highly complex multicellular organisms where intercellular communication via secreted proteins and their receptors is essential for overall fitness, quality control is a key functional role of the ER. The basic logic of ER quality control is threefold (Figure 11.5). First, the cell must have mechanisms to recognize a misfolded, misassembled, or otherwise damaged protein. This task is thought to be carried out by chaperones, which as discussed above, and can discriminate between mature and non-native protein structures. Second, once a potentially misfolded protein is identified,

Figure 11.5 The major steps in ER quality control. Schematic depiction of the overall logic of quality control: recognition of misfolded or unassembled polypeptides, typically by chaperone(s); targeting of the substrate to a retrotranslocation site; retrotranslocation (or extraction from the membrane) to the cytosol; and proteasomal degradation (usually via poly-ubiquitination)

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it must be delivered to degradation machinery. The ER lumen does not appear to contain any obvious mechanisms for protein degradation, necessitating the retrotranslocation of misfolded proteins back to the cytosol. And third, retrotranslocated proteins must be degraded in the cytosol by the major proteolytic system there, the proteasome. These latter two steps are often referred to as endoplasmic reticulum-associated degradation (ERAD). Although less common, there may be situations where misfolded proteins in the ER can be routed via alternative pathways for degradation in lysosomes. The entire process of quality control, and particularly the retrotranslocation step, is only partially understood and remains an intense area of current research.

Misfolded proteins are recognized by chaperones Nascent proteins entering the ER typically undergo repeated cycles of chaperone binding and release in an attempt to achieve their final folded and assembled state. If maturation nonetheless fails, the cell must eventually decide between continued attempts at folding versus triage for degradation. Although the mechanisms underlying this decision remain unclear, it appears that chaperone binding is a key event. Indeed, most mutant or otherwise misfolded proteins are typically found associated with chaperones, suggesting that this interaction is involved in their eventual degradation. This is likely to be important for three reasons. First, the chaperone prevents inappropriate interactions between a misfolded protein and other cellular factors. Second, the chaperone maintains the protein in a largely unfolded state (or facilitates unfolding in some cases), which may be important for its retrotranslocation out of the ER (see Section “A retrotranslocation pathway for misfolded protein degradation in the cytosol”). And third, the chaperone may interact with or deliver proteins to the machinery for retrotranslocation. For example, the chaperones BiP and PDI have been found in association with ER membrane factors implicated in retrotranslocation [55, 56]. In the case of glycoproteins, sugar trimming and ER lectins again play a key role, as they do in the folding cycle (Figure 11.4). After repeated de-glucosylation and re-glucosylation cycles through the CNX/CRT system, mannosidases (including a protein termed EDEM) remove the terminal mannose on which the glucosylation normally occurs [57, 58]. This allows the substrate to exit the CNX/CRT cycle, and then makes the mannose-trimmed glycan a substrate for binding by another lectin, termed OS-9 [59]. Mannose trimming by EDEM and binding by OS-9 routes substrates into degradation pathway, presumably via OS-9 associations with the retrotranslocation machinery [60, 61].

A retrotranslocation pathway for misfolded protein degradation in the cytosol The export of misfolded secretory or membrane proteins to the cytosol for degradation involves components within both the membrane and cytosol [62]. A critical step in this export process, which is nonetheless the least well understood, is the mechanism by which proteins in the ER are first exposed to the cytosol. It has long been postulated that a process analogous to forward translocation must occur in reverse, with a “retrotranslocation channel” acting as a conduit for substrate transport. However, the precise identity and definitive functional demonstration of such a component remains elusive. That notwithstanding, once misfolded proteins become cytosolically exposed they are poly-ubiquitinated. Ubiquitin is a small protein whose covalent attachment to substrates marks them for degradation [63]. The task of ubiquitination in the ERAD process is carried out by any of several ubiquitin ligases located at the cytosolic face of the ER membrane. Upon ubiquitination, additional components are recruited to facilitate the retrotranslocation and extraction of substrate from the ER. For most substrates, retrotranslocation involves a large ATPase complex containing a protein termed p97 (also called VCP or Cdc48) [64]. Together with associated co-factors, the p97 complex appears to recognize both substrate and ubiquitin, and uses the energy of ATP hydrolysis to mediate extraction. The p97-associated poly-ubiquitinated substrate is then delivered to the proteasome, a large multi-catalytic enzyme responsible for most protein degradation in the cytosol. In some cases, the proteasome may directly mediate both extraction and degradation, thereby obviating the need for p97 [65]. While the general scheme of recognition, retrotranslocation, ubiquitination, and degradation is likely to apply for essentially all misfolded secretory and membrane proteins, the details may well be different in a substrate-specific manner. For example, it is already clear that glycoproteins and non-glycoproteins utilize different (but potentially overlapping) pathways. Similarly, soluble lumenal proteins have different requirements for their degradation than integral membrane proteins. Furthermore, the specific location of a misfolded domain (whether in the lumen, the TMD, or cytosol) may influence which retrotranslocation machinery is utilized. Thus, just as multiple parallel and partially overlapping pathways operate during protein maturation of different types of substrate, it seems likely that quality control and retrotranslocation pathways will be equally complex. The diversity of substrates is simply too vast for a single unifying machinery to handle all potential cases.

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Physiological uses of quality control

The multi-faceted mammalian UPR

Although quality control and degradation are typically considered pathways of aberrant protein disposal, they are physiologically exploited for regulatory control in some cases. Two examples of central importance to liver physiology are the regulation of apolipoprotein B and of HMG-coA-reductase. Both proteins are extensively degraded at the ER by the same quality control and ERAD machinery used for misfolded proteins [66, 67]. Yet changes in the environment or physiological state of a cell can rapidly and selectively alter their fate from degradation to maturation. For apolipoprotein B, triglyceride abundance negatively regulates its ER-associated degradation, leading to greater secretion of lipoprotein particles [68]. Analogously, reduced cholesterol levels prevent HMG-coA-reductase degradation, allowing its increased functional expression in the cholesterol biosynthetic pathway [69]. Thus, general quality control pathways have been exploited for highly selective physiological regulation in these and other situations.

Non-native proteins, whether in the process of folding or in preparation for degradation, are typically chaperone-bound. Hence, exceeding the protein processing capacity of the ER has two consequences: the availability of unoccupied chaperones decreases, and at least some nascent polypeptides will not have chaperones bound to them. Both of these consequences seem to be utilized for sensing ER stress and activating the UPR. There are three known ER transmembrane proteins that act as UPR sensors: Ire1, PERK, and ATF6 [70]. The lumenal domains of each of these proteins associate with BiP, which is thought to maintain each in an inactive state. Thus, when BiP is otherwise occupied with folding substrates, it is titrated from the UPR sensors, leading to their activation [72]. In the case of Ire1, misfolded proteins may also directly bind to the lumenal domain, acting as a “ligand” for its activation [73]. Each UPR sensor functions in a different manner to initiate different downstream responses (Figure 11.6). Ire1 activation (via auto-phosphorylation) causes its cytosolic domain to act as an endonuclease to mediate the splicing of mRNA coding for Xbp1 [74, 75]. Upon Ire1-mediated removal of an intron from Xbp1 mRNA, a functional transcription factor can be translated. PERK activation leads to its cytosolic kinase domain phosphorylating the translation initiation factor eIF2α. Phosphorylation of eIF2α leads to a general reduction in overall translation (thereby reducing the burden of new proteins requiring ER function), while allowing increased translation of a select few messages, including the transcription factor ATF4 [76]. The UPR sensor ATF6, upon release from BiP, trafficks to the Golgi, where it is proteolytically processed within its TMD [77]. This cleavage releases the cytosolic domain of ATF6, which functions as a transcription factor. Thus, UPR activation leads to the production of at least three transcription factors (Xbp1, ATF4, and ATF6), which together initiate complex changes in gene expression involving hundreds of factors [78, 79]. The most obvious adaptations are the upregulation of chaperones needed for protein maturation in the ER. In addition, ER components involved in protein translocation, protein degradation, lipid biosynthesis, and others are also increased. The net result is, at the least, an increase in the protein processing capacity of the ER. In some cases, the ER itself may expand markedly, as occurs during the differentiation of pre-B-cells into B-cells. In addition to these transcriptional changes, other adaptations are also induced in the more acute time frame. For example, certain ER-bound mRNAs may be rapidly degraded during ER stress, perhaps by the nuclease activity of Ire1 [80]. Furthermore, the import of some secretory or membrane proteins into the ER may be attenuated as a consequence of reduced availability of

PHYSIOLOGICAL REGULATION OF THE ER The ER is not a static organelle. Its abundance, composition, and functions adapt to changes in functional need. Highly secretory cells and tissues (such as hepatocytes, the exocrine pancreas, and antibody-secreting B-cells) are packed with ER in amounts that can exceed those found in non-secretory cells by more than an order of magnitude. In fact, this correlation between ER abundance and secretory capacity first led to the hypothesis that this organelle was involved in secretion. It is therefore not surprising that cells have mechanisms to sense changes in the load of secretory and membrane proteins transiting the ER, and to adapt accordingly. The discovery of these pathways came about through the study of how cells respond to excessive misfolded proteins in the ER. The “unfolded protein response” (UPR) is activated whenever the load of secretory and membrane proteins exceeds the capacity of the ER to properly mature and/or metabolize them, a situation generally termed “ER stress” [70]. The net result of UPR activation is the initiation of signaling pathways that simultaneously alleviate the load on the ER (temporarily) and upregulate the biosynthetic and maturation machinery to expand ER processing capacity. Failure to effectively adapt to ER stress leads to chronic UPR activation, cellular dysfunction, and at some point, apoptotic cell death [71]. It is increasingly appreciated that chronic ER stress and the consequences arising from it play central roles in various diseases, including many afflicting the liver.

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Figure 11.6 The multifaceted unfolded protein response to ER stress. Upon the generation of excessive misfolded proteins in the ER, the chaperone BiP is titrated from three different signaling molecules (Ire1, PERK, and ATF6) that serve as UPR sensors. Upon release from BiP, each UPR sensor is activated, leading to different downstream changes that adapt to the stress, or in the case of excessive stress, lead to apoptosis

chaperones needed for translocation [12]. This translocational regulation is substrate-selective, and may serve to reduce the burden during acute stress of non-essential proteins that are particularly prone to misfolding. Thus, a combination of general and specific responses operating on multiple time frames provides cells with remarkable flexibility in adapting to changes in substrate flux through the ER. Very similar pathways are likely to also be involved in the physiological regulation of ER abundance that is necessitated during the differentiation of highly secretory tissues [81, 82].

CONCLUSION It has been almost 50 years since the ER was discovered as the site of secretory and membrane protein biosynthesis. In that time, a cohesive molecular framework has been developed to explain the mechanisms of selective segregation of these proteins to the ER, their translocation across or insertion into the ER membrane, and their maturation by various chaperones and processing enzymes. More recently, the major pathways for ER-based quality

control and maintenance of ER homeostasis have been elucidated. While the broad concepts and core machinery for most of these pathways are now in hand, a remarkably large number of gaps exist in our knowledge. How are the processes of targeting and translocation to the ER regulated, especially for complex proteins or under different conditions? How are multi-spanning membrane proteins reliably inserted in the precisely desired topology and subsequently folded and assembled into a functional product? What kind of specialized maturation and quality-control pathways operate in different cell types, and how do they differ from the universal ones that have been studied so far? How are the various steps in a protein’s folding and maturation coordinated? How does the cell distinguish between folding proteins en route to a mature product, and a misfolded protein that should be degraded? What is the mechanism of retrotranslocation? How is the response to ER stress and the choice between adaptation and death tuned differently among different cell types? And what is the role of ER stress in the various diseases associated with the liver? The answers to these and numerous other questions await further study and should occupy cell biologists for the foreseeable future.

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74. Yoshida, H., Matsui, T., Yamamoto, A., Okada, T. and Mori, K. (2001) XBP1 mRNA is induced by ATF6 and spliced by IRE1 in response to ER stress to produce a highly active transcription factor. Cell , 107, 881–91. 75. Shen, X., Ellis, R.E., Lee, K. et al. (2001) Complementary signaling pathways regulate the unfolded protein response and are required for C. elegans development. Cell , 107, 893–903. 76. Harding, H.P., Novoa, I., Zhang, Y. et al. (2000) Regulated translation initiation controls stress-induced gene expression in mammalian cells. Mol Cell , 6, 1099–108. 77. Ye, J., Rawson, R.B., Komuro, R. et al. (2000) ER stress induces cleavage of membrane-bound ATF6 by the same proteases that process SREBPs. Mol Cell , 6, 1355–64. 78. Travers, K.J., Patil, C.K., Wodicka, L. et al. (2000) Functional and genomic analyses reveal an essential

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12

Protein Degradation and the Lysosomal System Susmita Kaushik and Ana Maria Cuervo Department of Developmental and Molecular Biology, Marion Bessin Liver Research Center, Albert Einstein College of Medicine, Bronx, New York, NY, USA

INTRODUCTION TO INTRACELLULAR PROTEOLYSIS In all organs and tissues, intracellular proteins are continuously synthesized and degraded [1, 2]. This constant renewal of the proteome assures its stability and contributes to regulate its function. Altered or damaged proteins are eliminated by the proteolytic systems before their accumulation inside cells interferes with normal cell function [1, 3]. Damaged proteins are first recognized by molecular chaperones, which facilitate protein refolding/repairing (Figure 12.1). However, if the damage is too extensive, or under conditions unfavorable for protein repair, damaged proteins are targeted for degradation. The proteolytic systems thus constitute, along with intracellular chaperones, essential components of the surveillance mechanisms responsible for cellular quality control. Furthermore, the coordinated balance between protein synthesis and degradation also allows cells to rapidly modify intracellular levels of particular subsets of proteins in order to accommodate to a changing extracellular environment or to particular cellular conditions. Increased protein degradation can enhance the effect of reduced protein synthesis to decrease the levels of particular proteins inside cells. Conversely, shutdown of protein degradation is often used by The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

cells to augment the cellular protein pool. Added to these roles in cellular quality control and protein homeostasis, proteolysis is also often used by cells as an additional source of energy when nutrients are scarce [2, 4]. Thus, the free amino acids resulting from the breakdown of cellular proteins can be utilized for the synthesis of essential proteins during starvation, but also as an additional source of energy through their breakdown in the urea and creatinine cycles, or by their transformation into glucose through gluconeogenesis in the case of glucogenic amino acids. Despite the associated expenditure of energy required to break down intracellular products, this continuous recycling of the essential building blocks makes protein degradation a very conservative and economic process with a positive net energetic outcome. Lastly, protein degradation is also necessary during major cellular remodeling (i.e. embryogenesis, morphogenesis, cell differentiation), and as a defensive mechanism against harmful agents and pathogens [1, 5–7]. Protein degradation is an essential cellular process active and functional in all types of cells and conserved throughout evolution. Due to the very active metabolic nature of liver and the unique properties of this organ as experimental tool—accessibility, relative cellular uniformity, and ease for morphological analysis—most of the early studies in protein degradation were performed in liver. In fact, the major proteolytic systems, their essential

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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Life cycle of proteins (c) (a) Cytosol

Folding/ assembling

CHAPERONES REPAIR ENZYMES

(b) Protein

Aggregation Chaperone Ribosomes

TRANSLATION MACHINERY

Refolding/ Repair CHAPERONES

(d)

Degradation

Proteasome

Disaggregation ER

Lysosomes

PROTEOLYTIC SYSTEMS

Figure 12.1 Quality control mechanisms in liver. Two major systems contribute to quality control in liver: chaperones and the proteolytic systems. Chaperones guarantee proper folding of all proteins after synthesis (a) and act as their companions throughout their biological cycle, assisting in the unfolding/refolding required for translocation into different subcellular compartments or for assembly into protein complex. Alterations in protein folding (b,c) are first detected by chaperones, which often facilitate their refolding back into a functional protein. However, if proper refolding is not possible, the altered proteins are eliminated from the cell by degradation through the proteolytic systems, namely the ubiquitin-proteasome system and the lysosomes (d)

components, and many of the regulatory mechanisms of protein degradation were first identified in liver [8–12]. The initial interest in liver proteolysis revolved around the role of this process as a source of energy, in particular during starvation. However, the importance of the proteolytic systems in quality control in this organ has become increasingly appreciated in recent years both in normal liver physiology and in certain pathological conditions.

compartments that participate in complete degradation of proteins into their constitutive amino acids. Two major proteolytic systems are responsible for most of the intracellular protein degradation: the ubiquitin-proteasome system (UPS) and the lysosomes [2]. Although the main focus of this chapter is on the lysosomal system, we briefly review here the characteristics of the UPS and refer interested readers to recent reviews on this topic for details.

INTRACELLULAR PROTEOLYTIC SYSTEMS

THE UBIQUITIN-PROTEASOME SYSTEM

Hepatocytes, like most cells, contain a large variety of proteases both in the cytosol and confined in intracellular compartments. Some of these proteases, such as caspases, calpains, or different secretases, contribute to partial cleavage of cellular proteins, for the most part with regulatory purposes (reviewed in [13, 14]). For example, many enzymes are synthesized as inactive prozymogens that need to undergo partial cleavage in order to become active. Similarly, cleavage of particular regions in certain proteins results in changes in their intracellular location while still preserving full functionality. However, the term “proteolytic system”—main focus of this chapter—has been reserved to define the group of intracellular proteases, assisting components, and intracellular

The proteasome is a multicatalytic complex with a proteolytic core, known as the 20S proteasome, which in high eukaryotes results from the association of 28 subunits in 4 rings stacked as a cylinder-like structure [3, 15, 16]. Three major types of proteolytic activity have been described in 20S proteasomes, but because some of the subunits of this core are exchangeable, proteasomes with different catalytic activities coexist in all cells. The activity of the catalytic core is modulated through the assembly of different regulatory subunits (19S or 11S) that dock on one or both sides of the 20S proteasome to form several proteasome species, likely to participate in different cellular processes (Figure 12.2) [3, 15]. The components of the regulatory subunits are primarily

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E4

175

polyub 26S 19S

α β

E3

β

20S

α

Ubiquitination Substrate ubiquitin

19S

UBIQUITIN/PROTEASOME SYSTEM E1 E2

Figure 12.2 The ubiquitin-proteasome system. The proteasome is a multicatalytic complex formed by a catalytic core (20S) and regulatory subunits (19S and 11S). The catalytic core contains four rings of alpha and beta subunits assembled as a cylinder. Ubiquitin is attached to proteasome substrates through a process mediated by regulatory enzymes known as ubiquitin ligases (E). E1 activates ubiquitin for assembly to the substrates, while E2 transfers the activated ubiquitin to the substrate presented by E3. Multiple rounds through this cycle lead to the formation of polyubiquitin chains covalently linked to the targeted protein. Removal of the ubiquitin tag by the de-ubiquitinating subunits located in the regulatory 19S particle is required for internalization of the substrate into the catalytic core. Subunits of the regulatory 19S also modulate the aperture of the cylinder-like structure of the 20S proteasome to facilitate substrate access

chaperones, ATPases, and enzymes able to remove degradation tags from the substrate proteins delivered to the proteasome [17, 18]. Although degradation of substrate proteins directly by the nude 20S proteasome has been described, degradation of a vast number of substrates involves the participation of the regulatory complexes [16, 19, 20]. The subunits of this complex mediate substrate recognition and unfolding, and often act as the driving force that opens the proteasome barrel and “pushes” substrate proteins into the catalytic core [21]. The proteasome can degrade untagged proteins, but most substrates of this proteolytic system are selectively targeted for degradation through the covalent linkage of ubiquitin, a small (8 kDa) heat-stable protein that also undergoes self-conjugation, resulting in the formation of poly-ubiquitin chains bound to a lysine in the candidate substrate [16, 19, 20]. Linkage of ubiquitin to the cargo proteins is mediated by a series of enzymes—generically known as E ligases—that act sequentially to activate ubiquitin, present it to the substrate, and catalyze the conjugation [22]. Repeated cycles of ubiquitinization result in the formation of the poly-ubiquitin chain recognized by the chaperones and the ubiquitin binding subunits of the regulatory complex of the proteasome. In many instances, ubiquitinization is preceded by phosphorylation

of the substrate protein, which often favors the exposure of the lysine residue that will be used for conjugation of the ubiquitin by specific E-recognizing enzymes [16, 19, 20]. Ubiquitinization is a universal form of protein tagging used not only as a marker for protein degradation through the proteasome but also for degradation through other proteolytic systems, intracellular targeting of proteins to particular cellular compartments, signaling, enzyme activation, and regulation of membrane dynamics, among other things [22]. How the same tagging mechanism could be involved in so different cellular processes has been, for a long time, one of the burning questions in the field. Recent studies have started to reveal that the way in which ubiquitin is conjugated to the proteins is decisive for the function of the tag [23, 24]. Ubiquitin contains seven different lysine residues that can potentially be used for conjugation to the substrates. Linkage through particular residues has been shown now to determine their recognition by different auxiliary proteins involved in different cellular pathways, and hence to mediate the different fates of the conjugated proteins [24]. The UPS plays a critical role as part of the cellular protein quality control system for both cytosolic and secretory proteins [3, 25]. Unfolded proteins, largely newly synthesized cytosolic proteins that cannot reach

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their proper folded conformation, and a large subset of post-translationally damaged proteins (oxidized, glycated, etc.) are removed from the cytosol via the UPS system [26, 27]. Added to the participation of the UPS system in cellular homeostasis and protein quality control by the selective removal of altered proteins, this proteolytic complex also contributes to the regulatory degradation of essential intracellular proteins, usually in a very rapid manner. This fast degradation of factors involved in cell-cycle progression, cell division, transcription, and cell signaling confers the UPS a very critical regulatory role in multiple essential cellular processes [28]. Malfunctioning of the UPS severely impairs cell viability. Primary defects of components of this system have been identified in different types of aggregopathies, establishing a link between the UPS and cellular degeneration [30–33]. The toxic consequences of proteasome blockage have been extensively explored as anti-cancer treatment since functional proteasome is required for cell-cycle progression and cell division [34, 35].

THE LYSOSOMAL SYSTEM Lysosomes are single-membrane organelles containing a wide variety of hydrolases including proteases, lipases, glycosidases, and nucleotidases, which makes them able to degrade all kinds of macromolecules. This high concentration of enzymes was actually what determined their original discovery as a cellular fraction with high acid phosphatase activity [36]. These initial biochemical observations by the group of de Duve were shortly followed by the ultrastructural studies of Novikoff and colleagues, who performed the first electron microscopy study in the isolated fraction and confirmed the presence of single-membrane vesicles of 0.1–0.5 µm diameter and spherical shape [37]. Most of the main criteria that defined a lysosome have persisted despite advances in the understanding of the molecular components and functional relevance of this lytic compartment. The analysis of the different routes followed by the cargo—components to be degraded by the lysosomes—in order to reach the lysosomal lumen has given rise to the description of a series of lysosome-related compartments, such as endosomes, phagosomes, and autophagosomes, all of which will be described in the following sections. Cytosolic vesicles are catalogued as lysosomes, also referred to as secondary lysosomes once they have received cargo, and distinguished from the other lysosome-related vesicles when they fulfill the following criteria: single membrane, pH in the range of 4.5–5.5, active (cleaved) hydrolases, presence of lysosome-membrane proteins, and absence of typical endosomal markers such as mannose-6-phosphate receptor or particular endosome-associated Rab proteins [14, 36, 38, 39].

The enzymatic machinery Lysosomal hydrolases are synthesized as pre-proenzymes that get activated through proteolytic cleavage as the pH decreases. After synthesis in the endoplasmic reticulum (ER), all lysosomal enzymes traffic through the Golgi, where they become glycosylated and covalently tagged with a mannose-6-phosphate residue. This tag is selectively recognized by the mannose-6-phosphate receptor at the trans-Golgi network (TGN), which helps concentrate lysosomal enzymes inside small vesicles budding from the Golgi and targeted to endosomes. Sorting at the Golgi requires the participation of adaptor proteins such as AP1 and of the GGAs or Golgi-localized gamma-adaptin ear homology domain proteins [40, 41]. As the acidification of this compartment increases with maturation, the hydrolases dissociate from the receptor and are delivered to lysosomes, whereas the receptor is recycled back to the Golgi. A percentage of lysosomal enzymes escape this sorting step, despite the tagging, and follow the secretory pathway to be released in the extracellular media. Even though there is growing evidence that lysosomal enzymes can play important functions in extracellular matrix remodeling, cell defense, and maintenance of the extracellular environment [42], some of the released enzymes are internalized back into the cell via endocytosis after interacting with a variant of the mannose-6-phosphate receptor present at the plasma membrane. Double knockdown of the two mannose-6-phosphate receptors has revealed the presence of an as yet poorly understood mannose-6-phosphate-independent targeting [43]. Although more than 50 different lysosomal hydrolases have been described in the lysosomal lumen, lysosomal proteases—also known as cathepsins—are of particular interest for this chapter. Cathepsins can act both as endoand exopeptidases (cleaving directly the internal residues of the amino acid sequence of the cargo proteins, or only N-terminal or C-terminal residues, respectively) and belong to the families of serine, cysteine, and aspartic proteases [44–46]. It is generally accepted that degradation of substrate proteins is initiated by endoproteolytic cleavage that generates peptides amenable to exoprotease degradation. Lysosomal proteases reach maximal activity at the very acidic pH of the lysosomal lumen, but many of them still have considerable residual activity at neutral pH. The particular redox conditions of the lysosomal lumen and the low pH facilitate unfolding of internalized cytosolic proteins, which allows proteases to gain access to internal residues. Changes in the redox potential and in the lumenal pH modulate the proteolytic activity inside lysosomes [47, 48]. Defective lysosomal hydrolysis has been associated with severe human disorders known generically as lysosomal storage disorders (LSDs). These disorders differ in the particular hydrolase affected, but all of them share the presence of engorged lysosome-related compartments in the cytosol of the cell, with the consequent cell and

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organ expansion (i.e. hepatomegaly and splenomegaly are features common to many LSDs). Pathology arises both as a result of the defective processing of the substrate for the hydrolase affected in each LSD—which will limit recycling of particular essential components—and as a consequence of an abnormally expanded lysosomal system filled with un-degraded products that alter the lysosomal lumenal conditions. Thus, accumulation of theses products inside lysosomes reduces their pH, changes their redox status, and alters the dynamics of proteins between lysosomal lumen and membrane, as well as the ability of lysosomes to fuse with other lysosomes (homotypic fusion) or with other vesicular compartments in the cell (heterotypic fusion). Readers are referred to recent reviews for detailed descriptions of the different LSDs and current therapeutic advances for these disorders [49–54].

Proteins at the lysosomal membrane The lysosomal membrane is far from a mere static barrier to contain the highly active lumenal hydrolases away from the cytosol. On the contrary, proteins at the lysosomal membrane mediate the essential functions of this organelle. Thus, transporters at the membrane allow trafficking of cytosolic components in and out of the lysosomal compartment, a proton pump maintains the low pH lumenal lumen, and integral membrane proteins and the membrane’s associated partners facilitate the fusion of lysosomes with other vesicular compartments [40, 47, 55, 56]. The most abundant of the integral proteins at the lysosomal membrane are the lysosome-associated membrane proteins (LAMPs) [55–58]. Two different but highly homologous proteins, LAMP-1 and LAMP-2, are the most prominent members of this family. They are single-span membrane proteins with a very heavily glycosylated lumenal region and a very short cytosolic tail. Glycosylation constitutes about 60% of the mass of these proteins and is required to preserve their stability, likely by preventing lumenal hydrolases from gaining access to their peptide core. Despite their high homology, studies in animal models knocked out for LAMP-1 or LAMP-2 have revealed marked functional differences between these two proteins. Knockout of LAMP-1 has a very mild phenotype and upregulation of LAMP-2 [59], supporting the supposition that deficiency of LAMP-2 can be compensated for by LAMP-2. In contrast, loss of LAMP-2 expression results in a dramatic phenotype with major alterations in lysosomal biogenesis, problems in clearance of autophagic compartments, and altered vesicular trafficking [60]. This diversity of effects can be attributed to the existence in most cells of three isoforms of LAMP-2 resulting from the alternative splicing of the Lamp2 gene [61]. These isoforms—LAMP-2A, LAMP-2B, and LAMP-2C—have identical lumenal regions but distinctive transmembrane and cytosolic tail. As described below, LAMP-2A is

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essential for a selective type of autophagy of cytosolic proteins [62], whereas LAMP-2B seems to play a role in vesicular fusion between lysosomes and autophagic vesicles [60, 63]. Lastly, LAMP-2C has been proposed as a linker between lysosomes and the microtubule cytoskeleton, which explains its participation in different forms of vesicular trafficking. The other abundant type of lysosomal membrane protein is the lysosomal integral membrane protein (LIMP). The two members of this family described to date, LIMP1 and LIMP2, insert into the lysosomal membrane in a hairpin fashion, with N- and C-termini exposed in the cytosol. LIMP1 has been shown to contribute to fusion events in secretory organelles, whereas LIMP2 participates in lysosomal biogenesis through its interactions with vesicle fusion and fission components [64]. Although less abundant than LAMPs and LIMPs, the conserved vacuolar proton pump (V-type H+ ATPase) has been extensively studied for its essential role in maintaining the acid lysosomal lumen [39]. This pump uses the energy derived from ATP hydrolysis to promote translocation of protons through the coordinated action of a cytosolic region with ATPase activity and a transmembrane region that acts as a translocon. Different transporters are present at the lysosomal membrane to mediate translocation of hydrolysis byproducts from the lysosomal lumen toward the cytosol for recycling [65]. The only amino acid transporter cloned to date is the cysteine transporter or cystinosin, a seven-span transmembrane protein that, when mutated, gives rise to the LSD cystinosis [66]. A monosaccharide transporter has also been identified at the lysosomal membrane and mutations leading to functional loss of this transporter have been found in the LSD Salla disease. Lysosomal efflux of other products such as oligosaccharides, small peptides, and free fatty acids has been shown experimentally, but little is known about the transporters that modulate their exit from the lysosome. Lastly, some lysosomal enzymes are located at the lysosomal membrane rather than the lumen. Typical example of these membrane enzymes are the 70 kDa lysosomal apyrase-like protein, which contributes to the metabolism of tri- and diphosphate nucleotides, and the acetyl-CoA:α-glucosaminide N-acetyltransferase, which transfers acetyl groups to heparin sulfate. Part of the lysosomal acid phosphatase, originally used to identify this compartment in the early studies by de Duve, is also localized in the lysosomal membrane as an inactive single-span protein that is released into the lumen by proteolytic processing. As better understanding is gained on the function of different lysosomal membrane proteins, the number of diseases related to alterations in these proteins keeps growing. In addition to the above-mentioned LSD due to mutations in transporters, genetic alterations in the lamp2 gene have also been connected to a muscle vacuolopathy known as Danon disease [63]. Danon

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disease has been classified as a lysosomal glycogen storage disorder with normal acid maltase activity in which autophagic vacuoles (AV) accumulate in all tissues, including liver. Patients suffer fatal cardiomyopathy due to the weakening effect of the vacuoles on the cardiac muscle. LAMP-2 deficiency is the primary defect in this disease, which is associated with altered lysosomal biogenesis and malfunctioning of the autophagic system.

Lysosomal pathways for proteolysis Lysosomes are the common final compartment for degradation of components that originate from both the inside of the cell (autophagy) and from the plasma membrane or extracellular media (heterophagy). Because of the focus of this chapter on intracellular protein degradation, only a brief description of the heterophagic pathways is presented in this section. Interested readers are referred to recent reviews on endocytosis for more details about this fundamental process [67–72].

Heterophagic pathways Hepatocytes, like any other cell type, require continuous sampling of their surroundings in order to accommodate changes in the extracellular environment. This constant

Fluid phase

exchange of information is attained through endocytosis, which often leads to activation of complex signaling mechanisms originating from the plasma membrane or from the membrane of the internalized vesicular compartments (endosomes) [73, 74]. The extracellular components, or cargo, can be internalized through different endocytic mechanisms, which mainly differ in the manner in which this cargo is recognized (Figure 12.3). Through fluid-phase endocytosis or pinocytosis, small portions of the extracellular media, including mainly soluble macromolecules and micronutrients as well as small particles, are continuously internalized into small vesicles. The capability of this process is relatively low but because of its constant nature it can result in the internalization of as much as 30% of the plasma membrane per minute. This forces a coordinate balance between the endocytic process and the hepatocyte secretory mechanisms that will return most of this membrane back to the cell surface. Fluid-phase endocytosis is a non-saturable process with relatively low capacity, used often for internalization of molecules such as sucrose [75, 76]. One further step in the selectivity and efficiency of cargo internalization is achieved through a second type of endocytosis known as absorptive endocytosis [77]. In this case, the internalized molecules interact weakly with the plasma membrane before being endocytosed, resulting in some degree of cargo concentration in the particular region of the plasma membrane that undergoes invagination. In contrast to fluid-phase, absorptive endocytosis is saturable as

Absorption

Receptor-mediated

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Cargo interacts weakly with the plasma membrane

Cargo binds to specific receptors

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Saturable

Saturable

Low capacity

2–100 times higher

106–109 higher

hepatocytes

hepatocytes

hepatocytes

sucrose

aldolase

insulin-like growth factor

Figure 12.3 Types of endocytosis. Hepatocytes, like most cells in the organism, continuously sample and receive contents and information from the extracellular environment via endocytosis. The three types of endocytosis simultaneously active in hepatocytes are displayed here. Left: Fractions of the extracellular media are internalized in a non-selective manner through fluid-phase endocytosis, a very low-capability endocytic type. Middle: The interaction of extracellular components with the membrane prior to internalization explains the higher capability of absorptive endocytosis. This type of endocytosis can be saturated once all the surface of the cell membrane is occupied by interacting molecules. Right: Receptor-mediated endocytosis is the most selective type of endocytosis, in which extracellular components interact with particular receptors at the cell surface. The high capability of this pathway is attained through the concentration of the cargo-loaded receptor proteins in particular regions of the plasma membrane where internalization occurs. Examples of extracellular molecules internalized by each pathway are depicted at the bottom

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it depends on the amount of cellular surface available for cargo interaction, but the concentration step makes it about 100 times more efficient than fluid-phase endocytosis [78]. Many proteins in the blood are internalized in hepatocytes through this mechanism. The maximal example of selectivity and efficiency in cargo internalization occurs through receptor-mediated endocytosis, where cargo molecules first bind directly to particular receptor proteins at the plasma membrane, which triggers a series of events leading to the concentration of receptor-cargo in particular regions of the plasma membrane, internalization in vesicles that form through a complex process of adaptor protein assembly, and their delivery to lysosomes through a regulated series of vesicular fusion and fission events [71, 74, 79–82]. Like absorptive endocytosis, receptor-mediated endocytosis is a saturable process, but its capability is 106 –109 times higher than the former. Independent of the mechanisms that mediate internalization, lysosomes constitute the final destination of the cargo-containing vesicles. These endocytic vesicles, or endosomes, serve also as sorting compartments for molecules such as the receptors that are spared from degradation and are instead delivered back to the plasma membrane to continue their function [71]. Endosomes mature into acidic lysosomes through fusion and fission events regulated by pairs of proteins located in the membranes of both acceptor and donor vesicles [80]. Once a certain degree of acidification is attained, recycling is no longer possible and the cargo is completely degraded by the lysosomal hydrolases. Readers are referred to recent comprehensive reviews for a detailed description of the molecular components that participate in endocytosis [74, 80–82]. Lastly, macrophagic cells in the liver such as Kupffer cells are capable of internalization of extracellular pathogens and other large-sized particulate components by phagocytosis, a specialized form of heterophagy [80, 81]. In contrast to the discrete invaginations of the plasma membrane characteristic of endocytosis, during phagocytosis these professional defense cells undergo major deformity of their plasma membrane and cytoskeletal rearrangements to assure engulfment and internalization of the oversized cargo. Phagocytic vesicles or phagosomes fuse with secondary lysosomes through similar mechanisms to the ones followed by endocytic vesicles for complete degradation of their cargo. Readers are referred to recent reviews and other chapters in this book addressing the importance of the phagocytic activity of Kupffer cells in chronic liver inflammation and in the innate immune response [82, 83].

Autophagic pathways The term autophagy is commonly reserved for the group of different pathways that mediates the lysosomal degradation of components present inside cells, including

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all soluble proteins, organelles, cellular subcompartments, and particulate protein deposits. However, autophagic cargo does not always belong to the cell, as extracellular components internalized by phagocytosis can also be the target of degradation by the autophagic system through a process described as xenophagy (to point out the exogenous nature of the cargo) [84, 85]. Although autophagic degradation has been known since the early discovery of lysosomes and its morphological features and hormonal regulation have been extensively analyzed using liver as a study model, only recently has a complete molecular dissection of some of the autophagic pathways been obtained. The main drive for this “rediscovery” of autophagy has been the three different mutagenesis screenings in yeast, which led to the identification of more than 30 genes generically known as autophagy-related genes (ATG) [86]. Genetic manipulations of these genes—knockouts, knockdowns, and overexpressions—have helped in identifying novel physiological roles for autophagy and have linked dysfunction of this lysosomal pathway with prominent human diseases such as cancers, neurodegenerations, myopathies, and different metabolic disorders [1, 5]. Three major types of autophagy have been described in liver and in almost all types of mammalian cells: macroautophagy, microautophagy, and chaperone-mediated autophagy (CMA) [1, 5] (Figure 12.4). Since the molecular mechanisms and physiological relevance of each autophagic pathway are different, the following sections provide separate reviews of the major advances in the current understanding of each.

Macroautophagy The original description of a type of non-selective degradation in which whole regions of the cytosol sequestered into double-membrane vesicles were delivered to lysosomes upon fusion of the two vesicular compartments gave rise to the term “macroautophagy,” to differentiate it from a smaller-scale degradation resulting from small invaginations in the lysosomal membrane, or “microautophagy” [8, 11] (Figure 12.4). Macroautophagy is quantitatively the most important form of autophagy and also the best characterized as a result of the recent identification of the ATG genes. The protein products of these genes organize in four major groups: two conjugation cascades, an initiation complex, and a negative regulatory complex [1]. The conjugation events are required for the formation of the limiting membrane that elongates and seals itself, forming a double-membrane vesicle or autophagosome [87] (Figure 12.5). This is the only example so far in cells of a membrane forming de novo in the cytosol through a complex conjugation of a protein (Atg5) to another protein (Atg12) and of a protein (Atg8 or LC3 in mammals) to a lipid (phosphatidylethanolamine) [88]. The series of events required for conjugation and the similarity of the

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Chaperone- mediated Autophagy

Microautophagy

LAMP2A

Macroautophagy

CMA substratechaperone Lysosome

Autophagosome

Limiting membrane

Figure 12.4 Autophagic pathways in liver. Three major types of autophagy coexist in hepatocytes and almost all mammalian cells: macroautophagy, microautophagy, and chaperone-mediated autophagy (CMA). In macroautophagy, whole regions of the cytosol, including organelles and soluble proteins, are sequestered by a limiting membrane that seals to form an autophagosome. This double-membrane vesicle delivers cargo to lysosomes for degradation through vesicle-to-vesicle fusion. In microautophagy, whole regions of the cytosol are internalized into the lysosomal lumen in invaginations or projections of the lysosomal membrane that seal to form single-membrane tubules and vesicles. CMA allows selective degradation of soluble cytosolic proteins. Substrates are identified by a chaperone–cochaperone complex that delivers them to the membrane of the lysosome where they interact with a receptor protein, the lysosome-associated membrane protein type 2A or LAMP2A. Substrate proteins unfold and are translocated into the lumen, assisted by a lumenal chaperone

enzymes that modulate this process to E ligases make autophagic conjugation resemble, to some extent, the tagging of proteasome substrates by ubiquitinization. Activation of conjugation, required for the formation of the autophagosome, occurs through the activation complex. The main components of this complex are Atg6/beclin-1, a member of the phosphatidyl-inositol-3-kinase type III (PI3K-III), and several other exchangeable proteins that allow this complex to regulate both autophagic and endocytic pathways [89]. Although levels of the components of the initiation complex were initially proposed as good markers for autophagic activity, recent studies have shown that changes in the stoichiometry of their interactions, as well as in the intracellular location of these complexes, actually determine macroautophagy activity [90]. Once the autophagosome is formed, sequestering inside the cytosolic cargo, it is delivered to lysosomes in a microtubule-dependent manner. Both vesicles fuse through mechanisms yet to be elucidated, allowing lysosomal enzymes to gain access to the cargo (Figure 12.5). Although most of the autophagosomes fuse directly to secondary lysosomes, forming autophagolysosomes, in almost all cells direct fusion of autophagosomes to late endosomes, forming amphisomes, has also been described [91–93]. Although this interaction of the autophagic and

endocytic pathways does not usually contribute much to the autophagic degradation, under certain conditions when autophagosome–lysosome fusion is impaired, it becomes the main route for autophagosome clearance. The fourth subset of Atg proteins acts as a negative regulator of macroautophagy and has as its central component mTOR, one of the major nutrient-sensing intracellular kinases [94–96]. mTOR integrates the information received through the insulin-signaling pathway and the amino acid receptors, which informs this kinase of the energetic status of the cell. In the presence of nutrients or energy excess, mTOR is activated to exert a negative effect on macroautophagy. However, when nutrients are sparse, mTOR is inactivated, resulting in activation of macroautophagy. The downstream effectors of mTOR on the autophagic pathway are well characterized in yeast, but their mammalian homologues are yet to be identified. The hormonal regulation of macroautophagy in liver was well characterized even before the molecular components of this pathway were identified. The insulin/glucagon balance in blood has a direct effect on macroautophagic activity in liver [8, 12, 97]. In the post-prandial period, the high levels of insulin circulating in blood repress activation of macroautophagy, whereas the gradual increase in circulating levels of glucagon, as

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(a)

(b)

(c)

Figure 12.5 Ultrastructure of the autophagic system. (a) Electron micrographs of sections of mouse livers after six hours of starvation. Autophagic vacuoles—autophagosomes and autophagolysosomes—are indicated with arrows. Bottom panels show details of different stages of maturation of autophagic vacuoles, from early or immature vesicles (left) when cargo is still recognizable, to late or mature vesicles (right) where only degraded material and membranous structures are observed in the lumen. (b,c) Electron micrographs of lysosomal fractions isolated from the same livers. Autophagic vacuoles (b) and secondary lysosomes (c) are shown. Insets show representative examples of the type of autophagic/lysosome structure present in each fraction

the time after nutrient consumption progresses, results in activation of macroautophagy to provide the energy and the essential components required for the synthesis of proteins necessary during the nutritional stress. Besides an alternative source for cellular energy, macroautophagy is also a major component of the systems responsible for intracellular quality control [1]. Although selective removal of soluble proteins cannot be attained through this pathway, macroautophagy contributes to the continuous turnover of organelles, proteins adopting irreversible insoluble conformations (such as protein inclusions and aggregates), and it is at the forefront of the cellular response to organelle stress, from ER stress to mitochondrial or Golgi stress [46, 98, 99]. In contrast to the lack of selectivity of this pathway for soluble cytosolic proteins, removal of organelles or particulate structures from the cytosol via macroautophagy is a discriminatory process. Thus, only those structures identified as non-functional or damaged

are sequestered inside autophagosomes for their lysosomal delivery. Although the mechanisms mediating this selective recognition remain, for the most part, unknown, the importance of macroautophagy in quality control has been underscored as a result of recent studies with conditional mouse model knockout for macroautophagy genes in different tissues. Conditional knockout of the essential macroautophagy gene ATG7 in liver [100], the first established model of this type, revealed, as expected, a lower ability of the liver of these animals to accommodate proteolysis rates to the cellular energetic needs during nutrient deprivation. Unexpectedly, damaged organelles and protein inclusions accumulate in the livers of these animals even when maintained under normal nutritional conditions, supporting a critical role for macroautophagy in the maintenance of cellular homeostasis through the continuous removal of intracellular components [100]. Similar consequences have been observed upon conditional blockage of macroautophagy in other tissues

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and organs [101–103]. This subset of studies underscores a prevalent role for the basal activity of macroautophagy, previously classified as only a stress-induced pathway. Whether or not basal and inducible macroautophagy have similar molecular requirements and regulatory mechanisms is currently under intensive investigation. The high degrading capability of macroautophagy is the basis for its participation in processes requiring major cellular remodeling, such as embryogenesis, development, cell differentiation, wound healing, and tissue regeneration [1, 104]. In addition, as explained above, macroautophagic sequestration of pathogens, free in the cytosol after escaping the endocytic system or still contained in compartments of the endocytic and phagocytic pathway (xenophagy), confers macroautophagy an important role in cellular defense against external aggressors [105]. The wide variety of functions attributable to macroautophagy explains the negative consequences of the malfunctioning of this pathway and its association with different human pathologies. Recent connections established between macroautophagy failure and liver diseases will be reviewed in later sections.

Microautophagy Described initially as a constitutive mechanism for cytosolic content turnover in lysosomes, microautophagy remains poorly understood today. The classic definition of this autophagic pathway resulted from the morphological observation of secondary lysosomes containing multiple vesicular structures in their lumen filled with cytosolic content [11, 106] (Figure 12.4). Further studies, mainly in yeast, have demonstrated that the vacuole, equivalent to the lysosome in yeast, can trap whole regions of cytosol, including both soluble proteins and organelles, through invaginations, tubulations, and projections from this compartment’s membrane. This uptake of cytosolic content has been reproduced recently in vitro using isolated yeast vacuoles [107, 108]. These studies have revealed that deformation of the membrane requires cytoskeletal components and is regulated by a subset of gene products specific to this process. In addition, blockage of particular ATG genes also inhibits microautophagy in yeast, suggesting that macro- and microautophagy share some common components. Unfortunately, most of the macroautophagy-specific genes do not seem to be conserved in higher species, limiting the information about this pathway in mammals. Based on the mechanism of cargo sequestration, microautophagy has been considered a non-selective type of autophagy, but, as in the case of macroautophagy, some level of selectivity has been described in relation to the removal of organelles. Thus, after exposure to conditions associated with peroxisome proliferation in yeast, microautophagy contributes to the restoration of cellular homeostasis by selectively removing the excess of

peroxisomes [109]. Lysosomal elimination of peroxisomes has also been described in liver after clofibrate-induced proliferation of these organelles [110]. Whether or not this process takes place via microautophagy requires future elucidation. Microautophagy of complete nuclear regions by the vacuole has recently been described under the name “piecemeal microautophagy of the nucleus” [111]. In this case, targeted interactions of proteins in the nuclear envelope and in the vacuole membrane induce the formation of a nuclear bleb that is captured, pinched off, and degraded by the vacuole. Interestingly, genetic material is consistently excluded from the protruding regions of the nucleus. The identification of a similar process in mammalian cells is intensively sought after, since activation of such process could assist in the elimination of nuclear inclusions common to degenerative disorders and many protein conformational diseases.

Chaperone-mediated autophagy In contrast to the autophagic pathways described in previous sections, in CMA, soluble cytosolic proteins are delivered to the lysosomal lumen in a selective manner after crossing the lysosomal membrane [112, 113] (Figure 12.4). The selectivity of this pathway is conferred by a cytosolic chaperone, hsc70, the constitutive member of the hsp70 family of chaperones, which identifies in the substrate proteins a pentapeptide motif—biochemically-related to the pentapeptide KFERQ—and targets them to the lysosomal surface [114]. The substrate proteins bind there to monomeric forms of LAMP-2A [62], thus driving the multimerization of this receptor into the high-molecular-weight protein complex required for translocation [115]. After unfolding, and assisted by a chaperone located in the lysosomal lumen, substrates cross the lysosomal membrane and are rapidly degraded [116, 117]. Membrane levels of the lysosomal receptor are limiting for this pathway and determine overall the capability of particular lysosomes to perform CMA [118]. Transcriptional changes, regulated cleavage, and membrane subcompartmentalization of the lysosomal receptor contribute to regulate the lysosomal levels of LAMP-2A and consequently the cellular CMA activity [118]. Basal levels of CMA can be detected in almost all mammalian cells, but this pathway is maximally activated under conditions of stress [112, 113]. Activation of CMA fulfills two types of cellular needs: maintenance of a supply of amino acids necessary for protein synthesis under conditions of prolonged starvation and selective removal of damaged proteins from the cytosol. Although macroautophagy is activated in liver during the first hours of starvation to provide the amino acids and other macromolecules required for different anabolic processes under this condition, as starvation persists, macroautophagic

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activity decreases concomitant with a progressive increase in CMA activity. Activation of CMA late in starvation likely allows selective degradation of non-essential proteins to provide amino acids for the synthesis of proteins essential under this stress condition. CMA is also upregulated during mild oxidative stress and after exposure of cells to toxic compounds known to induce irreversible conformational changes in proteins [119]. Altered proteins can thus be eliminated selectively via CMA without affecting nearby functional proteins. Blockage of CMA in cultured cells does not result in major changes in cellular homeostasis under basal conditions, as proteolysis rates are maintained by a compensatory activation of macroautophagy [120]. However, although these pathways can compensate for one another, they are not redundant, and the lack of CMA becomes noticeable during stress conditions. Exposure of cells with impaired CMA to different types of stressors reveals their higher sensitivity to stress and a marked increase in apoptotic cell death, despite macroautophagy being perfectly functional [120]. These results support the theory that the selectivity of CMA might be important under these conditions.

PROTEIN DEGRADATION IN LIVER DISEASE AND AGING The intrinsic high metabolic activity of the liver makes fast adaptation to changes in nutritional conditions and tight quality control indispensable for its proper functioning. Consequently, deregulation of the major proteolytic systems described in this chapter results inevitably in altered liver function and has been shown to contribute to the pathogenesis of common liver diseases (Figure 12.6). Altered proteasome function contributes, at least in part, to the formation of Mallory bodies, one of the best-studied types of hepatic protein inclusion, observed in diverse chronic liver diseases such as alcoholic and non-alcoholic steatohepatitis, chronic cholestasis, metabolic disorders, and hepatocellular neoplasms [121]. Direct inhibition of the proteasome by different agents, such as ethanol in the case of alcoholic steatohepatitis, favors the accumulation of cytokeratins, different pro-apoptotic factors, and negative regulators of cytokine signaling, which contribute to the chronic disease [122].

Alpha-1-antitrypsin deficiency

HCV

• Soluble protein degraded by proteasome • Aggregates in the ER degraded by macroautophagy • Altered mitochondria

Hepatitis C Virus • Promotes ER stress • Blocks autophagosome– lysosome fusion

ER

Fatty liver • Lipid droplets degraded by macroautophagy • Chronic lipid stimulus inhibits macroautophagy leading to fatty liver

183

LYS MIT

LD

Cancer

AV

• Decreased macroautophagy in many cancers

Aging • Decreased macroautophagy • Decreased CMA

Figure 12.6 Autophagy and liver disease. Recent studies have revealed numerous connections between autophagy malfunctioning and liver pathology. Examples of some of these liver disorders and the changes in the autophagic system under these pathological conditions are shown here. In certain pathologies, for example alpha-1-antitrypsin deficiency, the soluble protein is degraded by the proteasome, while once the protein forms aggregates—in this case in the ER—macroautophagy is more adept at degrading these. Sustained lipid challenge leads to blockage of macroautophagy, which normally degrades the lipid stores, resulting in fatty liver disease. Decreased activity of the proteolytic systems is responsible, in part, for the adverse abnormalities observed in aging liver, while it is beneficial for the cancer cells. Bacteria and viruses subvert the autophagy machinery for their own replication. AV, autophagic vacuoles; ER, endoplasmic reticulum; HCV, hepatitis C virus; LD, lipid droplets; LYS, lysosomes; MIT, mitochondria

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Dysfunction of both the proteasome and the autophagic system has been described in alpha-1-antitrypsin deficiency, a protein conformational disorder resulting from point mutations in a secretory liver protein that accumulates in hepatocytes, leading to chronic liver inflammation and carcinogenesis [123]. The soluble form of the mutant protein is usually degraded by the proteasome, whereas the semi-aggregate and aggregate forms that accumulate in the ER depend on macroautophagy for their removal [124]. Furthermore, macroautophagy also contributes to the removal of the altered mitochondrion abundant in the alpha-1-antitrypsin-deficient liver [125]. A direct connection between the carcinogenesis associated with this disorder and autophagy has not been established yet. However, there is extensive evidence for an involvement of autophagy in oncogenesis [126]. A decrease in macroautophagic activity is often found in many types of cancer cell shown to be haploinsufficient for essential ATG genes [89]. Because reduced protein degradation and upregulation of protein synthesis are favorable conditions for rapid cell division, induction of macroautophagy in cancer cells often slows down their replication. However, macroautophagy is not completely shut down in these cells because it still constitutes a major defense against adverse conditions such as the poor nutritional environment of the center of hypovascularized tumors or the damage induced by anti-oncogenic treatments [126]. In fact, modulators of macroautophagy are currently being considered as possible enhancers of anti-cancer therapies [127, 128]. As in other cells, macroautophagy also plays a defensive role against liver pathogens. Bacteria and viruses are internalized and degraded in lysosomes. However, some pathogens can develop ways to overcome the lysosomal system. For example, infection by the hepatitis C virus has been shown to block macroautophagy, but not at the level of autophagosome formation [129]. Instead, this virus induces ER stress, which promotes formation of a large number of autophagosomes but at the same time blocks their fusion with lysosomes. Blockage of autophagosome formation prevents virus replication, supporting the theory that the virus might use these double-membrane vesicles for assembly, as previously described for other viruses in other cell types. Connections between liver disease and alterations in autophagy are not limited to the role of this pathway in quality control, but instead extend to the function of this catabolic process in the maintenance of the cellular energetic balance. Recent studies support the idea that a defect in macroautophagy could contribute to the pathogenesis of fatty liver [130]. Thus, basal macroautophagy activity has been demonstrated to contribute to the regulation of lipid stores in all cells. Macroautophagy of lipid droplets or macrolipophagy is particularly important in liver as this is the organ that receives the highest content of circulating lipids from lipolysis in the adipose tissue. However, a maintained lipogenic challenge, such as that

induced by high-fat diet, has a negative effect on liver macroautophagy [130]. The inability of the liver under these conditions to reduce the size of the lipid stores through macroautophagic degradation results in abnormal growth of lipid droplets inside hepatocytes, initiating the progression toward non-alcoholic fatty liver. It is also likely that defective macroautophagy as a result of insulin resistance in conditions such as aging may be the basis of fatty liver being associated with the metabolic syndrome of aging. A decrease with age in total rates of protein degradation has been described in liver of almost all organisms analyzed, supporting a negative effect of aging on the intracellular proteolytic systems [131]. The degradation of short-lived proteins, the main substrate of the UPS, is better preserved in old livers, suggesting that the impairment of this system is less pronounced than that of the lysosomal system. In fact, early studies in livers from young and old rodents showed no net differences in proteasome-dependent degradation [132, 133]. Moreover, the analysis of the three different proteolytic activities of the 20S proteasome revealed a decrease in some of its activities but an increase in others, suggesting that rather than dramatic quantitative changes, qualitative changes account for most of the altered degradation of proteasome substrates with age [132–134]. Changes in the 20S proteolytic activities with age may result in part from changes in their subunit composition [135]. Proteomic analyses have revealed the coexistence of different subpopulations of proteasomes as cells age. Defects in ubiquitination steps or in the complex processes that mediate recognition of ubiquitinated proteins by the proteasome might also occur with age, but there is still poor understanding of these changes. Age-related changes in the liver lysosomal system at the morphological level have been extensively reported in the literature. Expansion of the lysosomal vacuolar compartment and accumulation of undigested products inside lysosomes in the form of an autofluorescent pigment known as lipofuscin are often used as typical biomarkers of aging [136]. Accumulation of lipofuscin originates from changes in the proteolytic susceptibility of the intracellular components and from defects in autophagy [137]. A reduction in the induction of macroautophagy in the period in between meals, as well as problems with the clearance of the already-formed AV by lysosomes, seem to be behind the failure of this autophagic pathway with age. Due to the important role of this autophagic pathway in the maintenance of organelle homeostasis, the decline with age in macroautophagy has been proposed to be behind the increased number of dysfunctional mitochondria with age; this organelle is tightly linked to aging [138]. Defective macroautophagy induction results from the negative effect on this pathway of the constitutive signaling through the insulin receptor, even in the absence of insulin, characteristic of the metabolic syndrome of age [139, 140]. Although the molecular basis

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of this defect remains unknown, which prevents from any targeted restorative effort over macroautophagy, certain interventions have proven beneficial in the prevention of the age-dependent macroautophagic decline. Thus, caloric restriction, the only intervention known to slow down aging, and anti-lipolytic drugs preserve macroautophagy activity in livers of old rodents at levels comparable to those in young livers [140, 141]. A decrease in CMA activity has been well characterized in livers of old rodents [142]. The main reason for the CMA decline is a reduction in the lysosomal levels of the LAMP-2A receptor with age. Reduced levels of LAMP-2A are not caused by a transcriptional decrease, defects in translation, or altered trafficking to lysosome. Instead, low lysosomal levels of LAMP-2A originate from increased instability of this protein once it reaches the lysosomes [143]. Changes in the lipid composition of the lysosomal membrane seem to be behind the reduced stability of LAMP-2A. This decrease is initially compensated for by an increase in the amount of lysosomal chaperone that promotes translocation across the membrane but at late stages this compensation is not enough and CMA activity is severely impaired. A better understanding of the consequences of the decrease in CMA activity with age in liver was obtained recently through studies in a transgenic mouse model in which the decrease in levels of LAMP-2A in liver was prevented. Old mice with preserved CMA activity contain lower levels of oxidized and aggregate proteins than the livers of their wild-type littermates [144]. They also show improved ability to respond to stress and preserved liver function until late in life. These studies support the theory that a decline in CMA activity with age could be responsible, at least in part, for the accumulation of damaged proteins with age, lower resistance to stress, and functional decline in this organ [144]. On a positive note, restoration of only one of the different proteolytic systems seems enough to induce a major improvement, probably because of the existing cross-talk among these systems. As more insights are gained into the reasons for the functional decrease in other proteolytic pathways and as further attempts to correct these defects are undertaken, it may be possible to prevent other age-related changes in liver and utilize them in the fight against diseases of the aging liver.

CONCLUSION The proteolytic systems play a pivotal role in the maintenance of liver homeostasis and its energetic balance, and in the fight against intracellular and extracellular aggressors. The different mechanisms for protein degradation can no longer be considered as independent units. Increasing numbers of reports support the existence of continuous cross-talk among the different autophagic pathways and with the proteasome system. Compensation of one system for another is not complete but it is

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often enough to preserve cellular homeostasis at least under basal non-stressed conditions. Failure of the proteolytic systems has been identified as the basis of the pathogenesis of different liver diseases. The recent advances in our understanding of the molecular mechanisms behind the different intracellular proteolytic systems, their regulation, and the cellular consequences of their blockage or altered function could set the basis for novel therapeutic approaches for the treatment of liver diseases.

ACKNOWLEDGMENTS We thank the numerous colleagues in the field of autophagy who through their animated discussions have helped shape this chapter. We are in particular debt to the other members of our group for their insightful comments. Work in our laboratory is supported by NIH/NIA (AG021904, AG031782), NIH/NIKDD (DK041918), NIH/NINDS (NS038370), and a Glenn Foundation Award.

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13

Peroxisome Assembly, Degradation, and Disease Peter K. Kim Program in Cell Biology, The Hospital for Sick Children, Toronto, Ontario, Canada

INTRODUCTION Peroxisomes are single-membrane-bound organelles that are ubiquitous to most eukaryotic cells (Figure 13.1). Since their initial description by Rhodin in 1954 and biochemical characterization by de Duve in 1966, much more has become known about their physiological role in various organisms [1, 2]. Enclosed by a single membrane, peroxisomes contain a diverse array of enzymes involved in both lipid metabolism and biosynthesis. In all organisms, both alpha and beta oxidation of fatty acids occurs in peroxisomes. In yeast and plants, beta oxidation occurs exclusively in peroxisomes, whereas in mammalian cells beta oxidation occurs in both peroxisomes and mitochondria. However, alpha oxidation (the metabolism of branch-chain fatty acids) is believed to occur exclusively in peroxisomes in all organisms [3]. Peroxisomes are also involved in the biosynthesis of a number of biological products, such as penicillin in film fungi, and phytohormones in plants. In mammals, peroxisomes are required for the biosynthesis of bile acid, cholesterol, and plasmalogen (an essential ether lipid required for the development of white matter in the brain and myelin sheath) [4]. Another key function of peroxisomes is the metabolism of reactive oxygen species produced by the cell [5].

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

The importance of peroxisomes in both lipid metabolism and the biosynthesis of lipid-related products is seen in the various inheritable diseases that are associated with peroxisome function and formation. These peroxisomal genetic disorders are categorized into two general groups: single metabolic enzyme disorder, which affects only a single peroxisome function [1], and peroxisome biogenesis disorder (PBD), which results from a defect in peroxisome formation [6]. The characterization of these defects has aided in the identification of both enzymatic components involved in peroxisomal function and factors involved in peroxisome assembly. The peroxisomal factors involved in peroxisome biogenesis are called peroxins (PEXs). The characterization of PEX has been greatly assisted by both genetic and biochemical studies of peroxisome biogenesis in yeast and Chinese hamster ovary (CHO) mutant cells [7, 8]. In mammalian cells, 16 PEXs have been identified, whereas there are at least 32 and 22 in yeasts and plants respectively (see Table 13.1) [9]. Most research on peroxisomes has focused on its function and formation, mainly due to the growing number of genetic diseases involving function and formation. Recently, however, there has been a growing interest in peroxisome degradation, mainly due to its relationship to

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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(a)

(b)

Figure 13.1 Peroxisomes in mammalian cells. (a) Live cell confocol image of COS7 cells expressing a peroxisome marker, RFP-skl (light areas) and an ER marker (shaded). (b) An electron micrograph of a hepatocyte from rat liver. P, peroxisome; M, mitochondria; ER, endoplasmic reticulum. Note the close proximity of peroxisomes to the ER. Reproduced courtesy of Rachid Sougrat, Cell Biology and Metabolism Program at Eunice Kennedy Shriver National Institute of Child Health and Human Development

the degradation pathway, autophagy. Thus far, about 30 genes that are involved in the degradation of peroxisomes (a process called pexophagy) have been identified [2, 10]. Studies have shown that pexophagy is closely related to the autophagy pathway. This chapter will focus largely on our understanding of peroxisomes in the mammalian system. In particular, the chapter will cover the formation of the peroxisomal membrane; the import of both its membrane and matrix (peroxisomal lumen) proteins; and the degradation of peroxisomes. However, in order to appreciate the importance of peroxisome regulation (formation and degradation), the function of and diseases related to peroxisomes will be first discussed.

PEROXISOME FUNCTION AND DISORDER The incidence of all peroxisomal disorders is in excess of 1 : 20 000 individuals. PBD, also known as Zellweger spectrum disorder, is the most severe form of all the peroxisomal disorders due to the organism’s inability to form functional peroxisomes. PBD, which is estimated to occur in 1 in 50 000–100 000 live births, has a life expectancy of one to two years [11]. There is a wide range of symptoms associated with peroxisomal disorders. The most common physiological characteristics of patients with peroxisome disorder include: failure to thrive, enlarged liver, abnormalities in liver enzymes, facial abnormalities, degradation of motor skills, and mental retardation. The current knowledge about peroxisomal function has been fueled by the various disorders associated with peroxisomes. Therefore, each known function of peroxisomes will be discussed in relation to its associated disease.

Alpha oxidation Phytanic acid (3,7,11,15-tetramethylhexadecanioc acid) is a branched-chain fatty acid. This fatty acid is found in almost all animals including humans, however it is not endogenously synthesized in animals but rather is a constituent of their diet [12]. Phytanic acid in animals originates mainly from plants as phytol (3,7,11,15tetramethylhexadec-trans-2 ene-1-ol), which differs from phytanic acid by a double bond at its first carbon and an alcohol group instead of a carboxyl group. Phytol, found in the chlorophyll of plants, cannot be digested by mammalians until it is first liberated from chlorophyll by bacteria found in the rumen of ruminant animals [13]. Animals can efficiently absorb both free phytol and phytanic acid. Phytol is converted to phytanic acid in animals. The methyl-branch group on the third carbon prevents the metabolism of phytanic acid by beta oxidation. In order to initiate the metabolism of phytanic acid, the first alpha carbon unit is removed as formic acid (HCO3 H), and rapidly converted to carbon dioxide (CO2 ). Hence this mechanism is called alpha oxidation. The resulting product, pristanal, is converted to pristanic acid, which is further metabolized via the beta-oxidation pathway. Dysfunction in alpha oxidation in humans leads to a genetic disease called Refsum disease [3, 6]. This is characterized by an increase in phytanic acid in the plasma. It results from a non-functional mutation in the alpha-oxidation enzyme phytanoyl-CoA hydroxylase (PhyH) [14, 15], or in PEX7, a protein required to import peroxisomal targeting sequence 2 (PTS2) proteins such as PhyH [16]. Compared to other peroxisomal diseases, the onset of symptoms occurs later in children. Some clinical features include retinitis pigmentosa, chronic polyneuropathy, and cerebellar signs. Refsum is also one of the few peroxisomal disorders for which a therapy has been

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Table 13.1 Peroxin proteins Gene

Y/M

Function in biogenesis

Molecular function

Matrix protein import Matrix protein import PMP targeting; de novo formation Matrix protein import Matrix protein import Matrix protein import Matrix protein import Matrix protein import Matrix protein import Matrix protein import Proliferation Matrix protein import Matrix protein import Matrix protein import Matrix protein import PMP-targeting; proliferation De novo formation Matrix protein import Matrix protein import PMP-targeting; de novo formation Matrix protein import Matrix protein import Matrix protein import Proliferation Proliferation Proliferation Matrix protein import Proliferation Proliferation Proliferation Proliferation Proliferation Proliferation

Dislocation of PEX5 ? Membrane anchor of PEX19 Mono-ubiquitination of PEX5 PTS1-receptor/PEX7 accessory protein ATP-dependent dislocation of PEX5p PTS2-receptor Connects docking and translocon (?) complex ? PEX5 recycling Elongation of peroxisomes PEX5 recycling Docking complex Docking complex Membrane anchor of Pex1p/Pex6p

PEX PEX PEX PEX PEX PEX PEX PEX PEX PEX PEX PEX PEX PEX PEX

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15

Y/M Y/M Y/M Y/M Y/M Y/M Y/M Y Y Y/M Y/M Y/M Y/M Y/M Y

PEX PEX PEX PEX PEX PEX PEX PEX PEX PEX PEX PEX PEX PEX PEX PEX PEX

16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32

Y/M Y Y Y/M Y Y Y Y Y Y M Y Y Y/M Y Y/M Y

Class II mPTS importor (?) Docking complex PTS2-co-receptor Class II mPTS receptor and chaperone PTS2-co-receptor PTS2-co-receptor in yeast Membrane anchor of PEX4 Growth regulation Separation of peroxisomes Elongation of peroxisomes Membrane anchor of PEX1/PEX6 Elongation of peroxisomes Separation of peroxisomes Separation of peroxisomes Growth regulation Growth regulation Growth regulation

Y, yeast; M, mammals.

developed. The current therapy calls for controlling or eliminating phytanic acid from the diet.

Beta oxidation Beta oxidation is the process of metabolizing fatty acid to acetyl-CoA for ATP synthesis. Peroxisomes are the sole site of beta oxidation in plants and yeast. However, beta oxidation occurs in both mitochondria and peroxisomes in mammalian cells [17]. Both long and medium unsaturated and saturated fatty acids are metabolized in both organelles, with the mitochondria taking the lion’s share of this task. However, there is a subset of lipids that can only be beta-oxidized in peroxisomes, which include very-long-chain fatty acids, long-chain dicarboxylic acids, leukotriens, prostaglandins, iosprenoid-derived fatty acids, and pristainic acid (byproduct of alpha oxidation). The steps involved in beta oxidation in peroxisomes are similar to those found in mitochondria; however, the specific enzymes involved are different. Thus far, there are five different defects in peroxisomes that lead to beta oxidation dysfunction [1]. These are

X-linked adrenoleukodystorphy, acyl-CoA oxidase deficiency, D-bifunctional protein deficiency, sterol carrier protein X deficiency, and 2-methylacyl-CoA racemase deficiency. X-linked adrenoleukodystorphy is the most common of all single enzyme peroxisomal disorders, followed by D-bifunctional protein deficiency. The build-up of fatty acids in the central nervous system, liver, kidneys, and plasma causes several different types of symptoms depending on the severity of the mutation. The most common symptoms are behavioral, cognitive, and neurological deterioration.

Bile acid synthesis Cholesterol is converted to bile acids by multiple pathways [18]. All of these involve a large number of steps in several different subcellular locations. Several pathways involve the beta-oxidation steps found in peroxisomes. In one main pathway, peroxisomes are necessary to convert C27-bile acid intermediates to the mature C24-bile acid. The importance of peroxisomes in bile acid biosynthesis is seen in the increase of bile acid intermediate in the

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plasma and urine from children with PBD. In addition, a number of single-enzyme-deficiency peroxisomal disorders have also been reported. The consequences of bile acid abnormalities are believed to contribute to both liver disease and neuropathy.

Ether lipid biogenesis Mammalian ether phospholipids contain a single alkyl chain at the sn-1 position linked to the glycerol backbone via an ether group. The most common ether phospholipids are those containing vinyl ethers with either a phosphoethanolamine or phosphocholine head group. These vinyl-ether-linked ether lipids are collectively called plasmalogens. Although plasmalogens are minor constituents of phospholipids, they are highly concentrated in the heart, nervous tissues, testes, and kidneys [19]. In the heart, approximately 50% of phosphatidylcholines are plasmalogens. Plasmalogen is essential for the formation and function of gray/white matter and myelin. It is also necessary for the formation of sperm in male mice and the lens in the eye, and is a precursor for platelet-activating factor [20]. Plasmalogen is synthesized in concert between peroxisomes and the endoplasmic reticulum (ER) [20]. The initial steps occur in the peroxisomes and the process is then completed in the ER. The absence of plasmalogens in humans has been identified with a deficiency or mutation in either of these two enzymes.

PEROXISOME BIOGENESIS Since its first discovery, the origin of the peroxisomal membrane has been the subject of a much-contested debate. Based on electromicrographs of hepatic cells, peroxisomes were reported to be associated with the ER [21], suggesting the involvement of ER as the origin of peroxisomes. This view was soon smudged by the authoritative review by Lazarow and Fujiki [22], who suggested that peroxisomes are autonomous organelles similar to mitochondria and chloroplasts. This postulation was largely based on the evidence that many of the peroxisomal proteins (membrane and matrix) are synthesized in the cytosol and imported directly into peroxisomes. Based on these observations, Lazarow and Fujiki suggested that peroxisomes are autonomous organelles that propagate from pre-existing peroxisomes by growth and division. This view of the peroxisome as an autonomous organelle came under question due to a number of key observations. First, a number of peroxisomal proteins were observed in the ER of yeast, and in a peroxisomal–endoplasmic reticulum intermediate structure called pER in arabidopsis and mouse cells [23–25]. Gene complementation in peroxisomeless mutant cells also resulted in peroxisome formation, thus suggesting that peroxisome can be formed

de novo [26, 27]. Mutations in Pex3p and Pex9 in yeast and in PEX3, PEX19, and PEX16 in mammalian cells result in cells without any detectable peroxisomal structures. However, complementing the cells with the wild-type gene results in the formation of peroxisomes in a matter of days [26, 27]. These observations lead opponents of the autonomous model to suggest that peroxisomes can be formed de novo from the ER. Indeed, time-lapse imaging of live cells, along with biochemical approaches, has demonstrated that peroxisomes are indeed formed de novo from the ER [28–31]. Both Pex3p in yeast and PEX16 in mammalian cells are targeted initially to the ER before being routed to peroxisomes, suggesting that the ER is the membrane source of peroxisomes. Due to this evidence, peroxisomes are now considered to be semi-autonomous organelles of the secretory pathways. It is generally accepted that peroxisomes are formed by two mechanisms: de novo biogenesis from the ER; and fission of pre-existing peroxisomes (Figure 13.2) [32]. Of these two mechanisms, much more is known about the latter. Peroxisome fission involves three basic steps: elongation, constriction, and division [2, 32]. In the mammalian cells, the PEX11 genes mediate the elongation of peroxisomes. There are three subtypes of the PEX11 genes (˛, ˇ, and ), which are located on different chromosomes. These integral membrane proteins have been proposed to regulate peroxisome division under both induced and basal conditions [33–35]. PEX11α, and PEX11γ proteins are mainly expressed in the liver, whereas PEX1β is ubiquitously expressed. Two proteins, the dynamin-like protein 1 (DLP1) and Fission1 (Fis1), mediate fission of the elongated peroxisome fission. Gene silencing and dominant negative mutant studies have shown that DLP1 is required for peroxisome division [36], while Fis1, a tail-anchor protein, is required to recruit DLP1 onto the peroxisome membrane [37]. The component involved in the constriction of elongated peroxisome is not yet known. The de novo biogenesis of peroxisomes is less well understood. Peroxisomes are formed de novo from the ER; a small number of PEX proteins are recruited to the ER and localize to a subcompartment called the preperoxisomal reticulum. In S. cereviasiae, Pex3p is one of the PEXs that is initially targeted to the pre-peroxisomal reticulum [28–30]. Pex3p is a docking protein for the peroxisomal membrane protein (PMP) receptor Pex19p; thus targeting Pex3p to the pre-peroxisomal reticulum is thought to allow the recruitment of PMPs to form the peroxisomal membrane [28–30]. In contrast, mammalian homologue of Pex3p, PEX3, cannot target to the ER by itself [31, 38]. Instead it is recruited to the ER by another PMP called PEX16, which is directly targeted to the ER via the signal recognition particle(SRP)-dependent pathway [31]. The mechanism of the formation and the nature of the peroxisomal vesicle emerging from the ER are yet to be elucidated. The reason for two different mechanisms for peroxisome biogenesis also remains to be discovered. A second

13: PEROXISOME ASSEMBLY, DEGRADATION, AND DISEASE

ER

Fusion?

PMP & Matrix

Maturation of Fusion?

De novo biogenesis

PEX16 PEX3

Fission biogenesis

Maturation

Elongation (PEX11)

Fission (DLP/Fis1)

A third hypothesis is that both mechanisms work in concert with each other [2]. One of the key questions with the autonomous models regards the source of the peroxisomal lipid. In this model, the ER is the source of the membrane required for growth and division of peroxisomes. The pre-peroxisomal membrane emerges from the ER and fuses with pre-existing peroxisomes to allow for their growth and elongation. In the absence of any peroxisomes, these pre-peroxisomal membranes fuse with each other to from mature peroxisomes.

PEROXISOMAL PROTEIN IMPORT

Mature peroxisome

Constriction (?)

195

Figure 13.2 Peroxisome biogenesis. Peroxisomes are formed by two pathways: de novo biogenesis from the ER, and fission of a pre-existing peroxisome. In mammalian cells PEX3 and PEX16 are initially targeted to the ER, where they accumulate in a sub-compartment called the pre-peroxisomal reticulum. A mature peroxisome may form on this pre-peroxisomal reticulum before being released from the ER, or it may be released before it matures. Such pre-peroxisomes mature by importing all the peroxisomal components directly from the cytosol. These pre-peroxisomal vesicles may also fuse with each other or with peroxisomes formed by fission. Peroxisome fission is a three-step process of elongation, constriction, and fission.

possible hypothesis is that both mechanisms exist and are crucial for maintaining peroxisomes within the cell, but they are activated by different metabolic signals. Based on this model, the constitutive formation of peroxisomes may be served by de novo biogenesis, whereas the rapid peroxisome proliferation is mediated by peroxisome fission. The peroxisomes in rodent hepatic cells can respond rapidly to metabolic need by doubling and in some cases quadrupling the number of peroxisomes within days. This rapid proliferation of peroxisomes is mediated by the activation of peroxisome proliferator-activated receptor α (PPARα) [39]. PPARα is a member of a family of ligand-dependent nuclear transcription factors that regulate lipid metabolism. One gene that is upregulated by PPARα activation is PEX11 [40]. Therefore it is possible that peroxisome fission mainly occurs during rapid peroxisome proliferation, while de novo biogenesis is the constitutive formation of peroxisome during cell growth.

The peroxisome is unique among single-membrane-bound organelles in that it is able to import most of its protein components directly from the cytosol, yet like other single-membrane-bound organelles it has its origin in the ER. Similar to the targeting of mitochondrial proteins, both matrix and membrane proteins are targeted post-translationally to the membrane. This targeting is mediated by cytosolic receptors that recognize and bind a specific sequence on the peroxisomal protein and deliver it to the peroxisome. However, unlike mitochondria, peroxisomes do not have a stable translocon complex. Furthermore, the mechanism of protein translocation across the lipid bilayer may differ between the peroxisomal membrane and matrix proteins.

Targeting of peroxisomal membrane protein There are two classes of PMP. The first group, class I PMPs, is targeted post-translationally to peroxisomes and requires both PEX3 and PEX19 (Figure 13.3). Genetic studies of these three proteins show that mutation of these components results in the absence of any detectable peroxisomal membrane. PEX19 acts as the chaperone/receptor, which recognizes and binds the membrane peroxisome targeting sequence (mPTS). No consensus sequence exists for mPTS, however it usually consists of a putative transmembrane domain and some flanking sequence. The PEX19–PMP complex is recruited to the peroxisomal membrane by the docking of PEX19 with PEX3 [38]. The mechanism of PMP insertion into the bilayer is not yet known; it is generally believed however that PEX19 is recycled after the insertion of its cargo [41]. The majority of the PMPs are class I PMPs. The class II mPTS targeting is not dependent on PEX3 and PEX19, but rather is targeted initially to the ER before being routed to peroxisomes [28–31]. However, the mechanism of targeting of the class II mPTS is still unknown. Some evidence suggests that PEX16 is inserted into the ER in a co-translational fashion, presumably via

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PMP

Cargo binding 19 PMP

19 Docking

19 3

PMP

Translocation Release PMP Cytosol

16

Matrix

Figure 13.3 mPTS1 import. mPTS1 peroxisomal membrane proteins (PMPs) are targeted to peroxisomes in a three-step process. Cargo binding: PMPs are synthesized on free ribosomes and recognized and bound by PEX19. The docking of a PMP–PEX19 complex on a peroxisomal membrane is mediated by the interaction of PEX19 with PEX3 on the peroxisomal membrane. PEX19 is farneslyated, which may aid in its binding to membrane. PMPs are translocated into the lipid bilayer and PEX19 is released back to the cytosol

proteins exhibit PTS1, which consists of a tripeptide sequence (S/A/C)(K/R/H)(L/M) at the extreme C-terminal end of the protein [42, 43]. PTS1 is recognized and bound by the cytosolic receptor PEX5. PEX5 consists of two domains: a helix bundle at the N-terminus, and seven tetratricopeptide repeats (TPRs) at the C-terminus half of the protein. The TPRs are involved in binding PTS1, while the helix bundle is involved in binding to a docking complex on peroxisomes. The less common PTS2 is a nona-peptide with the consensus sequence (R/K)(L/V/I)X5 (H/Q)-(L/A) located on the N-terminus of the protein upon import into the matrix [44]. The cytosolic receptor for PTS2 is PEX7, which contains multiple WD40 domains involved in the binding to PTS2. Unlike PEX5, PEX7 requires accessory proteins to be able to target to peroxisomes. The accessory proteins for PEX7 for S. cerevisiae are PEX18 and PEX21. In mammalian cells, the PEX7 accessory protein is the spliced isoform of PEX5 called PEX5L, a longer form of PEX5, while in plants, PEX7 binds directly to PEX5. Since PEX5L still maintains its binding affinity to PTS1 proteins, it is believed that the import of PTS2 in both mammals and plants is coupled to the import of PTS1 proteins [2].

Docking

the SRP-dependent pathway, and that PEX16 acts as a component to recruit PEX3 onto either the ER or the peroxisome membrane [31].

The docking complex on the peroxisomal membrane consists of two proteins, PEX13 and PEX14. These transmembrane proteins form a complex that has several binding sites to both cytosolic receptors, suggesting a dynamic step-wise binding of the receptor–cargo complex. PEX14 is believed to contain the initial binding site for the receptor–cargo complex as it has a higher binding affinity than PEX13 [45].

Matrix protein targeting

Translocation and release

Unlike the import of ER, and mitochondria proteins, peroxisomal matrix proteins translocate across the lipid bilayer as fully-folded proteins, and in some cases as oligomeric complex [42]. Therefore, the mechanism of matrix protein import is more related to the nucleus, except that it does not have a stable translocon or channel. The import of matrix proteins can be divided into a four-step process: (i) cargo binding; (ii) docking; (iii) translocation and release; and (iv) receptor recycling (Figure 13.4) [2, 42].

The precise mechanism of matrix protein translocation and release across the peroxisomal membrane is not known. There are two hypotheses on how matrix proteins are translocated into the matrix: the Shuttle hypothesis and the Extended Shuttle hypothesis [46, 47]. The main difference between the two hypotheses is the location of the receptor protein (PEX5 and PEX7) in the peroxisomes. The Shuttle hypothesis suggests that the receptors are embedded in the membrane bilayer in topology such that the cargo can be released into the matrix. In the Extended Shuttle hypothesis, whole receptors enter into the matrix along with the cargo. Similarly, there are several hypotheses describing how folded proteins and protein complexes in particular are translocated across the peroxisomal membrane. One such postulation is the Transient Pore hypothesis,

Cargo binding Peroxisomal matrix proteins contain one of two targeting signals called PTS1 or PTS2. Most peroxisomal matrix

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197

PTS1

1) Cargo binding

PTS2

PTS2 5

7

5L 5

PTS1

7

UB 2) Docking

5L

4) Receptor recycling

UB 1

4

6

UB

Cytosol

22 2

14 13

12 10

5

26 Matrix

PTS1 PTS2

5 7

5L

3) Translocation & release

Figure 13.4 Matrix protein import. The import of matrix protein is divided into a four-step process. (i) Cargo binding: PTS1 proteins are bound by the cytosolic receptor PEX5, and PTS2 by PEX7. (ii) Docking: PEX14–PEX13 complex mediates the docking of the PTS-receptor complex onto the peroxisomal membrane. (iii) Translocation and release: the PTS–receptor complex is translocated across the lipid bilayer and the matrix protein is released from the receptor. (iv) Receptor recycling: the receptor is then recycled back into the cytosol by a retro-translocation mechanism. This involves the ubiquitination of the receptor on the membrane and then removal by the AAA–ATPase complex (PEX1–PEX6). PTS, peroxisome targeting sequence

which suggests that large pores rapidly assemble and disassemble based upon the need for the pores [48]. Another hypothesis is the Complex Disassembling model, where large complexes are disassembled before being imported through translocons which are large enough to import monomeric folded proteins but not oligomers [48]. The third hypothesis is the Invagination model, where proteins docked on the peroxisomal membrane are internalized in a similar mechanism as seen for multivesicular bodies in endosomes [49].

which is anchored to the peroxisomal membrane through its interaction with PEX22, ubiquitinates PEX5 with a single ubiquitin moiety [51]. The mono-ubiquitinated PEX5 is then removed from the peroxisomal membrane by the AAA ATPase (PEX1 and PEX6), which is then de-ubiquitinated by a yet-to-be-determined process [52, 53]. PEX5 can also be removed from the peroxisomal membrane via degradation by the ubiquitin proteasome system. For PEX5 degradation, PEX1 and PEX6 remove PEX5 that has been poly-ubiquitinated by Ubc4 from the membrane to the cytosol, where it is degraded by proteasome [54].

Receptor recycling After cargo release, both PEX5 and PEX7 are recycled back to the cytosol. This process requires a RING-finger protein complex, an E2-like protein, and an AAA–ATPase complex. The exact function of the RING-finger complex is not known but a mutation in any one of the proteins that makes up the three RING-finger protein complexes (PEX2, PEX10, and PEX12) results in the accumulation of PEX5 in peroxisomes, suggesting its role in receptor recycling [50]. The second component required in the removal of the receptors is an E2-like protein, PEX4. PEX4,

PEROXISOME DEGRADATION: PEXOPHAGY Peroxisome numbers are maintained by both biogenesis and degradation of peroxisomes. An autophagy-related process called pexophagy degrades peroxisomes. Like autophagy, there are two modes of pexophagy: macropexophagy and micropexophagy [10]. Macropexophagy occurs in all the eukaryotic cells that have been examined, including human; whereas micropexophagy has only been

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observed in two fungi species: P. pastoris and Aspergillus nidulans [10]. Therefore, only marcopexophagy will be discussed in this section. Macropexophagy is a substrate-selective form of macroautophagy. Although peroxisomes may be randomly degradated by bulk sequesteration of the cytosol via macroautophagy, under conditions of excess peroxisomes they are degraded by the peroxisome selective macropexophagy [55]. As in macroautophagy, in macropexophagy a double-membrane structure wraps around mature peroxisomes to form a new structure called pexophagosome. Pexophagosomes then fuse with lysosomes to degrade the sequestered peroxisomes. Since the formation of the pexophagosome is thought to be similar to the formation of autophagosome (which is discussed in Chapter 12), the mechanism of pexophagosomes will not be discussed here. Instead, this section will focus on the mechanism of peroxisome selection for degradation. The majority of the knowledge on the molecular mechanism of peroxisome selectivity comes mainly from the work done on H. polymorpha. In this yeast, there are two PEXs that appear to be involved in signaling peroxisomes for pexophagy; they are PEX3 (a protein important for PMP import) and PEX14 (a protein involved in the docking of matrix proteins). One of the initial steps in designating peroxisomes for pexophagy is the removal and degradation of PEX3. By a yet-to-be-determined mechanism, PEX3 is removed from the membrane and degraded in the cytosol [56]. PEX14, on the other hand, is not removed, but rather may be required to recruit specific autophagy factors such as Atg11 and Atg30 onto peroxisomes [57, 58]. The recruitment of the autophagy factors Atg11 and Atg30 is then believed to target the peroxisomes to autophagosomes in order to form pexophagosomes. The mechanism by which PEX14 recruits the autophagosomal factors, however, is not known. One potential clue appears in the recent findings in mammalian cells that ubiquitination of peroxisome proteins may promote its selection for degradation. In mammalian cells ubiquitination of PMPs appears to target peroxisomes to pexophagosomes for degradation. This targeting is mediated by the recruitment of a ubiquitin-binding protein, p62, onto peroxisome by the ubiquitination of PMPs. p62 (also known as Sequestosome 1) then targets ubiquitinated peroxisomes to newly-forming nascent autophagosomes through its interaction with the autophagosomes factor LC3. This mechanism is similar to the targeting of inclusion bodies to autophagosomes [59, 60].

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14

Gap and Tight Junctions in Liver: Composition, Regulation, and Function Takashi Kojima1, Norimasa Sawada1, Hiroshi Yamaguchi1, Alfredo G. Fort2 and David C. Spray2 1 Department

of Pathology, Sapporo Medical University School of Medicine, Sapporo, Japan 2 Department of Neuroscience, Albert Einstein College of Medicine, Bronx, New York, NY, USA

STRUCTURE OF JUNCTIONS IN THE LIVER Coordinated function of the liver depends on both longand short-range signaling among the numerous cell types that make up this tissue: hepatocytes, Ito cells, cholangiocytes, and endothelial cells. Such signaling is accomplished by hormones and transmitters moving through extracellular space, as well as by direct intercellular diffusion of ions and messenger molecules. Elaborate junctional complexes decorate the appositional surfaces of hepatocytes, separating apical from basolateral membrane domains and directly linking the cytoplasm of one cell with that of another. The junctional types that are the subject of this review are tight junctions or zonula occludens (ZOs), which separate extracellular bulk fluid The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

phases and maintain the segregation between apical and basolateral domains of the plasma membrane, and gap junctions or macula communicans, which function to allow direct intercellular communication from one cell to its neighbors. This chapter is an update that integrates new findings on both junctional types; for more detailed citations on gap and tight junctions, the reader is referred to earlier editions [1, 2]. Thin sections and freeze-fracture images of the junctions between dissociated hepatocytes reveal structures that are similar to those seen in vivo (Figure 14.1). In vertebrates, tight junctions play a central role in regulating the movement of solutes, ions, and water through the extracellular space in epithelial or endothelial sheets. In hepatocytes, tight junctions are found surrounding the bile canaliculi, where they seal the paracellular spaces between hepatocytes and maintain cellular polarity. By

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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(a)

(b)

(c)

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Figure 14.1 Fine structure of appositional membranes between rat hepatocytes. (a,c) Freeze-fracture; (b,d) thin sections. (a) Below the bile canaliculi, tight junction webs seen in this freeze-fracture electron micrograph seal off the canalicular surfaces from the rest of the appositional membrane. (b) Thin-section micrograph illustrates tight junctions (Tj) sealing the canaliculus (arrowhead) and a large gap junction (Gj) below it. Freeze-fracture (c) and thin-section (d) views show larger gap junctions below and in isolation from tight junctional strands. Bars = 0.1 µm Modified from [36], with permission

contrast, gap junctions control cell-to-cell movement of solutes, ions, and water, providing an intercellular pathway not accessible to dilution by extracellular fluid. In the hepatic acinus, bile flow in hepatocyte canaliculi is thought to require the organized and periodic contraction of bile canaliculi, where cell-to-cell spread of contraction occurs through signaling via gap junction channels. In the perfused rat liver, bile secretion induced by glucagon or vasopressin has been shown to be modulated by gap junction communication. Gap junctions occupy a large fraction, as much as 3%, of the total surface area of hepatocytes. In stained thin sections of liver tissue and of hepatocyte cell pairs, gap junctions are recognized as septilaminar linear

membrane appositions separated by an extracellular gap (Figure 14.1b,d). The seven lamina are the transparent extracellular gap sandwiched on either side by each cell’s membrane, consisting of two stained and thus electron-opaque lipid head groups on each side of the electron-lucent membrane interior. The overall thickness of this double membrane specialization is approximately 15–18 nm, and the extracellular space in the region of gap junctional contact is approximately 2–4 nm wide. In freeze-fracture images, gap junctions of liver and hepatocyte cell pairs are recognizable as arrays or plaques of approximately 8–9 nm intramembranous particles present in the P-fracture face in vertebrate tissues; complementary pits appear on the E-fracture face. These plaques are generally round or oval and can be quite large in hepatocytes (Figure 14.1c,d), commonly exceeding 1 µm in diameter and containing more than 10 000 particles. The particles seen in freeze-fracture replicas and the bridges across the extracellular space in thin section are believed to represent channels with hydrophilic walls extending from the cytoplasmic aspect of one cell to that of another (Figure 14.2a). High-resolution ultrastructural studies on isolated liver gap junctions using techniques of X-ray diffraction and low-angle scattering have added detail to the model of the gap junction channel. It is now generally accepted that liver gap junction channels are composed of 12 subunits, 6 of which are contributed by each cell; the assemblage of the 6 subunits contributed by each cell have been termed a connexon or hemi-channel (Figure 14.2a). The subunits are radially symmetrical around the central pore, and each is believed to be tilted slightly relative to the plane of the membrane. Cryoelectron microscopy and computer modeling have provided evidence that the formation of rat liver gap junctions requires a 30◦ rotation between hemi-channels for proper docking. The ultrastructure of tight junctions is quite different from gap junctions. In freeze-fracture, tight junctions appear as a set of continuous, anastomosing strands in the P-fracture face (Figure 14.1a), with complementary grooves in the E-face and in thin section appearing as very close membrane appositions. The role of tight junctions is to provide a high-resistance barrier to leakage of water and solutes into and from the bile canaliculus (the resistance to current flow has been measured as approximately 50 k by insertion of a microelectrode into the canalicular space). The number of tight junctional strands encircling the bile canaliculus is normally high, indicating a tight permeability barrier. As a consequence, the composition of the canalicular fluid can be quite different from that of the other pericellular fluid, allowing the accumulation of high concentrations of preferentially secreted organic anions within this intercellular compartment.

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Figure 14.2 Molecular components of liver gap junction channels. (a) Each gap junction channel consists of two connexons or hemi-channels, each containing six protein subunits (connexins, one of which is shaded in cross section). These connexons pair across extracellular space, forming the walls of the gap junction channel. (b) Membrane topologies of the two liver gap junction proteins Cx26 and Cx32. Note four transmembrane domains (rectangles), the third of which (filled) is the most amphipathic and is thought to line the pore; extracellular loops with symmetrical sulfhydryl (SH) groups, which are believed to play a role in channel formation; and the cytoplasmic location of amino and carboxy termini. (c) Organization of connexins into connexons and intercellular gap junction channels. Connexin proteins are oligomerized into connexons that are termed homomeric if they have one type of connexin or heteromeric if they contain multiple connexins. Different types of connexon give rise to either homotypic, heterotypic, or heteromeric intercellular channels

MOLECULAR COMPONENTS AND GENES OF GAP AND TIGHT JUNCTIONS Gap junctions The first gap junction protein isolated from detergent solubilized gap junctions displayed an apparent molecular weight of 27 kDa, although a 21 kDa protein was also detected (for historical overview, see [1]). Cloning of the cDNA encoding the more slowly migrating protein indicated that its predicted molecular weight is about 32 000 Da, and this protein is generally referred to as connexin32 (Cx32; in an alternative nomenclature this gap junction protein is referred to as β1 and its gene as Gjb1 in rodents and as GJB1 in man). The cDNA encoding the 21 kDa protein was subsequently cloned and found to encode a protein of predicted molecular weight of

26 000 Da; this connexin is now termed Cx26 (or β2 and its gene as Gjb2 or GJB2 ). Cx32 and Cx26, the components of hepatocyte gap junctions, are also found in a variety of other cell types; moreover, another gap junction protein, Cx43 (or α1; gene Gja1 or GJA1 ) is prominent between other liver cell types, including Ito cells, Kupffer cells, and endothelial cells (for reviews, see [3, 4]). The connexin gene family now comprises at least 20 proteins in vertebrates; connexins are apparently absent in invertebrates, where gap junction channels are formed by a separate gene family encoding innexin proteins [5]. Although vertebrate homologues of innexin proteins have been identified (see [6, 7]), these pannexin proteins are likely involved in formation of non-junctional, rather than junctional, channels (see [8, 9]). Hydropathy plots, confirmed by protease cleavage and epitope mapping studies, indicate that connexins are tetraspan membrane proteins with a very high degree of

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homology in extracellular and transmembrane domains (see Figure 14.2b). Homologous extracellular domains are thought to account for the ability of many connexins to form heterotypic channels (i.e. for the fact that cells expressing one type of connexin can often form gap junctions with cells expressing a different connexin). The greatest dissimilarities among connexins occur in the sequence and length of the carboxyl terminus, which is thought to play a major role in regulation of gap junction channels. It is within this cytoplasmic region of the proteins that most of the potential phosphorylation sites are found and against which most of the useful connexin-specific antibodies have been raised. Moreover, the carboxyl termini of several connexins have been shown to contain binding sites for kinases and scaffolding molecules, so that gap junction channels consist of a protein complex or Nexus [10]. Most cells express more than one type of connexin, leading to the possibility that both homomeric and heteromeric connexons may exist in vivo (Figure 14.2c). The existence of heteromeric connexons in the liver has been supported by biochemical experiments that have fractionated detergent-solubilized gap junctions and electrophysiological properties on hepatocytes isolated from Cx32-deficient and wild-type mice [11]. Experiments examining the movement of ions and dyes between cells coupled by different connexins have revealed that there are connexin-dependent differences in the permeation of intercellular channels. One striking example of possible importance to hepatocytes is that homomeric connexons made of Cx32 are permeable to both cAMP and cGMP, whereas heteromeric connexons composed of Cx32 and Cx26 lose permeability to cAMP but not to cGMP (Figure 14.2c). Characterization of the genomic sequence corresponding to Cx32 defined a gene structure of Gjb1 that is now appreciated as being common to almost all other members of the connexin gene family. The Cx32 coding sequence is included within one open reading frame on a single exon, which is separated from a tiny upstream exon (∼100 bp) by about 6 kb of intervening intronic sequence. Promoter mapping in the human liver-derived HUH7 cell line localized a minimal basal promoter within a 70 bp region immediately upstream of the mRNA start sites, and DNase hypersensitive sites 1.2 kb downstream of the Cx32 open reading frame. More recent experiments performed on MH1C1 rat hepatoma cells have identified positive and negative regulatory domains of the Cx32 promoter and nuclear proteins (HNF1) that bind to them [12]. The other hepatocyte gap junction protein, Cx26, has been proposed to be a tumor suppressor gene, on the basis of subtractive hybridization using normal and malignant human mammary epithelial cells. Cx26 expression in human mammary epithelial cells induced by 12-O-tetradecanoylphorbol-13-acetate (TPA) treatment is controlled at the level of transcriptional modification. Kiang et al. have cloned and sequenced the 5′ portion

of human Cx26 gene, revealing that the promoter region contains six GC boxes, two GT boxes, a TTAAAA box, and a TPA-induced DNase I hypersensitivity region (see [13, 14]), providing a site by which TPA may exert transcriptional control over Cx26. Nevertheless, induction of exogenous Cx26 in neuroblastoma cells by TPA treatment appears to be controlled by post-translational mechanisms [15].

Tight junctions Tight junctions, the most apically located of the intercellular junctional complexes, inhibit solute and water flow through the paracellular space (termed the “barrier” function (Figure 14.3a) (see [16]). They also separate the apical from the basolateral cell surface domains to establish cell polarity (termed the “fence” function) (Figure 14.3a) (see [17]). Recent evidence suggests that tight junctions also participate in signal transduction mechanisms that regulate epithelial cell proliferation, gene expression, differentiation, and morphogenesis (Figure 14.3a) [18]. Tight junctions have a complex molecular composition and the integral membrane components of tight junctions include occludin, the claudin family (see [19]), junctional adhesion molecules (JAMs) [20], coxsackie adenovirus receptor (CAR) [21], the 1G8 antigen of endothelial cells [22], and tricellulin [23]. The roles of these proteins in tight junction function are only just beginning to be clarified in detail (Figure 14.3b). The integral proteins occludin, claudin, and JAM bind to the domains of scaffold protein ZO1, GUK, PDZ1, and PDZ3, respectively (Figure 14.3b) [24]. As with the connexins, the major tight junction components occludin and the claudins are tetraspan proteins with intracellular amino and carboxyl termini. The detailed analyses of the manner of interaction of heterogeneous claudin species within and between tight junction strands suggest that distinct species of claudins are copolymerized linearly to form tight junction strands as homopolymers or heteropolymers and that the claudins interact between each of the paired strands in a homophilic or heterophilic manner, including both other claudins and/or occludin (Figure 14.3b). The claudin family at present consists of 24 members, including several previously identified junction-associated proteins such as RVP1 (claudin-3), CPE-R (claudin-4), TMVCF (claudin-5), and OSP (claudin-11). The claudins are solely responsible for forming tight junction strands and show tissue- and cell-specific expression of individual members (see [19]). Several lines of evidence point to claudins as the basis for the selective size, charge, and conductance properties of the paracellular pathway [24]. The claudins have two extracellular loop domains like connexins and occludin (Figure 14.3c), with the first loop (∼53 residues) twice the length of the second loop. The

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Figure 14.3 Function and molecular components of tight junctions. (a) Three major tight junctional functions (termed “fence,” “barrier,” “signaling”). (b) The major molecular components (left) and the organization of tight junction proteins with respect to binding of domains of ZO1. (c) The structural and functional features of claudins. (d) The known protein binding partners of occludin

first extracellular loop influences the paracellular charge selectivity, the second extracellular loop acts as a receptor for a bacterial toxin, and the C-terminus binds cytoplasmic proteins via a PDZ motif (Figure 14.3c) [25]. Claudins are between 20 and 27 kDa in molecular weight and amino sequences vary most in their C-terminus. Recently it was found that discrete residues within the first extracellular loop of claudin-1 in liver are critical for hepatitis C virus (HCV) entry (Figure 14.3c) [26]. In murine livers, claudin-1, -2, -3, -5, -7, -8, -12, -14 are detected together with occludin, JAM-A, CAR, and tricellulin, and claudin-1, -2, -3 are expressed in the bile canaliculus region of hepatocytes (see [27]). In the rat liver, claudin-2 immuno-localization shows a lobular gradient increasing from periportal to pericentral hepatocytes, whereas claudin-1 and -3 are expressed in the whole liver lobule. Furthermore, claudin-2 expression induces cation-selective channels in tight junctions of epithelial cells, as does expression of claudin-7 and -15.

Occludin, the first-discovered integral membrane protein of tight junctions, is most ubiquitously expressed at the apical-most basolateral membranes, and is the most reliable immunohistochemical marker for tight junctions. By contrast to the diversity of claudin isoforms, only a single occludin transcript has been described, although an alternatively spliced form of occludin (termed occludin 1B) has been reported. Like claudins, occludin is a tetraspan-membrane protein, with both termini in the cytoplasm, although it is twice as heavy (56 kDa) and it is predicted to have equally long extracellular loops (44 residues). Like claudins, occludin also contains PDZ binding domains for scaffold association proteins such as ZO1. JAMs (JAM-A, -B, -C, -4) are immunoglobulin superfamily proteins expressed at cell junctions in epithelial and endothelial cells as well as on the surface of leukocytes, platelets, and erythrocytes [20]. They are important for a variety of cellular processes, including tight

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junction assembly, leukocyte transmigration, platelet activation, angiogenesis, and adenovirus binding. Recently, in HepG2 cells and WIF-B cells which have a hepatic cell polarity, depletion of the integral tight junction protein JAM-A, which directly binds to the cell polarity protein PAR3, was found to inhibit formation of bile canaliculi (see [28, 30]). Tight junctions are formed not only by the integral membrane proteins claudins, occludin, and JAMs but also by many peripheral membrane proteins, including the scaffold PDZ proteins ZO1, ZO2, ZO3, multi-PDZ domain protein-1 (MUPP1), membrane-associated guanylate kinase with inverted orientation (MAGI)1, MAGI2, MAGI3, and cell polarity molecules ASIP/PAR3, PAR6, PALS1, and PALS1-associated tight junction (PATJ), and the non-PDZ-containing proteins, cingulin, symplekin, ZONAB, GEF-H1, aPKC, protein phosphatase 2A (PP2A), Rab3b, Rab13, PTEN, and 7H6 (see [24, 27]). ZO1, ZO2, and ZO3 are members of the membrane-associated guanylate kinase (MAGUK) family of proteins, displaying a characteristic multidomain structure comprising SH3, GUK, and multiple PDZ (PSD95-Dlg-ZO1) domains. ZO1 and ZO2 are also closely associated with polymerization of claudins [31]. Actin filaments have been prominently localized near the cytoplasmic region of tight junctions using electron microscopy (Figure 14.7b). ZO1 and ZO2 bind to actin filaments, suggesting a role of tight junctions in the organization of the actin cytoskeleton (Figure 14.3b).

Interaction between gap and tight junctions In hepatocytes, ZO1 is detected not only in the tight junction zone but also at adherens junction [32], whereas in intestinal epithelial cells bearing well-developed tight junctions, cadherins and ZO1 are clearly segregated into adherens and tight junctions, respectively. It has become established that Cx43 interacts with ZO1 following transfection with Cx43, both in normal fibroblasts and in cardiac myocytes (see [10]). This interaction is direct, through binding of the extreme carboxyl terminus of Cx43 and the second PDZ domain of ZO1. Furthermore, it is well known that small gap junction plaques are associated with tight junction strands in some cell types including hepatocytes (Figure 14.4a,b). In primary cultured rat hepatocytes, Cx32 is partly colocalized with occludin and claudin-1, which form tight junction structures (Figure 14.4c) [33]. In order to examine the roles of gap junctions in regulating the expression and structure of tight junctions, we transfected human Cx32 cDNA into immortalized mouse hepatocytes (CHST8 cells as well as hepatocytes derived from Cx32 null mice) which lack endogenous Cx32 and Cx26. In Cx32 transfectants, induction of tight junction strands and the integral tight

junction proteins occludin and claudins were observed, and small gap junction plaques appeared within the induced tight junction strands [34]. The induced endogenous occludin protein in the transfectants was found to bind to the exogenously expressed Cx32 protein. Furthermore, in the transfectants, upregulation of tight junction-associated protein MAGI1 was observed by cDNA microarray analysis [35]. These results indicate that gap junction and tight junction expression are closely correlated in hepatocytes, and we speculate that through this association gap junction expression may play a crucial role in the establishment of cell polarity via regulation of tight junction proteins. This finding supports previous studies in which gap junctions have been associated with other components of intercellular junctional components. More importantly, however, the demonstration of a direct linkage between occludin and Cx32 and previous studies showing high-affinity interactions between ZO1 and Cx43 indicate that connexins may form selective associations with specific components of adhesive or occluding junctions. More recently, Cx32 has been shown to interact with a PDZ-containing scaffolding protein, Discs Large homolog 1 (Dlgh 1) in the liver [36]. Through binding of connexins to cytoskeletal, adhesion, and signaling proteins, we have proposed that these proteins may promote the aggregation of connexin-specific scaffolds at junctional regions, providing not only intercellular signaling but also sites (the Nexus: 30) where intracellular signaling is transduced.

FUNCTIONS OF GAP JUNCTIONS IN THE LIVER Gap junctions are found between each of the cell populations of the liver, where the type of connexin expressed is cell-type and region specific (Figure 14.5). For example, the cells of the liver capsule, Ito (fat-storing) cells, cholangiocytes, and endothelial cells lining the venules express Cx43 as a major gap junction protein (see [3, 4]), whereas hepatocytes express only Cx26 and/or Cx32. Physiological consequences of the high level of gap junction expression between hepatocytes appear to include signal transfer and growth control. The relay of metabolic signals is important not only because of the anatomical arrangement of the hepatic acini but also because certain phenotypic features of hepatocytes are graded from periportal regions (characterized by the portal triad that includes the terminal portal venule) to pericentral regions (surrounding the terminal hepatic venule). For example, hepatocytes with higher glucogenic activity are found periportally rather than pericentrally. Nevertheless, the strong electrical coupling mediated by gap junctions between hepatocytes (Figure 14.6a,c) serves to equilibrate the membrane potentials among hepatocytes. In isolated rat liver, perfusion with glucagon leads to hyperpolarization of all hepatocytes across the hepatic acinus. However, in the presence of octanol, a gap junction blocker,

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Basal Apical Z-section (c) Figure 14.4 Interaction between gap junctions and tight junctions in primary cultured rat hepatocytes. (a,b) Freeze-fracture; (c) double staining of Cx32 and claudin-1. In freeze-fracture analysis of the adluminal plasma membrane, tight junction strands form well-developed networks (a) and many small gap junction plaques are observed peripherally or within the tight junction network (b). Cx32 immunoreactive lines are observed on the most subapical plasma membrane at cell borders, while Cx32-positive spots can be observed on the basolateral membrane (c). Claudin-1 immunoreactive lines are observed on the most subapical plasma membrane of the cell borders and are colocalized with Cx32-immunoreactive lines (arrows in (c))

glucagon-induced hyperpolarization is higher in periportal than in pericentral hepatocytes. Furthermore, heptanol blockade of gap junctions has been shown to abolish metabolic and hemodynamic effects of nerve stimulation within the liver. These findings are consistent with the notion that gap junctions play important functional roles during responses of liver cells to agents affecting the metabolic state of this organ. An unresolved question is the precise identity of the molecule(s) that provide the gap junction-mediated signaling. Gap junction channels are permeable to ions and molecules up to a molecular weight cutoff of about 1 kDa (Figure 14.6a), allowing possible involvement of

both ions and second messengers in intercellular signaling. Imaging studies on hepatocyte pairs demonstrated that injection of Ca2+ into one cell leads to diffusional spread to the adjacent cell [38]. This intercellular spread was blocked by treatment of the cells with the gap junction inhibitor heptanol, indicating that movement is by way of gap junction channels. Intracellularly-injected inositol 1,4,5-triphosphate (IP3 ) was also shown to lead to localized rises in intracellular Ca2+ in the injected cell and in its coupled partner. Because the Ca2+ rise in the recipient cell can occur far from the junctional region, diffusion is likely to be of IP3 or its metabolite, but not of Ca2+ itself. This study and others in which

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entral Vein

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Figure 14.5 Communication compartments in liver. (a) Cx32 staining in rat liver. (b) Cx26 staining in rat liver (C: central vein, P: periportal vein). Cx32-positive spots are constant across the acinar unit, while Cx26-positive spots decrease in a gradient from high levels periportally to low levels pericentrally. (c) The major hepatic cell type is the parenchymal cell or hepatocyte, which extends in branched arrays from the portal triad to the central vein. Each hepatocyte is coupled to all adjacent hepatocytes by generally massive gap junction plaques consisting of Cx32 and Cx26. In the rodent liver, Cx32 expression is constant across the acinar unit, while Cx26 expression decreases in a gradient from high levels periportally to low levels pericentrally (graphed schematically in the lower portion of the figure). Other cell types in the liver include endothelial cells, which line the portal vein and sinusoids, as well as fat-storing Ito cells. Both of these cell populations express Cx43, with the most intense staining within the portal triad (distribution graphed below the diagram)

synchronous Ca2+ oscillations have been shown to occur in hepatocyte couplets indicate that intercellular diffusion of Ca2+ and IP3 can lead to regenerative Ca2+ release within hepatocytes. Such regenerative Ca2+ elevations may serve the role of long-range signaling in propagation of canalicular contractions. Intercellular Ca2+ signaling is commonly used by cells to coordinate and regulate a wide range of cellular functions including cell growth and differentiation. Ca2+ waves may also closely contribute to the various functions of liver and have been shown to occur in the intact liver, following vasopressin stimulation. Ca2+ wave signaling among cultured hepatocytes may use two parallel pathways (Figure 14.6d) (see [39]). One of these is the intercellular propagation of Ca2+ waves initiated by the generation of IP3 , which diffuses to adjacent neighboring cells via gap junctional channels and are inhibited by gap junction blockers (Figure 14.6d). The other pathway involves extracellular Ca2+ wave propagation to both adjacent and non-adjacent cells, through the activation of purinergic receptors by ATP released from the stimulated cell, and is inhibited by purinergic receptor blockade (see [39]). Intercellular Ca2+ wave propagation using the multiplets of connected hepatocytes and the perfused liver may be useful as a model in which to study coordination and regulation of the cellular functions via gap junctional channels.

Growth control As in many other cell types, one of the roles that has long been proposed for gap junctions lies in the control of normal tissue growth. Evidence supporting this association includes reduced gap junction expression in hepatomas compared with that in adjacent normal tissue, inhibition of growth of tumors constitutively expressing Cx32, and stimulation of hepatocyte growth by tumor promoters that reduce functional coupling, including TPA, phenobarbital, benzoyl peroxide, DDT, lindane, arachidonic acid, and polyethylene glycols. Furthermore, the liver tumor promoters phenobarbital, PCB, DDT, and clofibrate decrease dye coupling in rat liver slices in vivo. These changes appear to involve aberrant expression of gap junction proteins, as searches for linkage between Cx32 mutations and hepatocarcinogenesis have been largely unsuccessful. Models that have been useful for clarifying the association between gap junction expression and growth control in liver include the regeneration of this tissue following surgical or chemical insults. Following partial hepatectomy, gap junctions between cells in the undamaged liver disappear over a time course of approximately 24 hours. Cells then undergo mitosis as the tissue regenerates and gap junctions reappear, all within 48–72 hours after injury. Electrical measurements from the tissue suggest that junctional conductance changes with

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Figure 14.6 Communication through gap junction channels. (a) Light and epifluorescence photos showing that Lucifer Yellow CH (5% wt/vol) injected iontophoretically into one cell of a pair of hepatocytes passes detectably to a coupled neighboring cell within 30 seconds. (b) Voltage-clamp experiment on a pair of hepatoma cells stably transfected with Cx32. A 50 mV hyperpolarization (V2) of one cell produces an initial current in the other cell (I1), which decreases over a period of seconds. This voltage sensitivity is characteristic of liver gap junction channels. (c) Single gap junction channel openings and closures can be seen in this example of a high-again recording from a poorly coupled pair of SKHep1 cells transfected with rat Cx32. In response to a driving force of 50 mV, channels continuously open and close, as indicated by abrupt transitions of equal size but opposite polarity recorded in each cell’s voltage-clamp circuit. In these initial studies [32], unitary conductance of Cx32 channels was calculated as about 120 pS; more recent studies in N2A cells transfected with human Cx32 or Cx26 revealed channels of about 55 and 135 pS G, which are consistent with conductances recorded from hepatocytes obtained from wild-type and Cx32 knockout mice [11]. (d) Mechanical induced Ca2+ wave propagation in mouse hepatocyte cell line stable transfected with Cx32. Ca2+ is measured in fluorescence of cells loaded Indo-1-AM (10 µM) and the changes in [Ca2+ ]i are indicated by the pseudocolored images. (e) Time course of changes of fluorescence ratios in the cells treated with gap junction blocker (18µ-glycyrrhetinic acid)

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a similar time course, although increased non-junctional resistance partially compensates in electrical coupling measurements. Using immunoblot techniques, the total amount of Cx32 in livers following partial hepatectomy has been found to undergo a similar cycle of decrease that coincides with time of maximal DNA synthesis and then reappearance; Cx26 is regulated in parallel. At times after partial hepatectomy or bile duct ligation when levels of gap junction protein in plasma membranes are reduced to their lowest, Cx26 and Cx32 expression may be regulated by post-transcriptional modification. In the regenerating liver of rats treated with a p38 mitogen-activated protein kinase (MAPK) inhibitor, the disappearance of Cx32 is inhibited without affecting hepatocyte proliferative activity (see [40]). This indicates that cellular proliferation can occur independent of downregulation of gap junction channels. Parallel measurements of steady-state Cx32 mRNA and protein levels and junctional conductance show that partial hepatectomy and acute hepatotoxicity cause similar changes. However, Cx26 mRNA was selectively increased before the onset of S-phase after partial hepatectomy and after bile duct ligation. By means of double immunolabeling using the nucleotide analog bromodeoxyuridine as an S-phase marker, Dermietzel et al. [41] showed a reciprocal correlation between the expression of Cx32 and the mitotic activity of hepatocytes during liver regeneration. The conclusion reached by this work indicated a significant reduction of Cx32 expression in S-phase cells. From these findings it seems reasonable to conclude that quantitative changes in gap junction expression play an important role in the control of proliferation in liver. Studies on chemically induced rat hepatocarcinomas also revealed a dramatic reduction in Cx32 expression under proliferative conditions [42]. It is plausible that the partial loss of gap junctions provides a selective advantage for those preneoplastic liver cells that develop into rapidly proliferating tumor cells. Dissociated hepatocytes undergo changes in gap junctions, expression of gap junction proteins, and electrical coupling that are temporally similar to those following tissue injury in situ; gap junctions initially disappear and then reappear over a time course of days. The disappearance of gap junctions can be delayed by treatment with cAMP or glucagon, or with protein synthesis inhibitors, indicating that mRNA stability may be responsible. The re-expression phase of gap junctions in cell culture is associated with renewed synthesis of gap junction mRNA and recovery of gap junction protein levels and electrotonic coupling. Transcription of tissue-specific mRNA including Cx32 was enhanced when hepatocytes were cultured in hormonally defined medium (HDM) in combination with certain glycosaminoglycans (GAGs) and proteoglycans (PGs). Moreover, Cx32 re-expression in primary cultured rat hepatocytes was dramatically enhanced in medium containing epidermal growth factor (EGF) and 2% dimethyl sulfoxide (DMSO), and Cx26 could also be induced when glucagon was added together with 2%

DMSO. These latter studies suggest that expression of gap junctions in hepatocytes may be closely related to oxidative stress such that oxygen radical scavengers such as DMSO or melatonin might be important inducing substances. In a DMSO-containing long-term primary mouse hepatocyte culture system, expression of Cx26 and Cx32, and functional coupling were also well maintained.

Properties of livers in Cx32 mutant animals A human genetic disease, the X-linked form of Charcot–Marie–Tooth disease (CMTX), involves more than 300 distinct mutations of Cx32 (see [43]). In an attempt to mimic CMTX disease in a rodent model, Cx32-deficient (Cx32 knockout (KO)) mice were generated through homologous recombination [44]. Although these mice did demonstrate progressive demyelination typical of the human disease, they also showed two deficiencies that have not been observed in CMTX patients. First, liver function was compromised, as evidenced by reduced glucose release from intact liver in response to sympathetic nerve stimulation or induction by hormones [44]. In Cx32 KO livers, Cx26 levels in Cx32 KO livers were decreased compared to wild types [44], and electrophysiological properties and permeability to IP3 in Cx32 KO hepatocytes were decreased compared to wild types. Second, proliferation rate of Cx32 KO hepatocytes in vivo was high, and spontaneous and chemically induced liver tumors were reported to be more prevalent in Cx32 KO than wild-type livers [45–47]. Furthermore, when we compared cultured hepatocytes from Cx32 KO and wild-type mice using serum free medium, proliferation rate of Cx32 KO hepatocytes was markedly higher than that of wild type in vitro [48]. Interestingly, however, proliferation rate of Cx32 KO hepatocytes after partial hepatectomy was significantly lower than in wild types [49]. More recent experiments on transgenic mice expressing a dominant-negative mutant of Cx32 have found delayed liver regeneration and an increase of susceptibility to chemical hepatocarcinogen [50]. Moreover, Cx32 dominant-negative mutant transgenic rats are resistant to hepatic damage by chemicals [52]. The Cx32 mutant animal models thus are extraordinarily promising for understanding various functions of liver gap junctions during cell growth and differentiation, including hepatocarcinogenesis.

REGULATION OF COUPLING BETWEEN HEPATOCYTES For detailed citations, see [1, 2]. The extent to which signals spread in the liver depends on the number of open gap junction channels.

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Gap junction channels between hepatocytes are closed by cytoplasmic acidification, cell injury, exposure of the cytoplasmic aspect to very high Ca2+ concentrations, CCl4 , and certain alcohols, and by large voltage gradients imposed across the junctional membrane [37]. Sensitivity of hepatocyte gap junctions to cytoplasmic acidification is low at near-normal pHi , but channel closure becomes profound at acidic pHi values. Strong impermeant acids and buffers do not act on conductance of the junctional membrane when applied to the cell exterior, indicating that the pH gating domain is on the cytoplasmic aspect of the channel. Treatments that elevate cytoplasmic free Ca2+ have long been known to decrease intercellular coupling, but sensitivity of gap junctional membranes to calcium is actually very low (see [38]). In hepatocytes, elevating cytoplasmic calcium to levels adequate to contract the bile canaliculus does not block electrical or dye coupling. Moreover, the gap junction-mediated diffusion of calcium ions demonstrated in hepatocytes limits the role of Ca2+ in closing liver gap junctions to gross tissue insult involving exposure of cytoplasmic aspects of the gap junction to extracellular Ca2+ concentrations. Modulation of signal spread through gap junction protein phosphorylation may be a common type of regulation. Cx32 is phosphorylated on SER233 in the cytoplasmic tail of this protein by cAMP-dependent protein kinase, which is correlated with increased conductance. Numerous other protein kinases are found in liver, including protein kinase C, Ca2+ -calmodulin (CaM)-dependent kinase, as well as several tyrosine kinases (including the insulin growth factor receptor (IGFR) and epidermal growth factor receptor (EGFR)). Ca2+ -CaM-dependent protein kinase II and protein kinase C phosphorylate Cx32 in isolated junctions, producing tryptic fingerprints distinct from those obtained with cAMP-dependent protein kinase, but their relevance has not yet been shown in vivo. Although in other cell types inhibition of junctional communication has been associated with the expression of tyrosine kinases, Cx32 in isolated rat liver gap junctions is not phosphorylated by purified pp60v-src tyrosine kinase or the insulin receptor, and the functional state of exogenously expressed Cx32 channels in Xenopus oocytes is not affected by the expression of pp60v-src. EGF has been shown to stimulate phosphorylation of Cx32 on at least one tyrosine residue, and the EGFR tyrosine kinase directly phosphorylates Cx32. Like other transmembrane proteins, gap junction proteins are synthesized by the translation of mRNA on ribosomes and are co-translationally inserted into the endoplasmic reticulum (ER). The nascent connexin protein must fold correctly and oligomerize with five other subunits to form a hexameric complex termed a connexon. For most connexins this process occurs after exit from the ER (see [53]). These studies reveal that Cx32 and Cx43 remain mainly as monomers in the ER and oligomerize in the Golgi apparatus, although within different subcompartments (ER/Golgi intermediate compartment:

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Cx32; trans-Golgi network: Cx43). The same has not been observed for Cx26, where oligomeric states are found in ER fractions and following in vitro microsomal expression. A large portion of Cx26 may sidestep Golgi and traffic directly to the plasma membrane as, in contrast to Cx32, Cx26 has been reported to still reach the plasma membrane upon application of brefeldin A (a Golgi disrupting drug); however, others have reported inhibition of Cx26 oligomerization by brefeldin A treatment. Careful scrutiny of the oligomerization process is necessary given that premature oligomerization has been observed in overexpression systems, resulting in misleading assumptions of the biosynthetic pathway of connexins [54]. The use of diverse cell systems may also explain some of the differing results observed by various groups. Studies aimed at determining the minimal regions required for oligomerization have mainly focused on Cx43, highlighting transmembrane regions TM2 and TM3, as well as the cytoplasmic loop (CL) and extracellular loop 2 (E2), as necessary for ER-inhibition of oligomerization (see [54]). Liver connexin domains responsible for oligomerization and trafficking still require further study; however, in the case of Cx32, an essential region for trafficking to the surface is contained between residues 207 and 219 on the cytoplasmic carboxyl terminus, and may also play a role in oligomerization [55]. It is unknown whether oligomerization and trafficking through the various compartments in the cell are assisted by chaperones, although the regulation of oligomerization in the ER and Golgi apparatus suggests the presence of patrolling complexes that prevent inter-connexin association. There is some evidence that the calcium-sensing molecule CaM may interact with Cx32 to permit its proper oligomerization and trafficking to the surface. Gap junction trafficking pathways outside of the ER and Golgi compartments are not clearly defined and there is some speculation whether connexons made up of different connexins use alternate routes, mainly due to equivocal effects of microtubule or microfilament depolymerization on connexin trafficking. Growing evidence is solidifying the view that microtubules play a major role in regulation of connexin trafficking in vesicular bodies (see [56, 58]). Although most connexin biosynthesis studies have not focused on liver, nocodazole, vinblastine, and colchicine treatments (which disrupt microtubule networks) have been shown to affect the regulation of gap junction levels in primary cultured rat hepatocytes, and mouse and rat livers, the latter showing an inhibition of estrogen-mediated Cx26 increase with microtubule disruption. Once within the plasma membrane, connexons are believed to freely diffuse laterally and dock with an opposing connexon from another cell. It is unclear whether this process is active or passive; however, there is evidence that cell adhesion molecules could play a role in the formation of gap junctions. Studies performed by Gaietta et al. demonstrated that the insertion of FlAshand ReAsh-labeled Cx43 to membrane plaques occurs

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laterally onto the edges, whereas removal takes place from the center of the plaque (see [59]) in a process that may involve clathrin/actin-mediated endocytosis [60]. Because docked connexons cannot be separated by most physiological conditions, junctional removal is a phagocytic-like event which forms double-cell membrane vesicles, termed annular junctions. It is unclear whether this is the only form of internalization for gap junctions since annular membrane profiles are not commonly observed in normal liver. Connexin half-lives are unusually short for transmembrane proteins, around 3–6 hours for mouse liver connexins. It has been shown that internalized gap junctions are degraded by lysosomes and proteasomes, and that this degradation of connexins might also depend on the integrity of microtubules. Similarly, microfilament disruption with cytochalasin affects endocytosis into double-membrane annular gap junction profiles and abolishes clustering of gap junctions at the plasma membrane. The dependence of gap junction regulation on cytoskeletal components such as microtubules and microfilaments offers a plausible mechanism for directed trafficking within the cell, which suggests two possibilities for the presence of polarized distribution of gap junctions in liver hepatocytes: directed delivery to lateral membranes or indiscriminate delivery to all membranes coupled to active removal of connexons from apical surfaces. Breidert et al. have observed for Cx49, a lens gap junction protein, vesicular trafficking and localization to lateral and basal but not apical membranes in the polarized Madin–Darby canine kidney (MDCK) cell line [61]. Furthermore, Shaw et al. have shown that microtubules are essential for delivery of connexin-positive vesicles to sites of cell–cell adhesion [58], which, in hepatocytes, are present in the lateral membranes.

FUNCTION AND REGULATION OF TIGHT JUNCTIONS Tight junctions inhibit solute and water flow through the paracellular space (barrier function) and also separate the apical from the basolateral cell surface domains to establish cell polarity (fence function). To investigate the barrier function of tight junctions, three approaches are commonly used to evaluate the permeability of the paracellular pathway to ions and uncharged hydrophilic macromolecules: passage of electron-dense dyes, transepithelial electrical resistance (TER), and transepithelial flux of substances lacking affinity for membrane transporters [62]. Barrier function can be visualized by the ability of tight junctions to block the passage of electron-dense molecules, such as ruthenium red, lanthanum (Figure 14.7b), and cationic ferritin. A modification of this method combining biotinylated compounds and fluorescence microscopy has been used. Measurement of TER is the simplest and most rapid

(a)

(b)

Figure 14.7 Tight junctions in rat liver in vivo. (a) Distribution of occludin in rat liver. Occludin is observed in bile canalicular regions. (b) Thin-section image in which the tight junctions can be seen flanking the microvillus-filled canaliculus. Lanthanum was introduced into the extracellular spaces via the vascular system and is seen to penetrate the extracellular spaces but is denied access to the canalicular lumen by the tight junctions

approach, in which electrodes placed on opposite sides of an epithelial sheet are used to pass current pulses and measure resultant voltage deflections. The third approach to determining paracellular permeability is to measure the passive movement of flux of hydrophilic uncharged molecules, commonly radioactively labeled D-mannitol, raffinose, polyethylene glycol, and various methylated dextrans across an epithelial sheet as a function of time. To investigate the fence function of tight junctions, fluorescent lipids or lipid probes (such as boron-dipyrromethene (BODIPY)-labeled sphingomyelin) are inserted into the outer leaflet of the plasma membrane and the ability of the tight junction to restrict the dye localization to the cell surface domain is examined (Figure 14.8d,e) (see [63]). Disruption of actin filaments deteriorates the barrier function of tight junctions, although the protein interactions among tight junction proteins have not been fully clarified. Polarized epithelial cell, such as MDCK, Caco-2, and T84 cell lines, have been successfully exploited to

14: GAP AND TIGHT JUNCTIONS IN LIVER: COMPOSITION, REGULATION, AND FUNCTION

(a)

213

(b)

(d)

(c)

(e)

Figure 14.8 Tight junctions in primary cultured rat hepatocytes in vitro in the presence of DMSO/glucagon medium. (a) In freeze-fracture of the adluminal plasma membrane, tight junction strands form well-developed networks. (b) Thin-section image in which the tight junctions can be seen with circumferential actin filaments. (c) Occludin-immunoreactive lines are observed on the most subapical plasma membranes at cell borders. (d,e) Fence function images of tight junctions. The hepatocytes are labeled with BODIPY-shingomyelin. The fluorescent probe was effectively retained in the apical domain (d). In the hepatocytes treated with calcium chelator (3 mM EGTA) for five minutes, the probe diffused through the tight junction, labeling the basolateral face, and therefore appeared to penetrate the cells (e)

study the regulations of tight junctions in vitro. However, for hepatocytes that display a complex polarity and deliver apical proteins via an indirect route, in vitro studies have been hampered by lack of well-polarized hepatic lines, although it has been reported that phalloidin and vasopressin induce dysfunction of tight junctions and the cytoskeleton of rat hepatocyte couplets. In primary rat hepatocytes cultured with DMSO/glucagon, tight junction strands form well-developed networks in freeze-fracture at the adluminal plasma membrane (Figure 14.8a) and circumferential actin filaments are observed near the tight junction regions (Figure 14.8b). Moreover, occludin-, claudin-1-, ZO1-, and ZO2-immunoreactive lines were strongly observed on the most subapical plasma membrane of the cell borders (Figure 14.8c) (see [64]), and the fence function of tight

junctions in the cells, as examined by diffusion of labeled sphingomyelin, was well maintained (Figure 14.8d,e). Treatment of the cells with the actin polymerizing drug mycalolide B caused disappearance of both the circumferential actin filaments and occludin, while tight junction strands remained virtually intact, leading to the hypothesis that occludin, but not other transmembrane proteins, plays a role in the linkage between the actin cytoskeleton and tight junctions in hepatocytes. Changes of tight junction strands and expression of occludin and claudin-1 are observed during DNA synthesis and redifferentiation in cultured rat hepatocytes. Furthermore, downregulation of claudin-1 and upregulation of claudin-2 by growth factors, TGF-β, HGF, EGF, and by cytokines, IL-1β, and oncostatin M are also observed in cultured rat hepatocytes

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Non-parenchymal cells

Hepatocyte

Oncostatin M

TNF-α

OSMR gpl30 IL-l-R TNFR

IL-6

gp130

EGF

IL-6R EGFR c-met TGFβRI

IL-1β

HGF TGF-β

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TJ

TGFβRII

3

*

* *

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*

2.5 Relative expression of proteins

MAPK p38 MAPK PI3K PKC STAT Smad

*

*p < 0.01 vs control 2.0

1.5

1.0 *

* *

0.5

M

β On

co

sta

tin

-1 IL

Fα TN

-6 IL

F EG

F HG

Fβ TG

co

ntr

ol

0

Figure 14.9 Effectors (growth factors and cytokines) of tight junction proteins in hepatocytes. (a) Growth factors and cytokines derived from non-parenchymal cells and the receptors and signaling pathways in hepatocytes. (b) The changes in expression of claudin (CL)-1 and -2 proteins in hepatocytes after treatments with the cytokines and growth factors in (a)

(Figure 14.9) (see [64]). These findings suggest that the growth factors and cytokines may affect bile canalicular sealing by tight junctions during regeneration and inflammation of livers. Although tight junctions in these differentiated cultured hepatocytes assume the distribution seen in simple polarized epithelial cells, this culture system provides a useful model in which to study hepatocyte tight junctions. Recent evidence indicates that the transcription repressor Snail plays a central role in the epithelium– mesenchyme transition (EMT), by which epithelial cells lose their polarity. When Snail was overexpressed in cultured mouse epithelial cells, EMT was induced with concomitant repression of the expression of claudins and occludin not only at the mRNA but also at the

protein level; the mechanism appears to involve direct binding of Snail to the E-boxes of the promoters of claudin/occludin genes, similar to E-cadherin [65]. In the oncogenic Raf1-transfected mouse hepatic cell line, expression of occludin and claudin-2 and barrier function were downregulated during EMT, during which downregulation of E-cadherin and upregulation of Snail occurred (see [66]). In mature rat hepatocytes in vitro, TGF-β induces EMT by downregulation of claudin-1 and the fence function via Snail [63]. A role for tight junctions in intracellular signaling has been proposed. To elucidate the mechanisms of signal transmission required for the regulation of the tight junction, researchers have examined the effects of signaling pathways such as MAPK and PI3-kinase on the regulation

14: GAP AND TIGHT JUNCTIONS IN LIVER: COMPOSITION, REGULATION, AND FUNCTION

of tight junctions (see [67]). In primary rat hepatocytes, the changes of claudin-1 and -2 by growth factors and cytokines are regulated via distinct signal transduction pathways (Figure 14.9) (see [63, 64]). Furthermore, in the regenerating liver and the proliferative hepatocytes of rats, treatment with a p38 MAPK inhibitor prevents the changes of claudin-1 (see [40]). Occludin is an important regulatory component of signal transduction associated with tight junctions [68, 69]. The long carboxy-terminal region of occludin is rich in serine, threonine, and tyrosine residues and possesses a coiled-coil structural domain, which interacts with c-Yes, the mu isoform of protein kinase C (PKC), Cx26, the regulatory subunit of PI3-kinase, and PP2A, respectively (Figure 14.3d) [69, 70]. In primary cultures of occludin-deficient mouse hepatocytes, claudin-2 expression and apoptosis were induced by downregulation of the activation of MAPK and Akt. In the hepatic cell lines derived from occludin-deficient mice, claudin-2 expression and serum-free induced apoptosis were also increased by downregulation of the activation of MAPK and Akt. Furthermore, in the hepatic cell lines transiently transfected with mouse and rat occludin genes, induction of claudin-2 expression and apoptosis were inhibited with increases in activation of MAPK and Akt. These findings show that occludin plays a crucial role in claudin-2-dependent tight junction function and the apoptosis involving MAPK and Akt signaling pathways in hepatocytes [71]. Furthermore, a number of signaling molecules have been associated with the regulation of tight junctions, including tyrosine kinases, cAMP, Ca2+ , PKC, heteromeric G proteins, and phospholipase C (see [72]). The protocol of switching between Ca2+ -containing and Ca2+ -free solutions has demonstrated that in monolayers incubated in low-calcium medium, tight junction proteins are disassembled and dephosphorylated, and less tightly associated with actin filaments (see [72]). This indicates that the function of tight junctions may be locally regulated by signaling events within the tight junction plaque or may regulate some aspect of intracellular signaling. Phosphorylation of the tight junction proteins may be important to coordinating tight junction assembly or regulating tight junction function [73]. It is also thought that cytoplasmic signaling pathways can affect the barrier function of tight junctions. ATP depletion experiments result in altered tight junction structure, decreased TER, and an increased association of tight junction proteins with the actin filaments (see [72]). The PKC-activating phorbol esters induce a rapid decrease in tight junction permeability and alter perijunctional actomyosin (see [74]). Changes in paracellular permeability induced by the Vibrio cholerae toxin (so-called zonula occludens toxin, ZOT) are also associated with a reorganization of perijunctional actin [75]. The small GTP-binding

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protein Rho regulates actin filament organization and influences the organization and permeability of associated tight junctions (see [76]). Because hepatocyte tight junctions are the only intercellular barrier between the sinusoidal and the canalicular spaces, the major consequences of the liver disease jaundice may result from the disturbance of tight junctions. In tissue thin sections, the tight junctions of hepatocytes can be seen to block the access of the experimentally administered extracellular tracer (lanthanum) to the canalicular lumen (Figure 14.6b); irregularities in the structure and distribution of tight junctions, accompanied by increased permeability, have been observed during extrahepatic cholestasis after common bile duct ligation, intrahepatic cholestasis after ethinylestradiol treatment (see [77]), partial hepatectomy (see [78]), and hepatocyte injury repair following hepatotoxic drug administration or experimental colitis (see [64, 79, 80]). Loss of gap junctions, leaky tight junctions, and disorganized actin bundles are considered to be sufficient aberrations to cause cholestasis and the eventual development of jaundice. With advances in studies of tight junctions, it has become clear that these three causes are never independent of each other. Disruption of actin filaments clearly deteriorates the barrier function of tight junctions, although the protein interaction among tight junction proteins has not been fully clarified. Mechanisms of tight junction protein trafficking are an underexplored area of study and much is still unknown about the roles played by cellular compartments or cytoskeletal proteins in delivery to the surface membrane. Pathways of occludin targeting to the tight junction complex remain obscure; however, domains within the molecule and binding partners responsible for localization to tight junctions are beginning to be recognized, such as cingulin, F-actin, and ZO1 (see [81]). Although ZO1 is known to bind to occludin through the carboxyl terminus and has been proposed to mediate tight junction localization, it is unclear whether it is essential since domain and truncation mutants are still found at tight junctions (see [82]). The carboxyl terminus contains several phosphorylation sites and phosphorylation has been shown to increase junctional localization, demonstrating the regulatory aspect of this domain. However, Subramanian et al. demonstrated that carboxyl terminus deletions did not affect targeting, whereas removal of residues from the fourth transmembrane domain resulted in only cytoplasmic occludin localization [80]. They further showed that microtubule disruption affected delivery to the plasma membrane, but not maintenance, while microfilaments were required for maintenance of localization and organization at the surface. Other groups have also observed the need for an intact actin microfilament system for proper occludin localization [81]. Removal of occludin may occur through ubiquitination [82], caveolae-mediated endocytosis [83], and degradation by proteasomes [84],

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THE LIVER: EXPRESSION AND FUNCTION OF TIGHT JUNCTIONS IN HUMAN LIVERS

although recycling back to the plasma membrane in a Rab13-dependent pathway is also observed [85]. Like occludin, claudin carboxyl terminus-deletion mutants are still able to form tight junctions, and may also depend on the formation of adhesion junctions and actin cytoskeleton to form proper tight junction strands [86]. Claudin-3 has been shown to internalize in a similar fashion to connexins, together with the opposed claudin in a double-membrane vesicular body [87]. Interestingly, it has been observed that claudin-3 is sorted into different compartments to occludin, ZO1, and JAMs, suggesting selective mechanisms of vesicle sorting post-endocytosis. Little is known about JAM trafficking, and much may be similar to occludin and claudin pathways. For example, JAMs depend on actin cytoskeleton for proper localization to the plasma membrane [81].

EXPRESSION AND FUNCTION OF TIGHT JUNCTIONS IN HUMAN LIVERS In human livers, occludin, JAM-A, ZO1, ZO2, claudin-1, -2, -3, -7, -8, -12, -14, and tricellulin are detected together with well-developed tight junction structures (Figure 14.10a,c), and claudin-2 shows a lobular gradient increasing from periportal to pericentral hepatocytes as in the livers of rat and mouse, whereas claudin-1 is expressed throughout the whole liver lobule (Figure 14.10b). As a genetic disease of human tight junction protein, missense mutations in ZO2 have been identified in patients with familial hypercholanemia [88]. In neonatal ichthyosis-sclerosing cholangitis (NISCH) syndrome, mu-

Oc JAM-A ZO-1 ZO-2

CL-1 CL-2 CL-3 CL-4 CL-5 CL-6 CL-7 CL-8 CL-9 CL-12 CL-14

TRIC

(a)

P

P

C

C

CL-1

CL-2 (b)

BC BC

(c)

Figure 14.10 Expression of tight junction proteins and their structures in human livers. (a) RT-PCR for tight junction molecules in human liver. (Oc, occludin; CL, claudin; TRIC, tricellulin). (b) Staining of claudin (CL)-1 and -2 in human liver. (C, central vein area; P, periportal vein area). (c) Tight junction structure in human liver. (Thin-section image at left; freeze-fracture image at right; BC, bile canaliculus)

14: GAP AND TIGHT JUNCTIONS IN LIVER: COMPOSITION, REGULATION, AND FUNCTION

tations of claudin-1 are also observed and the lack of claudin-1 may lead to increased paracellular permeability between bile duct epithelial cells [89]. However, biliary disease in the mouse claudin-1 KO model was not detected, as the KO mice died at birth [26]. HCV is an enveloped positive-stranded RNA hepatotropic virus and three host cell molecules are important entry factors or receptors for HCV internalization: scavenger receptor BI (SR-BI), the tetraspanin CD81, and claudin-1 [90]. CD81 and claudin-1 act as co-receptors during late stages in the entry process. Furthermore, not only claudin-1 but also claudin-6 and -9 are entry cofactors for HCV [90]. The claudin tight junction proteins are novel key factors for HCV and may provide new targets for antiviral drug therapy. Other tight junction components may also bind viruses. For example, binding of a reovirus coat protein to JAM-A has been reported, involving the extracellular JAM-A domains responsible for the homodimerization [91, 92]. Moreover, CAR is related to immunoglobulin JAMs and is localized to junctional domains [93]. Human hepatic stem cells, which are pluripotent precursors of hepatoblasts and thence of hepatocytic and biliary epithelia, are located in ductal plates in fetal livers and in canals of Hering in adult livers [94]. The stem cell phenotype comprises expression of EpCAM, NCAM, CK19, c-kit, claudin-3, and weak levels of albumin, but no expression of AFP or adult liver-specific proteins such as transferrin, Cx26, Cx32, PEPCK, DPPVI, or P450s [95]. It is thought that claudin-3 is a unique and novel hepatic stem cell surface marker.

CONCLUSION The two most prominent junctional types in liver are the gap junctions, which provide direct intercellular communication, and the tight junctions, which serve to partition membrane domains of individual cells and to occlude extracellular space, restricting pericellular diffusion. The integral membrane proteins of gap junctions, the connexins, are well characterized and it has recently been realized that they bind to cytoplasmic proteins, suggesting that the connexins may be a nucleation site for intracellular as well as intercellular signaling molecules. But the peripheral tight junction proteins were identified first; the finding of most intense interest is the participation of a novel family of integral membrane proteins, the claudins. Although these junctional structures perform different functions, there are numerous points at which functional studies overlap. Indeed, the findings that the traditionally tight junction-associated protein ZO1 also binds to a connexin, and that occludin is colocalized with Cx32 in transfectants, indicate the possibility for either coordinate or reciprocal regulation of macromolecular complexes containing

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gap and tight junction proteins. Studies of protein–protein interactions and of coordinate and subordinate regulation of gene families are expected to soon disclose the intricacies of inter- and intracellular signaling and growth control (at gap junctions and the regulatory mechanisms of the “blood-biliary barrier” formed by tight junctions (see [96])). Furthermore, as with the HCV receptor CD81, the major gap junction component (the connexins) and the major tight junction components (occludin and the claudins) are tetraspan proteins. Future studies are likely to reveal new functions of connexins, occluding, and claudins, which may be associated not only with HCV but with the other virus entry processes and downstream signaling.

ACKNOWLEDGMENT The authors greatly thank Dr M. Murata, Dr T. Yamamoto, Dr M. Lan, Dr M. Imamura, Dr S. Son, and Miss E. Suzuki (Sapporo Medical University, Japan). The research by our laboratories is largely supported by NIH Grant DK-41918 and by Grants-in-Aid from the Ministry of Education, Culture, Sports Science, and Technology, and the Ministry of Health, Labour and Welfare of Japan.

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26. Evans, M.J., von Hahn, T., Tscherne, D.M. et al. (2007) Claudin-1 is a hepatitis C virus co-receptor required for a late step in entry. Nature, 446, 801–5. 27. Sawada, N., Murata, M., Kikuchi, K. et al. (2003) Tight junctions and human diseases. Med Electron Microsc, 36, 147–56. 28. Konopka, G., Tekiela, J., Iverson, M. et al. (2007) Junctional adhesion molecule-A is critical for the formation of pseudocanaliculi and modulates E-cadherin expression in hepatic cells. J Biol Chem, 282, 28137–48. 29. Braiterman, L.T., Heffernan, S., Nyasae, L. et al. (2008) JAM-A is both essential and inhibitory to development of hepatic polarity in WIF-B cells. Am J Physiol Gastrointest Liver Physiol , 294, G576–88. 30. Umeda, K., Ikenouchi, J., Katahira-Tayama, S. et al. (2006) ZO-1 and ZO-2 independently determine where claudins are polymerized in tight-junction strand formation. Cell , 126, 741–54. 31. Itoh, M., Nagafuchi, A., Yonemura, S. et al. (1993) The 220-kD protein colocalizing with cadherins in non-epithelial cells is identical to ZO-1, a tight junction-associated protein in epithelial cells: cDNA cloning and immunoelectron microscopy. J Cell Biol , 121, 491–502. 32. Kojima, T., Kokai, Y., Chiba, H. et al. (2001) Cx32 but not Cx26 is associated with tight junctions in primary cultures of rat hepatocytes. Exp Cell Res, 263, 193–201. 33. Kojima, T., Spray, D.C., Kokai, Y. et al. (2002) Cx32 formation and/or Cx32 mediated intercellular communication induce expression and function of tight junctions in hepatocytic cell line. Exp Cell Res, 276, 40–51. 34. Murata, M., Kojima, T., Yamamoto, T. et al. (2005) Tight junction protein MAGI-1 is up-regulated by transfection with connexin 32 in an immortalized mouse hepatic cell line: cDNA microarray analysis. Cell Tissue Res, 319, 341–47. 35. Duffy, H.S., Iacobas, I., Hotchkiss, K. et al. (2007) The gap junction protein connexin32 interacts with the Src homology 3/hook domain of discs large homolog 1. J Biol Chem, 282, 9789–96. 36. Spray, D.C., Ginzberg, R.D., Morales, E.A. et al. (1986) Physiological properties of dissociated rat hepatocytes. J Cell Biol , 103, 135–44. 37. S´aez, J.C., Connor, J.A., Spray, D.C. et al. (1989) Hepatocytes gap junctions are permeable to the second messengers inositol 1,4,5-trisphosphate and calcium ions. Proc Natl Acad Sci U S A, 86, 2708–12. 38. Scemes, E., Suadicani, S.O. and Spray, D.C. (2000) Intercellular calcium wave communication via gap junction-dependent and independent mechanisms, in Current Topics in Membranes, Vol. 49 (ed. C. Peracchia), Academic Press, San Diego, pp. 145–73. 39. Kojima, T., Yamamoto, T., Murata, M. et al. (2003) Role of the p38 MAP-kinase signaling pathway for Cx32 and claudin-1 in the rat liver. Cell Commun Adhes, 10, 1–7. 40. Dermietzel, R., Yancey, S.B., Traub, O. et al. (1987) Major loss of the 28 kDa protein of gap junction in proliferating hepatocytes. J Cell Biol , 105, 1928–34.

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41. Janssen-Timmen, U., Traub, O., Dermietzel, R. et al. (1986) Reduced number of gap junctions in rat hepatocarcinomas detected by monoclonal antibody. Carcinogenesis, 7, 1475–82. 42. White, T.W. and Paul, D.L. (1999) Genetic diseases and gene knockouts reveal diverse connexin functions. Annu Rev Physiol , 61, 283–310. 43. Nelles, E., Butzler, C., Jung, D. et al. (1996) Defective propagation of signals generated by sympathetic nerve stimulation in the liver of connexin32-deficient mice. Proc Natl Acad Sci U S A, 93, 9565–70. 44. Temme, A., Buchmann, A., Gabriel, H.D. et al. (1997) High incidence of spontaneous and chemically induced liver tumors in mice deficient for connexin32. Curr Biol , 7, 713–16. 45. Moennikes, O., Buchmann, A., Ott, T. et al. (1999) The effect of connexin32 null mutation on hepatocarcinogenesis in different mouse strains. Carcinogenesis, 20, 1379–82. 46. Moennikes, O., Stahl, S., Bannasch, P. et al. (2003) WY-14,643-mediated promotion of hepatocarcinogenesis in connexin32-wild-type and connexin32-null mice. Carcinogenesis, 24, 1561–65. 47. Kojima, T., Fort, A., Tao, M. et al. (2001) Gap junction expression and function in primary cultures of Cx32 deficient (KO) mouse hepatocytes. Am J Physiol , 281, G1004–13. 48. Temme, A., Ott, T., Dombrowski, F. et al. (2000) The extent of synchronous initiation and termination of DNA synthesis in regenerating mouse liver is dependent on connexin32 expressing gap junctions. J Hepatol , 32, 627–35. 49. Dagli, M.L., Yamasaki, H., Krutovskikh, V. et al. (2004) Delayed liver regeneration and increased susceptibility to chemical hepatocarcinogenesis in transgenic mice expressing a dominant-negative mutant of connexin32 only in the liver. Carcinogenesis, 25, 483–92. 50. Asamoto, M., Hokaiwado, N., Murasaki, T. et al. (2004) Connexin32 dominant-negative mutant transgenic rats are resistant to hepatic damage by chemicals. Hepatology, 40, 205–10. 51. Sarma, J.D., Wang, F. and Koval, M. (2002) Targeted gap junction protein constructs reveal connexin-specific differences in oligomerization. J Biol Chem, 277, 20911–18. 52. Maza, J., Das Sarma, J. and Koval, M. (2005) Defining a minimal motif required to prevent connexin oligomerization in the endoplasmic reticulum. J Biol Chem, 280, 21115–21. 53. Martin, P.E., Steggles, J., Wilson, C. et al. (2000) Targeting motifs and functional parameters governing the assembly of connexins into gap junctions. Biochem J , 349, 281–87. 54. Shaw, R.M., Fay, A.J., Puthenveedu, M.A. et al. (2007) Microtubule plus-end-tracking proteins target gap junctions directly from the cell interior to adherens junctions. Cell , 128, 547–60. 55. Lauf, U., Giepmans, B.N., Lopez, P. et al. (2002) Dynamic trafficking and delivery of connexons to the

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SECTION A : HEPATOCYTE

15

Copper Metabolism and the Liver Michael L. Schilsky1 and Dennis J. Thiele2 1 Department

of Medicine, Yale University Medical Center, New Haven, CT, USA 2 Department of Pharmacology and Cancer Biology, Duke University Medical Center, Durham, NC, USA

PHYSIOLOGY AND PATHOPHYSIOLOGY

Copper transport proteins

Overview Copper plays a critical biochemical role in the function of many enzymes and proteins that contain this essential element. Consequently, copper deficiency leads to loss of the catalytic function of copper-dependent enzymes and structural changes in proteins with prosthetic copper, while excess metal is toxic and leads to cell injury and death. Homeostatic control mechanisms have evolved to coordinate a healthy balance for copper, allowing cells to accumulate sufficient copper to support essential biochemical reactions and yet prevent toxicity due to an excess of this metal. Much of our knowledge of these metabolic pathways is derived from the study of the inherited metabolic disorders of copper metabolism and from microbial and mouse models that have deciphered new copper transporters, metallochaperones, and signaling pathways. This in turn has led to a better understanding of the pathophysiology of each of the metabolic disorders of copper metabolism, aiding in diagnosis and treatment and in the design of future therapies.

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

A key aspect of maintaining appropriate copper balance is the regulation of copper uptake into cells and its efflux. Cellular copper acquisition is accomplished with great specificity through the ubiquitously expressed copper transporter, Ctr1. Ctr1 is an integral membrane protein localized predominantly to the plasma membrane, but also to endosomal compartments. Ctr1 is structurally and functionally conserved from yeast to man [1–3]. The protein has three transmembrane domains, a methionine-rich extracellular amino terminus, a cysteine-histidine cluster in the cytosolic carboxyl terminus, and an MX3 M motif (where M is methionine) in the second transmembrane domain (Plate 15.1). While the MX3 M motif is essential for copper uptake, the methionine-rich ectodomain is thought to play a role in copper uptake only under copper-limiting conditions [3]. The functional unit of the protein is likely a homotrimer, with a pore that forms between the subunit transmembrane domain interfaces [4–6]. Experimental evidence strongly suggests that Ctr1 transports Cu+ across membranes, with Cu+2 likely interconverted to Cu+ by cell-surface metalloreductases prior to transport by Ctr1 [7]. Unlike many metal transporters, Ctr1 has no ATP hydrolysis domain, but experimental

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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evidence suggests that membrane electrochemical potential may be an important contributor to the driving force behind copper movement across the plasma membrane. Ctr1 also traffics from the cell surface to internal vesicles under elevated copper conditions, a function that is dependent on the MX3 M motif [8–11]. Since this same motif is essential for copper transport by Ctr1, it is likely that the physiological signal for this trafficking is the transport of Cu+ itself. This post-translational relocalization of Ctr1 in response to changing copper abundance would permit rapid adaptation to high copper states, an important adaptation as Ctr1 mRNA levels themselves do not change markedly as a function of copper status [12]. Genetic studies in mice have demonstrated that Ctr1 is essential during development, as loss of both Ctr1 alleles results in embryonic lethality midway through gestation [13, 14]. Copper transport within cells is accomplished with the aid of metallochaperones that permit copper to move preferentially, and in a controlled manner, to other copper transporters, to compartments, or to copper-dependent enzymes. Atox1 was identified as the chaperone that directs copper to the copper-transporting ATPases ATP7A or ATP7B, depending on the cell type [15, 16] (see below). This protein binds Cu+ near its surface, for the specific and tightly controlled transfer to other proteins. Copper is transferred from the surface of the protein to repeated cysteine- and methionine-rich metal binding domains (MBDs) within the amino terminus of the cytosolic domain of the ATP7A and ATP7B copper-transporting ATPases [17, 18]. Studies have demonstrated direct interactions between Atox1 and ATP7A/ATP7B, lending further evidence for the role of this protein in the controlled transfer of copper to the copper-transporting ATPases [19]. Cu, Zn superoxide dismutase (Cu, Zn SOD) is a homodimeric enzyme that is abundantly expressed in liver and which functions in the disproportionation of superoxide anion, a potentially toxic reactive oxygen species generated from the mitochondrial electron transport chain and from other metabolic reactions. Cu, Zn SOD activity is absolutely dependent on the presence of copper to drive redox chemistry. Copper is supplied by the copper chaperone for SOD (CCS) [20]. While CCS structurally resembles SOD, it lacks key catalytic residues required for superoxide disproportionation, but the structural similarity allows it to form a complementary interface with SOD for copper transfer via direct CCS–SOD interactions. While Cu, Zn SOD and CCS were initially thought to reside strictly in the cytosol, recent studies demonstrate a significant concentration of the enzyme, and its copper chaperone, in the mitochondrial intermembrane space [21, 22]. The generation of CCS knockout mice has demonstrated that CCS plays a primary role in copper loading onto Cu, Zn SOD, although there appears to be a CCS-independent mechanism for copper loading onto mammalian Cu, Zn SOD [23, 24]. Interestingly, mice lacking the Cu, Zn SOD gene develop hepatocellular carcinoma, underscoring the importance of having sufficient

copper to activate this critical enzyme for oxidative stress defenses. A third arm for intracellular delivery targets the mitochondria, where the multi-subunit enzyme cytochrome c oxidase is located and which functions in the generation of ATP through oxidative phosphorylation. While only two of the dozen or so subunits of cytochrome oxidase bind copper—Cox1 and Cox2—these two subunits are encoded in the mitochondrial genome. This requires that a copper delivery system bring copper not only to the mitochondria but also into the mitochondrial lumen. While it is currently unclear how copper is delivered from its site of import by Ctr1 to mitochondria, proteins have been identified that are conserved from yeast to humans and that function in the assembly of copper onto mitochondrial cytochrome oxidase [25]. These include Cox17, which functions in the mitochondrial intermembrane space, and Sco1, Sco2, and Cox11, which play more proximal roles in copper delivery to Cox1 and Cox2 [25–28]. Moreover, it appears that mitochondrial copper is in vast excess over the demands of cytochrome oxidase, suggesting that mitochondria may be important homeostatic organelles for copper [29, 30]. While the copper-transporting ATPases ATP7A and ATP7B each pump copper from two strategic points, their distinct physiological functions are primarily determined by their patterns of tissue- and cell-type-specific expression. ATP7A is predominantly expressed in epithelial cells under normal conditions, where it serves to deliver copper, donated by Atox1 to the secretory compartment. Under elevated copper conditions in intestinal enterocytes, ATP7A traffics to the basolateral membrane, where it pumps copper into the portal circulation for delivery to the liver [31]. ATP7B is primarily expressed within hepatocytes, though several other cell types are known to express this protein [32–34]. In hepatocytes, the delivery of copper to the secretory compartment by ATP7B is critical for copper loading onto ceruloplasmin, which is thought to occur in the trans-Golgi [32], thereby moving this abundant copper-loaded protein into the circulation, where it is thought to function as a ferroxidase involved in Fe loading onto transferrin [33]. ATP7A and ATP7B are typical metal-transporting ATPases that are conserved from microbes to humans [34]. ATP7A and ATP7B each harbor cytosolic amino termini containing six repeats of cysteine-rich MBDs that accept copper from Atox1 [35, 36] (Plate 15.2). These MBDs bind copper and likely transfer it to a membrane channel lined with a critical cysteine-proline-cysteine motif that is present in most metal-transporting ATPases. Conformational changes in the copper-transporting ATPases that occur with this process are driven by cycles of phosphorylation and dephosphorylation, ultimately moving the copper across the membrane [30]. The importance of this phosphorylation cycle, and the critical function of the ATP binding region of these proteins, is underscored by the frequency of gene mutations that affect this site. The carboxyl termini, and other regions of these ATPases, likely

15: COPPER METABOLISM AND THE LIVER

contain signals that drive protein localization to and from the Golgi and relocate the proteins to the plasma membrane [37]. Hepatocytes and other cells are protected from the toxicity associated with high copper levels by metallothioneins (MTs), small cysteine-rich proteins that bind to copper, zinc, cadmium, and other metals through cysteine-thiolate bonds [38]. MT gene transcription is potently activated in response to elevated copper through the action of the MTF1 transcription factor, a protein with multiple zinc-finger DNA binding domains that binds to metal response elements (MREs) in the promoters of target genes [38]. While in response to elevated copper levels cytosolic MTF1 accumulates in the nucleus, the precise mechanisms whereby MTF1 senses and responds to changes in the levels of copper and other metals are not completely understood. Consistent with the liver as a central organ for copper metabolism, MTF1 knockout mice die in utero at ∼14 days of gestation and MTF1 appears to play an essential role in embryonic liver development [39]. Indeed, MTF1 knockout cells are defective in the expression of mRNAs encoding MT proteins and other proteins that play critical roles in the liver.

225

The digestive system Like other essential elements, the daily dietary intake of copper must meet losses in order to maintain homeostasis. Most diets contain between 2 and 10 mg of copper, and about 10% of the ingested copper is absorbed by duodenal enterocytes [40–42]. Daily losses occur from cellular turnover, from enteral secretion of fluids and urinary excretion. The renal excretion of copper, typically 10–20 µg per day, is a small component of the overall daily excretion of copper in man, which is mainly accomplished by the gut in the form of non-absorbed copper and copper excreted into bile by the liver [41, 42]. Most of the excess copper that is absorbed in the gut is excreted in bile, and urine copper excretion is maintained more or less constant in amount. Figures 15.1 and 15.2 highlight these processes and the role of the enterocyte in the gut and hepatocytes in the liver in copper metabolism. Dietary copper is the sole source of this metal. Given that Ctr1 is specific for Cu+ uptake, it is likely that dietary copper is transformed from Cu+2 to Cu+ by one or more cell-surface metalloreductases and transported across the plasma membrane by Ctr1 [43]. It is unclear if Ctr1 moves from the cell surface in hepatocytes in

Dietary Cu

Absorption stomach/duodenum

Non-absorbed Cu

Portal circulation Hepatic uptake ATOX1 transfer Ceruloplasmin synthesis Mitochondrial uptake/ SOD Cu acquisition Storage

L I V E R

Biliary excretion

Ceruloplasmin Circulating copper Tissue

RBC(SOD)

Renal excretion Urine

Feces

Tissue storage

Figure 15.1 Copper metabolism. Shown are the major pathways for copper metabolism in the body. In Wilson disease, the pathway for biliary copper excretion is defective due to lack of functional ATP7B. This leads to an increase in hepatic copper accumulation and subsequently an increase in copper exported from the liver and delivered to other tissues. Renal excretion is a minor pathway for copper excretion in comparison. However, chelating agents increase renal copper excretion while zinc blocks enterocyte copper transfer to the circulation and increases the fecal excretion of copper. In Menkes disease there are general deficits in copper transfer between cells, but the lack of functional ATP7A in enterocytes results in enterocyte copper hyper-accumulation, poor copper absorption due to failure to export copper from the basolateral membrane, and the generation of a copper-deficient state in the periphery

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THE LIVER: PHYSIOLOGY AND PATHOPHYSIOLOGY Dietary Cu Cu+2 Apical

Enterocyte

Cu+

Basolateral Ctr1

Cu,Zn SOD +1

Cu+2 Cu

Ctr1

reductase

Hepatocyte

mitochondria Cytochrome oxidase

CCS

Cu-GSH

Golgi

Golgi

ATP7B

Peri-canalicular vesicle

Cu in bile Apical

Cu-ATOX1 Ctr1

ATP7A

Basolateral

Ceruloplasmin

Basolateral

Cu-Albumin, other ligands

Figure 15.2 A model for copper metabolism in the enterocyte and hepatocyte. Enterocytes: The Ctr1 high-affinity Cu+ transporter is present on the apical membrane of enterocytes as well as in intracellular vesicles. A putative metalloreductase reduces Cu2+ to Cu+ for import by Ctr1. Ctr1 and a metalloreductase likely function in the export of Cu+ from endosomal compartments. In enterocytes, Cu+ is transported into the secretory compartment for loading onto Cu-dependent enzymes, or exported via the basolateral membrane by the P-type Cu+ -transporting P-type ATPase ATP7A. After release from the enterocyte into the bloodstream, Cu is transported in the serum, bound to ligands that may include histidine, albumin, and other uncharacterized molecules, via the portal vein to the liver. Hepatocytes: Ctr1 present on the basolateral membrane of hepatocytes imports Cu+ primarily from the portal circulation. Cytosolic copper is carried by Atox1 to the P-type Cu+ transporting ATPase, ATP7B, which resides mainly in the trans-Golgi machinery. Here copper is incorporated into ceruloplasmin or trafficked to vesicles for export into bile. Copper acquired by hepatocytes is also routed for incorporation into Cu, Zn SOD or transported to mitochondria for incorporation into cytochrome oxidase. When copper is abundant, the excess copper is excreted by this vesicular pathway to the apical membrane and into bile. When copper is limiting, ATP7B remains mainly in the trans-Golgi and ceruloplasmin biosynthesis continues. Copper is excreted via complexing with glutathione and other uncharacterized ligands

response to changes in environmental copper as it has been reported to do in other cell types. Copper in the enterocyte that is not required for metabolic function is bound by Atox1, MTs, and glutathione or, alternatively, is transferred out of cells to the bloodstream by ATP7A at the basolateral membrane. In Menkes disease enterocyte copper absorption is reduced due to the absence or loss of fully functional ATP7A. Copper that enters the bloodstream in the portal circulation is thought to be bound by albumin at its amino-terminal MBDs, or to histidine, but the precise identity of other copper ligands that function in copper delivery to the periphery is still under investigation. The main site for uptake from the circulation is the liver, where the metal is avidly extracted by hepatocytes. The membrane transporter for uptake into hepatocytes is likely to be Ctr1, as Ctr1 is strongly expressed in liver. Cu+ is transported into the hepatocytes and is bound to the metallochaperone Atox1, which delivers copper to ATP7B, the Wilson disease gene product [17, 18]. ATP7B is located mainly in the trans-Golgi and also in endosomal or pre-lysosomal vesicles destined for export via the bile canaliculus into bile [31, 42]. ATP7B is thought to

cycle between the trans-Golgi and the endosomal vesicles depending upon the amount of copper present [31]. When copper is abundant there is an increased presence of ATP7B in the vesicular compartment, and when copper is not present in excess, ATP7B is mostly localized to the trans-Golgi network. Some investigations have suggested that ATP7B reaches the apical membrane directly [44, 45], however this remains a point of controversy. Ceruloplasmin is a serum glycoprotein ferroxidase that is synthesized mainly by hepatocytes and functions in systemic iron distribution. Ceruloplasmin contains six copper atoms per molecule of protein and the nascent protein acquires its copper in the trans-Golgi secretory network [32, 36]. In Wilson disease, where ATP7B transport of copper into the trans-Golgi is defective, ceruloplasmin is synthesized but does not acquire copper properly. The process of ceruloplasmin copper loading within the secretory compartment does not appear to require metallochaperones, as copper has been shown to bind cooperatively to the protein [36]. Apo-ceruloplasmin (without copper) has a distinct conformation from that of the copper-filled protein. This sometimes results in a protein that has a

15: COPPER METABOLISM AND THE LIVER

shortened half-life, resulting in a reduced steady state of ceruloplasmin in the circulation [46]. However, in other instances the protein is relatively stable but lacks its typical copper-dependent ferroxidase activity. In Menkes disease and in other states of severe copper deficiency the amount of copper available for transport into hepatocytes and into the trans-Golgi network is low, leading to lack of incorporation of copper into ceruloplasmin. Therefore ceruloplasmin levels are also often low in patients with Menkes disease. In the genetic disorder aceruloplasminemia there is a defect in the gene for ceruloplasmin that causes a lack of ceruloplasmin protein production or, alternatively, alters copper binding by the protein, with the result that levels of the apoprotein in the circulation are very low [47]. In non-hepatic cells, the ATP7A protein serves to transport excess copper from the cell. The adaptive response of cell trafficking of ATP7A in response to excess copper was described before it was recognized that ATP7B could undergo similar intracellular relocalization [31]. ATP7A also transports copper into the Golgi network of these cells, and is important for copper incorporation into other metalloproteins such as lysyl oxidase and dopamine beta hydroxylase.

Renal copper metabolism The renal excretion of copper, typically 10–20 µg per day, increases often to above 100 µg daily in untreated Wilson disease, in acute copper poisoning, and in cholestatic liver diseases where hepatic copper levels are also markedly elevated [41, 48]. Little was known about the renal copper transport prior to the discovery of Ctr1, ATP7A, and ATP7B and many questions still remain about their expression and integrated function in this organ. Ctr1 expression was demonstrated in the kidney [49, 50], where it transports circulating copper from as yet poorly identified carriers. Some of the transported copper entering the kidney tubules is returned to the bloodstream by the copper-transporting ATPases ATP7A and ATP7B. While the exact cellular expression of these proteins in the kidney in man is not clearly defined, the murine orthologues ATP7A and ATP7B are expressed in glomeruli, and ATP7B is expressed in the kidney medulla [50]. It has been suggested that ATP7A and ATP7B may also be involved in copper re-absorption in the loops of Henle. Copper-dependent intracellular trafficking of ATP7B and ATP7A appears to also occur in kidney cells. As in other cell types, this intracellular trafficking from the trans-Golgi network to intracellular vesicles or proximal to the cell surface in response to increased environmental copper permits the protein to function in copper export back into the bloodstream or into intracellular storage pools. Linz et al. in 2008 [50] demonstrated intracellular

227

trafficking of ATP7B and ATP7A in renal cells as well as in hepatocytes in response to copper. The different localization of the two copper-transporting proteins is critical to their function. ATP7A traffics to the basolateral membrane, where it transports copper destined for re-absorption into the blood, ridding renal cells of excess copper. While ATP7A moves from the Golgi to the plasma membrane in response to copper, ATP7B likely redistributes excess copper into vesicles in cells rather than directly exporting the copper, which could account for the increase in renal tubular copper content in response to excess copper presented to the kidneys in patients with Wilson disease [41].

Copper toxicity Excess copper is toxic to cells. Oxidative injury to membranes and DNA, and altered protein synthesis are amongst the many toxic effects of excess copper [51, 52]. Hepatocellular necrosis results from copper accumulation in hepatocytes. However, in the setting of acute liver failure, apoptotic injury may predominate [53]. Recently an interesting link between copper accumulation and copper-induced apoptosis was recognized. The protein XIAP (X-linked inhibitor of apoptosis) is inhibited by increases in cellular copper content, and this in turn lowers the threshold to copper-induced injury [54]. Studies suggest that XIAP directly binds to copper, altering its stability. The tri-peptide glutathione, present in high concentration in the cytosol of liver cells, provides the necessary reducing potential to protect proteins and to aid with conjugation and transport processes. Some biliary copper excretion is likely due to copper transport via copper glutathione complexes that are transported by proteins such as the canalicular multi-organionic transporter (cMOAT) present on the canalicular membrane of liver cells [55]. Animals that lack cMOAT have functional ATP7B-dependent hepatic copper transport but lack the high-capacity, low-affinity transport of glutathione copper complexes. Similarly, ATP7B-dependent transport is absent in the LEC rat, where cMOAT likely functions to excrete copper that accumulates [56]. As occurs in patients with Wilson disease, this cMOAT function does not appear adequate for eliminating all of the excess copper, and cellular damage ensues over time. It is thought that in some patients with acute liver failure due to Wilson disease there is a secondary injury that results in a reduced antioxidant capacity, thereby triggering a sudden decompensation. Certainly any insult that reduces either cellular reducing capacity—glutathione being the largest contributor to this—or other protective mechanisms for antioxidant injury (i.e. SOD) will result in a more rapid cellular injury.

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THE LIVER: HUMAN DISORDERS OF COPPER METABOLISM

HUMAN DISORDERS OF COPPER METABOLISM Wilson disease Wilson disease is an autosomal recessive disorder caused by mutation of ATP7B that occurs in ∼1/30 000 individuals (see [57] for review). The presence of two ATP7B mutations, one on each allele, or homozygosity for a single mutation, leads to toxic copper accumulation in hepatocytes and later in the central nervous system. There are over 500 different reported mutations of ATP7B as well as many polymorphisms of the gene, and most are cataloged by the laboratory of Dr Diane Cox [58]. The most common of these, H1069Q in exon 14, is present most often in the European Slavic population; the R778L missense mutation in exon 8 is most often absent in Europe but is present more commonly in Asians [59]. However, given the genetic diversity of many populations, such as that in North America, most patients are actually compound heterozygotes with a different mutation on each allele of the ATP7B gene [60]. The disorder typically presents as clinical liver disease earlier on or as neuropsychiatric disease in the second and third decades of life, though abnormalities in liver tests and liver histology may be silent in some. Rarely, neurological deficits can occur before the age of 10 years [61]. Untreated, patients will progress to cirrhosis and liver failure or to severe neurological disease with dystonia and Parkinsonism. Timely treatment can arrest disease progression and reverse some of the copper-induced injury [41, 57]. The diagnosis of Wilson disease rests upon either the demonstration of specific phenotypical markers such as corneal Kayser-Fleischer rings, elevated urinary copper excretion, a reduced level of ceruloplasmin in the circulation, and increased hepatic copper, as well as appropriate histology, or on molecular genetic evidence for the presence of two abnormal alleles of ATP7B [57]. Improvements in technical aspects of DNA sequencing and analysis have enabled the screening of the complete ATP7B gene for mutations and this test has become commercially available. No single test other than the molecular genetic testing can firmly establish the diagnosis, so often a combination of the above testing is necessary. At the last International Meeting on Wilson disease a scoring system was proposed to help clinicians determine the probability for pursuing a diagnosis of Wilson disease [62]. This scoring system accounts for the clinical and biochemical testing noted above, but also includes a heavy weighting for confirmatory molecular testing. More recently an algorithm for approaching patients who are asymptomatic or have liver disease and those who have neuropsychiatric symptoms has been published as part of diagnosis and treatment guidelines by the American Association for the Study of Liver Disease (AASLD) [57].

Treatment for Wilson disease involves either lifelong medical therapy or liver transplantation. Medical therapies designed to chelate copper and promote urinary copper excretion include the initially developed drug that was parenterally administered, dimercaptopropanol (British anti-lewisite), but this has been relegated to a mainly historical role and has been supplanted by the orally-administered D-penicillamine and trientine [57]. The compound tetrathiomolybdate is another drug that avidly chelates copper and has been tried experimentally for the early phase of treatment for neurologically-affected patients [63]. Treatment with copper chelators that increase cupriuresis has been the mainstay of treatment for Wilson disease, with trientine and penicillamine being the available agents [41, 57, 64–66]. Higher dosages are used for initial therapy for symptomatic patients; for asymptomatic patients and those on maintenance, treatment can be accomplished with lower daily doses. Zinc salts have also been used for treatment of Wilson disease, but are best used for initial therapy for asymptomatic patients or for maintenance therapy of symptomatic patients previously treated with chelators [67]. Zinc’s predominant mode of action is to block copper absorption by the enterocyte by stimulating the endogenous intracellular chelator MTs [40, 67]. The enterocytes take up copper that displaces the zinc from the MTs in the enterocyte, and the cells are routinely shed, thereby increasing the fecal excretion of copper. Evidence suggest that dietary Zn might also compete for copper uptake via an as yet unidentified Ctr1-independent copper transporter [68]. Zinc can also stimulate MTs in liver cells and other cells throughout the body, and perhaps may help protect these cells from further copper toxicity [40, 56]. Liver transplantation is curative for Wilson disease [69], effectively amounting to gene therapy by replacing the site of the abnormality of the disorder in the liver cells. Liver transplant is reserved for those patients with acute liver failure and those with advanced liver disease with chronic liver failure not amenable to pharmacological therapy [57, 70, 71]. Outcomes for transplantation of patients with Wilson disease have generally been excellent. However, there may be reduced survival and complications in patients with neurological features at the time of their transplant [72]. Living donor liver transplant using the liver of parents who are obligate heterozygotes for Wilson disease have been reported to be successful, with the recipient developing no signs or symptoms of liver disease [73]. Experimental hepatocyte cell transplantation in animal models of Wilson disease have demonstrated proof of principle that replacement of an adequate cell volume with normal hepatocytes can successfully reverse the phenotypical manifestations of the disorder, which include reducing hepatic copper load, increasing circulating ceruloplasmin, and increasing biliary copper excretion [74–76]. However, the use of allogeneic cells would require systemic immune suppression and would not eliminate portal hypertension if cirrhosis were advanced.

15: COPPER METABOLISM AND THE LIVER

Other adjunctive treatments include reduction of dietary copper and treatment of the complications of portal hypertension, such as with beta blockers for the prevention of variceal bleeding and diuretics for ascites [57]. Other specific treatments may be directed at neurological symptoms and include the use of muscle relaxants or injection of botulinum toxin for dystonia or antidepressant, and anxiolytic or antipsychotic medication for psychiatric symptoms. Gene therapy approaches for Wilson disease have also been tested in animal models. Both adenoviral and lentiviral vectors have been used to transduce liver cells with the full-length ATP7B gene, with some significant but transient success [77, 78]. As with cell transplant, many of the phenotypical changes in the transduced cells result in demonstrated increases in the levels of holo-ceruloplasmin and biliary copper excretion [77]. Further improvements in gene delivery and expression may some day enable this modality of treatment for patients. However, if disease is discovered at the advanced stage then it is likely that transplantation will still be needed for some individuals.

Menkes disease Menkes disease is an X-linked recessive disorder that results from mutations of the gene for the P-type copper-transporting ATPase, ATP7A [79]. In Menkes disease there is a progressive neurodegeneration that presents in infancy [80]. ATP7A is important for copper absorption from the enterocyte as well as copper excretion from most cell types other than hepatocytes, under normal conditions. The clinical manifestations of Menkes disease are due to a decrease in the activities of copper-dependent enzymes with prosthetic coppers like dopamine-β-hydroxylase, cytochrome c oxidase, and lysyl oxidase. Affected infants appear healthy at birth but develop hypotonia, seizures, and failure to thrive by ∼two months of age. Further progression of neurodegeneration typically leads to death by three years of age. Treatment with daily copper supplementation, parenteral copper administration with copper-histidinate, or subcutaneous delivery of other copper salts may improve the outcome if therapy is started soon after birth, before there is significant neurodegeneration [81, 82]. Pre-emptive treatment is typically only possible when the diagnosis is made prior to symptom development, after the diagnosis is established in another affected sibling [82]. However, newborn screening is not routinely available, and early detection is difficult because clinical abnormalities in affected newborns are absent or subtle. Furthermore, the usual biochemical markers—low serum copper and ceruloplasmin—are unreliable in the neonatal period. Molecular diagnosis can potentially be performed, but the gene is relatively large (150 kb) and there are many different mutation types, including point mutations, large deletions, and chromosomal rearrangements.

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A test for the neonatal diagnosis of Menkes disease that uses the measurement of metabolites of serum neurotransmitters was recently developed [81]. The copperdependent enzyme dopamine-β-hydroxylase converts dopamine to norepinephrine, and these are in turn metabolized to dihydroxyphenylacetic acid and to dihydroxyphenylglycol, respectively. Deficiency of dopamine-β-hydroxylase, as occurs in Menkes disease, leads to a high ratio of dopamine to norepinephrine as well as of dihydroxyphenylacetic acid to dihydroxyphenylglycol. These abnormal ratios can be used to identify presymptomatic disease, enabling pre-emptive therapy with copper supplementation by copper-histidine or other copper compounds in an attempt to ameliorate the disease and improve clinical outcomes.

Idiopathic copper toxicosis The development of copper overload in Indian and non-Indian children is now referred to as idiopathic copper toxicosis [83]. In afflicted patients, copper overload typically occurs in infancy and is progressive in nature. Patients clinically present with jaundice, ascites, and hepatosplenomegaly. Though liver copper may be markedly elevated, serum ceruloplasmin is not lower and there is no central nervous system deposition. The current hypothesis regarding the pathogenesis of this disorder is that there is environmental exposure to excess copper in addition to a genetic defect leading to excess stores of hepatic copper. This is consistent with the clustering of cases in Tyrolia, where elevated copper in well water was found [84]. Histologically there is liver injury with progressive fibrosis and an abundance of Mallory bodies. Attempts to link this to defects in ATP7B or MURR1 (COMMD1) have been unsuccessful [85]. Treatment for this disorder includes dietary modification to avoid copper intake, and the use of chelating agents such as d-penicillamine [83]. If found early enough, this treatment is effective in preventing disease progression. Recognition of water contamination directly or through the use of copper cooking vessels has led to a reduced incidence of this disorder [83].

Copper deficiency syndromes The initial recognition of copper deficiency came from the use of parenteral nutrition that lacked trace metal supplementation. These patients developed anemia and neutropenia, and sometimes thrombocytopenia. Since the addition of trace metals, this problem has been corrected. More recently there was a recognized syndrome of acquired neurological disorders linked to copper deficiency [86–89]. The disease in these patients often mimicked B12 deficiency with subacute combined degeneration. In some

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there were operative interventions that changed copper absorption from the upper intestine. In other patients, exogenous zinc administration that led to reduced copper absorption was uncovered, while some appeared to have hyperzincemia and hyperzincuria. The cause of the elevated zinc in these individuals remains obscure. The clinical picture of myelopathy was associated with a low serum copper and reduced ceruloplasmin. Copper supplementation results in stabilization but not always clinical improvement in the neurological symptoms in these individuals. The neutropenia and anemia due to the effect of copper deficiency on the bone marrow typically reverses with adequate replacement therapy.

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ACKNOWLEDGMENTS In memory of our mentor and friend, Dr Irmin Sternlieb; thanks to the Wilson’s Disease Association, the Yale New Haven Transplantation Center, and the Yale Liver Research Center (MLS); work on copper homeostasis is supported by US National Institutes of Health grants GM41840 and DK074192, and by a grant from the International Copper Association, Ltd to D.J.T.

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53. Strand, S., Hofmann, W.J., Grambihler, A., Hug, H., Volkmann, M., Otto, G., Wesch, H., Mariani, S.M., Hack, V., Stremmel, W., Krammer, P.H. and Galle, P.R. (1998) Hepatic failure and liver cell damage in acute Wilson’s disease involve CD95 (APO-1/Fas) mediated apoptosis. Nat Med , 4, 588–93. 54. Mufti, A.R., Burstein, E., Csomos, R.A., Graf, P.C., Wilkinson, J.C., Dick, R.D., Challa, M., Son, J.K., Bratton, S.B., Su, G.L., Brewer, G.J., Jakob, U. and Duckett, C.S. (2006) XIAP Is a copper binding protein deregulated in Wilson’s disease and other copper toxicosis disorders. Mol Cell , 21 (6), 775–85. 55. Dijkstra, M., Kuipers, F., van den Berg, G.J., Havinga, R. and Vonk, R.J. (1997) Differences in hepatic processing of dietary and intravenously administered copper in rats. Hepatology, 26, 962–66. 56. Schilsky, M.L., Stockert, R.J. and Sternlieb, I. (1994) Pleiotropic effect of the LEC mutation: a rodent model of Wilson’s disease. Amer J Physiol , 266, G907–13. 57. Roberts, E. and Schilsky, M.L. (2008) A practice guideline on Wilson disease: an update. Hepatology, 47, 2089–11. 58. Wilson Disease Mutation Database, http://www.medical genetics.med.ualberta.ca/wilson/index.php. 59. Ferenci, P. (2006) Regional distribution of mutations of the ATP7b gene in patients with Wilson disease: impact on genetic testing. Hum Genet , 120, 151–59. 60. Shah, A.B., Chernov, I., Zhang, H.T., Ross, B.M., Das, K., Lutsenko, S. et al. (1997) Identification and analysis of mutations in the Wilson disease gene (ATP7B): population frequencies, genotype-phenotype correlation, and functional analyses. Am J Hum Genet , 61, 317–28. 61. Singh, S., Dilawari, J.B., Chawla, Y. and Walia, B.N. (1989) Wilson’s disease in young children from northern India. Trop Gastroenterol , 10, 46–50. 62. Ferenci, P., Caca, K., Loudianos, G., Mieli-Vergani, G., Tanner, S., Sternlieb, I., Schilsky, M., Cox, D. and Berr, F. (2003) Diagnosis and phenotypic classification of Wilson disease. Liver Int , 23, 139–42. 63. Brewer, G.J., Askari, F., Lorincz, M.T., Carlson, M., Schilsky, M., Kluin, K.J., Hedera, P., Moretti, P., Fink, J.K., Tankanow, R., Dick, R.B. and Sitterly, J. (2006) Treatment of Wilson disease with ammonium tetrathiomolybdate: IV. Comparison of tetrathiomolybdate and trientine in a double-blind study of treatment of the neurologic presentation of Wilson disease. Arch Neurol , 63, 521–27. 64. Walshe, J.M. (1988) Diagnosis and treatment of presymptomatic Wilson’s disease. Lancet , ii, 435–37. 65. Walshe, J.M. (1973) Copper chelation in patients with Wilson’s disease. A comparison of penicillamine and triethylene tetramine dihydrochloride. Q J Med , 42, 441–52. 66. Sternlieb, I. and Scheinberg, I.H. (1968) Prevention of Wilson’s disease in asymptomatic patients. N Engl J Med , 278, 352–59. 67. Brewer, G.J., Yuzbasiyan-Gurkan, V., Lee, D.Y. and Appelman, H. (1989) Treatment of Wilson’s disease with zinc. VI. Initial treatment studies. J Lab Clin Med , 114, 633–38.

68. Lee, J., Petris, M.J. and Thiele, D.J. (2003) Characterization of mouse embryonic cells deficient in the Ctr1 high affinity copper transporter: identification of a Ctr1-independent copper transport system. J Biol Chem, 277, 40253–59. 69. Groth, C.G., Dubois, R.S., Corman, J., Gustafsson, A., Iwatsuki, S., Rodgerson, D.O., Halgrimson, C.G. and Starzl, T.E. (1973) Metabolic effects of hepatic replacement in Wilson’s disease. Transplant Proc, 5, 829–33. 70. Schilsky, M.L., Scheinberg, I.H. and Sternlieb, I. (1994) Hepatic transplantation for Wilson’s disease: indication and outcome. Hepatology, 19, 583–87. 71. Emre, S., Atillasoy, E.O., Ozdemir, S., Schilsky, M., Rathna Varma, C.V., Thung, S.N., Sternlieb, I., Guy, S.R., Sheiner, P.A., Schwartz, M.E. and Miller, C.M. (2001) Orthotopic liver transplantation for Wilson’s disease: a single-center experience. Transplantation, 72, 1232–36. 72. Stracciari, A., Tempestini, A., Borghi, A. and Guarino, M. (2000) Effect of liver transplantation on neurological manifestations in Wilson disease. Arch Neurol , 57, 384–86. 73. Asonuma, K., Inomata, Y., Kasahara, M., Uemoto, S., Egawa, H., Fujita, S. et al. (1999) Living related liver transplantation from heterozygote genetic carriers to children with Wilson’s disease. Pediatr Transplant , 3, 201–5. 74. Malhi, H., Irani, A.N., Volenberg, I., Schilsky, M.L. and Gupta, S. (2002) Early cell transplantation in LEC rats modeling Wilson’s disease eliminates hepatic copper with reversal of liver disease. Gastroenterology, 122, 438–47. 75. Malhi, H., Joseph, B., Schilsky, M.L. and Gupta, S. (2008) Mechanisms to repopulate liver with healthy donor cells for phenotypic correction in the LEC rat model of Wilson disease. Regen Med , 2, 165–73. 76. Irani, A.N., Malhi, H., Slehria, S., Gorla, G.R., Volenberg, I., Schilsky, M.L. and Gupta, S. (2001) Correction of liver disease following transplantation of normal hepatocytes in LEC rats modeling Wilson’s disease. Mol Ther, 3, 302–9. 77. Terada, K., Nakako, T., Yang, X.L., Iida, M., Aiba, N., Minamiya, Y., Nakai, M., Sakaki, T., Miura, N. and Sugiyama, T. (1998) Restoration of holoceruloplasmin synthesis in LEC rat after infusion of recombinant adenovirus bearing WND cDNA. J Biol Chem, 273, 1815–20. 78. Merle, U., Encke, J., Tuma, S., Volkmann, M., Naldini, L. and Stremmel, W. (2006) Lentiviral gene transfer ameliorates disease progression in Long-Evans cinnamon rats: an animal model for Wilson disease. Scand J Gastroenterol , 41, 974–82. 79. Seidel, J., Møller, L.B., Mentzel, H.J., Kauf, E., Vogt, S., Patzer, S., Wollina, U., Zintl, F. and Horn, N. (2001) Disturbed copper transport in humans. Part 1: mutations of the ATP7a gene lead to Menkes disease and occipital horn syndrome. Cell Mol Biol (Noisy-le-grand), 47, 141–48.

15: COPPER METABOLISM AND THE LIVER

80. Menkes, J.H. (1999) Menkes disease and Wilson disease: two sides of the same copper coin. Part I: Menkes disease. Eur J Paediatr Neurol , 3, 147–58. 81. Kaler, S.G., Holmes, C.S., Goldstein, D.S., Tang, J., Godwin, S.C., Donsante, A., Liew, C.J., Sato, S. and Patronas, N. (2008) Neonatal diagnosis and treatment of Menkes disease. N Engl J Med , 358, 605–14. 82. Kaler, S.G., Buist, N.R., Holmes, C.S., Goldstein, D.S., Miller, R.C. and Gahl, W.A. (1995) Early copper therapy in classic Menkes disease patients with a novel splicing mutation. Ann Neurol , 38 (6), 921–28. 83. Tanner, M.S. (1998) Role of copper in Indian childhood cirrhosis. Am J Clin Nutr, 67, 1074S–81S. 84. M¨uller, T., Sch¨afer, H., Rodeck, B., Haupt, G., Koch, H., Bosse, H., Welling, P., Lange, H., Krech, R., Feist, D., M¨uhlendahl, K.E., Br¨amswig, J., Feichtinger, H. and M¨uller, W. (1999) Familial clustering of infantile cirrhosis in Northern Germany: a clue to the etiology of idiopathic copper toxicosis. J Pediatr, 135, 189–96. 85. M¨uller, T., van de Sluis, B., Zhernakova, A., van Binsbergen, E., Janecke, A.R., Bavdekar, A., Pandit,

86.

87.

88.

89.

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A., Weirich-Schwaiger, H., Witt, H., Ellemunter, H., Deutsch, J., Denk, H., M¨uller, W., Sternlieb, I., Tanner, M.S. and Wijmenga, C. (2003) The canine copper toxicosis gene MURR1 does not cause non-Wilsonian hepatic copper toxicosis. J Hepatol , 38 (2), 164– 68. Kumar, N., Gross, J.B. Jr. and Ahlskog, J.E. (2004) Copper deficiency myelopathy produces a clinical picture like subacute combined degeneration. Neurology, 63, 33–39. Kumar, N., McEvoy, K.M. and Ahlskog, J.E. (2003) Myelopathy due to copper deficiency following gastrointestinal surgery. Arch Neurol , 60, 1782–85. Tan, J.C., Burns, D.L. and Jones, H.R. (2006) Severe ataxia, myelopathy, and peripheral neuropathy due to acquired copper deficiency in a patient with history of gastrectomy. JPEN J Parenter Enteral Nutr, 30, 446–50. Kelkar, P., Chang, S. and Muley, S.A. (2008) Response to oral supplementation in copper deficiency myeloneuropathy. J Clin Neuromuscul Dis, 10, 1–3.

16

The Central Role of the Liver in Iron Storage and Regulation of Systemic Iron Homeostasis Tracey A. Rouault1, Victor Gordeuk2 and Gregory Anderson3 1 National

Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD, USA 2 Howard University Medical Center, Washington, D.C., USA 3 Queensland Institute of Medical Research, PO Royal Brisbane Hospital, Brisbane, Queensland, Australia

INTRODUCTION The liver is a major iron repository in mammals, and advances in the last decade have revealed that the liver has an important role in sensing and regulating systemic iron homeostasis. Hepatocytes secrete a peptide hormone, hepcidin, which regulates efflux of iron from macrophages and absorption of iron across the intestinal mucosa. Signals from a variety of sources activate or repress the promoter of hepcidin to determine how much hepcidin mRNA is made, which in turn determines how much hepcidin peptide is synthesized and secreted into the circulation. When systemic iron levels are high, hepcidin levels are high, while low systemic iron levels are associated with diminished hepcidin secretion. Other variables such The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

as inflammation can increase hepcidin expression, which is important in the pathogenesis of the anemia of chronic disease. Thus, net expression of hepcidin represents integration at the hepcidin promoter of many important variables. In this chapter, we will review the role of the liver in systemic iron metabolism and discuss the signaling pathways unique to hepatocytes that enable the liver to coordinate regulation of overall systemic iron homeostasis.

THE LIVER AS A MAJOR IRON REPOSITORY In healthy adult humans, the liver is estimated to store from 0.07 to 0.4 g of iron. A combined analysis of

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previous studies using quantitative phlebotomy indicated that total body median storage iron was approximately 0.8 g among 39 normal men predominantly in the third and fourth decades of life, and 0.4 g among 20 normal women of similar age [1–5]. Based on the assumption that one-third of iron stores are normally in the liver, this would translate to a normal median hepatic iron content of 0.27 g for men and 0.13 g for women. Iron is sequestered within the iron storage protein, ferritin, a large 24 subunit protein composed of ferritin L (for ‘predominant in liver’) and H (for ‘predominant in heart’) chains (Figure 16.1). The ferritin subunits form a spherical protein shell that contains numerous channels. On the inside of the spherical protein, iron is oxidized from the ferrous (2+) form to the ferric (3+) form by the ferroxidase activity of the ferritin H chain, and insoluble ferric iron deposits grow from their initial deposition sites on side-chains of ferritin L subunits [6, 7]. As ferritin iron uptake and oxidation continue, thousands of ferric iron atoms accumulate within the ferritin sphere, which has the capacity to store up to 4500 iron atoms. As the iron contained within ferritin is not very bioavailable, ferritin sequesters iron and reduces the availability of cytosolic iron to interact with oxygen and generate harmful reactive oxygen species. Ferritin not only sequesters iron, but also serves as a source of iron for the metabolic needs of the cell. Iron can be released from intact ferritin that has been monoubiquitinated and will be ultimately degraded by the proteasome [8], or else ferritin multimers can undergo degradation in lysosomes and release iron after the ferritin heteropolymer has been degraded in the lysosome [9].

SERUM TRANSFERRIN IS THE MAJOR SOURCE OF IRON FOR TISSUES Transferrin (Tf) is a high-affinity binding protein for ferric iron that circulates in the plasma. It is synthesized mainly by hepatocytes in the liver and is secreted into the circulation. The plasma usually contains a significant excess of Tf, such that only about one-third of the iron binding sites on Tf are occupied by iron. Tf contains two lobes, each of which contains a ferric (Fe3+) iron-binding site [11]. Cells in tissues throughout the body can obtain iron from circulating Tf by increasing expression of transferrin receptors (TfRs). TfR1 is the major receptor for Tf and binds diferric Tf with high affinity (107 –109 M−1 ) at physiological pH. Monoferric and apoTf are bound with much lower affinity (106 and 10%) increases and that of bilirubin

diglucuronide decreases. In all disorders caused by UGT1A1 deficiency, biliary excretion of unconjugated bilirubin increases, resulting in increased incidence of pigment stones, as well as cholesterol stones with a bile pigment nidus. Both conjugated and unconjugated bilirubin accumulate in plasma in acquired inflammatory or cholestatic diseases of the liver. In these cases, other components of the bile, especially bile salts, also accumulate. In DJS, a benign condition caused by inherited ABCC2 deficiency, conjugated bilirubin accumulates in plasma, but the total bile salt concentrations remain normal. In addition, a dark-brown pigment accumulates in hepatocytes, giving the liver a characteristic black appearance. Urinary coproporphyrin excretion remains normal in DJS, but the ratio of isoform I–III is reversed, so that isoform I becomes more abundant [25]. Rotor syndrome is a genetically unrelated autosomal recessive disorder in which hepatic bilirubin storage is abnormal. There is no accumulation of pigments in hepatocytes. Urinary porphyrin excretion is increased and the ratio of isoforms I–III is approximately 1 : 1, which is similar to that found in many acquired liver diseases. In addition to DJS, which is a harmless disorder, genetic lesions of several other canalicular export pumps are associated with various degrees of hepatobiliary injury and consequent accumulation of both conjugated and unconjugated bilirubin in plasma. Three types of severe progressive familial intrahepatic cholestasis (PFIC I, II and III) are caused by genetic lesions of both alleles of the ATP8B1 (also termed FIC-1), ABCB11 (bile salt export pump (BSEP)), and ABCB4 (multidrug resistance protein 3 (MDR3)), respectively [26–28] (for details, see Chapter 42 by Gissen and Knisely). While PFIC syndromes present in infancy and require liver transplantation as a life-saving procedure, heterozygous lesions of the ATP8B1 and ABCB4 genes, or genetic lesions that only partially affect the function of these proteins, can cause more chronic and relatively benign disorders presenting in adolescents or adults. These include benign recurrent intrahepatic cholestasis type I (BRIC-I) and benign recurrent intrahepatic cholestasis type II (BRIC-II), which are associated with mutations of ATP8B1 and ABCB11 , respectively [29, 30]. Pancreatitis is an extrahepatic component of BRIC-I, while BRIC-II is associated with a high risk of cholelithiasis. Heterozygous mutations of ABCB4 are found in approximately one-third of patients with unexplained recurrent cholestasis, cholesterol gall stones, or biliary sludge [31]. Low biliary phospholipids-associated cholelithiasis, which is recognized as a clinical syndrome, may be caused by absent or reduced ABCB4 activity. Two other genetic lesions are associated with bile canalicular and structural anomalies: Alagille syndrome and abnormalities of Villin gene expression. Alagille syndrome is characterized by the paucity of interlobular bile ducts and chronic cholestasis, cardiac anomalies

17: DISORDERS OF BILIRUBIN METABOLISM

(most commonly peripheral pulmonic stenosis), butterfly vertebrae, prominent Schwalbe line of the eye, and abnormal facies. Liver transplantation may be required in patients with progressive liver synthetic dysfunction, intractable pruritus, osteodystrophy, or massive variceal bleeding. Genetic lesions of JAG1 are responsible for Alagille syndrome [32] (for details, see Chapter 42).

ACKNOWLEDGMENT This work was supported in part by NIH grants DK-46057 and DK-39137.

REFERENCES 1. London, I.M., West, R., Shemin, D. and Rittenberg, D. (1950) On the origin of bile pigment in normal man. J Biol Chem, 184, 351. 2. Kappas, A. and Drummond, G.S. (1995) Direct comparison of Sn-mesoporphyrin, an inhibitor of bilirubin production, and phototherapy in controlling hyperbilirubinemia in term and near-term newborns. Pediatrics, 95, 468. 3. Valaes, T., Petmezaki, S., Henschke, C., Drummond, G.S. and Kappas, A. (1994) Control of jaundice in preterm newborns by an inhibitor of bilirubin production: studies with tin-mesoporphyrin. Pediatrics, 93, 1. 4. Westlake, D.W.S., Roxburgh, J.M. and Talbot, G. (1961) Microbial production of carbon monoxide from flavonoids. Nature, 189, 510. 5. Elbirt, K.K. and Bonkovsky, H.L. (1999) Heme oxygenase: recent advances in understanding its regulation and role. Proc Assoc Am Physicians, 111, 438. 6. Hayashi, S., Takamiya, R., Yamaguchi, T., Matsumoto, K., Tojo, S.J., Tamatani, T., Kitajima, M., Makino, N., Ishimura, Y. and Suematsu, M. (1999) Induction of heme oxygenase-1 suppresses venular leukocyte adhesion elicited by oxidative stress: role of bilirubin generated by the enzyme. Circ Res, 85, 663. 7. Roy Chowdhury, J., Roy Chowdhury, N. and Arias, I.M. (1980) Bilirubin conjugation in the spiny dogfish Squalus acanthias, the small skate Raja erinacea and the winter flounder Pseudopleuronectes americanus. Comp Biochem Physiol , 668, 523–28. 8. Bonnet, R.J., Davis, E. and Hursthouse, M.B. (1976) Structure of bilirubin. Nature, 262, 326. 9. Kuenzle, C.C., Weibel, M.H. and Pelloni, R.R. (1973) The reaction of bilirubin with diazomethane. Biochem J , 133, 357. 10. Breimer, L.H., Wannamethee, G., Ebrahim, S. and Shaper, A.G. (1995) Serum bilirubin and risk of ischemic heart disease in middle-aged British men. Clin Chem, 41, 1504. 11. Temme, E.H.N., Zhang, J., Schouten, E.G. et al. (2001) Serum bilirubin and 10-year mortality risk in a Belgian population. Cancer Causes Control , 12, 887.

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12. Zucker, S.D., Horn, P.S. and Sherman, K.E. (2004) Serum bilirubin levels in the US population: gender effect and inverse correlation with colorectal cancer. Hepatology, 40, 827. 13. Shapiro, S.M., Bhutani, V.K. and Johnson, L. (2006) Hyperbilirubinemia and kernicterus. Clin Perinatol , 33, 387. 14. Lee, K.-S. and Gartner, L.M. (1983) Management of unconjugated hyperbilirubinemia in the newborn. Semin Liver Dis, 3, 52. 15. Wang, P., Kim, R.B., Roy-Chowdhury, J. and Wolkoff, A.W. (2003) The human organic anion transport protein SLc21A6 is not sufficient for bilirubin transport. J Biol Chem, 278, 20695. 16. Bosma, P.J., Seppen, J., Goldhoorn, B. et al. (1994) Bilirubin UDP-glucuronosyltransferase 1 is the only relevant bilirubin glucuronidating isoform in man. J Biol Chem, 269, 17960. 17. Ritter, J.K., Chen, F., Sheen, Y.Y., Tran, H.M., Kimura, S., Yeatman, M.T. and Owens, I.S. (1992) A novel complex locus UGT1 encodes human bilirubin, phenol, and other UDP-glucuronosyltransferase isozymes with identical carboxyl termini. J Biol Chem, 267, 3257. 18. Kadakol, A., Ghosh, S.S., Sappal, B.S., Sharma, G., RoyChowdhury, J. and Roy-Chowdhury, N. (2000) Genetic lesions of bilirubin uridinediphosphoglucuronate glucuronosyltransferase causing Crigler-Najjar and Gilbert’s syndromes: correlation of genotype to phenotype. Hum Mutat , 16, 297. 19. Gantla, S., Bakker, C.T.M., Deocharan, B., Thummala, N.R., Zweiner, J., Sinaasappel, M., Roy Chowdhury, J., Bosma, P.J. and Roy Chowdhury, N. (1998) Splice site mutations: a novel genetic mechanism of Crigler-Najjar syndrome type 1. Am J Hum Genet , 62, 585. 20. Bosma, P.J., Roy Chowdhury, J., Bakker, C., Gantla, S., DeBoer, A., Oostra, B.A., Lindhout, D., Tytgat, G.N.J., Jansen, P.L.M., Oude Elferink, R.P.J. and Roy Chowdhury, N. (1995) A sequence abnormality in the promoter region results in reduced expression of bilirubin-UDP-glucuronosyltransferase ∼1 in Gilbert syndrome. N Engl J Med , 333, 1171. 21. Stoll, M.S., Lim, C.D. and Gray, C.H. (1977) Chemical variants of the urobilins, in Bile Pigments, Chemistry and Physiology (eds P.D. Berk and N.I. Berlin), US Government Printing Office, Washington, DC, p. 483. 22. Huang, W., Zhang, J., Chua, S.S., Qatanani, M., Han, Y., Granata, R. and Moore, D.D. (2003) Induction of bilirubin clearance by the constitutive androstane receptor. Proc Natl Acad Sci U S A, 100, 4156. 23. Xie, W., Yeuh, M.F., Radominska-Pandya, A., Saini, S.P., Negishi, Y., Bottroff, Cabrera, G.Y., Tukey, R.H. and Evans, R.M. (2003) Control of steroid, heme, and carcinogen metabolism by nuclear pregnane X receptor and constitutive androstane receptor. Proc Natl Acad Sci U S A, 100, 4150. 24. Roy-Chowdhury, J., Locker, J. and Roy-Chowdhury, N. (2003) Nuclear receptors orchestrate detoxification pathways. Dev Cell , 4, 607.

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25. Kondo, T., Kuchiba, K. and Shimizu, Y. (1976) Coproporphyrin isomers in Dubin-Johnson syndrome. Gastroenterology, 70, 1117. 26. Eppens, E.F., van Mil, S.W., de Vree, J.M. et al. (2001) FIC1, the protein affected in two forms of hereditary cholestasis, is localized in the cholangiocyte and the canalicular membrane of the hepatocyte. J Hepatol , 35, 436. 27. Wang, L., Soroka, C.J. and Boyer, J.L. (2002) The role of bile salt export pump mutations in progressive familial intrahepatic cholestasis type II. J Clin Invest , 110, 965. 28. Jacquemin, E., De Vree, J.M., Cresteil, D. et al. (2001) The wide spectrum of multidrug resistance 3 deficiency: From neonatal cholestasis to cirrhosis of adulthood. Gastroenterology, 120, 1448. 29. Bull, L.N., van Eijk, M.J., Pawlikowska, L., DeYoung, J.A., Juijn, J.A., Liao, M., Klomp, L.W., Lomri, N., Berger, R., Scharschmidt, B.F., Knisely, A.S., Houwen,

R.H. and Freimer, N.B. (1998) A gene encoding a P-type ATPase mutated in two forms of hereditary cholestasis. Nat Genet , 18, 219. 30. Van Mil, S.W., Van Der Woerd, W.L., Van Der Brugge, G. et al. (2004) Benign recurrent intrahepatic cholestasis type 2 is caused by mutations in ABCB11. Gastroenterology, 127, 379. 31. Ziol M., Barbu, V., Rosmorduc, O., Frassati-Biaggi, Barget, N., Hermelin, B., Scheffer, G.L., Bennouna, S., Trinchet, J., Beaugrand, M., Ganne-Carrie, N. (2008) ABCB4 heterozygous gene mutations associated with fibrosing cholestatic liver disease in adults. Gastroenterology, 135, 131. 32. Oda, T., Elkahloun, A.G., Meltzer, P.S. and Chandrasekharappa, S.C. (1997) Identification and cloning of the human homolog (JAG1) of the rat Jagged1 gene from the Alagille syndrome critical region at 20p12. Genomics, 43, 376.

18

Hepatic Fatty Acid Metabolism and Dysfunction David L. Silver Department of Biochemistry, Albert Einstein College of Medicine, Bronx, New York, NY, USA

INTRODUCTION The liver is central to the maintenance of whole-body energy balance through its regulated supply of glucose, ketone bodies, and lipoprotein triglycerides (TGs) to peripheral tissues. The liver can be considered a central processing station for the storage and metabolism of dietary-derived lipids and glucose. In conditions of energy excess, dietary-derived and adipose-derived fatty acids as well as excess dietary glucose are taken up by the liver and stored in the form of glycogen or metabolized to TGs. However, under conditions of energy insufficiency, such as during a fast, adipose-derived fatty acids flux back to the liver and are esterified to glycerol and transiently stored in TGs, leading to a concomitant increase in the synthesis of very low density lipoprotein(VLDL) production and secretion, oxidation of fatty acids, and ketogenesis. Thus, proper metabolic handling of fatty acids by the liver is essential to maintaining organismal energy balance. Impairment in fatty acids storage in the liver may pose an important risk factor for insulin resistance, type 2 diabetes, non-alcoholic fatty liver disease (NAFLD), and atherosclerosis. This review will provide a current overview of hepatic fatty acid and TG metabolism and storage, mechanisms regulating hepatic lipogenesis and oxidation, and mechanisms giving rise to dysfunction of hepatic TG and fatty acid metabolism and their pathophysiological consequences.

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

HEPATIC TRIGLYCERIDE BIOSYNTHESIS AND METABOLISM Under energy-sufficient conditions, such as in the post-prandial state, TG biosynthesis begins with the ATP-dependent carboxylation of acetyl-CoA by acetyl-CoA carboxylase (Plate 18.1) (ACC). Two genes encode ACC, ACC1 and ACC2. ACC1 is highly expressed in the liver, while ACC2 is highly expressed in skeletal muscle [1]. The product of the ACC reaction is malonyl-CoA, which exerts allosteric inhibition on carnitine palmitoyl-CoA transferase, the rate-limiting enzyme in mitochondrial long-chain fatty acid import and hence plays a pivotal role in inhibiting mitochondrial fatty acid oxidation in energy-sufficient states [2]. Malonyl-CoA is the substrate for fatty acid synthase (FAS), which mediates the enzymatic condensation of malonyl-CoA with acetyl-CoA in the first round of 2-carbon addition, followed by consecutive additions of malonyl-CoA to the growing fatty acyl chain. Thiolase activity of FAS releases palmitate (C16:0), the first product of long-chain fatty acid biosynthesis. Palmitate can be modified by endoplasmic reticulum (ER) or mitochondrial fatty acid elongases (e.g. elov5, elov6) and desaturated at the delta 9 position by stearoyl-CoA desaturase (e.g. SCD-1). The first step in de novo TG biosynthesis involves

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the esterification of acyl-CoAs to glycerol-3-phosphate associated with mitochondria or ER, by a family of enzymes called acyl-CoA:glycerol-sn-3-phosphate acyltransferase (GPAT) to produce lysophosphatidic acid (LPA). LPA is also produced by the peroxisomeassociated dihydroacetyl-3-phosphate reductase. LPA is further esterified with acyl-CoA by the action of acyl-CoA:1-acylglycerol-sn-3-phosphate acyltransferases (AGPATs) localized to the ER and mitochondria to produce phosphatidic acid (PA). Since glycerolipid biosynthesis constitutes a branched pathway, PA can be utilized for the biosynthesis of phospholipids. PA is dephosphorylated to produce diacylglycerol (DAG) by a family of PA phosphatases belonging to the lipin gene family [3]. The role of lipin in lipogenesis and gene expression has recently been thoroughly reviewed [3]. The committed step in TG biosynthesis utilizes DAG by the diacylglycerol acyltransferase (DGAT) enzymes DGAT1 and DGAT2 to produce TG [4, 5]. Because of the anhydrous nature of TG, it is rapidly stored within cytoplasmic lipid droplets [6]. Under energy-limiting conditions, such as during a fast, the liver transiently acquires fatty acids released from stored TG in lipid droplets (LDs) by catecholamine activation of hormone-sensitive lipase (HSL) in adipose tissue. Fatty acids taken up by the liver are thio-esterified to coenzyme-A by a family of acyl-CoA synthetases, some of which have been found on cytosolic LDs [7, 8]. Excessive amounts of acyl-CoAs are transiently stored as TG through the action of the acyl-CoA transferases described above. Acyl-CoA import in the mitochondria through CPT1 and CPT2 is rate-limiting for the oxidation of long-chain fatty acids. Acetyl-CoA produced from the β-oxidation of fatty acids can be enzymatically converted to HMG-CoA through the action of HMG-CoA synthetase in mitochondria. HMG-CoA is decarboxylated to produce acetoacetyl-CoA. Acetoacyl-CoA can be interconverted through reduction of the beta-carbonyl group to beta-hydroxybuterate-CoA, and the CoA removed through thiolysis. Both of these products of fatty acid metabolism are classified as ketone bodies. Ketone bodies are not utilized by the liver as an energy source because the liver does not express succinyl-CoA transferase, but are released from the liver for use as energy by heart, skeletal muscle, and brain. In addition to β-oxidation of fatty acids during a fast, the liver will export fatty acids in the form of core TG on newly synthesized VLDL into plasma, which delivers the fatty acids to tissue through the hydrolysis of core TG by lipoprotein lipase, or through endocytic uptake of whole VLDL particles [9].

CYTOPLASMIC TRIGLYCERIDE LIPID DROPLETS The biochemical steps involved in TG synthesis are evolutionarily conserved from S. cerevisiae to humans; so

too is the process of TG storage within cells, in structures commonly referred to as cytoplasmic LDs. Since LDs are the intracellular depots for storage of TG, they play an important role in maintaining whole-body energy balance through the regulated hydrolysis of LD TG by hydrolases, resulting in increased fatty acid availability. However, excessive storage of LD TG in liver and skeletal muscle in humans is associated with insulin resistance [10]. LDs are composed of a monolayer of phospholipids with acyl chains of greater saturation than that found in cellular membranes, surrounding a neutral lipid core composed of combinations of TGs, ether lipids, cholesteryl esters, and retinyl esters, depending on cell type [11–13]. Interestingly, LDs have proportionately more phosphatidylcholine and lysophsopholipids but lower amounts of sphingolipids, phosphatidylethanolamine, and phosphatidylserine than cellular membranes [12]. In addition to the unique lipid composition of LDs, they are composed of a unique proteome. The most well characterized LD-associated proteins are members of the PAT family, with perilipin being the first family member identified and the best characterized of the five known members: perilipin, ADRP, TIP47, S3-12, and OXPAT [14] (Plate 18.2). Perilipin is highly expressed in adipocytes, where it surrounds the LD and is important for both the inhibition of lipolysis of LD TGs, and the catecholamine-induced activation of LD lipolysis by HSL [15]. HSL-deficient mice have decreased levels of TG in adipose tissue, but increased DAG, indicating the existence of a second lipase that preferentially utilizes TG as substrate [16]. This second lipase was identified and named adipose tissue triglyceride lipase/desnutrin (ATGL, also called PNPLA2, iPLA2, or TTS2.2)[17–19]. ATGL is a member of the patatin-domain-containing family of lipases that interacts with LDs [20]. Interestingly, ATGL-deficient mice have significant increases in adipose and liver TG, but massive increases in cardiac TG, which leads to cardiac hypertrophy and failure [21]. This finding may be interpreted as indicating that other lipases, such as HSL in adipose, only partially compensate for the absence of ATGL in adipose, skeletal muscle, and liver. In addition, these results indicate that LD storage and turnover is highly active in heart, with the latter pathway dependent on ATGL activity [22]. Similar to ATGL deficiency in mice, humans carrying homozygous inactivating mutations in ATGL present with multisystem TG accumulation, including hepatomegaly and cardiac dysfunction [23]. Perilipin is not expressed in the liver. ADRP is ubiquitously expressed in tissues and has been shown to be important for maintaining hepatic TG levels. Mice expressing mutant ADRP have a 57% reduction in hepatic TG compared to control mice, and are still able to produce LDs, albeit in significantly reduced amounts, when fed a high-fat diet. ADRP-deficient hepatocytes have an upregulation of TIP47, raising the possibility that TIP47 may functionally compensate for the absence of ADRP to regulate LD accumulation [24, 25]. Unlike in adipose tissue, the identity

18: HEPATIC FATTY ACID METABOLISM AND DYSFUNCTION

259

Lipid droplets Lipid droplets

DGAT

DGA

FIT

FIT

cytosol

cytosol

ER lumen ER lumen

Figure 18.1 A model for lipid droplet biogenesis. Triglycerides that are synthesized at the ER by the enzymatic action of diacylglycerol acyltransferase enzymes (DGAT) can be directed into cytosolic lipid droplets. TAG expands between the leaflets of the ER (shown as white sphere in membrane) and bud off into cytosolic lipid droplets, potentially requiring the action of FIT proteins. Lipid droplets then acquire PAT family proteins and additional lipid droplet-associated proteins (shown as shaded ring around droplets)

of the lipases controlling hepatic LD turnover has not been determined. Moreover, the physiological signals mediating TG storage and turnover in liver are different than in adipose. Fasting in mice induces LD lipolysis in adipose tissue, while LD biosynthesis is transiently increased in the fasted liver of mice [26]. A major question with regard to the biology of LDs concerns the mechanisms responsible for their biogenesis. A favored model of LD formation is that TGs synthesized from DGATs in the ER accumulate between the leaflets of the ER membrane, which eventually buds off the ER into the cytoplasm [14] (Figure 18.1). This model satisfies two important criteria for LD biogenesis. First is the location of LD biosynthesis to the ER, because the ER is the site of TG biosynthesis where DGAT enzymes are localized. Indeed, LDs are commonly visualized by EM methods to be in close association with ER membranes [27]. Second is the requirement for a phospholipids monolayer, which could be derived from the outer leaflet of the ER membrane. However, no experimental evidence is known that strictly supports this model. A different model for LD formation has been proposed, which shares similarities with the mechanism by which specific prokaryotes, such as nocardioforms, produce LDs that use soluble wax ester synthases and DGAT-like enzymes to synthesize TG in close association with the plasma membrane [28]. EM studies in Rhodococcus opacus have shown LDs forming on the inner leaflet of the plasma membrane without evidence for LD budding [29]. Recent studies using EM immunogold localization of ADRP in macrophage have suggested that specific ER domains are sites of LD formation and that LD formation may occur on the outer leaflet of the ER membrane [30]. Recently, several forward genetic screens in S. cerevisiae using gene-targeted mutant libraries, and in drosophila cells using genome-wide RNAi knockdowns have uncovered a large array of functionally diverse proteins that appear to be involved in controlling

LD morphology and lipolysis [31–33]. It has been estimated that 1.5% of the drosophila genome encodes for proteins that play a role in LD biology [33]. This estimate is staggering and underscores the multiple functions and complexity of LDs within cells not limited to fat storage. Thus far, these genetic screens have not identified proteins that are sufficient or necessary for the formation of LDs. Recently, an evolutionarily conserved family of ER-resident membrane proteins named fat-inducting transcripts (FITs) 1 and 2 has been identified that meets several criteria for proteins that may constitute part of the machinery necessary for the formation of LDs [34]: (i) FIT genes are evolutionarily conserved from S. cerevisiae to humans; (ii) FIT proteins are exclusively localized to the ER, the proposed subcellular site for LD biosynthesis; (iii) one of the FIT proteins, FIT2, is ubiquitously expressed in mouse and human tissues; and (iv) overexpression of FIT proteins induces LD formation without affecting TG biosynthesis. The molecular mechanism by which FIT proteins mediate LD formation, as well as their physiological and pathophysiological role in TG storage in tissues such as the liver, remains to be determined. Several other proteins have recently been implicated in the regulation of LD formation or metabolism, such as Fsp27 [35, 36], seipin [31, 32], prp19 [37], and an Arf1 family member, Arf79F [33].

REGULATION OF HEPATIC TRIGLYCERIDE BIOSYNTHESIS The regulation of the TG content of the liver is under tight control by nutritional signals such as carbohydrates, fatty acids, and insulin. The liver has two main mechanisms to reduce hepatocyte TG content: VLDL biosynthesis and export into plasma, and mitochondrial and peroxisome oxidation of fatty acids. Both of these mechanisms

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are upregulated in fasting and downregulated in feeding. Fasting-induced upregulation requires two transcription factors, Foxo1 and peroxisome proliferators-activated receptor alpha (PPARα). Foxo1 is a member of the forked head–winged X factor that is a direct substrate of Akt/PKB phosphorylation, such that in the fed state, insulin signaling induces the activation of Akt and phosphorylation of Foxo1 and nuclear exclusion [38]. In the fasted state, with low levels of insulin, non-phosphorylated Foxo1 localizes to the nucleus and activates genes important in gluconeogenesis, such as pyruvate carboxylase and phosphoenolpyruvate carboxykinase (PEPCK), but also plays an important role in positively regulating hepatic VLDL secretion [39]. During fast, VLDL secretion by the liver provides heart and skeletal muscle with energy in the form of fatty acids. In addition to VLDL secretion, fatty acids are released from adipocytes and taken up by the liver, or hydrolyzed from LDs within hepatocytes, oxidized to ketone bodies, and released into blood. Oxidation of fatty acids for ATP production or ketogenesis is under tight regulation by PPARα [40]. Unsaturated fatty acids are activators of PPARα [41], although the identity of endogenous PPARα ligands is still under debate [42]. Recently, it has been demonstrated that induction of ketogenesis by the liver requires fibroblast growth factor 21 (FGF21). Importantly, FGF21 is a direct target of PPARα and may constitute a feed-forward regulatory loop in which FGF21 may serve to sustain ketogenesis during fast, but the mechanism by which FGF21 signals to activate ketogenesis remains to be determined [43, 44]. Hepatic TG biosynthesis in the fed state is primarily regulated through two other transcription factors, sterol regulatory element binding protein 1c (SREBP1c) and carbohydrate response element binding protein (ChREBP) (Plate 18.3). Insulin induces hepatic lipogenesis through the upregulation of SREBP1c. SREBP1c activates genes involved in fatty acid biosynthesis, such as ACC and FAS, the two rate-limiting enzymes in fatty acid biosynthesis [45]. The effect of insulin on SREBP1c expression is mediated in large part by liver X receptor (LXR) [46]. In the liver, LXR controls important steps in the oxidative conversion of cholesterol to bile acids by regulating the expression of cyp7alpha, the rate-limiting enzyme in bile acid synthesis, high density lipoprotein (HDL) biosynthesis, and cholesterol excretion into bile [47]. Mice treated with LXR agonists show increased biliary cholesterol excretion and plasma HDL, but markedly enhanced hepatic TG content as a result of enhancement of SREBP1c expression [47]. Thus, LXR plays a central role in regulating hepatic cholesterol metabolism and lipogenesis. SREBP1c-deficient mice have a 50% decrease in hepatic fatty acid synthesis, indicating the existence of another regulatory pathway for hepatic lipogenesis [48]. Interestingly, overexpression of a constitutively active form of SREBP1c in hepatocytes that lack glucokinase, the enzyme essential for the first step in glycolysis, have

a reduced induction of lipogenic gene expression in addition to reduced glycolytic gene expression, indicating that a second transcriptional mechanism that requires a product of glucose metabolism is important for hepatic lipogenesis in response to carbohydrate [49]. This second pathway was shown to be due to the activity of ChREBP [50]. ChREBP is activated by both glucose and xylulose-5-phosphate, a product of the pentose phosphate shunt, indicating that ChREBP maintains hepatic fatty acid biosynthesis gene expression in the face of excess calories in the form of carbohydrates, and is thus critical for hepatic glucose sensing [50, 51]. The role of ChREBP in hepatic steatosis was examined by shRNA knockdown in ob/ob mice. ob/ob mice are leptin deficient, insulin resistant, and hyperglycemic, and have severe hepatic steatosis and overexpress ChREBP and SREBP1c [52, 53]. Knockdown of ChREBP in ob/ob livers resulted in a significant decrease in hepatic steatosis, associated with decreased expression of ACC, FAS, and SCD-1 without a change in nuclear SREBP1c levels [53]. SCD-1 is rate limiting for the formation of mono-unsaturated fatty acids utilized for TG biosynthesis. Interestingly, hepatic knockout of SCD-1 reduces hepatic steatosis in mice fed high-fat or high-carbohydrate diets and improves hepatic insulin sensitivity [54–56]. These expected decreases in lipogenic gene expression in ChREBP knockdown livers were met with increases in β-oxidation. ChREBP knockdown livers had decreased levels of ACC1 and ACC2 protein, correlating with decreased malonyl-CoA and increased plasma beta-hydroxybuterate in fasted mice [53], likely resulting in de-inhibition of CPT1, the enzyme constituting the rate-limiting step in mitochondrial β-oxidation. These conclusions are supported by the finding that ACC2-deficient mice have increased rates of β-oxidation in liver [57]. While clearly SREBP1c and ChREBP contribute to regulating hepatic lipogenesis, the quantitative contribution for each transcription factor in this process has not entirely been defined. It may be that SREBP1c and ChREBP allow the liver to fine-tune hepatic lipogenesis by perceiving nutrient status through both insulin (via SREBP1c) and cellular glucose metabolism (via ChREBP).

PATHOBIOLOGY OF HEPATIC TRIGLYCERIDE Hepatic steatosis in NAFLD is highly associated with obesity, insulin resistance, and type 2 diabetes [58]. The cause–effect relationship of fatty liver and insulin resistance has not been entirely resolved. Donnelly et al. [59] determined that the majority (60%) of the fatty acids contributing to hepatic fat in NAFLD patients are plasma-derived, likely from lipolysis in adipose tissue, but a significant number are de novo synthesized by the liver. While it is clear that hyperinsulinemia in mice induces

18: HEPATIC FATTY ACID METABOLISM AND DYSFUNCTION

hepatic expression of SREBP1c, leading to enhanced lipogenic gene expression and fatty liver, fatty liver in the absence of hyperinsulinemia can be found in association with hepatic insulin resistance [46, 58]. Hepatic insulin resistance is characterized by enhanced hepatic glucose production and VLDL secretion; the former is a major cause in failure of glucose control over time, while the latter is a likely cause of increased incidence of atherosclerosis in most diabetic patients [60]. Hepatic deficiency of SREBP1 in ob/ob mice was found to significantly ameliorate steatosis and hepatic TG levels, but exacerbated hepatic insulin resistance in this mouse model [61]. These findings support the idea that hyperinsulinemia drives hepatic lipogenesis in a SREBP1c-dependent fashion in the milieu of hepatic insulin resistance manifested by increased glucose production, and suggests that hepatic fat per se is not directly causal in hepatic insulin resistance. The idea that the diabetic liver has a mixture of insulin resistance and sensitivity [62] has gained new momentum with recent reports describing a liver-specific insulin receptor knockout (LIRKO) mouse model. LIRKO mice have hepatic insulin resistance, as shown by increased gluconeogenic gene expression and fasting hepatic glucose production, with decreased lipogenic gene expression associated with decreased SREBP1c expression and a subset of SREBP1c targets such as FAS and SCD-1 in non-fasted mice [63]. Interestingly, LIRKO livers, similarly to type 2 diabetics, have increased VLDL and apoB secretion [63]. However, TG content in LIRKO livers is similar to that in wild-type mice, which may be explained by the observation that LIRKO mice have increased hepatic expression in DGAT and GPAT [63]. Contrary to type 2 diabetics, LIRKO livers secrete an unusual VLDL particle that is TG poor and enriched in cholesterol [63]. Nonetheless, the LIRKO study clearly defines insulin signaling as a lipogenic inducer in the liver. An important question that has been raised is which components of insulin signal transduction regulate glucose production (gluconeogenesis and glycogenolysis) and lipogenesis in the liver? The hepatic control of glucose production and lipogenesis by insulin signaling has previously been viewed as being mediated by separate branches of the insulin signal transduction pathway, with insulin receptor substrate (IRS) 1 primarily controlling VLDL and TG production, and IRS2 regulating hepatic glucose production (Figure 18.2). This view has recently been refuted by elegant studies utilizing liver-specific IRS1, IRS2, and double IRS1/2 knockout (KO) mice. Liver-specific IRS1 KO mice have enhanced gluconeogenesis in the fed state, but normal gluconeogenesis in the fasted state [64]. Liver-specific IRS2 KO mice show an opposite effect, with enhanced gluconeogenesis in the fasted state, but unchanged gluconeogenesis in the fed state [64]. SREBP1c expression was downregulated in fed liver-specific IRS1 KO mice, and fasted liver-specific IRS2 KO mice. Double liver-specific IRS1/2 KO mice have severe hepatic insulin resistance, with decreased hepatic glucose production, SREBP1c expression,

(a)

261

(b)

Figure 18.2 A revised model of hepatic Irs function. (a) The prevailing model dictated that Irs2 and Foxo1 regulated glucose production, while Irs1 and Foxa2 regulated lipogenic gene expression and lipoprotein production. (b) The revised model indicates that Irs1 and Irs2 essentially regulate similar pathways, but under different physiological conditions (fasting or feeding). Mice doubly deficient in hepatic Irs1 and Irs2 have decreased triglycerides and plasma VLDL, supporting the idea that excessive insulin signaling leads to plasma lipid abnormalities seen in type 2 diabetes. Reproduced from Haeusler & Accili, Cell Metab 8:7–9, Copyright (2008), with permission from Elsevier

and VLDL secretion, supporting the model that IRS1 and IRS2 do not regulate separate metabolic pathways in response to changing insulin levels, but rather a single linear pathway that responds to feeding (IRS1) or fasted (IRS2) conditions [64]. Moreover, IRS signaling appears to converge on Foxo1, defining Foxo1 as the major regulator of hepatic gluconeogenic and lipogenic gene expression, although the exact role of Foxo1 in regulating lipogenesis remains to be determined [65]. These studies utilizing liver-specific IRS KO mice further support the notion that excessive hepatic insulin signaling, and not insulin resistance, is a major force in the etiology of dyslipidemia found in type 2 diabetes.

ADIPOSE FAT STORAGE AND HEPATIC INSULIN RESISTANCE Excessive storage of TG in white adipose tissue is highly associated with insulin resistance and type 2 diabetes in humans [66]. Insulin resistance increases the risk of premature mortality from heart failure and atherosclerosis [67]. The occurrence of obesity and consequently insulin resistance is becoming a worldwide problem [68]. The mechanisms to explain the propensity of obese humans for having insulin resistance and co-morbidities (e.g. hypertension, dyslipidemia, cardiac dysfunction) are not entirely understood. Adipose tissue plays an essential role in whole-body glucose homeostasis through its ability to store excessive calories as TGs in cytosolic LDs,

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as well as to act as an endocrine organ to secrete insulinsensitizing hormones, known as adipokines, and as an organ that regulates immune functions (cytokine secretion and tissue macrophage functions), while skeletal muscle is the principal tissue accounting for whole-body glucose disposal through the translocation of the Glut4 glucose transporter to the plasma membrane [69–71]. The TG stored in cytosolic LDs in adipose can have a major impact on whole-body glucose metabolism and insulin sensitivity because the free fatty acids (FFAs) stored as triacylglycerol (TAG) in LDs can be released from adipocytes into the circulation to influence insulin sensitivity and therefore glucose metabolism in skeletal muscle and liver, to regulate insulin secretion by the pancreas, and to enhance lipogenesis and lipoprotein secretion by the liver [69]. Therefore, adipose tissue is highly integrated into the control of whole-body glucose and lipid metabolism. It is currently thought that at some point in the development of obesity, adipose tissue no longer has the ability to safely store TG, resulting in the accumulation of TG in peripheral organs such as the liver, skeletal muscle, heart, and pancreas [72]. Such peripheral accumulation of TG is correlated with tissue dysfunction, also known as “lipotoxicity”, leading to decreased glucose uptake in skeletal muscle, hepatic steatosis, and cardiac dysfunction [73, 74]. An emerging view explaining correlations of cellular TAG with insulin resistance is that lipid metabolites such as acyl-CoAs, ceramide, and DAG antagonize insulin signal transduction [58, 75]. In the case of DAG, data indicate that it activates the novel protein kinase c (PKC)s PKCθ (in skeletal muscle) and epsilon (in liver) directly to inhibit insulin signaling, and that the storage of lipid metabolites in cytosolic LDs, through conversion of fatty acyl-CoAs and DAG to TAG, is beneficial to insulin sensitivity [58]. In a recent published study supporting this idea, overexpression of DGAT1, constituting the committed step in TG biosynthesis (DGAT2 has a similar function), in skeletal muscle increased intramyocellular triglycerol (IMTG) levels and reduced DAG and ceramide, while at the same time enhancing insulin sensitivity in mice on a high-fat diet [76]. In a separate model of fatty muscle, overexpression of DGAT2 in skeletal muscle increased IMTG, fatty acyl-CoAs, and ceramides, and impaired insulin signaling and sensitivity in skeletal muscle, supporting the idea that lipid metabolites, rather than IMTG accumulation, are likely causative factors in insulin resistance [77]. Hepatic TG storage, like in skeletal muscle, may be a surrogate marker for increased lipid metabolites. Transgenic overexpression of DGAT2 in livers of mice fed a standard chow diet resulted in fatty liver without hepatic insulin resistance. A similar finding was recently reported, in which liver-specific microsomal triglyceride transfer protein (MTP) KO mice fed a high-fat diet developed fatty liver, but maintained normal hepatic insulin sensitivity, supporting the idea that hepatic TAG alone is not sufficient to cause insulin resistance [78]. Shulman and co-workers have recently shown that

PKCε impairs insulin receptor activity and that transient knockdown of PKCε in rats fed a high-fat diet improves hepatic insulin signaling [79, 80]. By examining mice deficient in mitochondrial gpat, catalyzing the production of LPA, Shulman and co-workers have shown that gpat KO mice have decreased hepatic TAG accumulation and increased hepatic insulin sensitivity on a high-fat diet [81]. Gpat KO mice had significant decreases in hepatic DAG but increased fatty acyl-CoAs [81]. In an opposite experiment, overexpression of gpat transiently using adenovirus in rats resulted in fatty liver associated with hepatic insulin resistance (increased hepatic glucose production). Gpat overexpression in this model also resulted in increased levels of DAG and enhanced activation of PKCε, possibly providing a mechanism to explain hepatic insulin resistance (Figure 18.3). Together these studies with gpat overexpression and knockdown rodent models support the conclusion that DAG plays more of causative role in hepatic insulin resistance than do fatty acyl-CoAs, and that TAG is not causative in hepatic insulin resistance. However, studies in cell culture have indicated that hepatic fatty acids, particularly saturated fatty acids like palmitate, are lipotoxic and may contribute to hepatic insulin resistance [82, 83]. One explanation for the effects of saturated fatty acids on hepatic insulin resistance may be that saturated fatty acids like stearate and palmitate are preferentially utilized for ceramide synthesis, while mono-unsaturated fatty acids like oleate are utilized for phospholipid and TG biosynthesis [84]. A recent and interesting example that is counter to the idea that saturated Fatty acid

glucose Insulin receptor

AKT2 PKC-ε LCFA-CoA

P

DAG

FAS Foxo1

Foxo1

Gluconeogenesis

Gluconeogenic gene expression

nucleus

Figure 18.3 Model for the mechanism of fatty acidinduced hepatic insulin resistance. Increased hepatic pools of DAG derived from fatty acid uptake or de novo fatty acid biosynthesis mediated by fatty acid synthase (FAS) activate PKC-ε, which binds to and inactivates the insulin receptor kinase, resulting in reduced insulin signaling to AKT2. Reduced AKT2 activity results in decreased phosphorylation of Foxo1 and increased nuclear levels of Foxo1. Nuclear Foxo1 induces gluconeogenic gene expression, such as phosphoenolpyruvate carboxykinase, glucose-6-phosphatase, resulting in increased hepatic glucose production

18: HEPATIC FATTY ACID METABOLISM AND DYSFUNCTION

fatty acids are causative in hepatic insulin resistance is seen in studies involving elov6-deficient mice [85]. Elov6 KO mice fed a lipogenic diet (low fat, high carbohydrate) are protected from the development of hepatic insulin resistance, but paradoxically have increased hepatic levels of palmitate. Increased palmitate in elov6 mice may be explained by the decreased expression of SCD-1 found in these mice, and since SCD-1 hepatic deficiency protects rodents from hepatic insulin resistance, the protection seen in elov6 KO mice may be attributed to decreased SCD-1.

PPARγ AND NAFLD The idea that enhancing the capacity of adipose to store TG is important for maintaining insulin sensitivity is further supported by findings that humans and mice given the insulin-sensitizing drugs thiazolidinedione drugs (TZDs) to activate PPARγ have an increase in body weight due to expansion of adipose and increased TAG storage [86]. PPARγ is a member of the nuclear hormone receptor superfamily and is a master regulator of adipogenesis that is essential for the insulin sensitizing effects of TZDs, high-affinity PPARγ ligands that enhance adipocyte differentiation [86]. PPARγ1 and γ2 are encoded by the same gene but arise from alternative promoter usage, giving rise to an additional 28 or 30 kDa N-terminal extension of PPARγ2 [86]. PPARγ2 is adipose-specific in its expression, and increases during the first several days of adipogenesis of NIH 3T3-L1 cells [86, 87]. PPARγ activates an array of target genes that play roles in establishing the adipocyte phenotype (TG accumulation) and endocrine functions (adipokine expression) [88]. Mice with a whole-body deficiency of PPARγ or the adipose-specific isoform PPARγ2 have decreased adipose mass but are insulin resistant on a high-fat diet [89–92]. Similar findings in humans with PPARγ-activating and -inhibiting mutations support a beneficial role for PPARγ in regulating adipogenesis and insulin sensitivity [86, 93]. With regard to the role of PPARγ in liver insulin sensitivity and TG accumulation (fatty liver), liver-specific deficiency of PPARγ in ob/ob mice dramatically reduces hepatosteatosis seen in this model, but paradoxically enhances insulin resistance in muscle and adipose tissue [94]. Surprisingly and somewhat paradoxically, PPARγ +/− mice are protected from high-fat diet-induced weight gain and insulin resistance [95, 96]. In line with observations in PPARγ +/− mice, inhibition of PPARγ activity in wild-type mice using several antagonists resulted in decreased adiposity and enhanced insulin sensitivity on a high-fat diet, indicating a complex relationship between adipose TAG stores (i.e. body weight) and insulin sensitivity [86]. Nonetheless, treatment of PPARγ +/− mice with PPARγ antagonists resulted in the re-emergence of insulin resistance associated with ectopic deposition of TAG in liver and skeletal

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muscle [95]. Together, these studies imply that the expansion of adipose stores of TG following TZD treatment is part of the insulin-sensitizing effect of PPARγ activation. Overexpression of Pref-1, an inhibitor of adipogenesis, in mouse adipose in vivo resulted in insulin resistance, glucose intolerance, and hypertriglyceridemia in the presence of decreased adiposity [97]. Indeed, the observations that lipodystrophic humans and mice are insulin-resistant and have increased levels of TGs in non-adipose tissues further support the importance of adipose tissue in regulating whole-body insulin sensitivity [58]. The mechanisms by which PPARγ activation reduces hepatic steatosis are likely manyfold, including increasing subcutaneous adipose depots, enhancing fatty acid oxidation in muscle and liver, decreasing adipose cytokine expression, and enhancing expression of adiponectin implicated in directly sensitizing the liver to insulin [86]. A recent revealing study that connects the hepatic effects of adiponectin with regulation of body adiposity showed that overexpression of adiponectin in adipose tissue of insulin-resistant ob/ob mice reduced TAG and DAG levels in liver, increased glucose tolerance and insulin sensitivity, and decreased adipose tissue macrophage levels and inflammatory markers, all in the face of a paradoxical expansion of adipose mass (hyperplasia) and increased markers of adipose lipogenesis [98].

REGULATING THE BALANCE BETWEEN HEPATIC LIPOGENESIS AND FATTY ACID OXIDATION The development of hepatic steatosis may be partially attributed to an imbalance between hepatic fatty acid oxidation and biosynthesis that is tightly regulated by the activity of ACC. Two isoforms of ACC are expressed in humans and animals, ACC1 and ACC2. Both are encoded by separate genes, and ACC2 has an additional hydrophobic 136aa N-terminal extension [1]. ACC1 is cytosolic while ACC2 is associated with the mitochondria [1]. Mitochondrial localization of ACC2 has been shown to require 114aa of the N-terminal hydrophobic domain [1]. ACC1 is highly expressed in liver and adipose tissue, while ACC2 is primarily expressed in oxidative tissues such as skeletal muscle and heart. Consistent with its tissue expression pattern, ACC1 is primarily involved in the production of malonyl-CoA for fatty acid biosynthesis, while it has been thought that ACC2 is critical for the allosteric inhibition of CPT1 activity through the local production of malonyl-CoA at the outer mitochondrial membrane, which is also consistent with it being highly expressed in skeletal muscle [99]. ACC2 KO mice have decreased adiposity compared to wild-type controls due to increased β-oxidation in skeletal muscle, heart, and also liver, supporting the idea that decreased malonyl-CoA at

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the mitochondria is important for regulating CPT1 activity [100]. Moreover, ACC2 KO mice were found to be protected from diet-induced obesity and diabetes after high-fat/high-carbohydrate feeding, likely due to enhanced fatty acid oxidation [57]. Not surprisingly, a gene deletion of ACC1 on the other hand resulted in embryonic lethality, similar to deletion of FAS [101], indicating that de novo fatty acid biosynthesis is critical for embryogenesis [102]. Two liver-specific ACC1 KO mice have been independently generated to determine the contribution of ACC1 to de novo hepatic fatty acid biosynthesis. Both liver ACC1 KO mouse models showed a similar compensatory increase in ACC2 expression, making the assessment of ACC1 in de novo fatty acid biosynthesis difficult to determine [103, 104]. However, Mao et al. [104] did find in their ACC1 liver-specific KO mouse reduced hepatic TG levels (40% decrease), explained by a 50% decrease in de novo fatty acid biosynthesis in isolated hepatocytes and a larger decrease in hepatic malonyl-CoA concentrations (70% decrease). However, despite these decreases in de novo fatty acid biosynthesis, these liver-specific ACC1 KO mice were not protected from the development of fatty liver or insulin resistance from high-fat/high-carbohydrate feeding. The liver-specific ACC1 KO mice generated by Harada et al. [103] had no significant change in rates of hepatic lipogenesis, and no significant decrease in malonyl-CoA concentration, suggesting that malonyl-CoA synthesized by ACC2 can be utilized by FAS for de novo fatty acid biosynthesis. Whether ACC2 can contribute to hepatic de novo fatty acid biosynthesis in livers expressing endogenous ACC1 is not clear. In support of the idea that ACC1 activity is important for de novo fatty acid biosynthesis, transient knockdown of ACC1 and ACC2 using antisense oligonucleotides (ASOs) in primary rat hepatocytes in vitro showed that ACC1 knockdown was sufficient to inhibit de novo lipogenesis, but ACC2 knockdown was without effect [105]. A combined knockdown of ACC1 and ACC2 using ASO in vivo in rats resulted in protection from hepatic steatosis induced by high-fat feeding associated with reduced hepatic malonyl-CoA levels [105]. Importantly, and indicating a potential clinical benefit of targeting ACC, amelioration of hepatic steatosis was associated with decreased hepatic glucose production and increased phosphorylation of both Akt and Foxo1, indicating increased hepatic insulin sensitivity [105].

METABOLIC FATE OF “OLD” AND “NEW” FAT During fast, hepatocytes adapt to changing substrate availability by controlling the balance of glucose oxidation and fatty acid oxidation. PPARα plays a central role in upregulating genes involved in β-oxidation [106], ketogenesis [43, 107], and genes that negatively regulate

insulin sensing [108] (e.g. Trbl3) and glucose oxidation [109] (e.g. PDK4). PPARα is highly expressed in fatty acid oxidative tissues such as skeletal and cardiac muscle, but also in liver, supporting the idea that the liver plays an important role in fat oxidation. PPARα-deficient mice when fasted for prolonged periods of time are hypoglycemic and show decreased ketogenic and hepatic fatty acid oxidative gene expression, underscoring the critical role of PPARα in maintaining energy balance during fast [110]. Although endogenous ligands for PPARα remain to be definitively identified, polyunsaturated fatty acids are potent activators of PPARα [41], and presumably endogenous tissue ligands for PPARα are polyunsaturated fatty acids. Fatty acids in the liver can be derived from either de novo lipogenesis driven by dietary carbohydrates, dietary fat carried in the blood by chylomicrons, or fat stored in adipocytes. Semenkovich and co-workers recently proposed that dietary-derived fat—dietary carbohydrates for driving lipogenesis—is “new” fat, and fat stored in adipose tissue is “old” fat, with both having different metabolic fates or effects on hepatic lipid and glucose metabolism [42] (Figure 18.4). To test this hypothesis, Chakravarthy et al. generated liver-specific FAS KO mice (FASKOL) [42]. In addition to ACC, FAS is one of the rate-limiting enzymes in de novo fatty acid biosynthesis. FAS primed with acetyl-CoA catalyzes the repetitive condensation of malonyl-CoA to produce the saturated fatty acid palmitate (C16 : 0). Unexpectedly, the absence of FAS in liver did not protect FASKOL mice from hepatic steatosis when fed a lipogenic diet (zero fat/high carbohydrate), but exacerbated it compared to control mice. FASKOL mice had increased levels of hepatic malonyl-CoA, consistent with the possibility of

DIET

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Figure 18.4 Model for the distinct effects of hepatic fatty acids on PPAR activity. It is hypothesized that two metabolically distinct pools of fatty acids are stored in hepatocytes-“new fat” and “old fat.” “New fat” is derived from dietary absorption or de novo synthesis in the hepatocyte, while ”old fat” is derived from peripherally stored fatty acids in adipose. Both dietary and de novo synthesized fatty acids can activate PPARα, leading to increased fatty acid β-oxidation (FAO), gluconeogenesis (GNEO), and maintenance of cholesterol homeostasis (CHOL). Fat derived from peripheral stores does not activate PPARα as effectively as ”new fat,” leading to the development of fatty liver. Reprinted from [42], Copyright  2005, with permission from Elsevier

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decreased hepatic β-oxidation. Importantly, hepatic steatosis in FASKOL mice was reversed by PPARα activation with a synthetic ligand, raising the possibility that FAS directly provides fatty acid ligands for PPARα activation. In addition, prolonged fasting (24 hours) of FASKOL mice resulted in a similar phenotype to fasted PPARα KO mice, with hypoglycemia, hypoketonemia, and exacerbated fatty liver, that was reversible by PPARα activation with synthetic ligand. Taken together, the study of FASKOL mice shows that not all fatty acids are metabolically equal, such that fatty acid generated by FAS (“new” fat) provides ligands for PPARα, while “old” fat does not and is preferentially stored in LDs. Their work also raises the intriguing possibility that palmitate is an endogenous PPARα ligand.

LINKS BETWEEN UPR AND HEPATIC LIPOGENESIS The unfolded protein response (UPR) is an evolutionarily conserved pathway that allows cells to respond to unfolded and misfolded proteins in the ER by arresting ongoing protein synthesis, upregulating the synthesis of protein chaperones in the ER to assist with protein folding, and eliminating unfolded and misfolded proteins in the ER [111]. The UPR is comprised of three primary pathways: PERK, IRE1, and ATF6. ATF6 is a transcription factor that is proteolytically cleaved in response to stress signals by site-2 protease in the Golgi, the same protease that liberates the transcription factor domain of IRE-1α

ER

265

SREBPs. ATF6 activates XBP1, in addition to other targets, resulting in increased phospholipids synthesis, presumably allowing the cell to generate new ER membranes and cope with protein load. PERK and IRE1 (α and β) are ER-resident transmembrane kinases that are responsible for translational inhibition through phosphorylation of eukaryotic translation initiation factor eIF2α and production of the transcription factor XBP1, respectively. The activation of XBP1 by IRE1 occurs through a post-transcriptional mRNA splicing of the XBP1 mRNA by IRE1α, resulting in translation of XBP1 and its nuclear import [112, 113]. An initial link between UPR and lipid metabolism, in addition to commonalities between SREBPs and ATF6 proteolytic processing by S2P [114, 115], was indicated by the observation that ob/ob mice, which are leptin deficient, insulin resistant, and have severe fatty liver, also have increased molecular markers for UPR activation in liver such as PERK and IRE1α [116]. IRE1α also activates the c-jun amino-terminal kinase signaling pathway, leading to attenuation of insulin signaling in liver [116], supporting the concept that ER stress, TG accumulation in liver, and hepatic insulin resistance are connected. An unexpected link between XBP1 and hepatic lipogenesis was recently demonstrated, further strengthening the link between ER stress and regulation of hepatic lipid metabolism (Figure 18.5). Lee et al. [117] generated an inducible deletion of XBP1 in mouse liver and discovered that hepatic fatty acid and cholesterol synthesis was dramatically reduced to 85–90% of wild-type levels, likely explaining significant hypolipidemia in XBP1 liver-deficient mice. However, XBP1-deficient mice did

ATF6 SREBP

lum

en

SCAP

Regulate gene expression for lipid synthesis

XBP-1

nucleus

S2P

i olg

n me

SCAP

lu

G

Figure 18.5 Convergence of regulators of lipid synthesis and the unfolded protein response. The membrane-tethered transcription factors SREBP and ATF6 share a common processing pathway involving site 2 protease (S2P) in the Golgi, where the transcription factor domain is released for import into the nucleus. XBP1 requires processing in order to enter the nucleus and activate gene expression. Both SREBPs and XBP1 control the expression of genes involved in lipogenesis. However, unlike SREBPs, the activation of ATF6 and XBP1 in response to stress is independent of cellular cholesterol. Arrows eminating from S2P indicate proteolytic processing events by S2P. Grey arrows indicate the trafficking of transcription factors to the nucleus

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not have decreased apoB secretion, indicating that the hypolipidemia was a consequence of decreased hepatic lipogenesis. Decreased lipogenesis in XBP1 KO livers was also surprisingly not due to decreased expression of SREBP1c, SREBP2, or ChREBP, but rather a peculiar decrease in a subset of lipogenic genes, namely ACC2, DGAT2, and SCD-1. Changes in other lipogenic genes such as ACC1, FAS, and HMG-CoA synthase and reductase were not decreased in XBP1 KO livers, despite the fact that hepatic long-chain fatty acid synthesis and cholesterol were decreased. Decreases in hepatic lipogenesis in XBP1-deficient livers were independent of ER stress. XBP1 expression is upregulated in mouse liver after fructose feeding and in isolated hepatocytes by glucose, raising the possibility that ChREBP regulates XBP1. The mechanism by which XBP1 connects ER stress to the control of hepatic lipogenesis and the relative contributions of SREBPs and XBP1 to this task in physiological and pathophysiological states remain to be determined.

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88. Evans, R.M., Barish, G.D. and Wang, Y.X. (2004) PPARs and the complex journey to obesity. Nat Med , 10, 355–61. 89. Duan, S.Z., Ivashchenko, C.Y., Whitesall, S.E., D’Alecy, L.G., Duquaine, D.C., Brosius, F.C. III, Gonzalez, F.J., Vinson, C., Pierre, M.A., Milstone, D.S. et al. (2007) Hypotension, lipodystrophy, and insulin resistance in generalized PPARgamma-deficient mice rescued from embryonic lethality. J Clin Invest , 117, 812–22. 90. He, W., Barak, Y., Hevener, A., Olson, P., Liao, D., Le, J., Nelson, M., Ong, E., Olefsky, J.M. and Evans, R.M. (2003) Adipose-specific peroxisome proliferator-activated receptor gamma knockout causes insulin resistance in fat and liver but not in muscle. Proc Natl Acad Sci U S A, 100, 15712–17. 91. Imai, T., Takakuwa, R., Marchand, S., Dentz, E., Bornert, J.M., Messaddeq, N., Wendling, O., Mark, M., Desvergne, B., Wahli, W. et al. (2004) Peroxisome proliferator-activated receptor gamma is required in mature white and brown adipocytes for their survival in the mouse. Proc Natl Acad Sci U S A, 101, 4543–47. 92. Zhang, J., Fu, M., Cui, T., Xiong, C., Xu, K., Zhong, W., Xiao, Y., Floyd, D., Liang, J., Li, E. et al. (2004) Selective disruption of PPARgamma 2 impairs the development of adipose tissue and insulin sensitivity. Proc Natl Acad Sci U S A, 101, 10703–8. 93. Guilherme, A., Virbasius, J.V., Puri, V. and Czech, M.P. (2008) Adipocyte dysfunctions linking obesity to insulin resistance and type 2 diabetes Nat Rev Mol Cell Biol May;9(5): 367–77. 94. Matsusue, K., Haluzik, M., Lambert, G., Yim, S.H., Gavrilova, O., Ward, J.M., Brewer, B., Reitman, M.L. and Gonzalez, F.J. Jr. (2003) Liver-specific disruption of PPARgamma in leptin-deficient mice improves fatty liver but aggravates diabetic phenotypes. J Clin Invest , 111, 737–47. 95. Kubota, N., Terauchi, Y., Miki, H., Tamemoto, H., Yamauchi, T., Komeda, K., Satoh, S., Nakano, R., Ishii, C., Sugiyama, T. et al. (1999) PPAR gamma mediates high-fat diet-induced adipocyte hypertrophy and insulin resistance. Mol Cell , 4, 597–609. 96. Miles, P.D., Barak, Y., He, W., Evans, R.M. and Olefsky, J.M. (2000) Improved insulin-sensitivity in mice heterozygous for PPAR-gamma deficiency. J Clin Invest , 105, 287–92. 97. Lee, K., Villena, J.A., Moon, Y.S., Kim, K.H., Lee, S., Kang, C. and Sul, H.S. (2003) Inhibition of adipogenesis and development of glucose intolerance by soluble preadipocyte factor-1 (Pref-1). J Clin Invest , 111, 453–61. 98. Kim, J.Y., van de Wall, E., Laplante, M., Azzara, A., Trujillo, M.E., Hofmann, S.M., Schraw, T., Durand, J.L., Li, H., Li, G. et al. (2007) Obesity-associated improvements in metabolic profile through expansion of adipose tissue. J Clin Invest , 117, 2621–37. 99. Kim, K.H. (1997) Regulation of mammalian acetylcoenzyme A carboxylase. Annu Rev Nutr, 17, 77–99. 100. Abu-Elheiga, L., Matzuk, M.M., Abo-Hashema, K.A. and Wakil, S.J. (2001) Continuous fatty acid oxidation

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109. Wende, A.R., Huss, J.M., Schaeffer, P.J., Giguere, V. and Kelly, D.P. (2005) PGC-1alpha coactivates PDK4 gene expression via the orphan nuclear receptor ERRalpha: a mechanism for transcriptional control of muscle glucose metabolism. Mol Cell Biol , 25, 10684–94. 110. Kersten, S., Seydoux, J., Peters, J.M., Gonzalez, F.J., Desvergne, B. and Wahli, W. (1999) Peroxisome proliferator-activated receptor alpha mediates the adaptive response to fasting. J Clin Invest , 103, 1489–98. 111. Ron, D. and Walter, P. (2007) Signal integration in the endoplasmic reticulum unfolded protein response. Nat Rev Mol Cell Biol , 8, 519–29. 112. Calfon, M., Zeng, H., Urano, F., Till, J.H., Hubbard, S.R., Harding, H.P., Clark, S.G. and Ron, D. (2002) IRE1 couples endoplasmic reticulum load to secretory capacity by processing the XBP-1 mRNA. Nature, 415, 92–6. 113. Lee, K., Tirasophon, W., Shen, X., Michalak, M., Prywes, R., Okada, T., Yoshida, H., Mori, K. and Kaufman, R.J. (2002) IRE1-mediated unconventional mRNA splicing and S2P-mediated ATF6 cleavage merge to regulate XBP1 in signaling the unfolded protein response. Genes Dev , 16, 452–66. 114. Ye, J., Rawson, R.B., Komuro, R., Chen, X., Dave, U.P., Prywes, R., Brown, M.S. and Goldstein, J.L. (2000) ER stress induces cleavage of membrane-bound ATF6 by the same proteases that process SREBPs. Mol Cell , 6, 1355–64. 115. Goldstein, J.L., Rawson, R.B. and Brown, M.S. (2002) Mutant mammalian cells as tools to delineate the sterol regulatory element-binding protein pathway for feedback regulation of lipid synthesis. Arch Biochem Biophys, 397, 139–48. 116. Ozcan, U., Cao, Q., Yilmaz, E., Lee, A.H., Iwakoshi, N.N., Ozdelen, E., Tuncman, G., Gorgun, C., Glimcher, L.H. and Hotamisligil, G.S. (2004) Endoplasmic reticulum stress links obesity, insulin action, and type 2 diabetes. Science, 306, 457–61. 117. Lee, A.H., Scapa, E.F., Cohen, D.E. and Glimcher, L.H. (2008) Regulation of hepatic lipogenesis by the transcription factor XBP1. Science, 320, 1492–96.

19

Lipoprotein Metabolism and Cholesterol Balance David E. Cohen Department of Medicine, Gastroenterology Division, Brigham and Women’s Hospital, Harvard Medical School and Harvard-Massachusetts Institute of Technology Division of Health Sciences and Technology, Boston, MA, USA

INTRODUCTION Lipids are insoluble or sparingly soluble molecules that are essential for membrane biogenesis and maintenance of membrane integrity. They also serve as energy sources, hormone precursors, and signaling molecules. In order to facilitate transport through the relatively aqueous blood, non-polar lipids, such as cholesteryl esters or triglycerides, are packaged within lipoproteins. Increased concentrations of certain lipoproteins in the circulation are associated strongly with atherosclerosis. Much of the prevalence of cardiovascular disease, the leading cause of death in the USA and most Western countries, can be attributed to elevated concentrations of cholesterol-rich low-density lipoprotein (LDL) particles, as well as lipoproteins that are rich in triglycerides. Epidemiologically, decreased concentrations of high-density lipoproteins (HDL) also predispose to atherosclerotic disease. This chapter highlights the biochemistry and physiology of cholesterol and lipoproteins. Because abundant clinical outcomes data have proven that morbidity and mortality from cardiovascular disease can be reduced by the use of lipid-lowering drugs, mechanisms of pharmacological intervention that can ameliorate hyperlipidemia will be discussed.

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

BIOCHEMISTRY AND PHYSIOLOGY OF CHOLESTEROL AND LIPOPROTEIN METABOLISM Lipoproteins are macromolecular aggregates that transport triglycerides and cholesterol through the blood. Circulating lipoproteins can be differentiated on the basis of density, size, and protein content (Table 19.1). As a general rule, larger, less dense lipoproteins have a greater percentage composition of lipids; chylomicrons are the largest and least dense lipoprotein subclass, whereas HDL are the smallest lipoproteins, containing the lowest lipid content and the highest proportion of protein. Structurally, lipoproteins are microscopic spherical particles ranging from 7 to 100 nm in diameter (Figure 19.1). Lipoprotein particles consist of a monolayer of polar, amphipathic lipids that surrounds a hydrophobic core [1]. Each lipoprotein particle also contains one or more types of apolipoprotein (Table 19.1). The polar lipids that compose the surface coat are unesterified cholesterol and phospholipid molecules, which are arranged in a monolayer [2]. The hydrophobic core of a lipoprotein contains the cholesteryl esters (cholesterol molecules

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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Table 19.1 Characteristics of plasma lipoproteinsa. Reproduced from Jonas & Phillips, Chapter 17, Biochemistry of Lipids, Lipoproteins & Membranes, 5e, Copyright (2008), with permission from Elsevier ml−1 )

Density (g Diameter (nm) Total lipid (% wt) Composition (% dry weight) Protein Triglycerides Unesterified cholesterolCholesteryl esters Phospholipids (% wt lipid) Electrophoretic mobilityb Major apolipoproteins

CM

VLDL

IDL

LDL

HDL

< 0.95 75–1200 98

0.95–1.006 30–80 90

1.006–1.019 25–35 82

1.019–1.063 1.063–1.210 18–25 5–12 75 67

2 83 8

10 50 22

18 31 29

25 9 45

33 8 30

7 None B48, A-I, A-IV, E, C-I, C-II, C-III

18 Preß B100, E, C-I, C-II, C-III

22 ß B100, E, C-I, C-II, C-III

21 ß B100

29 α or Preß A-I, A-II, C-I, C-II, C-III, E

CM, chylomicron; VLDL, very low-density lipoprotein; IDL, intermediate-density lipoprotein; LDL, low-density lipoprotein; HDL, high-density lipoprotein. a Adapted with permission from Jonas, A. (2002) Lipoprotein structure, in Biochemistry of Lipids, Lipoproteins and Membranes (eds D.E. Vance and J.E. Vance),

4th edn, Elsevier, Amsterdam, pp. 483–504. b Electrophoretic mobility of lipoprotein particles is designated according relative to migration of plasma α- and β-globulins.

Unesterified Cholesterol

Cholesteryl Ester Triglyceride Apolipoprotein Phospholipid

Figure 19.1 Structure of lipoprotein particles. Lipoproteins are spherical particles (7–100 nm in diameter) that transport hydrophobic molecules, principally cholesterol and triglycerides, as well as fat-soluble vitamins. Each type of lipoprotein particle has a surface that is composed of a monolayer of phospholipid and unesterified cholesterol molecules. These polar lipids form a coating that shields a hydrophobic core of non-polar triglyceride and cholesteryl esters from interacting with the aqueous environment of plasma. Lipoproteins contain amphipathic apolipoproteins (also called apoproteins), which associate with the surface lipids and hydrophobic core. Apolipoproteins provide structural stability to the lipoprotein particle and act as ligands for specific cell-surface receptors or as cofactors for enzymatic reactions. Unless otherwise indicated in subsequent figures, phospholipid and unesterified cholesterol molecules are not shown for clarity. Figure and legend modified from Scapa, Kanno and Cohen [84] with permission from Blackwell

linked by an ester bond to a fatty acid) and triglycerides (three fatty acids esterified to a glycerol molecule). The apolipoproteins (also referred to as apoproteins) are amphipathic proteins that intercalate into the lipid membrane of lipoproteins. In addition to stabilizing the structure of

lipoproteins, apolipoproteins engage in biological functions. They may act as receptor ligands for lipoprotein particles or may activate enzymatic activities in the plasma. As will be discussed, the apolipoprotein composition determines the metabolic fate of the lipoprotein. From a metabolic perspective, lipoprotein particles can be divided into lipoproteins that participate in the delivery of triglyceride molecules to muscle and fat tissue (the apolioprotein B (apoB)-containing lipoproteins, chylomicrons, and VLDL (very low-density lipoprotein)s) and lipoproteins that are involved primarily in cholesterol transport (HDL and the remnants of apoB-containing lipoproteins). HDL also serve as a reservoir for exchangeable apolipoproteins in the plasma, including apoA-I, apoC-II, and apoE. The following discussion presents each lipoprotein class in the context of its function.

Metabolism of apoB-containing lipoproteins The primary function of apoB-containing lipoproteins is to deliver fatty acids in the form of triglycerides to muscle tissue for ATP biogenesis and to adipose tissue for long-term storage. Chylomicrons are formed in the intestine and transport dietary triglycerides, whereas VLDL particles are formed by the liver and transport triglycerides that are synthesized endogenously. For conceptual purposes, the metabolic life span of apoB-containing lipoproteins can be divided into three phases: assembly, intravascular metabolism, and receptor-mediated clearance. This is also a convenient categorization because pharmacological agents are available that influence these different phases.

19: LIPOPROTEIN METABOLISM AND CHOLESTEROL BALANCE

Assembly of apoB-containing lipoproteins The cellular mechanisms by which chylomicrons and VLDL are assembled are quite similar. Regulation of the assembly process depends upon the availability of apoB and triglycerides, as well as the activity of microsomal triglyceride transfer protein (MTP) [3]. The gene that encodes apoB is transcribed principally in the intestine and the liver. Apart from this tissue-specific expression, there is little transcriptional regulation of the apoB gene. By contrast, a key regulatory event that differentiates chylomicron from VLDL metabolism is editing of apoB mRNA (Figure 19.2). Within enterocytes but not hepatocytes, a protein named apoB editing complex-1 (apobec-1) is expressed. This protein constitutes the catalytic subunit of the apoB editing complex, which deaminates a cytosine at position 6666 of the apoB mRNA molecule [4]. This converts the cytosine to a uridine base. As a result, the codon containing this nucleotide is converted from a glutamine to a premature stop. When translated, the intestinal form apoB48 is 48% as long as the full-length protein expressed in the liver, which is referred to as apoB100. As a consequence, chylomicrons, the apoB-containing lipoproteins produced by the intestine, contain apoB48. By contrast, VLDL particles produced by the liver contain apoB100. Figure 19.3a illustrates the cellular mechanisms for the assembly and secretion of apoB-containing lipoproteins. As the apoB protein is translated by ribosomes, it crosses into the endoplasmic reticulum (ER) [3, 5]. Within the ER, triglyceride molecules are added co-translationally to the elongating apoB protein (i.e. apoB is lipidated)

apoB Gene Transcription

Liver & Intestine

apoB mRNA Intestine

Editing

No Editing

Liver X

X

Translation apoB48

apoB100

Figure 19.2 Editing of ApoB mRNA. The apoB gene, with exons represented by squares and introns by lines, is transcribed in both the intestine and the liver. In the intestine, but not the liver, a protein complex containing apobec-1 modifies a single nucleotide in the apoB mRNA. As a result, the codon containing this nucleotide is converted to a premature stop codon, as indicated by the “X”. The protein that is translated in the intestine (apoB48) is only 48% as long as the full-length protein that is translated in the liver (apoB100). Figure and legend modified from Scapa, Kanno and Cohen [84] with permission from Blackwell

273

by the action of MTP [3]. Once apoB has been fully translated, the nascent lipoprotein is enlarged in the Golgi apparatus, during which MTP adds additional triglycerides to the core of the particle [5]. By unclear mechanisms, cholesteryl esters are also added to the core. This assembly process produces the lipoprotein particles, each of which contains a single molecule of apoB. The diet is the main source of triglycerides in chylomicrons (Figure 19.3b), so their assembly, secretion, and metabolism are collectively referred to as the exogenous pathway of lipoprotein metabolism. By contrast, cholesteryl esters in chylomicrons are derived mainly from biliary cholesterol (approximately 75%), with the remainder contributed by dietary sources. During digestion, cholesteryl esters and triglycerides in food are hydrolyzed to form unesterified cholesterol, free fatty acids, and monoglycerides [6]. Bile salts, phospholipids, and cholesterol are secreted by the liver into bile and are stored in the gall bladder during fasting as micelles and vesicles, which are macromolecular lipid aggregates that form due to the detergent properties of bile salt molecules. The stimulus of eating a meal promotes emptying of gall bladder bile into the small intestine, where micelles and vesicles solubilize the digested lipids. Lipid absorption into enterocytes of the duodenum and jejunum is facilitated mainly by micelles [7]. Long-chain fatty acids and monoglycerides are taken up separately into the enterocyte by carrier-mediated transport and then re-esterified to form triglycerides by the enzyme diacylglycerol acyltransferase (DGAT) [8]. By contrast, medium-chain fatty acids are absorbed directly into the portal blood and metabolized by the liver. Dietary and biliary cholesterol from micelles enter the enterocyte via a protein channel named Neimann-Pick C 1-like protein (NPC1L1) [9, 10]. Some of this cholesterol is immediately pumped back into the intestinal lumen by the ATP-dependent action of a heterodimeric protein, ATP binding cassette (ABC) G5/ABCG8 [9, 11]. The fraction of cholesterol that remains is esterified to a long-chain fatty acid by acyl-CoA : cholesterol acyltransferase (ACAT) [12]. Once triglycerides and cholesteryl esters are packaged together with apoB48, the chylomicron particles are exocytosed into the lymphatics for transport into the circulation via the thoracic duct (Figure 19.3c). The plasma concentration of triglyceride-rich chylomicrons varies in proportion to dietary fat intake. VLDL comprise triglycerides that are assembled by the liver using plasma fatty acids derived from adipose tissue synthesized de novo. For this reason, the assembly, secretion, and metabolism of VLDL are often referred to as the endogenous pathway of lipoprotein metabolism. Hepatocytes synthesize triglycerides in response to increased free fatty acid flux to the liver from adipose tissue. This typically occurs in response to fasting, thereby insuring a continuous supply of fatty acids for delivery to muscle in the absence of triglycerides from the diet. Interestingly, dietary saturated fats [13] as well as carbohydrates [14, 15]

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Enterocyte or Hepatocyte

Cytosol

Presence of Triglycerides Lipidation

Ribosome

CM MTP

MTP

VLDL

ApoB Absence of Triglycerides Degradation ApoB

Endoplasmic Reticulum

(a) Triglyceride Absorption

LYMPH

Cholesterol Absorption ENTEROCYTE

LYMPH

INTESTINAL LUMEN

INTESTINAL LUMEN 2 Fatty Acid + Monoglyceride

Cholesterol ACAT

ENTEROCYTE

NPC1L1

DGAT Cholesteryl Ester

ABCG5/G8

Exogenous Triglyceride

Bile salt

(b)

LYMPH CM apoB48

Chylomicron Formation ENTEROCYTE

INTESTINAL LUMEN

Exogenous Triglycerides

Cholesteryl Ester

(c)

Figure 19.3 Assembly and secretion of apoB-containing lipoproteins. (a) Chylomicron (CM) and VLDL particles are assembled and secreted by similar mechanisms in the enterocyte and hepatocyte, respectively. The apoB protein (i.e. apoB48 or apoB100) is translated by ribosomes and enters the lumen of the ER. If triglycerides are available, the apoB protein is lipidated by the action of microsomal triglyceride transfer protein (MTP) in two distinct steps, accumulating triglyceride as well as cholesterol ester molecules. The resulting CM or VLDL particle is secreted by exocytosis into the lymphatics by enterocytes or into the plasma by hepatocytes. In the absence of triglycerides, the apoB protein is degraded. (b) Triglycerides and cholesterol are simultaneously absorbed from the intestinal lumen by different mechanisms. Cholesterol is taken up from micelles across a regulatory channel named NPC1LI. A fraction of the cholesterol is pumped back into the lumen by ABCG5/G8, a heterodimeric ATP-dependent plasma membrane protein. The remainder of the cholesterol is converted to cholesteryl esters by ACAT. Triglycerides are taken up as fatty acids and monoglycerides, which are re-esterified by DGAT. (c) Exogenous triglycerides plus cholesteryl esters are assembled into CM within the enterocyte. Panel (a) and legend modified from Scapa, Kanno and Cohen [84] with permission from Blackwell. Panel (b) and legend modified from Cohen, D.E. and Armstrong, E.J. [85] with permission of Lippincott, Williams and Wilkins

19: LIPOPROTEIN METABOLISM AND CHOLESTEROL BALANCE

also stimulate the synthesis of triglycerides within the liver. By cellular mechanisms that are highly similar to those that produce chylomicrons, MTP in hepatocytes lipidates apoB100 to form nascent VLDL particles. Under the continued influence of MTP, the nascent VLDL particles coalesce with larger triglyceride droplets and are secreted directly into the circulation. VLDL particles may also acquire apoE, apoC-I, apoC-II, and apoC-III within the hepatocyte prior to secretion. However, these apolipoproteins may also be transferred to VLDL from HDL during their circulation through the bloodstream. The translation of apoB48 in the intestine and apoB100 in the liver is constitutive. This permits the immediate production of chylomicrons and VLDL particles when triglyceride molecules are available. In the absence of triglycerides, such as in enterocytes during fasting, apoB is degraded by a variety of cellular mechanisms [16, 17].

Intravascular metabolism of apoB-containing lipoproteins Within the circulation, chylomicrons and VLDL particles must be activated in order to target triglyceride delivery to muscle and fat tissues (Figure 19.4a). This requires the addition of an optimal compliment of apoC-II molecules, which occurs at least in part by aqueous transfer of apoC-II from HDL particles [18, 19]. Because there is an inherent delay in the transfer of apoC-II to chylomicrons and VLDL particles, this allows time for widespread circulation of triglyceride-rich particles throughout the body [19]. Lipoprotein lipase (LPL) is a lipolytic enzyme that is expressed on the vascular surface of the endothelium of capillaries in muscle and fat tissues [20]. It is a glycoprotein that is anchored in place by electrostatic interactions with a separate glycoprotein on the endothe-

VLDL apoB100

LIVER

275

CM apoB48

INTESTINE

apoC-II

apoE α-HDL

apoAI (a)

Capillary in Muscle or Adipose Tissue CM apoB48

VLDL apoB100

Fatty Acids

Fatty Acids

Lipoprotein Lipase

Endothelium

(b)

Figure 19.4 Intravascular metabolism of apoB-containing lipoproteins. (a) Following secretion, chylomicron (CM) and VLDL particles are activated for lipolysis when they encounter HDL particles in the plasma and acquire the exchangeable apolipoprotein apoC-II. (b) When CM and VLDL circulate into capillaries of muscle or fat tissue, apoC-II promotes binding of the particle to LPL, which is bound to the surface of endothelial cells. LPL mediates hydrolysis of triglycerides, but not cholesteryl esters, from the core of the lipoprotein particle. The resulting fatty acids are taken up into muscle or fat tissue. Figure and legend modified from Scapa, Kanno and Cohen [84] with permission from Blackwell

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THE LIVER: BIOCHEMISTRY AND PHYSIOLOGY OF CHOLESTEROL AND LIPOPROTEIN METABOLISM

lial cell membrane. Once chylomicrons and VLDL particles acquire apoC-II, they can bind to LPL, which hydrolyzes triglycerides from the core of the lipoprotein (Figure 19.4b). LPL-mediated lipolysis liberates free fatty acids and glycerol, which are then taken up by the neighboring parenchymal cells. Because the expression level and intrinsic activity of LPL in muscle versus fat tissue is regulated according to the fed/fasting state, this allows the body to direct the delivery of fatty acids preferentially to muscle during fasting and to fat post-prandially [21]. The rate of lipolysis of chylomicrons and VLDL triglycerides is also controlled by apoC-III, which is an inhibitor of LPL activity [18]. Antagonism of LPL activity by apoC-III may be an additional mechanism that promotes widespread distribution of triglyceride-rich particles in the circulation.

VLDL Remnant apoB100

Receptor-mediated clearance of apoB-containing lipoproteins As LPL continues to hydrolyze triglycerides from chylomicrons and VLDL, the particles become progressively depleted of triglycerides and relatively enriched with cholesterol. Once approximately 50% of triglycerides have been removed, the particles lose affinity for LPL and dissociate [19]. The exchangeable apolipoproteins apoA-I and apoC-II (as well as apoC-I and apoC-III) are then transferred to HDL in exchange for apoE (Figure 19.5a) [22], which serves as a high-affinity ligand for receptor-mediated clearance [23, 24]. Upon acquiring apoE, the particles are termed chylomicron or VLDL remnants [24]. As explained below, intermediate-density lipoproteins (IDL) are also remnant particles.

CM Remnant apoB48

apoCII apoE α-HDL

apoAI (a) CM or VLDL Remnant

Sinusoidal Endothelium HSPG

Space of Disse

Sequestration Hepatic Lipase Lipolysis

Uptake

LDLr

LRP

HSPG

Hepatocyte (b)

Figure 19.5 Formation and hepatic uptake of remnant particles. (a) Upon completion of hydrolysis, chylomicrons (CM) and VLDL lose affinity for lipoprotein lipase. When an HDL particle is encountered, apoC-II is transferred back to HDL particles in exchange for apoE. The resulting particles are CM and VLDL remnants and IDL, which are VLDL particles that interact for prolonged periods with LPL to become more dense. (b) The activity of LPL results in remnant lipoprotein particles that are small enough in size to enter the space of Disse. Remnant lipoproteins are sequestered in the space of Disse by binding to high-molecular-weight heparan sulfate proteoglycan (HSPG) molecules. This is followed by binding of hepatic lipase, which promotes lipolysis of some residual triglycerides in the core of the remnant lipoproteins and the release of fatty acids (as indicated by the dashed arrow). Uptake of remnant lipoprotein particles into hepatocytes is mediated by the LDL receptor (LDLr), the LDL-related protein (LRP), a complex formed between LRP and HSPG, or HSPG alone. Figure and legend modified from Scapa, Kanno and Cohen [84] with permission from Blackwell

19: LIPOPROTEIN METABOLISM AND CHOLESTEROL BALANCE

Remnants of chylomicrons and VLDL, as well as some IDL particles, are taken up by the liver in a three-step process (Figure 19.5b) [24, 25]. The first step is sequestration of the particles within the space of Disse between the fenestrated endothelium of the liver sinusoids and the sinusoidal (basolateral) plasma membrane of hepatocytes. Sequestration requires that the remnant particles become small enough during the process of lipolysis to fit in between the endothelial cells. Once in the space of Disse, remnants are bound and sequestered by large heparan sulfate proteoglycans (HSPGs). The next step is remodeling within the space of Disse by the action of hepatic lipase, a lipolytic enzyme that is similar to LDL but is expressed by hepatocytes. Hepatic lipase appears to optimize the triglyceride content of remnant particles, so they are efficiently cleared by receptor-mediated mechanisms. The final phase of remnant clearance is receptor-mediated uptake. This is accomplished by one of four pathways. At the sinusoidal hepatocyte plasma membrane, remnant particles may be bound and taken up by the LDL receptor (LDLr), the LDL receptor-related protein (LRP), or by HSPGs. A separate pathway is mediated by the combined activities of the LRP and HSPGs. These efficient and redundant mechanisms allow for efficient particle clearance, so that the half-life of remnants in the plasma is approximately 30 minutes.

Formation and clearance of LDL particles Whereas apoB48-containing chylomicron remnants are completely cleared from the plasma, the presence of apoB100 alters the metabolism of VLDL remnants so that only approximately 50% are cleared by pathways for remnant particles. The difference begins when the other 50% are metabolized to a greater extent by LPL, becoming an increment smaller and relatively deficient in triglycerides and enriched in cholesteryl esters. When converted to remnants following exchange of apolipoproteins with HDL, these more dense particles are called IDL. Because IDL contain apoE, a fraction of these particles may be cleared into the liver by remnant receptor pathways (Figure 19.5) [26]. However, the remainder are converted to LDL by hepatic lipase, which further hydrolyzes triglycerides in the cores of IDL (Figure 19.6). The further reduction size of the particle results in the transfer of apoE to HDL. As a result, LDL are distinct, cholesteryl ester-enriched lipoproteins with apoB100 as their only apolipoprotein [19]. The LDLr is the only receptor capable of clearing significant amounts of LDL from the plasma [27]. The LDLr is expressed on the surface of hepatocytes, macrophages, lymphocytes, adrenocortical cells, gonadal cells, and smooth muscle cells. Due to the lack of apoE, the LDL particles are relatively weak ligands for the LDLr [27]. As a result, the half-life of LDL in the circulation is markedly prolonged (two to four days). This explains

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LIVER IDL apoB100

LDL apoB100

Hepatic lipase

HDL

apoAI

Figure 19.6 Formation of LDL particles. Formation of LDL occurs when IDL particles interact with hepatic lipase to become more dense and cholesteryl ester-enriched. As a result, both apoE and apoC-II lose affinity for the particle and are transferred to HDL, leaving only apoB100. Figure and legend modified from Cohen, D.E. and Armstrong, E.J. [85] with permission of Lippincott, Williams and Wilkins

in part why LDL cholesterol accounts for approximately 65–75% of total plasma cholesterol. Interaction of apoB100 with the LDLr facilitates receptor-mediated endocytosis of LDL particles and subsequent vesicle fusion with a lysosome [27]. The LDLr is recycled to the cell surface, while the LDL particle is hydrolyzed to release unesterified cholesterol, which impacts three major homeostatic pathways. First, intracellular cholesterol inhibits HMG CoA reductase, the enzyme that catalyzes the rate-limiting step in de novo cholesterol synthesis. Second, cholesterol activates ACAT to increase esterification and storage of cholesterol in the cell. Third, LDLr expression is downregulated, reducing further uptake of cholesterol into the cells. The majority of LDLr (approximately 70%) are expressed on the surface of hepatocytes. As a result, the liver is primarily responsible for the removal of LDL particles from the circulation. Reductions in intracellular cholesterol levels, such as occur during therapy with statin drugs, lead to LDLr upregulation. This is due to the proteolytic processing and activity of sterol response element binding protein 2 (SREBP-2) [28].

HDL metabolism and reverse cholesterol transport Virtually all cells in the body are capable of synthesizing all of the cholesterol they require. However, only the liver has the capacity to eliminate cholesterol by secretion into bile in its unesterified form or following its conversion to bile salts. In addition to serving as a reservoir for exchangeable apolipoproteins for the metabolism

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of apoB-containing lipoproteins, HDL form a class of lipoproteins that plays a key role in cholesterol homeostasis by removing excess cholesterol from cells and transporting it through plasma to the liver. This process is often referred to as reverse cholesterol transport (Figure 19.7a) [29, 30]. The major apolipoproteins of HDL are apoA-I and apoA-II. ApoA-I is the main structural determinant of HDL, and participates in the formation, as well as interaction of the particle with its receptor, scavenger receptor class B, type I (SR-BI) [31]. The function of apoA-II is not well understood [32].

HDL formation HDL formation occurs mainly in the liver [33], although a fraction percentage is contributed by the small intestine. The earliest events occur when lipid-poor apoA-I is secreted by the liver or intestine [34], or dissociates from lipoprotein particles in the plasma [35, 36]. The amphipathic apoA-I molecules interact with ABCA1 [37], which is imbedded in the sinusoidal membrane of hepatocytes or the basolateral membrane of enterocytes [31]. ABCA1 incorporates a small amount of membrane phospholipid and unesterified cholesterol into the apoA-I molecule [31]. The resulting small disk-shaped HDL particle consists mainly of phospholipid and apoA-I. The particle that is formed is referred to as nascent or preß-HDL, due to its characteristic migration on agarose gels [36].

Intravascular maturation of HDL Because disk-shaped preß-HDL are relatively inefficient at removing excess cholesterol from cell membranes, these particles must mature into spherical particles within the plasma. This occurs due to the activity of two distinct circulating proteins (Figure 19.7b). Lecithin : cholesterol acyltransferase (LCAT) binds preferentially to disk-shaped HDL and converts cholesterol molecules within the particle to cholesteryl esters [31]. This is accomplished by transesterification of a fatty acid from a phosphatidylcholine molecule on the surface of the HDL to the hydroxyl group of a cholesterol molecule. The reaction also creates a lysophosphatidylcholine molecule, which departs from the particle upon binding to serum albumin [38]. Because they are so highly insoluble, cholesteryl esters spontaneously migrate into the core of the HDL particle [19]. The development of a hydrophobic core converts the preß-HDL to a spherical HDL particle, which exhibits α migration on agarose gels. The second important protein that contributes to HDL maturation in the plasma is phospholipid transfer protein (PLTP) [39]. PLTP transfers phospholipids from the surface coat of apoB-containing remnant particles to the

surface coat of an HDL. During LPL-mediated lipolysis of apoB-containing lipoproteins, the particles become smaller as triglycerides are removed from the core. This leaves a relative excess of phospholipids on the surface of the particle. Because phospholipids are highly insoluble and cannot otherwise dissociate from a particle, PLTP removes excess phospholipids and thereby maintains the appropriate surface concentration for the shrinking core. By transferring phospholipids to the surface of an HDL, PLTP also replaces the molecules that are consumed by the LCAT reaction. This allows the core of the HDL to continue to enlarge. Based on studies on mice lacking PLTP, it has been estimated that as much as 80% of HDL phospholipids are transferred from the surface coat of remnant particles [40].

HDL-mediated cholesterol efflux from cells Cellular cholesterol efflux is the mechanism by which excess insoluble cholesterol molecules are removed from cells. This occurs when unesterified cholesterol is transferred from the plasma membrane of cells to an HDL particle. The mechanism of cholesterol efflux varies depending upon the cell type and the type of HDL particle [37, 41, 42]. Lipid-poor preß-HDL particles can promote cholesterol efflux by interacting with ABCA1. In addition to beginning the process of HDL formation by the liver, this is also a mechanism for removing excess cholesterol from macrophages within the subendothelial space, thereby protecting them from cholesterol-induced cytotoxicity. Spherical HDL very efficiently stimulate cholesterol efflux, which occurs by several mechanisms. The particles may interact with SR-BI on the plasma membrane, which can promote cholesterol efflux. In addition to ABCA1 and SR-BI, macrophages express ABCG1, which also mediates efflux of cholesterol to spherical HDL. Finally, spherical HDL particles may promote cholesterol efflux in the absence of binding to a specific cell-surface protein. Although cholesterol has very low monomeric solubility [43], it can dissociate in appreciable amounts and travel short distances through the plasma to acceptor particles that are enriched with phospholipids on their surfaces. Quantitatively, efflux to spherical HDL particles accounts for most of the removal of excess cholesterol from cells. This capacity of HDL to remove cellular cholesterol is enhanced through the activities of LCAT and PLTP, which prevent the surface coat of the particle from becoming saturated with cholesterol from cells.

Delivery of HDL cholesterol to the liver When mature HDL particles circulate to the liver, they interact with SR-BI, the principal HDL receptor [44],

19: LIPOPROTEIN METABOLISM AND CHOLESTEROL BALANCE

LIVER

BLOOD

TISSUES LCAT

Cholesterol Phospholipid ABCA1

279

α-HDL

preβ-HDL apoA-I

Cholesterol

SR-BI LCAT PLTP Hepatic CETP Lipase (a) CM or VLDL Remnant

Albumin

PLTP

Lyso-PC LCAT

Cell CETP

α-HDL

(b)

Figure 19.7 Reverse cholesterol transport. (a) The process of reverse cholesterol transport begins when apoA-I is secreted from the liver. ApoA-I in plasma interacts with ATP binding cassette protein A1 (ABCA1), which incorporates a small amount of phospholipid and unesterified cholesterol from hepatocyte plasma membranes to form a discoidal-shaped preß-HDL particle. Due to the activity of LCAT in plasma, preß-HDL particles mature to form spherical α-HDL. Spherical α-migrating HDL particles function to accept excess unesterified cholesterol from the plasma membranes of cells in a wide variety of tissues. The unesterified cholesterol is transferred from the cell to nearby HDL particles by diffusion through the plasma. As explained in (b), LCAT and PLTP increase the capacity of HDL to accept unesterified cholesterol molecules from cells by allowing for expansion of the core and the surface coat of the particle. CETP exchanges some of the cholesteryl ester molecules from the cores of HDL for triglycerides from the cores of remnant particles. HDL particles interact with scavenger receptor, class B type I (SR-BI), which mediates selective hepatic uptake of cholesteryl esters, but not apoA-I. This process is facilitated when hepatic lipase hydrolyzes triglycerides from the core of the particle. The remaining apoA-I molecules may begin the cycle of reverse cholesterol transport again. In (a), solid arrows indicate metabolic events in HDL metabolism, whereas dashed arrows denote transfer of molecules. (b) LCAT, PLTP, and CETP promote the removal of excess cholesterol from the plasma membranes of cells. LCAT removes a fatty acid from a phosphatidylcholine molecule in the surface coat of α- (or preß-) HDL and esterifies an unesterified cholesterol molecule on the surface of the particle. The resulting lysophosphatidylcholine (lyso-PC) becomes bound to albumin in the plasma, whereas the cholesteryl ester migrates spontaneously into the core of the lipoprotein particle. The unesterified cholesterol molecules that are consumed by LCAT are replaced by unesterified cholesterol from cells. HDL phospholipids that are consumed by LCAT action are replaced with excess phospholipids from remnant particles by the activity of PLTP. As described in (a), CETP increases the efficiency of cholesterol movement to the liver by exchanging cholesteryl ester molecules from the core of α-HDL for triglycerides from the core of remnant particles. In (b), solid arrows denote protein-mediated lipid transfer, whereas dashed arrows indicate that lipids move by diffusion through the plasma. Figure and legend modified from Scapa, Kanno and Cohen [84] with permission from Blackwell

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which is expressed on the sinusoidal plasma membranes of hepatocytes [45]. Although it can mediate cholesterol efflux from cells with excess cholesterol, SR-BI in the liver promotes selective uptake of lipids, a process whereby the cholesterol and cholesterol esters of HDL particles are taken up into the cell, in the absence of uptake of apolipoproteins [31]. During SR-BI-mediated selective lipid uptake, apoA-I is liberated to participate in preß-HDL formation. As a result, the life span of an HDL particle is two to five days, suggesting that each apoA-I molecule can participate in many cycles of reverse cholesterol transport. Among the tissues that express high levels of SR-BI are the adrenal glands and gonads, presumably reflecting their requirement for cholesterol in order to support steroidogenesis [31]. Delivery of cholesterol from extrahepatic tissues to the liver is optimized by two additional proteins, cholesterol ester transfer protein (CETP) and hepatic lipase. CETP is a plasma protein that transfers cholesteryl esters from mature spherical HDL to the cores of remnant lipoproteins in exchange for triglyceride molecules, which are inserted into the cores of the HDL particles (Figure 19.7b) [46]. This process allows the body to utilize remnant particles that have completed their function of triglyceride transport for purposes of transporting cholesterol to the liver. Removal of cholesteryl ester molecules from HDL appears to serve two functions. First, it further increases the capacity of HDL to take on additional cholesterol molecules from cells. Second, it makes the process of selective uptake by SR-BI more efficient [46, 47]. This is because hydrolysis of triglycerides by hepatic lipase on the hepatocyte surface facilitates the activity of SR-BI (Figure 19.7a).

Biliary lipid secretion Once cholesterol is delivered to the liver by the process of reverse cholesterol transport, it is eliminated by biliary secretion. A key essential step occurs when a fraction of the cholesterol is converted to bile salts. Cholesterol 7α-hydroxylase (CYP7A1) is an enzyme that is expressed only in hepatocytes and catalyzes the rate-limiting step in the catabolism of cholesterol to bile salts [48]. Bile salts, unlike cholesterol, are highly soluble in water. Moreover, bile salts are biological detergents, which promote the formation of micelles. These macromolecular aggregates are rich in phospholipids, which are derived from hepatocyte membranes and solubilize cholesterol in bile for transport from the liver to the small intestine [49], essentially functioning as counterparts to HDL particles in plasma. Bile formation begins when ABCB11, an ATP-driven canalicular membrane transporter [50], functions to pump bile salts into bile [51]. Bile salts within bile then activate two other ATP-dependent transporters on the canalicular membrane of the hepatocyte [50]: ABCB4 [52] and a

heterodimer of ABCG5 and ABCG8 [53]. These promote the secretion of phospholipid and cholesterol molecules, which together with bile salts form mixed micelles. Biliary lipids are stored in the gall bladder during fasting. The stimulus of a fatty meal leads to gall-bladder contraction, which propels its contents into the small intestine. As described above, bile facilitates the digestion and absorption of fats, in addition to promoting the elimination of endogenous cholesterol.

Cholesterol balance Because cholesterol is converted by the liver to bile salts and is secreted unmodified into bile, overall cholesterol balance depends upon the disposition of both types of molecules. Rather than being lost in the feces after participating in cholesterol transport and fat digestion, most bile salt molecules are recycled when they are taken up by high-affinity transport proteins in the distal ileum. Bile salts enter the portal circulation and are transported back to the liver, where they are cleared by hepatocytes from the blood with high first-pass efficiency. Bile salts are then re-secreted into bile. This process of recycling bile salts between the liver and the intestine is referred to as the enterohepatic circulation [54]. The enterohepatic circulation is highly efficient, allowing < 5% of secreted bile salts to be lost in the feces. However, because bile salts are secreted in such large amounts, the small fractional loss of bile salts amounts to about 0.4 g per day [54]. Considering that cholesterol is the substrate for bile salt synthesis, fecal bile salts represent a source of cholesterol loss from the body. Sensitive nuclear hormone receptors within the liver are capable of detecting the rate of loss of bile salts into the feces [48, 55]. These receptors tightly regulate transcription of bile salt synthetic genes. As a result, the liver synthesizes precisely an amount of bile salts that is sufficient to replace what is lost in the feces. Whereas cholesterol is enzymatically converted into bile salt molecules mainly by CYP7A1 [48], an alternative pathway is initiated by sterol 27-hydroxylase (CYP27A). This contributes only a minor proportion of bile salts under normal circumstances [56]. Unlike cholesterol, amphiphilic bile salts, together with phospholipid molecules from the liver, forms mixed micelles that transport cholesterol within the biliary tree and small intestine [57]. On a daily basis, an average of 24 g of bile salts are secreted [54], together with 11 g of phospholipids [58] and approximately 1.2 g of cholesterol [59–62]. In addition to cholesterol that is secreted into bile each day, the average American diet contributes approximately 0.4 g per day to intestinal cholesterol. Therefore, dietary cholesterol represents only a minor fraction (25%) compared to the endogenous (i.e. biliary) cholesterol that passes through the intestine [63]. The extent to which intestinal

19: LIPOPROTEIN METABOLISM AND CHOLESTEROL BALANCE

cholesterol is absorbed appears to be genetically regulated. Each individual absorbs a fixed percentage of intestinal cholesterol. In the population, percentages range from as low as 20 to more than 80 [64, 65]. For example, when an average individual absorbs 50%, this will amount to half of the 1.6 g (i.e. 1.2 g of biliary cholesterol plus 0.4 g of dietary cholesterol). The other half (0.8 g) is lost in the feces. This, combined with a loss of 0.4 g per day of cholesterol in the form of fecal bile salts, yields a total cholesterol loss from the body of 1.2 g per day. Taking into account intestinal absorption of dietary cholesterol and reabsorption of biliary cholesterol, total body cholesterol synthesis is 0.8 g (i.e. cholesterol synthesis = fecal loss of cholesterol + bile salts − dietary cholesterol intake) [63]. This amounts to double what is consumed in the average diet.

PHARMACOLOGICAL CLASSES AND AGENTS The decision to treat dyslipidemia is largely dependent upon the calculated cardiovascular risk. A number of clinical algorithms exist for determining initiation of therapy. Goals for lipid lowering were established in the 2001 National Cholesterol Education Program Adult Treatment Panel III (ATP III) guidelines [66], which were updated in 2004 based upon the results of several additional large, randomized clinical trials [67]. These guidelines provide target LDL levels based on 10-year risk of death from cardiovascular disease. The guidelines stress that it is always important to first promote therapeutic lifestyle changes, including reduction of dietary saturated fat and cholesterol intake, weight reduction, increased physical activity, and possibly stress reduction. Successful dietary therapy can reduce total cholesterol by 5–25%, depending upon adherence and the metabolic basis for elevated cholesterol concentrations. If this approach is unsuccessful or insufficient to normalize lipid levels, drug therapy is generally recommended. Five classes of agents are available for pharmacological modification of lipid metabolism. Three of these classes (i.e. inhibitors of cholesterol synthesis, bile salt sequestrants, and cholesterol absorption inhibitors) have relatively well-defined effects on lipid metabolism. Whereas the overall effects of the other two classes (i.e. fibrates and niacin) are clear, their molecular mechanisms of action are diverse and still the subject of active investigation. The following is a condensed summary of the mechanisms of these agents.

Inhibitors of cholesterol synthesis HMG CoA reductase inhibitors, commonly known as statins, competitively inhibit the activity of HMG CoA reductase, the rate-limiting enzyme in cholesterol synthesis

281

[27]. Inhibition of this enzyme results in a transient, modest decrease in cellular cholesterol concentrations. This activates a cellular signaling cascade culminating in the activation of SREBP2, which is a transcription factor that upregulates expression of the gene encoding the LDLr. Increased LDLr expression causes increased uptake of plasma LDL and, consequently, decreases plasma LDL cholesterol concentrations [28]. Approximately 70% of LDLr are expressed by hepatocytes, with the remainder expressed by a variety of cell types in the body.

Inhibitors of bile salt absorption The bile salt sequestrants are cationic polymers that bind non-covalently to negatively charged bile salt molecules in the small intestine. The resin–bile salt complex cannot be re-absorbed in the distal ileum and is excreted into the stool. Decreased bile salt reabsorption by the ileum partially interrupts enterohepatic bile salt circulation, causing hepatocytes to upregulate 7α-hydroxylase, the rate-limiting enzyme in bile salt synthesis from cholesterol [68]. The increase in bile salt synthesis decreases hepatocyte cholesterol concentration, leading to increased expression of the LDLr and enhanced LDL clearance from the circulation [27]. The effectiveness of bile salt sequestrants in clearing LDL from plasma is partially offset by concurrent upregulation of hepatic cholesterol and triglyceride synthesis, which stimulates the production of VLDL particles by the liver [69]. As a result, bile salt sequestrants may also raise triglyceride levels and should be used with caution in patients with hypertriglyceridemia.

Inhibitors of cholesterol absorption Cholesterol absorption inhibitors reduce cholesterol absorption by the small intestine. This includes dietary cholesterol, but more importantly they reduce the reabsorption of biliary cholesterol, which makes up the majority of intestinal cholesterol [63]. Like statins and bile salt binding resins, cholesterol absorption inhibitors reduce LDL cholesterol by increasing clearance via the LDLr [70]. However, they may also inhibit hepatic production of VLDL. There are two available cholesterol absorption inhibitors: plant sterols/stanols and ezetimibe. Plant sterols and stanols are naturally present in vegetables and fruits. However, they may be consumed in larger amounts from nutritional supplements. Plant sterols and stanols are similar in molecular structure to cholesterol, but are substantially more hydrophobic [71]. As a result, plant sterols and stanols displace cholesterol from micelles, increasing the loss of cholesterol from the feces [6]. The plant sterols and stanols are themselves poorly absorbed. Based upon

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their mechanism of action, gram quantities of plant sterols and stanols are required to reduce plasma LDL cholesterol concentrations by approximately 15%. Because an average diet contains 200–400 mg of plant sterol and stanols, the molecules must be enriched in dietary supplements (approximately 2 g) to be effective [6]. Ezetimibe at very low concentrations reduces intestinal cholesterol absorption by about 50%, without reducing the absorption of triglycerides or fat-soluble vitamins. Ezetimibe decreases cholesterol movement from micelles into the enterocyte by inhibiting uptake through the brush border protein NPC1L1 (Figure 19.3b) [10]. A reduction in cholesterol absorption that is achieved by either plant sterols and stanols or ezetimibe would be expected to decrease the cholesterol content of chylomicrons and therefore the movement of cholesterol from the intestine to the liver. This is because chylomicrons retain the intestinal cholesterol that was originally incorporated into the particles after they are metabolized to remnants by the liver. Within the liver, cholesterol derived from chylomicron remnants contributes to the cholesterol that is packaged into VLDL particles. Therefore, inhibiting cholesterol absorption reduces cholesterol incorporation into VLDL, and decreases LDL cholesterol concentrations in the plasma. Reduced hepatic cholesterol contents [72] lead to upregulation of the LDLr [70], which also contributes to the mechanism of LDL lowering by cholesterol absorption inhibitors.

Fibrates Fibrates bind and activate peroxisome proliferator-activated receptor α (PPARα), a nuclear receptor expressed in hepatocytes, skeletal muscle, macrophages, and the heart [73]. Upon fibrate binding, PPARα heterodimerizes with the retinoid X receptor (RXR). This heterodimer binds to peroxisome proliferator response elements (PPREs) present in the promoter region of specific genes and activates transcription of the target genes [74]. Activation of PPARα by fibrates results in numerous changes in lipid metabolism [73, 74] that act to decrease plasma triglyceride levels and increase plasma HDL. The decrease in plasma triglyceride levels is caused by increased muscle cell expression of LPL, decreased hepatic expression of apoC-III, and increased hepatic oxidation of fatty acids. The increased muscle expression of LPL results in increased uptake of triglyceride-rich lipoproteins, with a resultant decrease in plasma triglyceride levels [75]. Because apoC-III normally functions to inhibit interaction of triglyceride-rich lipoproteins with their receptors [18, 76], the decrease in hepatic production of apoC-III may potentiate the increased LPL activity. The mechanisms by which fibrates raise plasma HDL levels are complex [77]. Whereas PPARα decreases hepatocyte production of apoA-I in the mouse, the opposite

appears be the case in humans. This contributes directly to increased plasma HDL. Upregulation of SR-B1 and ABCA1 in macrophages presumably promotes cholesterol efflux from these cells in vivo. Hepatocytes also reduce expression of SR-B1 in response to PPARα, which contributes to increased HDL levels in the plasma. Fibrates lower LDL levels modestly. The lower LDL levels result from a PPARα-mediated shift in hepatocyte metabolism toward fatty acid oxidation. PPARα increases the expression of numerous enzymes involved in fatty acid transport and oxidation. This increases fatty acid catabolism, leading to decreased triglyceride synthesis and VLDL production. PPARα activation also results in LDL particles of larger size, which appear to be taken up more efficiently by the LDLr. Many of these effects of PPARα on lipid metabolism are still the subject of basic and clinical investigation, which may lead to development of more selective PPARα agonists that are capable of targeting selective aspects of lipid metabolism [78].

Niacin Niacin (nicotinic acid, vitamin B3 ) is a water-soluble vitamin. At physiological concentrations, it is a substrate in the synthesis of nicotinamide adenine dinucleotide (NAD) and nicotinamide adenine dinucleotide phosphate (NADP), which are important cofactors in intermediary metabolism. The pharmacological use of niacin necessitates large doses (1500–3000 mg day−1 ) and is independent of the conversion of nicotinic acid to NAD or NADP. Niacin decreases plasma LDL cholesterol and triglyceride concentrations and increases HDL cholesterol [79]. Recent studies have identified a G-protein-coupled receptor on adipocytes, which appears to mediate the well-documented metabolic changes associated with niacin administration [80]. Niacin decreases adipose tissue hormone-sensitive lipase activity. Decreased hormone-sensitive lipase reduces peripheral tissue triglyceride catabolism, and therefore decreases the flux of free fatty acids to the liver. This, together with local effects of niacin in liver [79], decreases the rate of hepatic triglyceride synthesis and VLDL production. Niacin also decreases the fractional catabolic rate apoA-I [79, 81]. The increased plasma apoA-I increases plasma HDL concentrations and presumably augments reverse cholesterol transport.

Omega-3 fatty acids The omega-3 fatty acids eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA), also referred to as fish oil, are effective at reducing plasma triglycerides [82].

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Although the molecular mechanisms are incompletely understood, the effects are reduced hepatic triglyceride biosynthesis and increased fatty acid oxidation in the liver [83].

CONCLUSION The efficacy of available lipid-lowering drugs to reduce LDL cholesterol, particularly with statins, represents an important advance in reducing cardiovascular disease mortality. Future advances will build upon a rich knowledge base of lipoprotein metabolism in order to identify new biochemical targets for the management of dyslipidemias.

ACKNOWLEDGMENTS This work was supported in part by research grants DK56626 and DK48873 from the National Institutes of Health (US Public Health Service), and an Established Investigator Award from the American Heart Association.

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19: LIPOPROTEIN METABOLISM AND CHOLESTEROL BALANCE

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SECTION B : BILE SECRETION

20

Bile Acids and the Enterohepatic Circulation Alan F. Hofmann Division of Gastroenterology, Department of Medicine, University of California, San Diego, CA, USA

INTRODUCTION The mammalian hepatocyte performs myriad functions. One of these is the conversion of cholesterol to conjugated bile acids, followed by their secretion across the canalicular membrane into bile. At its simplest, bile acid formation can be considered the dominant chemical pathway for cholesterol elimination. The matter is made more complex for three reasons. First, efficient intestinal conservation of secreted bile acids and subsequent efficient plasma clearance of bile acids returning to the liver leads to the accumulation of a pool of recycling bile acids that moves between the liver and the intestine; the movement of this pool of bile acid molecules is termed the enterohepatic circulation. Second, bile acids are altered by intestinal bacteria during their enterohepatic circulation, leading to the accumulation of novel bile acids in the bile acids that circulate enterohepatically. Third, bile acids have multiple functions in the biliary tract and the intestine. Some of these functions relate to the amphipathic properties of bile acids that enable them to solubilize polar lipids in mixed micelles. Other functions relate to signaling properties of bile acids. Indeed, bile acids by being formed in the hepatocyte and acting on G-protein-coupled receptors in non-hepatic tissues fulfill the definition of being hormones. In this brief chapter, an attempt will be made to summarize current knowledge of bile acid chemistry and biology in mammals, with emphasis on bile acid metabolism in the The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

hepatocyte. Principles of therapy with bile acid agonists (exogenous bile acids or bile acid congeners) and bile acid sequestrants will be reviewed. There has been stunning progress in the past decade, evidenced by cloning of most of the major bile acid transporters, and identification of the nuclear transcription factors activated by bile acids. Recently, there has been emerging evidence that bile acids influence intermediary metabolism. For older references on the chemistry and biology of bile acids, two textbooks are available [1, 2], as well as several recent reviews of bile acid metabolism [3–5] which expand on some of the topics discussed only briefly here. Other chapters in this book deal with bile formation and secretion (Chapter 23), cholesterol (Chapter 18), basolateral transport by the hepatocyte (Chapter 21), apical transport (Chapter 24), biliary epithelial function (Chapter 25), and inheritable (Chapter 42) and acquired (Chapter 43) cholestasis. The Falk Foundation of Freiburg Germany hosts a conference on bile acids every two years, and the published proceedings of these conferences document the continuing progress in the field (see [6]).

BILE ACID CHEMISTRY Bile acid biosynthesis The biosynthesis of bile acids from cholesterol involves multiple enzymes—at least 15—and takes place in

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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Figure 20.1 Depiction of changes in the cholesterol molecule when it is biotransformed to chenodeoxycholic acid, the root C24 bile acid

multiple organelles [7, 8]. It is believed to occur in pericentral hepatocytes. Both the steroid nucleus and the side chain of cholesterol are modified during bile acid biosynthesis, and the result is a marked change in the physicochemical properties of cholesterol. Cholesterol is a flat membrane-loving molecule with a terminal 3β-hydroxy group facing the aqueous phase. In contrast, the bile acid nucleus is bent and amphipathic, as the hydroxyl groups are positioned on only one face (the α face) of the steroid nucleus; in addition, the terminal carbon atom of the side chain becomes functionalized, enabling conjugation (see below). Figure 20.1 shows the changes in chemical structure in the biotransformation of cholesterol to chenodeoxycholic acid (CDCA), the root bile acid. In the major pathway for bile acid biosynthesis, the first step is hydroxylation at the carbon atom C-7. Such 7α-hydroxylation is mediated by CYP7A1, a microsomal enzyme, and considerable evidence indicates such 7-hydroxylation is the rate-limiting enzymatic step in bile acid biosynthesis. The 3β-hydroxy group of cholesterol is oxidized to a 3-oxo group by a steroid dehydrogenase. The 5,6 double bond of cholesterol (in the B ring) is simultaneously isomerized to a 4,5 position (in the A ring). Two aldo-keto reductases then reduce the 3-oxo group to a 3α-hydroxy group and convert the 4,5 molecule to a saturated molecule in which the A and B rings are cis to each other, denoted by the C-5 hydrogen atom being in the β configuration. The result is a C27 sterol that has a 3α,7α-dihydroxy-5β (A/B cis ring juncture) but has maintained the side chain of cholesterol. Hydroxylation at C-12 may also have occurred, mediated by CYP7B1. Cholesterol has a branched C8 (isooctane) side chain. In bile acid synthesis in mammals, the C8 isooctane side chain undergoes a modified β-oxidation, resulting in the formation of a C5 isopentanoic side chain. Such biotransformation begins with a mitochondrial hydroxylation at the terminal C-27 carbon mediated by CYP27A1. The

oxidation may continue to the C-27 acid, mediated by the same hydroxylase. Eventually, in a modified β-oxidation, three carbons are cleaved from the side chain via a C-24-oxo intermediate. All of the steps in the oxidative cleavage of the cholesterol side chain acids occur in the peroxisome. The end result is a di- or trihydroxy 5β-C24 bile acid with a C19 nucleus and a C5 chain. The carboxyl group is present as the thio-ester of coenzyme A (CoA) and will then be conjugated with glycine or taurine (see below). In humans, two bile acids are synthesized from cholesterol: CDCA (3α,7α-dihydroxy) and cholic acid (3α,7α,12α-trihydroxy). CDCA inherits a 3-hydroxy group from cholesterol, and 7-hydroxylation is the rate-limiting step in bile acid biosynthesis. Thus, as noted above, CDCA may be considered the root bile acid. Its cumbersome name results from its being discovered some decades after cholic acid, its source (“the goose”, cheno- in Greek), and its chemical property of having one less oxygen atom (hence the term “deoxy”) when subjected to elemental analysis. Cholic acid is not formed by 12-hydroxylation of CDCA, but by microsomal 12-hydroxylation of an early intermediate in CDCA synthesis. In a few mammals (bears and some caviomorphs), the root bile acid is the 7β-hydroxy epimer of CDCA termed ursodeoxycholic acid (UDCA). Whether UDCA is formed by direct 7β-hydroxylation of cholesterol or by epimerization of a 7α-hydroxy group is not known. Some caviomorphs form bile acids with a 7-oxo group as their primary bile acid. In humans, the daily synthesis rate of cholic acid averages 200 mg per day and that of CDCA is about 100 mg per day. The fractional turnover rate of CDCA—about 20% per day—is less than that of cholic acid—about 30% per day. As a consequence, the pool sizes of each of the primary bile acids are roughly equal, and average about 1 g each [9].

20: BILE ACIDS AND THE ENTEROHEPATIC CIRCULATION

Presently, there are considered to be two pathways of bile acid biosynthesis. In the so-called “neutral” pathway, nuclear changes are completed before side chain oxidation begins. This pathway leads to both cholic acid and CDCA. In the “acidic” or “alternative” pathway, side chain hydroxylation at C-27, generally followed by oxidation to a carboxyl group, initiates bile acid biosynthesis. Subsequent steps involve nuclear changes. This pathway is considered to generate mostly CDCA. There are also additional minor routes of bile acid biosynthesis beginning with 24-hydroxycholesterol and 25-hydroxycholesterol. The individual steps in bile acid biosynthesis are important in that defects in any one of these enzymes lead to clinical manifestations. When a defect occurs, the intermediates accumulate and are excreted in urine, where they are readily detected by mass spectrometry. Biosynthesis of intermediates may be greatly increased because of loss of negative feedback regulation of bile acid biosynthesis by bile acids, the end product of the biosynthetic pathway. Treatment involves replacement with exogenous bile acids. Inborn errors of bile acid biosynthesis causing cholestatic liver disease are discussed in detail on chapter 42. A deficiency of CYP27A1 that mediates hydroxylation (followed in some cases by oxidation to a carboxyl group) causes the disease cerebrotendinous xanthomatosis. Exuberant hydroxylation of bile acid intermediates occurs, and these, in the form of their glucuronides, are excreted in urine and bile. Administration of CDCA represses bile acid synthesis and ameliorates the disease. Other mammals form trihydroxy bile acids by hydroxylation at a site other than C-12, and in general trihydroxy bile acids are more common in mammals than are dihydroxy bile acids. Pigs form a 6α-hydroxy-derivative of CDCA termed hyocholic acid. Mice and rats form 6β-hydroxy derivates of CDCA termed muricholic acids. In non-mammals, hydroxylation occurs at still additional sites: 1β, pigeon; 15α, swans and geese; 16α, birds and snakes. All of these hydroxylations occur on the α face of the steroid nucleus, with the result that the steroid nucleus has amphipathic properties. Hydroxylation may also occur on the side chain on the α-carbon (in the R configuration; such bile acids occur in marine mammals). A few species desaturate the side chain, forming a double bond between C-22 and C-23. The structures of C24 bile acids occurring in mammals are shown in Table 20.1. Bile acids are the only small molecules that vary widely in vertebrates. It appears that in evolution, there were many solutions to eliminating cholesterol in the form of a water-soluble, amphipathic molecule. In ancient fish (for example cartilaginous fish), some bony fish, a few ancient mammals (elephant, manatee, hyrax), and some amphibians, the C8 side chain undergoes only hydroxylation at C-27, with the result that a C27 bile alcohol rather than a bile acid is formed. (A few frog species have bile alcohols in which the side chain is shorter or longer than C8 .) In

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amphibians, ancient reptiles, and ancient birds, the side chain undergoes oxidation at the terminal carbon atom to form a C8 carboxylic acid. Such C27 bile acids are the dominant bile acids in these species. Side chain structure is used to divide “bile salts” into three great classes: C27 bile alcohols, C27 bile acids, and C24 bile acids. It is beyond the scope of this chapter to discuss the complex chemistry of bile acids and bile alcohols in non-mammals; a review of older work is available [2]. Figure 20.2 summarizes bile alcohol and bile acid formation in most vertebrates, showing the major sites of hydroxylation.

Bile acid conjugation After their biosynthesis from cholesterol, C24 bile acids are linked in amide linkage to glycine or taurine. The linkage is between the carboxyl group of the bile acid and the amino group of glycine or taurine. The term “conjugation” was used historically for this biotransformation, but in the past two decades it has been recognized that bile acids may undergo other modes of conjugation; that is, sulfation, glucuronidation, and N-acetylaminoglucosidation. Linkage to glycine or taurine may be termed N-acylamidation or simply “amidation,” and the glycine- and taurine-conjugated bile acids may be referred to as amidates. (The term amidation for bile acids should not be confused with that used for amidation (with ammonia) of a C-terminal carboxyl group of a peptide). The amide bond is highly stable and is not cleaved in tissues unless pathological levels of bile acids are reached. Glycine and taurine amidates are also resistant to pancreatic carboxypeptidases, whereas amidates in which bile acids are coupled to other amino acids (except for aspartate) are rapidly cleaved. Figure 20.3 illustrates the various types of bile acid conjugation. Conjugation (amidation) of bile acids has a number of biological and physicochemical consequences [10, 11]. First, the carboxyl group of a glycine amidate has a lower pKa (about 4.0) than that of its corresponding unconjugated precursor (about 5.0). The pKa of taurine conjugates is less than 2. As a result, conjugated bile acids are always fully ionized at physiological pH and therefore are impermeable to the apical cell membranes of the hepatocyte, cholangiocyte, and enterocyte. In addition, amidation makes bile acids soluble at acidic pH and resistant to precipitation by Ca2+ ions. As a result, high concentrations of bile acids can be maintained in the lumen of the biliary tract and small intestine. Bile alcohols cannot ionize and are likely to have an extremely low aqueous solubility. They are rendered water soluble by esterification with sulfate (sometimes termed “sulfonation”) at C-27, a biotransformation mediated by a cytosolic sulfotransferase. Such sulfation converts bile alcohols into water-soluble amphipaths that have the solubilizing properties of amidated bile acids.

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Table 20.1 C24 bile acids of healthy mammalsa Trivial name Cholic Allocholic Deoxycholic Allodeoxycholic Chenodeoxycholic Ursocholic Ursodeoxycholic Lithocholic (None proposed) Lagodeoxycholic α-Muricholic β-Muricholic ω-Muricholic Murideoxycholic Hyocholic Hyodeoxycholic Vulpecholic (None proposed) (None proposed) (None proposed) (None proposed) (None proposed) (None proposed) Phocaecholic (None proposed) Isolithocholic Isodeoxycholic Isochenodeoxycholic Isoursodeoxycholic Isocholic

A/B RJ, A ring substituentsb

B ring substituents

5β,3αOH 5α,3αOH 5β,3αOH 5α,3αOH 5β,3αOH 5β,3αOH 5β,3αOH

7αOH 7αOH — — 7αOH 7βOH 7βOH

C and D ring C5 side chain substituents substituents Source Occurrence

5β,3αOH — 5β,3αOH — 5β,3αOH — 5β,3αOH 6βOH,7αOH 5β,3αOH 6βOH, 7βOH 5β,3αOH 6αOH,7βOH 5β,3αOH 6βOH 5β,3αOH 6αOH,7αOH 5β,3αOH 6αOH 5β,1αOH,3αOH 7αOH 5β,1βOH,3αOH 7αOH 5β,3αOH 5β,3αOH 5β,3αOH 5β,3αOH 5β,3αOH 5β,3αOH 5β,3αOH 5β,3βOH 5β,3βOH 5β,3βOH 5β,3βOH 5β,3Boh

7-oxo 7-oxo 7-oxo 7βOH 7αOH 7αOH — — 12αOH 7αOH 7βOH 7αOH

12αOH 12αOH 12αOH 12αOH — 12αOH —

None None None None None None None

L L or I L or L L or L or

— 15αOH 12βOH — — — — — — — —

None None None None None None None None None None None

I I I I L I I L I L L

— 12αOH — — 12αOH — 12OH — — — — 12αOH

None None None (22+ ) None (22+ ) 23-(R)-OH 23-(R)-OH 23-(R)-OH — — — — —

L L L L L L L I I I I I

I I I I

Many species Minor BA in newborn rabbit Many species Minor BA in rabbit Many species Trace BA in human Major BA: nutria, bear, beaver Many species Wombat Trace BA in rabbit Rodents (minor bile acid) Rodents Rodents (minor bile acid) Rodents Pig Pig Australian opossum Minor BA in infant, mouse, sheep Caviomorphs, koala, sifaka Sloth Paca (caviomorph) Agouti Minor BA in sea mammals Major BA in sea mammals Minor BA in sea mammals Major fecal BA in human Major fecal BA in human Minor fecal BA in human Minor fecal BA in human Minor fecal BA in human

a Hydroxy-oxo bile acids, trace BA in both biliary and fecal BA are not listed.

Complex mixtures of C27 bile alcohol (sulfates) and C24 bile acids occur in the elephant, manatee, hyrax, rhinoceros, and some caviomorphs. The bile of horses contains a complex mixture of C27 alcohol sulfates, C27 bile acids, and C24 bile acids. Several primates contain a substantial proportion of C27 bile acids. Unsaturated derivatives of isolithocholic and isochenodeoxycholic acid have been reported in disease conditions [2]. b A/B RJ, A/B ring juncture; L, liver; I, intestinal bacteria.

Bacterial modification of bile acids Bile acids circulate enterohepatically. In the small intestine, conjugated bile acids are exposed to bacteria whose enzymes mediate hydrolysis of the amide bond connecting the bile acid with the amino group of glycine or taurine. Such deconjugation generates an unconjugated bile acid [12]. In the large intestine and possibly in the terminal ileum, bacterial enzymes also mediate removal of the 7α-hydroxy group to form 7-deoxy bile acids. Such 7-deoxy bile acids are termed secondary bile acids to indicate that they were formed by bacterial enzymes from primary bile acids. Primary bile acids are defined as those bile acids biosynthesized in the hepatocyte from cholesterol. Dehydroxylation at C-7 is a complex, multienzyme process mediated by anaerobic bacteria. It involves formation of a 3-oxo4,5 6,7 resonating intermediate [12]. In humans, when CDCA undergoes 7-dehydroxylation,

lithocholic acid (LCA) (3α-hydroxy) is formed. When cholic acid undergoes 7-dehydroxylation, deoxycholic acid (DCA) (3α,12α-dihydroxy) is formed. Both of these bile acids are absorbed passively to some extent from the colon and join the recycling primary bile acids. Bacteria also may dehydrogenate the nuclear hydroxyl groups to render hydroxy, oxo-bile acids, and may epimerize the 3α-hydroxy group of bile acids to a 3β-hydroxy group. Deconjugation and dehydroxylation decrease the solubility of bile acids, and the aqueous concentration of bile acids in the human cecum is 1/20th that of the proximal small intestine [13]. Bile acids have direct and indirect antimicrobial properties [14]. The decrease in bile acid concentration in the cecum may allow bacteria to flourish. Cecal bacteria produce large amounts of short-chain fatty acids from incompletely absorbed carbohydrate, and such short-chain fatty acids are an important source of calories in some species.

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293

(a) (b)

(c)

(d)

Figure 20.2 Biotransformation of cholesterol (a) to C27 bile alcohols (b), C27 bile acids (c) and C24 bile acids (d). Bile alcohols are shown as conjugated with sulfate; bile acids, with taurine. Sites of hydroxylation occurring in many species are shown by large arrows; those occurring in only a few species are shown by small arrows. New sites of hydroxylation may be discovered in the future

Figure 20.3 Conjugation of bile acids. N-acylamidation with taurine or glycine is by far the major mode of conjugation. Bile acid sulfation occurs with amidates of lithocholic acid in healthy humans, and with amidates of primary bile acids in cholestatic liver disease. Bile acid glucuronidation is rare except for administered hyodeoxycholic acid in humans or for C23 (nor) bile acids such as norursodeoxycholic acid

ENTEROHEPATIC CYCLING OF BILE ACIDS Overview When a meal is ingested, bile acids that are stored in the gall bladder are slowly discharged into the proximal small intestine. Efficient intestinal conservation of secreted bile acids in the distal small intestine (>95%) leads to the

accumulation of mass of bile acids that circulates between the liver and small intestine, as illustrated in Figure 20.4. The amount of bile acids, that is recycled is termed the bile acid pool and can be measured by an isotope dilution technique [9]. Small amounts of bile acids escape intestinal absorption but the pool of recycling bile acids is kept relatively constant by biosynthesis from cholesterol in the hepatocyte. In the few species lacking a gall bladder (rat, deer, horse, among others), the bile acid pool is stored in the small intestine.

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THE LIVER: ENTEROHEPATIC CYCLING OF BILE ACIDS

to by nal sion

ort transit to by mesen and hepa arterial b flow

fecal excretion (= biosynthesis)

Figure 20.4 Schematic depiction of the enterohepatic circulation of bile acids (and bile alcohol sulfates) in species possessing a gall bladder. Absorption from the small intestine in addition to that mediated by the ileal transport system may involve passive absorption of protonated glycine dihydroxy bile acids, paracellular absorption, or carrier-mediated absorption using transporters not yet identified. At present, the major transporters of the ileal transport system are apical ASBT and basolateral OSTα/OSTβ

The amount of bile acids entering the small intestine is termed bile acid secretion and is a flux; it can only be measured by an indicator dilution technique such as is used to measure cardiac output [15]. Division of bile acid secretion by the pool size gives the recycling frequency of the bile acid pool, probably a meaningless number physiologically. The bile acid pool of humans recycles about twice during digestion of an average meal, and averages about 12 g day−1 . In humans, biliary bile acids consist of approximately equal proportions of conjugates of CDCA and cholic acid. DCA conjugates constitute a smaller fraction that increases with age. LCA amidates (partly sulfated) and UDCA amidates constitute a small percentage of biliary bile acids [16].

Intestinal conservation of bile acids The efficient intestinal conservation of bile acids is mediated by active and passive absorption; active absorption is

likely to be the major mechanism. However, the efficiency of absorption and the proportion of active versus passive absorption is likely to be species-dependent. As conjugated bile acid anions are membraneimpermeable, they can only be absorbed by carriermediated mechanisms or by the paracellular route. Conjugated bile acid anions are usually considered too large to leak through the paracellular junctions of the healthy human small intestine, but whether they are absorbed paracellularly in subjects with increased intestinal permeability is not known. Glycine-conjugated dihydroxy bile acids, when protonated, are membrane-permeable, and passive absorption across the apical membrane can occur in principle when duodenal content is acidic [17]. The dominant mechanism for the absorption of conjugated bile acids in human and mouse is active absorption by the ileal bile acid transport system. This involves an apical sodium-dependent cotransporter, apical bile acid transporter (ASBT), whose synthesis is mediated by the gene SLC10A2 . Knockout of this gene in mice [18] or

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absence of a normal gene in humans [19] causes profound bile acid malabsorption. Basolateral transport is mediated by a heterodimer of two proteins—OSTα/OSTβ [20]. Bile acids largely exist in conjugated form in bile. Unconjugated bile acids are formed by bacterial enzymes. Such deconjugation begins in the ileum, at least in humans. In non-human species, it might well occur in the more proximal small intestine. In humans, and presumably in all species with a cecum, deconjugation goes rapidly to completion when bile acids enter the large intestine [12]. The resultant unconjugated bile acids have varying degrees of passive membrane permeability. LCA, a monohydroxy bile acid, and probably most dihydroxy bile acids, are highly membrane-permeable. Trihydroxy bile acids have much more limited membrane permeability and are absorbed more slowly. However, if formed in the distal small intestine, they may also be absorbed actively by the ileal transport system. The magnitude of unconjugated bile acid absorption in the distal small intestine is likely to exceed that of unconjugated bile acid absorption from the colon, based on modeling of the enterohepatic circulation of bile acids in humans [21]. It has always been assumed that unconjugated bile acids pass through the enterocytes of the small intestine without undergoing any biotransformation. Recently, a pathway involving luminal deconjugation of CDCA conjugates, passive absorption, sulfation, and extrusion of the CDCA sulfate back into the lumen presumably by the efflux pump MRP2, has been identified in a small group of children with functional constipation [22]. There is no a priori reason why this sequence of events cannot occur in the healthy individual.

Damage and repair of bile acids during enterohepatic cycling In humans, the end products of deconjugation and 7-dehydroxylation are LCA and DCA; these are the major fecal bile acids. In mice and rats, deconjugation and 7-dehydroxylation generate murideoxycholic acid (3α,6β-dihydroxy) as well as DCA. In addition to deconjugation and 7-dehydroxylation, bacterial enzymes may mediate additional changes in the bile acid molecule. A common change is epimerization of the 3α-hydroxy group to a 3β-hydroxy group; such 3β-hydroxy bile acids are termed iso-bile acids. In addition, the hydroxy groups may be oxidized to an oxo-group. Removal of the sulfate group at C-3 may generate a 3,4 compound. Isomerization of the A/B ring juncture from A/B cis to A/B trans may occur, presumably via a 4,5 and/or 5,6 intermediate to form 5α (allo) bile acids. In the side chain, desaturation at C-22–C-23 may occur. All of these changes result in an enormous complexity of unconjugated fecal bile acids [23]. Moreover, deconjugation and desulfation may not be complete, and esterification of bile

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acids with fatty acids and nucleic acids may occur. Thus, giving a complete description of fecal bile acids—by class and by individual bile acids within a class—is an enormously difficult challenge to an analytical chemist. The great complexity of fecal bile acids contrasts with the relative simplicity of biliary bile acids, which are usually predominantly two to four in number. There are two reasons for the relative simplicity of biliary bile acids as compared to fecal bile acids. First, not all fecal bile acids are absorbed from the colon. Second, and more importantly, “damaged” bile acids are repaired during their transport through the hepatocyte. In the repair processes, iso (3β-hydroxy) bile acids are epimerized to 3α-hydroxy bile acids [24]. 3-oxo bile acids are also reduced to 3α-hydroxy bile acids. 7-oxo bile acids are reduced to 7-hydroxy bile acids, but whether the reduction goes to a 7α-hydroxy or a 7β-hydroxy group (or both) is species-dependent. Whether or not the major bacterial biotransformation pathway of 7-dehydroxylation is reversed depends on the bile acid and the species. DCA is effectively rehydroxylated at C-7 to form cholic acid in some species, and partly rehydroxylated in others. However, in humans, no 7-rehydroxylation occurs, and absorbed DCA is conjugated and circulates with the primary bile acids. LCA is a toxic bile acid when administered orally in many species [25, 26] and its detoxification is mediated in a species-dependent manner. In rats, LCA undergoes hydroxylation at C-6; in wombats at C-15 [27]. In humans, LCA undergoes a unique mode of double conjugation [26]. It is not only amidated at its C-24 carboxyl group, but also undergoes esterification with sulfate at C-3. Such sulfated amidates of LCA are not substrates for the ileal transport system and as a consequence LCA is rapidly lost from the enterohepatic circulation. Sulfation of the amidates of LCA is not complete during transport through the hepatocyte, and only about half of the LCA amidates secreted into bile are sulfated [16]. However, the unsulfated half is absorbed from the small intestine, again partly sulfated during hepatocyte transport, and these double conjugates are rapidly excreted. Bile acid glucuronides are not formed to any extent. Ethereal glucuronides (at C-3) do not form as the charged amidates do not enter the interior of the endoplasmic reticulum (ER); ester glucuronides (at C-24) cannot occur because of the amide bond at that location.

Hepatic uptake of bile acids The majority of bile acids returning to the liver are in conjugated form and their composition is that of biliary bile acids. A minority are unconjugated. The unconjugated bile acids in turn have two sources: first, unconjugated bile acids formed in the distal small intestine by bacterial deconjugation of biliary primary and secondary bile acids,

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and second, a very complex mixture of unconjugated secondary bile acids formed by bacterial enzymes in the colon. Bile acids return to the liver in portal blood. Trihydroxy bile acids are bound to albumin with moderate affinity (50–80%); dihydroxy bile acids and LCA are more tightly bound (>95%) [28]. Uptake occurs in direct proportion to concentration, and is greatest in the most periportal hepatocytes. Uptake is highly efficient (50–90%) and independent of load. The efficiency of uptake depends on bile acid structure and the state of conjugation; it is also species-dependent. Plasma is enriched in those bile acids that are inefficiently extracted. Therefore plasma bile acids are enriched in dihydroxy conjugates (compared to trihydroxy conjugates), in unconjugated bile acids (compared to their corresponding conjugated bile acid derivatives), and possibly in iso (3β-hydroxy) bile acids. Those bile acids that spill over into the systemic circulation are presented once again to the hepatocyte in hepatic arterial blood flow as well as splanchnic blood flow. The T 1/2 of plasma disappearance in humans for all bile acids that have been studied is less than five minutes [28]. Because hepatic uptake of bile acids is efficient, because bile acids are bound to albumin in plasma, and because the small amount of bile acids that enters the glomerular filtrate is reabsorbed in the proximal tubule (mediated by ASBT), loss of bile acids in the urine is negligible. Uptake of conjugated bile acids is mediated by NTCP1. Probably one or more of the diverse OATPs (sodium-independent) also participate (see chapter 21) Uptake of conjugated bile acids by NTCP is unidirectional (because of the transmembrane sodium gradient), but there is a cotransporter (MRP4) which mediates co-efflux of conjugated bile acids and glutathione from the hepatocyte into the space of Disse. Therefore conjugated bile acid uptake may involve percolation down the plate of hepatocytes [29]. Uptake of unconjugated bile acids may proceed by multiple mechanisms. First, unconjugated bile acids may avail themselves of the transporters (NTCP and OATPs) that mediate conjugated bile acid absorption. Second, a fatty acid transporter, FATP5, mediates simultaneous transmembrane permeation and esterification with CoA [30]. Third, mono- and dihydroxy bile acids may be absorbed passively (that is, without the involvement of a carrier) by simple flipflop through the lipid domains of the basolateral membranes. Unconjugated bile acids, if not esterified with CoA during entry, will immediately undergo CoA formation. They then travel to the peroxisome, where amidation with taurine or glycine occurs. The bile acid CoA ligase accepts as substrate only bile acids with a chain length of C5 or greater. Bile acids with a C8 side chain are converted to C24 bile acids. Bile acids with a C6 side chain (homo-bile acids) undergo β-oxidation in peroxisomes to form bile acids with a C4 side chain (nor or C24 -nor bile acids). The subsequent

fate of these C23 molecules depends on their structure. Trihydroxy nor bile acids, for example norcholate, are secreted into bile as such. Dihydroxy nor bile acids are in part secreted as such into bile; another part partitions into the ER interior and undergoes ethereal and/or ester glucuronidation. C22 (dinor) bile acids are metabolized in the same manner as C23 (nor) bile acids [31]. Bile acids that enter the hepatocyte are rapidly removed by secretion into the canalicular space mediated by the ATP-stimulated transporter BSEP [32, 33]. Details of transhepatocyte transport have not yet been clarified. The working assumption is that transport is mostly by diffusion, and perhaps to a minor extent by vesicular transport. The monomeric concentration of bile acids in hepatocyte cytosol is believed to be very low—perhaps 1 µM. Canalicular transport of bile acids induces bile acid-dependent bile flow and biliary lipid secretion, as reviewed in Chapter 23. During late pregnancy, biosynthesized conjugated bile acids are transferred from the fetus to the mother across the placenta. The major transporters are members of the OATP and MRP families [34]. Unconjugated dihydroxy bile acids are membrane-permeable and may transfer passively from mother to fetus or fetus to mother [35].

Plasma bile acids In health, the predominant source of bile acids in the systemic circulation is spillover of bile acids returning from the intestine. In dogs with a biliary fistula, the concentration of plasma bile acids is too low to measure. In cholestatic liver disease, newly synthesized as well as returning bile acids are regurgitated from hepatocytes because of upregulation of efflux transporters [36]. In addition, bile acids may be secreted into canalicular bile and then reabsorbed by cholangiocytes. Plasma bile acids also increase because of decreased first-pass clearance by the hepatocyte. Ileal input of bile acids varies in the fasting state because of the infrequency and variable activity of the interdigestive motility complex. Because the level of plasma bile acids fluctuates during the fasting state, fasting-state plasma bile acid levels vary considerably, and their level has not been shown to be superior to conjugated bilirubin levels for the detection of liver disease or the assessment of hepatocyte function [37]. Plasma bile acid levels can be useful to stage cholestatic liver disease, for example, in cholestasis of pregnancy [38]. Whether the low level of plasma bile acids in the systemic circulation in health has any physiological function is not known. In mice, bile acids activate a G-coupled protein receptor in brown adipose tissue, leading ultimately to triiodothyronine formation and thermogenesis [39]. In humans, plasma bile is higher in the first few years of life when brown adipose tissue is present [40], and the possibility remains that this “function” occurs in infants.

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REGULATION OF THE ENTEROHEPATIC CIRCULATION Regulation of bile acid biosynthesis The enterohepatic circulation of bile acids has two modes of regulation. The first is negative feedback of bile acid biosynthesis from cholesterol in the hepatocyte. The second is negative feedback regulation of bile acid transport in the ileal hepatocyte. Both of these regulatory mechanisms are signaled by the intracellular concentration of bile acids acting on the nuclear receptor FXR. In health, bile acid biosynthesis occurs mostly in the pericentral hepatocytes [41]. Bile acid synthesis does not occur in the periportal hepatocytes as it is suppressed by the flux of bile acids returning from the intestine. In the pericentral hepatocytes, bile acids readily enter the nucleus as the molecules are sufficiently small to easily diffuse through the pores of the nuclear membrane. In the nucleus, bile acids bind to FXR. An FXR–RXR heterodimer acts to stimulate the synthesis of an inhibitory protein SHP, which in turn displaces a promotor factor (HNF4) from the promotor of the CYP7A1 gene, the rate-limiting enzyme in bile acid biosynthesis [42]. An additional essential negative regulatory factor is FGF15(19), a peptide secreted by the ileal enterocyte [43]. This peptide acts on a basolateral receptor FGF4 which ultimately leads via a JNK-mediated pathway to synergize suppression of CYP7A1 transcription. The result of these complex regulatory circuits is that bile acid biosynthesis is autoregulated by bile acids themselves in a negative-feedback manner. At present, a regulatory role for microRNAs has not been identified. Additional genes besides CYP7A1 are regulated by FXR. These include SLC10A1 , the gene encoding NTCP1, the sodium-dependent cotransporter involved in conjugated bile acid uptake; SLC10A1 is downregulated by FXR activation. A gene that is upregulated is ABCB11 , the gene encoding BSEP, the conjugated bile acid efflux pump. Therefore the end result of FXR activation is to decrease the concentration of bile acids in all hepatocytes. When bile acid return to the liver is decreased, for example by ileal dysfunction, bile acid synthesis increases. The increase is 10-fold in humans and 20-fold in mice. The site of the increased bile acid biosynthesis may involve not only pericentral cells but also hepatocytes located in the direction of the portal triad. Increased bile acid biosynthesis requires increased availability of cholesterol, the precursor. Cholesterol may be made available either by uptake of low-density lipoproteins (LDLs) from plasma or by increased biosynthesis of cholesterol, or both.

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Regulation of ileal transport Bile acid transport by the ileal enterocyte is also regulated by negative feedback [44]. Regulation is believed to be largely at the level of the sodium-dependent uptake transporter ASBT, which is downregulated by elevated bile acid concentrations in the enterocyte and upregulated by decreased concentrations. Regulation is considered to be modulated by activation of FXR, which in turn upregulates the synthesis of the negative regulatory protein SHP. SHP, in turn, displaces HNF1α from the promotor of the SLC10A2 gene (which encodes ASBT). There are, however, species differences in the response of ASBT to bile acid or bile acid sequestrant feeding. Moreover, bile duct ligation, which should upregulate ASBT, actually downregulates ASBT in rats [45]. Feeding of CDCA, a potent FXR agonist, causes relatively little increase in bile acid secretion in humans. In contrast, feeding of UDCA, which acts an FXR antagonist, increases bile acid secretion, suggesting upregulation of ileal transport [46]. Whether basolateral efflux (mediated by OSTα/OSTβ) is regulated is not known. Two groups working independently showed that enterally administered bile salts suppressed bile acid synthesis, but that intravenously administered conjugated bile acids did not [47, 48]. This puzzling observation suggested that enteral exposure to bile acids caused the release of a substance that was required for inhibition of bile acid synthesis. This factor was identified by the Kliewer group as FGF15(19), which was shown to be released via an FXR-mediated mechanism when bile acid concentrations were increased in the ileal enterocyte [43]. The released FGF then acted on a hepatocyte receptor, this in turn leading to a phosphorylation cascade that was required for the negative feedback inhibition of bile acid synthesis by bacteria. The importance of this peptide was shown by recent experiments from the Dawson group, in which basolateral transport via OSTα/OSTβ was genetically ablated in mice [49]. Because of the elevated bile acid concentration in the ileal enterocyte, FGF was released and no compensatory increase in bile acid synthesis occurred, despite animals having a decreased ileal transport of bile acids and bile acid malabsorption. The situation is to be contrasted with ileal resection, in which both bile acid return and FGF release are markedly decreased, followed by a marked increase in bile acid biosynthesis. In animals, administration of bile acid sequestrants upregulates bile acid uptake in more proximal enterocytes. Whether this occurs in humans is not known.

BILE ACID FUNCTIONS Bile acid functions are summarized in Table 20.2. The major function of bile acid is to sustain nutrition for the growing organism by solubilizing dietary lipids [9]. Bile acids also denature dietary proteins, enhancing their

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Table 20.2 Functions of bile acids in mammals

• Whole organism • Elimination of cholesterol • Liver • Hepatocyte • Insertion of canalicular bile acid and phospholipid transporters Induction of bile flow and biliary lipid secretion Promotion of mitosis during hepatic regeneration Regulation of gene expression by activation of FXR Endothelial cells Regulation of hepatic blood flow via activation of TGR5 Biliary tract • Lumen • Solubilization and transport of cholesterol and organic anions • Solubilization and transport of heavy metal cations • Cholangiocyte • Stimulation of bicarbonate secretion via CFTR and AE2 • Promotion of proliferation when obstruction to bile flow • Gall bladder • Modulation of cAMP-mediated secretion • Promotion of mucin secretion Small intestine • Lumen • Micellar solubilization of dietary lipids • Cofactor for bile salt dependent lipase • Antimicrobial effects • Ileal enterocyte • Regulation of gene expression via nuclear receptors • Ileal epithelium • Secretion of antimicrobial factors (FXR-mediated) Large intestine • Colonic epithelium and muscular layer • Promotion of defecation by increasing propulsive motility • Colonic enterocyte • Modulation of fluid and electrolyte absorption Brown adipose tissue • Promotion of thermogenesis via TGR5

• • • • •

•

•

•

•

susceptibility to tryptic digestion [50]. Bile acids also contribute to the relative sterility of the proximal small intestine by their direct and indirect antimicrobial effects [13]. The role of the liver is to provide those bile acids lost from the enterohepatic circulation. Bile acids may modulate the hepatic microcirculation by activating TGR5, a G-protein coupled receptor whose activation results in nitric oxide release [51]. In addition, the active secretion into the canaliculus of bile acids returning from the intestine generates a major fraction of bile flow in many species. In bile, bile acids form mixed micelles with phosphatidylcholine, and such mixed micelles solubilize cholesterol. They can also bind heavy metal cations

electrostatically and trap lipophilic drug metabolites. Bile acids may also modulate ductular bile flow by acting on apical G-protein coupled receptors. The process of biliary lipid secretion is very important in humans, in that about half of cholesterol elimination results from biliary secretion of cholesterol. However, the high proportion of cholesterol in bile (in relation to that of bile acids and phospholipid) is responsible for the high prevalence of cholesterol gallstone formation. Most vertebrate species have relatively little cholesterol in bile, and in these species cholesterol must be eliminated largely by conversion to bile acids [52]. Nonetheless, in mice, direct secretion of cholesterol into the lumen by the enterocyte may contribute to cholesterol elimination [53].

DISTURBANCES IN BILE ACID METABOLISM Disturbances in bile acid metabolism include defects in bile acid biosynthesis and conjugation, defects in membrane transport, defects in flow, and increased degradation of bile acids by the enteric flora. Each of the enzymes involved in bile acid biosynthesis and conjugation (amidation) may be absent at birth. Defects in A-ring modification may present as cholestatic liver disease or giant cell hepatitis (see Chapter 42). Defects in bile acid conjugation should result in lower bile acid concentrations in the proximal small intestine, because the unconjugated bile acids are absorbed passively. The result should be a syndrome of fat-soluble vitamin malabsorption that responds to conjugated bile acid administration [54]. Bile acid transport from the hepatocyte into the canaliculus may be inborn because of defective or absent BSEP [55]. Bile acid retention in the hepatocyte leads to apoptosis/necrosis and ultimately to liver failure. Drugs that interfere with BSEP transport function have now been identified; such drugs induce pruritus [56]. Bile acid retention in the hepatocyte leads to production of early growth response factor 1, a transcription factor that leads to leukocyte recruitment to the liver [57]. Bile acid regurgitation from the hepatocyte should lead to stellate cells being exposed to higher concentrations of bile acids. Whether bile acids play a role in stellate cell activation and subsequent fibrosis is unclear at present. Obstruction of bile flow, whether at a ductular or duct level, also leads to bile acid retention and induction of apoptosis/necrosis of hepatocytes. As cholestasis increases, there is sulfation of CDCA amidates and urinary excretion of the CDCA-amidated sulfates as well as conjugates of cholic acid. With complete biliary obstruction, DCA disappears from plasma bile acids, as no bile acids are exposed to the enteric flora. Bile acid synthesis decreases markedly with urinary excretion, balancing hepatic synthesis.

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Elevated plasma bile acid levels have long been thought to induce pruritus. Indeed, bile acids fulfill all of the criteria for cause and effect, in that bile acid removal ameliorates itching and bile acid administration induces itching. Nonetheless, in cholestatic liver disease there is an imperfect correlation between plasma bile acid levels and pruritus, in that plasma levels may be low and patients itch, and plasma levels may be high and patients not itch. These observations indicate that additional factors are involved in the etiology of pruritus. Removal of bile acids (and possibly other molecules) by dialysis against albumin is associated with a striking diminution in pruritus, which may be long-lasting [58]. Pruritus is also improved by aspirating the bile acid pool endoscopically [59], by partial biliary diversion [60], or by ileal bypass [61]. All of these procedures reduce the load of bile acids to the liver. Potent inhibitors of ileal bile acid transport have been developed [62] but these have not been tested clinically. There is some evidence for upregulation of ileal transport in cholestatic liver disease [63], which should make such agents highly useful. Cholestatic liver disease leads to a deficiency of bile acids in the small intestine. This in turn results in less efficient absorption of saturated fatty acids and lack of absorption of fat-soluble vitamins. In cirrhosis, there is decreased bile acid secretion and bacterial overgrowth in the small intestine. The bacterial overgrowth leads to increased bacterial deconjugation and passive absorption of unconjugated bile acids, causing a further deficiency of bile acids and possibly further bacterial overgrowth—a vicious cycle. In rats with induced cirrhosis, administration of conjugated bile acids corrects bacterial overgrowth and bacterial translocation to lymph nodes [64]. However, the clinical utility of this approach has not been tested. Ileal dysfunction causes bile acid malabsorption and a compensatory increase in bile acid biosynthesis. In this new steady state, an increased flux of bile acids pours into the colon, inducing secretion, clinically manifest as diarrhea. Patients respond symptomatically to bile acid sequestrant administration [65]. As yet defects in bile acid uptake by the hepatocyte have been identified only as a phenotype [66]. It could be that there is such a redundancy of uptake transporters that loss of any single transporter has little effect on bile acid uptake. When hepatic uptake of bile acids does not occur, the enterohepatic circulation is not complete. The only bile acids reaching the small intestine are those newly synthesized; the synthetic rate should be greatly increased because of loss of negative-feedback regulation. Bile acids returning from the intestine are diverted to the urine. However, there should be no liver damage. The syndrome should present with pruritus, elevated plasma bile acid concentrations, and normal liver tests.

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THERAPY WITH BILE ACID AGONISTS AND ANTAGONISTS Bile acid agonists: therapy with bile acids and bile acid congeners Therapy with natural bile acids arose in the 1970s when it was discovered that oral administration of CDCA would lower the cholesterol saturation of bile and induce the gradual dissolution of cholesterol gallstones [67]. CDCA was mildly hepatotoxic and induced dose-dependent diarrhea. When UDCA was shown to have similar efficacy but to be devoid of side effects, it replaced CDCA for medical dissolution of gallstones [68]. Nonetheless, medical dissolution of gallstones was imperfect therapy for three reasons. First, some stones did not dissolve because either they were not composed of cholesterol or they had a non-cholesterol surface layer. Second, complete dissolution took many months, as the rate of dissolution averaged only 1 mm per month decrease in stone diameter. Third, recurrence of stones (albeit without symptoms) occurred in about half of the patients within a few years. Laparoscopic cholecystectomy was introduced and soon became the standard of practice as it was safe, effective for all kinds of stones, and curative. Natural bile acids (cholic acid, UDCA, CDCA) have also been used as replacement therapy for inborn errors of bile acid biosynthesis, as discussed in Chapter 42. UDCA had a renascence when it was shown to improve liver tests in cholestatic liver disease. In primary biliary cirrhosis (PBC), UDCA delays the time to liver transplantation [69]. In primary sclerosing cholangitis (PSC), high doses of UDCA may lower disease progression, but efficacy remains uncertain [70]. UDCA was also shown in placebo-controlled trials to be effective in cholestasis of pregnancy [71]. UDCA may improve the clinical course of cystic fibrosis [72] and accelerate natural recovery from drug-induced cholestasis. The mechanism of action of UDCA is complex. It involves multiple effects such as induction of bile flow—because it is a naturally choleretic bile acid—and induction of bile acid synthesis and canalicular transporters [73]. UDCA also has antiapoptotic and anti-inflammatory effects [74]. Activation of FXR downregulates NTCP and upregulates BSEP; together these changes should decrease the concentration of bile acids in the hepatocyte, a goal of therapy for cholestatic liver disease. A potent FXR agonist, 6-ethylCDCA was synthesized by Pellicciari and his colleagues [75]; its efficacy is being explored in PBC. Potential uses of a potent FXR agonist in liver diseases [76] and metabolic diseases [77–79] have been proposed. Other FXR agonists have been reported [80], but no information is available on their therapeutic properties. NorUDCA is a C23 homolog of UDCA that differs from UDCA in having one less carbon atom in the side chain. During transport through the hepatocyte, norUDCA is not

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amidated, but is partly glucuronidated. Another fraction is excreted as such into bile and undergoes cholehepatic shunting, exposing cholangiocytes to multiple passes of the compound [81, 82]. NorUDCA has been shown to be much more effective than UDCA in decreasing periductular inflammation in the mdr2 knockout mouse [83]. Such mice lack phospholipid in bile and the simple bile acid micelles attack cholangiocytes, generating a histological lesion resembling that present in patients with PSC. NorUDCA may undergo clinical testing in PSC in the future, despite there being no abnormality in biliary phospholipid secretion in PSC. Patients with defective MDR3 are now being identified because of their phenotype of recurrent cholestasis or choledocholithiasis [84]; such patients may respond to norUDCA. Bile acid derivatives that activate TGR5 selectively are being synthesized, and may possibly have a role in the metabolic syndrome [85]. Cholylsarcosine is a synthetic conjugated bile acid analog in which cholic acid is coupled via an amide linkage to sarcosine (N-methylglycine). The compound is resistant to bacterial deconjugation and 7-dehydroxylation [86]. Cholylsarcosine has been used to restore the micellar bile acid concentration in patients with short bowel syndrome and thereby increase triglyceride absorption [87]. As noted above, it abolishes bacterial overgrowth, bacterial translocation to lymph nodes, and endotoxemia in rats with carbon tetrachloride-induced cirrhosis [64]. However, the compound has no patent protection and clinical trials in cirrhosis are unlikely. The compound should also be useful in inborn errors of bile acid conjugation, although taurine- or glycine-conjugated bile acids might be equally efficacious.

binding isotherm when compared to cholestyramine or colestipol, and this permits a lower dose in tablet form. All of these agents decrease pruritus in cholestatic liver disease, provided bile acid secretion into the intestine is sufficient. Colesevelam has been shown to increase insulin sensitivity in type II diabetes [88], possibly by promoting the release of glucagon-like peptide from the distal small intestine; it has recently been approved for this indication.

CONCLUSION The details of bile acid biosynthesis and transport are gradually being elucidated. Most of the possible inborn errors of bile acid biosynthesis have been identified, and therapy with primary bile acid replacement has been shown to be efficacious. UDCA has been shown to be safe and of some efficacy in PBC. However, PBC is an autoimmune disease and therapy with UDCA is palliative rather than curative. NorUDCA targets biliary ductules and its utility in PSC and MDR3 deficiency will be tested in the next decade. Bile acids activate the nuclear receptor FXR and the G-coupled protein TGR5. Time will also tell whether these are useful therapeutic targets for the potent agonists that are being developed by bile acid chemists.

ACKNOWLEDGMENTS The author thanks Claudio Schteingart, PhD for preparation of Figures 20.2 and 20.3.

REFERENCES Bile acid transport inhibitors Bile acid sequestrants were shown to be useful adjuncts to the statins for lowering plasma LDL cholesterol levels. The mechanism of action was induction of bile acid malabsorption, a compensatory increase in bile acid biosynthesis, and upregulation of LDL receptors on the hepatocyte. Potent, non-absorbable inhibitors of the ileal ASBT were developed [62] but have not yet been introduced into clinical practice. In principle, such agents may decrease cholestatic pruritus.

Bile acid sequestrants Three polymeric bile acid sequestrants—cholestyramine, colestipol, and colesevelam—are available in the United States. All bind bile acids and remove them from solution. With steady-state administration, bile acid synthesis (and loss) increases by a factor of 4. Colesevelam has a superior

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Hepatocyte Basolateral Membrane Organic Anion Transporters Jo H. Choi, John W. Murray and Allan W. Wolkoff Division of Hepatology, Marion Bessin Liver Research Center, and Department of Anatomy and Structural Biology, Albert Einstein College of Medicine, Bronx, NY, USA

excretion of a number of typical organic anions, including bile acids, for which the liver plays an essential role.

INTRODUCTION The hepatocyte removes a large number of organic anionic compounds from the blood, including various endogenous compounds such as bilirubin and bile acids, as well as xenobiotics. Binding to albumin enables these compounds to circulate in blood despite limited aqueous solubility, and in general they are cleared rapidly from the circulation by the hepatocyte despite being almost entirely protein-bound [1]. Although there were suggestions that the albumin–organic anion complex was taken up by the hepatocyte, a number of studies indicate that the organic anion is extracted from its protein carrier during the uptake process [2, 3]. The liver is ideally designed for extraction of protein-bound molecules. Unlike most other organs, which have a tight capillary endothelium, the hepatic capillary (sinusoidal) endothelium is fenestrated. This permits direct interaction of large molecular complexes with the hepatocyte plasma membrane [4, 5]. Although a particular organic anionic molecule may have lipophilic properties, its tight binding to albumin limits the extent of cellular uptake by simple dissolution across lipid bilayers. Although this process can occur [6], uptake of these compounds has been found to be facilitated by several hepatocyte plasma membrane transport proteins. This chapter will consider mechanisms for uptake and The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

HEPATOCYTE UPTAKE OF BILE ACIDS Sodium-dependent bile acid uptake across the hepatocyte basolateral membrane Uptake of conjugated bile acids from sinusoidal blood occurs against unfavorable electrical and chemical gradients and is mediated largely by a saturable, sodium-dependent symport system [7–9]. The marked dependency of bile acid uptake on extracellular sodium has been demonstrated in multiple experimental models, including the isolated, perfused rat liver, isolated and cultured hepatocytes, and basolateral plasma membrane vesicles [9–11]. However, there is a substantial sodium-independent fraction of bile acid uptake, representing approximately 20–25% of total uptake. The avid affinity of the hepatocyte for bile acids results in a high first-pass clearance from portal blood of as much as 90% [12–14]. The driving force for this largely sodium-dependent process is

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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the extracellular-to-intracellular sodium gradient, which is maintained by the action of the basolateral sodium pump, Na+ -K+ -ATPase [15]. In several experimental models, sodium-dependent taurocholate transport had an apparent Km of 6–56 µM, depending on the species and experimental system [16]. The efficiency of bile acid transport is dependent upon the position and number of hydroxyl groups on the steroid nucleus as well as the length and charge of the bile acid side chain [17–20]. 6-hydroxylated bile acid derivatives are preferred substrates for the hepatic sodium-dependent bile acid transporter [21]. In contrast, the 3α-hydroxy group present in all natural bile acids is not essential for high-affinity interaction with this transporter [22]. Furthermore, Na+ gradient-dependent sinusoidal uptake of taurocholate can be stimulated by low concentrations of albumin [23]. The process is electrogenic based upon electrophysiological measurements of transport-associated currents, as well as direct measurement of fluorescent bile acid transport rates in voltage-clamped cells [24, 25]. These studies support a Na+ : bile acid coupling stoichiometry of 2 : 1.

Na+ Taurocholate Cotransporting Polypeptide (Ntcp) A sodium-dependent bile acid transporter, now called the Na+ /taurocholate cotransporting polypeptide (ntcp), was isolated by expression cloning in Xenopus laevis oocytes by Hagenbuch and associates [26]. Ntcp is part of a new family of mammalian sodium-dependent bile acid transporters that also includes the brush border membrane transporter expressed in the ileum, renal proximal tubule cell, and cholangiocyte, with which it shares approximately 35% amino acid sequence identity [27]. The cDNA sequence of ntcp has an open reading frame of 1086 nucleotides encoding a protein of 362 amino acids (calculated molecular mass 39 kDa) with five potential N-linked glycosylation sites [19, 28]. Although initial studies were interpreted as consistent with an ntcp model consisting of nine transmembrane domains [30], more recent studies suggest that it has seven transmembrane domains [31], similar to what was suggested by initial computer-assisted models [26]. Northern blot analysis with the cloned rat ntcp probe revealed cross-reactivity with mRNA species from rat kidney and intestine as well as from liver tissues of mouse, guinea pig, rabbit, and man. A 1599 bp cDNA encoding the human NTCP was subsequently cloned [32]. This cDNA encodes a protein of 349 amino acids (calculated molecular mass of 38 kDa) and exhibits 77% amino acid homology with the rat ntcp [32]. In vitro translation experiments indicate that the transporter is glycosylated and that its polypeptide backbone has an apparent molecular mass of 33–35 kDa. Kinetic studies indicated that the human NTCP has a higher affinity for taurocholate (apparent Km = 6 µM) than the rat transporter (apparent

Km = 25 µM), although other studies reported a Km of 10 µM for rat ntcp-mediated transport of taurocholic acid [19]. Southern blot analysis of genomic DNA and immunofluorescence in situ hybridization have mapped the human NTCP gene to chromosome 14 and the rat ntcp gene to chromosome 2 [32, 33]. Ntcp2, encoding a truncated 317 amino acid protein, has been cloned from mouse liver [33]. It has a shorter C-terminal end and also mediates saturable Na+ -dependent transport of taurocholate when expressed in Xenopus laevis oocytes. Analysis of the gene revealed that ntcp2 is produced by alternative splicing where the last intron is retained. Its functional importance and cellular localization are unknown. Although a physiological role for ntcp in bile acid transport has been suggested [15], there are no studies that demonstrate this. As of this date, no ntcp knockout (KO) or knockdown models have been reported, and the major evidence supporting this suggestion is limited to finding similar Km s for bile acid transport in ntcp-transfected cells as compared to rat hepatocytes, and parallel decreases of ntcp expression and Na+ -dependent taurocholate transport in liver regeneration and in hepatocytes in culture [15]. It must be recognized that these physiological perturbations could result in altered function of other potential bile acid transporters, and it cannot be concluded that there is a direct cause and effect relationship between ntcp expression and bile acid transport in these models. An early study in which Xenopus laevis oocytes were microinjected with rat liver cRNA in the presence of ntcp antisense RNA found a 95% reduction in taurocholate uptake [15, 34]. However, now that we have more complete genomic databases, it is clear that the antisense oligonucleotide that was used to inhibit ntcp translation (TAACCCATCAGAAAGCCAGA) was not specific for ntcp and could potentially inhibit synthesis of other rat liver as well as Xenopus proteins [35]. There have been no inheritable disorders described in which expression of ntcp is affected, although disorders in which other members of the Na+ -dependent solute transport family are mutated have been described [36–44]. Several genetic polymorphisms of ntcp that affect its expression in vitro have been described [45], but there is no information regarding a clinical phenotype of those individuals who harbor any of these mutations. The mouse model of erythropoietic protoporphyria, in which the ferrochelatase gene was knocked out, has elevated plasma bile acid levels and absent plasma membrane expression of ntcp and organic anion transporting polypeptide 1 (oatp1/oatp1a1) [46]. These mice have massive bile duct proliferation and biliary fibrosis. However, plasma disappearance of a tracer dose of 3 H-taurocholate was near normal, with over 90% found in liver and bile of these mice within 30 minutes of injection [46]. This indicates that bile acids can still be cleared rapidly from serum even in the absence of ntcp. Another example in which ntcp expression was dissociated from bile acid

21: HEPATOCYTE BASOLATERAL MEMBRANE ORGANIC ANION TRANSPORTERS

uptake was described in rats seven days after bile duct ligation. Ntcp expression was virtually undetectable in livers from these animals, while hepatocytes isolated from these rats maintained initial Na+ -dependent uptake of 3 H-taurocholate at 30% of control levels [47].

Phylogeny and ontogeny of Ntcp mRNA encoding ntcp is detected in mammalian species, including rat, mouse, and man, but has not been found in livers from non-mammalian species, including chicken, turtle, frog, and small skate [48]. When expression of ntcp was studied in developing rat liver, mRNA was detected between 18 and 21 days of gestation, at the time when Na+ -dependent bile acid transport activity is first detected [49]. Interestingly, when mRNA from the small skate was injected into Xenopus oocytes, only sodium-independent, chloride-dependent bile acid uptake was expressed, consistent with prior functional studies of bile acid transport in this species. Of note is the fact that various mammalian hepatoma cell lines such as HuH7 and HepG2 do not express ntcp mRNA. These findings indicate that ntcp expression is a property of the well-differentiated mammalian hepatocyte. The mechanisms underlying ontogenic regulation of ntcp have been studied in detail [28]. Steady-state mRNA levels for the basolateral transporter were less than 20% of adult values prior to birth, increased to 35% on the first postnatal day, and reached adult levels by one week of age. This was paralleled by transcription rates that were low prior to birth, reached 47% of adult levels by day 1, and were maximal by one week of age. Surprisingly, the full complement of the ntcp protein was present well before adult levels of mRNA were reached. The basolateral protein was expressed at 82% of adult levels on the first day of life but was of lower apparent molecular mass (39 kDa), a difference that persisted until four weeks of age. N-glycanase digestion suggested that this difference could be fully accounted for by differences in extent of N-linked glycosylation. These results indicate that ontogeny of ntcp is complex and it appears to be regulated at transcriptional, translational, and post-translational levels.

Ntcp mediates uptake of conjugated bile acids and other organic anions It is now clear that ntcp is a multispecific organic anion transporter [16, 19]. In inhibition studies performed in primary cultured rat hepatocytes and ntcp-transfected COS7 cells, Na+ -dependent uptake of taurocholate was inhibited by nine bile acids and five non-bile acid organic anions in a concentration-dependent manner [50]. These inhibitors included BQ123 and indomethacin, neither of

307

which is transported by ntcp. In ntcp-transfected cells, Na+ -dependent uptake of taurocholate was strongly inhibited by bile salts as well as by estrone 3-sulfate, bumetanide, cyclosporin A, sulfobromophthalein, and oligomycin [51, 52]. Another study [19] examined transport of a series of C23, C24, and C27 bile acid derivatives in HeLa cells stably transfected with rat ntcp or oatp1a1, in which expression was under regulation of a zinc-inducible promoter. Similar uptake patterns were observed for both ntcp and oatp1a1, except that unconjugated hyodeoxycholate was a substrate of oatp1 but not ntcp. Conjugated bile acids were transported better than unconjugated bile acids, and the configuration of the hydroxyl groups (alpha or beta) had little influence on uptake. Although cholic and 23 norcholic acids were transported by ntcp and oatp1a1, other unconjugated bile acids (chenodeoxycholic, ursodeoxycholic) were not. In contrast to ntcp, oatp1a1-mediated uptake of the trihydroxy bile acids taurocholate and glycocholate was fourto eightfold below that of the corresponding dihydroxy conjugates. Additionally, ntcp mediated high-affinity, sodium-dependent transport of sulfobromophthalein (BSP) with a Km similar to that of oatp1a1-mediated transport of BSP (Km = 3.7 vs. 3.3 µM, respectively). Uptake of BSP and taurocholic acid showed mutually competitive inhibition. Taken together, all of these studies suggest that the substrate specificity of ntcp is broad, and its role in hepatocyte transport physiology remains to be elucidated fully.

Regulation of Ntcp expression by specific transcription factors The rat genomic DNA encoding ntcp has been cloned, and several cis elements and transactivating factors involved in expression of the gene have been identified [52, 53]. Transcriptional regulation of ntcp gene expression has been studied by a number of investigators [47, 53, 54]. Important roles in ntcp gene expression have been described for HNF1, HNF4, and the RXR/RAR heterodimer complex [52–54]. These transcription factors are not specific for modulation of ntcp expression, and regulate expression of multiple proteins in hepatocytes, including organic anion transporters [55]. Regulation of expression by transcription factors is a relatively slow process. Recent studies, described below, have elucidated novel mechanisms for the rapid regulation of transporter activity and subcellular distribution.

Regulation of Ntcp transport activity by altering its subcellular distribution Vmax for Na+ -dependent uptake of taurocholic acid increases substantially within minutes of incubation of

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hepatocytes in 3′ ,5′ -cyclic adenosine monophosphate (cAMP) [56, 58]. This involves redistribution of transporter from an intracellular pool to the plasma membrane, without synthesis of new transporter [59, 60]. Immunoblot analysis with phosphoamino acid-specific antibodies indicates that ntcp is a serine/threonine phosphoprotein, and that cAMP incubation results in reduced phosphorylation of the transporter through stimulation of phosphatase activity [61, 62]. It appears that this cAMP-mediated dephosphorylation of ntcp results in its increased retention in the plasma membrane. The mechanism underlying this phenomenon is not as yet clear. A study in which the human hepatoma cell line, HepG2, was transiently transfected with green fluorescent protein (GFP)-tagged ntcp showed that exposure to cAMP resulted within 2.5 minutes in a 40% increase in GFP fluorescence at the plasma membrane [63]. This recruitment of intracellular ntcp-GFP to the cell surface was prevented by cytochalasin D, a disruptor of the actin-based microfilament cytoskeleton. In contrast, perturbation of microtubules following incubation of cells in nocodazole prevented even baseline ntcp-GFP from reaching the plasma membrane. Interestingly, cells regained plasma membrane fluorescence within two hours after nocodazole removal. Based on these findings, it was suggested that targeting of ntcp to the plasma membrane is a two-step process in which the first step involves delivery of ntcp to a subplasma membrane location via microtubules, following which ntcp transfers to microfilaments that carry it to the plasma membrane. To examine the mechanism by which microtubules are required for trafficking of ntcp, an in vitro motility system was used. In preceding studies, this fluorescence microscopy system enabled study of mechanisms that mediate and regulate microtubule-based motility of endocytic vesicles prepared from rat liver [64–66]. Ntcp-containing vesicles are also present in this preparation, as shown by immunoblot and immunofluorescence microscopy [60]. To determine whether these ntcp-containing vesicles could bind to and move along microtubules, a microassay chamber was used in which the glass surface was coated with fluorescent microtubules [67]. Vesicles were flowed into this chamber and antibody against ntcp followed by fluorescent secondary antibody were added [60]. As seen in Figure 21.1, fluorescent antibody marked ntcp-containing vesicles that were attached to microtubules in the chamber. Following addition of 50 µM ATP, these ntcp-containing vesicles moved along the microtubules to which they were attached. Immunofluorescence analysis using specific antibodies revealed that ntcp-containing vesicles are associated with the minus-end-directed microtubule motor dynein, and with the plus-end-directed microtubule motors kinesin-1 and kinesin-2 (Figure 21.2a). Interestingly, antibodies to dynein and kinesin-1 inhibited motility of ntcp-containing vesicles, while antibody to kinesin-2 had no effect on

Figure 21.1 Characterization of bidirectional motility of ntcp-containing vesicles on microtubules in vitro. Polarity-marked fluorescent microtubules were attached to glass chambers, incubated with an endocytic vesicle preparation, and washed. Ntcp-containing vesicles were then visualized with primary and fluorescent secondary antibodies. 50 µM ATP was added to initiate motility. Time-lapse digital fluorescence images were captured in Cy2 (to detect ntcp) and Cy3 (to detect microtubules) channels. Seconds after addition of ATP are indicated at the upper left. The arrows follow two motile ntcp-containing vesicles (bright dots) moving in opposite directions. Arrowheads show the original location of the vesicles. “+” and “–” indicate the plus and minus ends of the microtubules. Bar = 5 µm Reprinted from [60] with permission

motility (Figure 21.2b). These results suggest that unlike the other two motors, kinesin-2 either requires activation or may not play a role in motility of these vesicles. This will require further study. It should be noted that this kinesin-2 antibody inhibits motility of late endocytic vesicles [66]. Previous studies in cells and animals suggested that trafficking of ntcp is regulated at least in part by the PI3 kinase/PKCζ pathway [68–70]. A simplified schematic of the PI3 kinase/PKCζ pathway and its relationship to vesicle motility is shown in Figure 21.3a. Addition of LY294002, an inhibitor of PI3 kinase, resulted in reduced vesicle motility, presumably from reduced formation of PIP3 from vesicle-associated PIP2 (Figure 21.3b). Addition of PIP3, an activator of PKCζ, enhanced vesicle motility (Figure 21.3c). Addition of PIP3 with LY294002 overcame its inhibition, while addition of PKCζ pseudosubstrate, a potent inhibitor of PKCζ, in the presence or absence of PIP3, virtually eliminated vesicle motility (Figure 21.3c). These data show that PKCζ is an important regulator of the motility of ntcp-containing vesicles [60].

21: HEPATOCYTE BASOLATERAL MEMBRANE ORGANIC ANION TRANSPORTERS

309

% of ntcp-containing vesicles

100

80 (1840)

(855)

60 (1190) 40

20 (351) 0

Dynein

Kinesin-2

Kinesin-1

mouse lgG

(a)

Motile vesicles (% of total)

50

Total motility minus-end directed plus-end directed

(415) (101)

40

(166) * (252)

30

* 20

** (316) **

10 ** 0

Buffer

Mouse lgG

Dynein

** Kinesin-1 Kinesin-2

(b)

Figure 21.2 Motility of ntcp-containing vesicles is mediated by cytoplasmic dynein and kinesin-1. (a) Endocytic vesicles were bound to microtubules. Ntcp-containing vesicles were then visualized with primary and fluorescent secondary antibodies as in Figure 21.1. They were further labeled with antibodies against dynein, kinesin-1, kinesin-2, or normal mouse IgG, followed by fluorescent-labeled secondary antibody. The percentage of ntcp-containing vesicles that co-localize with each motor protein is shown. Parentheses indicate the number of vesicles counted. (b) Motility of ntcp-containing vesicles toward the plus and minus ends of microtubules was scored following incubation with motor protein antibodies or non-immune mouse IgG. The black bars denote overall motility; the light gray and dark gray bars denote the proportion of minus-end- and plus-end-directed vesicles, respectively. *p < 0.01, **p < 0.0001 compared to the corresponding mouse IgG control. Reprinted from [60] with permission

Interestingly, the pseudosubstrate had no effect on motility of late endocytic vesicles containing asialoorosomucoid (ASOR), indicating that this regulatory pathway has specificity for ntcp-containing vesicles [60]. The requirement of PKCζ activity for motility of ntcp-containing vesicles was also examined in HuH7 cells that had been transiently transfected with rat ntcp-GFP [60]. This recombinant protein has been found to be functional for bile acid uptake and translocates

to the plasma membrane in response to cAMP [63]. Transfection with GFP alone yielded a diffuse pattern of cellular fluorescence (Figure 21.4a, left panel) [60]. In contrast, transfection with ntcp-GFP resulted in localization of fluorescence to the plasma membrane, as well as intracellular vesicles (Figure 21.4a, right panel) [60]. Motility of the intracellular vesicles containing ntcp-GFP was analyzed using time-lapse epifluorescence microscopy in the presence and absence of a cell-permeant,

310

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PIP2

activates

PIP3

LY294002

motile vesicles

PKCz

PKCzPS (a)

Motility of ntcp-containing vesicles (% of total MT-bound)

35

(289)

30 25

(234)

(251)

(294)

20 15 10

Regulation of Ntcp in cholestasis

5 0 0

50

100

150

200

[LY294002] (mm) (b)

45

Motile ntcp-containing vesicles (% of total)

specificity of this inhibition, motility of late endosomes in HuH7 cells was measured in the presence and absence of the inhibitor. As seen in Figure 21.4b, motility of ASOR-containing late endocytic vesicles was essentially unaffected by Myr-PKCζPS. This was identical to the result in vitro, where purified late endocytic vesicles were also resistant to the effects of the PKCζ inhibitor. Thus, these studies show that PKCζ is specifically required for the motility of ntcp-containing vesicles but not late endocytic vesicles both in vitro and in living cells [60].

** (317)

* (223)

40 35 30 25

(570) * (234)

20 15 10

** (221)

5 0

2 3 02 3+ fer 00 PIP 40 3 IP zPS Buf 94 9 P M 2 2 m LY LY PIP mM PKC 10 mM mM mm 10 mm 50 50 10 50 +

(c)

Figure 21.3 Involvement of the PI3K–PKCζ pathway in ntcp-containing vesicle motility. (a) A simplified PI3K–PKCζ pathway for regulation of ntcp-containing vesicle motility. PI3K converts PIP2 to PIP3, which then activates PKCζ, leading to vesicle motility through unknown substrates. LY294002 and PKCζPS are inhibitors of PI3K and PKCζ, respectively. (b) Motility of ntcp-containing vesicles on microtubules was scored following incubation with buffer, 50, 100, or 200 µM LY294002. (c) Motility of ntcp-containing vesicles was scored following incubation with buffer, 10 µM PIP3, 50 µM LY294002 with and without 10 mM PIP3, or PIP3 along with 50 mM PKCζPS. *p < 0.005, **p < 0.0001 compared to buffer control. Parentheses indicate the number of vesicles scored. Reprinted from [60] with permission

myristoylated version of PKCζPS (Myr-PKCζPS). These studies showed a dose-dependent inhibition of ntcp-GFP containing vesicle motility, with a nearly 80% reduction in the number of moving vesicles in the presence of 50 µM Myr-PKCζPS (Figure 21.4b). To determine the

Hepatocyte bile acid uptake is reduced in cholestasis [71, 72]. However, there is some lack of correlation with ntcp expression. It is clear from a number of studies that following common bile duct ligation in the rat, ntcp protein and mRNA expression are profoundly reduced [47, 73–75]. However, several studies suggest that bile acid uptake is relatively well maintained following common bile duct ligation. In one study, rats were subjected to five days of common bile duct obstruction, following which obstruction was removed [76]. Serum bile acids, which had been highly elevated, returned to normal within 60–90 minutes of relief of obstruction. This was accompanied by prompt excretion of bile acids into bile. Plasma clearance of a tracer dose of taurocholate injected at the time of relief of biliary obstruction was near normal. This normalization of transport function is faster than return of ntcp levels to normal would be expected to occur. In another study [47], hepatocytes were isolated from rats in which the common bile duct had been ligated for seven days. Ntcp protein levels were reduced by 90%, while the Vmax for Na+ -dependent taurocholate uptake was reduced by only 70%. When basolateral plasma membrane vesicles were prepared from livers of rats in which the common bile duct was ligated for 50 hours, Na+ -dependent taurocholate uptake remained normal [77]. Transport studies were also performed in isolated perfused livers from rats 24 and 72 hours following common bile duct ligation [78]. Quantitation of bile acid uptake following addition of 3 H-taurodeoxycholate to the perfusate (16 nmole min−1 g−1 ) showed little if any difference in uptake between control and bile duct-ligated livers. Perfusion with a larger dose of this bile acid (4000 nmole min−1 g−1 ) resulted in an approximately 35% reduction in maximal uptake by livers that had undergone common bile duct obstruction for either 24 or 72 hours. In the aggregate, these studies show at most a modest reduction of hepatocyte bile acid uptake following mechanical cholestasis in the face of a profound reduction in ntcp protein and mRNA. These data suggest that there may be other Na+ -dependent bile acid transporters in addition to ntcp [47].

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Vector

311

ntcp-GFP (a)

70

(161)

(171)

60 (224)

% motile vesicles

50

ntcp-GFP vesicles Late ASOR vesicles

40 (323)

30 20

(241) 10 (174)

0 0

25

50

75

100

[Myr-PKCzPS] (mm) (b)

Figure 21.4 PKCζ is required for the motility of ntcp-containing vesicles but not for late endosomes. (a) HuH7 cells transfected with ntcp-GFP and GFP vector control 24 hours post-transfection. The plane of focus of the epifluorescent microscope was selected to highlight the vesicular nature of ntcp-GFP. (b) Motility of ntcp-GFP vesicles (black symbols) and late endosomes (gray symbols) in these living ntcp-GFP-transfected HuH7 cells following incubation with 0–100 µM Myr-PKCzPS. Parentheses indicate the number of vesicles scored. Reprinted from [60] with permission

Ntcp expression is downregulated during hepatic regeneration Na+ -dependent uptake of taurocholic acid is markedly reduced 24 hours following two-thirds partial hepatectomy in the rat [79]. Serum bile acid levels are substantially increased for at least one week as compared to sham-operated controls [80, 81]. Expression of ntcp protein is reduced by over 90% at 24 hours of partial hepatectomy [79, 81] and normalizes by approximately one week [80]. In contrast, protein and mRNA expression of the canalicular ATP-dependent bile salt export pump (bsep) is essentially unchanged during liver regeneration [80]. It has been speculated that preservation of canalicular bile salt excretion by bsep in the face of reduced

uptake may protect hepatocytes during the regenerative process from potentially toxic bile salts.

Microsomal epoxide hydrolase Levy and colleagues suggested that microsomal epoxide hydrolase (mEH) may also mediate basolateral sodium-dependent bile acid transport [82]. They showed functional expression of sodium-dependent bile acid transport in the relatively well-differentiated MDCK dog kidney cell line that was transfected with mEH cDNA. They also provided evidence that mEH is inserted into the endoplasmic reticulum membrane with two topological orientations, one of which is targeted to the plasma

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membrane [81, 83]. Other investigators failed to show uptake of taurocholate following expression of mEH in Xenopus laevis oocytes and in a stably transfected fibroblast cell line [84]. They also showed that mEH is expressed in hepatoma tumor cell lines that show no bile acid transport activity [84]. However, evidence was presented indicating that in these situations, mEH is intracellular and not targeted to the plasma membrane [83]. The quantitative importance of mEH-mediated bile acid transport in hepatocytes remains to be determined.

HEPATOCYTE UPTAKE OF NON-BILE ACID ORGANIC ANIONS Sodium-independent organic anion transport Similar to hepatic uptake of bile acids, uptake of organic anions such as bilirubin and BSP is rapid and has carrier-mediated kinetics [1]. Studies performed in overnight cultured rat hepatocytes revealed saturable uptake of BSP that was inhibited by bilirubin [85, 86]. Ligand was taken up by cells, while albumin remained extracellular. Of interest was the finding that isosmotic substitution of NaCl in medium by sucrose resulted in an over 80% reduction of BSP uptake [85]. This was not due to a requirement for extracellular Na+ , as substitution of Na+ by K+ , Li+ , or choline had no effect. However, replacement of Cl− by HCO− 3 or gluconate reduced BSP uptake by approximately 40% [85]. Similar results were found for bilirubin uptake by cultured rat hepatocytes as well as by isolated perfused rat liver. The basis for this chloride-dependency is not clear. Studies performed with 36 Cl revealed that BSP uptake requires the presence of external Cl− and is not stimulated by unidirectional Cl− gradients, suggesting that BSP transport is not coupled to Cl− transport [86].

protein [90, 91]. However, further study showed that this protein was identical to the ß-subunit of the inner mitochondrial membrane protein, F1 -ATPase [91]. Although antigenic reactivity was present in both plasma membrane and mitochondrial domains, as seen by immunocytochemical electron microscopy as well as by immunoprecipitation of surface-iodinated hepatocytes [91], a role for this protein in transport has not been shown. Tiribelli, Sottocasa, and colleagues isolated a 170 kDa protein termed bilitranslocase [92–94]. This protein is composed of 37 and 35.5 kDa subunits. They reported that liposomes into which bilitranslocase was inserted were able to transport BSP [92]. They also reported that antibody to this protein inhibited transport of BSP into liver cell plasma membrane vesicles [92]. Bilitranslocase (accession number Y12178) was cloned from a λgt11 rat liver library utilizing a monoclonal antibody [93]. The reported derived protein sequence is unique and has been used in studies of potential bilirubin binding sites [94]. However, BLAST analysis reveals that the nucleotide sequence from which the protein sequence was derived is 93% identical to the inverse strand sequence of rat ceruloplasmin. This suggests that a cloning artifact likely occurred. Later studies used the transport assay developed in cultured rat hepatocytes as the basis for an expression cloning strategy in Xenopus laevis oocytes [95]. Injection of oocytes with total rat liver poly (A)+ RNA resulted in the functional expression of chloride-dependent BSP extraction from albumin [95]. Using a subselection cloning protocol, a single cDNA was isolated [96]. The encoded protein was named oatp1, now known as oatp1a1. This protein mediates uptake of BSP and also mediates transport of various bile salts such as taurocholate in an Na+ -independent fashion [97]. Since the initial description of oatp1a1, over 20 additional members of the oatp family have been described [98, 99]. Amino acid sequences of these proteins have a high degree of homology [98]. Table 21.1 lists the members of the oatp family that are expressed in hepatocytes from rats, mice, and humans. As indicated in the table, these hepatocyte oatps are distributed on the basolateral (sinusoidal) plasma membrane. They are of similar size and have similar predicted membrane topologies and biochemical characteristics.

Identification of sodiumindependent organic anion transporters

Function of Hepatocyte Oatps

The studies described above supported the existence of hepatocyte surface membrane organic anion transporter(s). Prior to development of strategies for expression cloning, a number of candidate transporters were proposed on the basis of studies that included affinity labeling and purification by affinity chromatography. Berk and colleagues isolated a 55 kDa protein termed BSP/bilirubin binding protein [87–89]. Wolkoff and colleagues isolated a different 55 kDa protein termed organic anion binding

While a great deal has been published regarding compounds that members of the oatp family can transport in vitro [99, 100], there has been less information regarding their function in vivo [101]. Several recent studies have examined the function of oatp1b2 in mice with a targeted disruption of the gene encoding this protein [102–104]. In each of these studies, there was no reported influence of disruption of this gene on breeding, development, or viability. One study [105] found that oatp1b2 KO mice had

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313

Table 21.1 Oatp family members found in rat, mouse, and human liver. (ND, not determined)

Species

Original name

Rat Rat Rat Rat Mouse Mouse Mouse Human Human Human Human

oatp1 oatp2 oatp4 oatp9 Oatp1 Oatp2 Oatp4 OATP-A OATP-C OATP8 OATP-B

New name

NCBI accession number

C-terminal sequence

Potential PDZ consensus

Plasma membrane localization

oatp1a1 oatp1a4 oatp1b2 Oatp2b1 Oatp1a1 Oatp1a4 Oatp1b2 OATP1A2 OATP1B1 OATP1B3 OATP2B1

NM_017111 NM_131906 NM_031650 NM_080786 NM_013797 NM_030687 NM_020495 NM_021094 NM_006446 NM_019844 NM_007256

KTKL VTED ETPL LQEL KTKL KTKL ETPL KTKL ETHC AAAN DSRV

Yes No Yes No Yes Yes Yes Yes No No Yes

Yes Yes Yes ND Yes ND Yes Yes Yes Yes Yes

Reprinted from [57] with permission.

reduced plasma clearance of dibromosulphthalein (DBSP) and were protected from the toxic effects of microcystin and phalloidin. A second study [103] showed that oatp1b2 KO mice had an approximately fourfold reduction in the liver-to-plasma ratio of rifampicin, smaller differences (less than twofold) for cerivastatin and lovastatin, and no difference for pravastatin or simvastatin. A third study [104] in which pravastatin was infused continuously to reach steady state showed a two- to threefold reduction in the liver-to-plasma ratio of pravastatin. It should be noted that there is no true homolog of this protein in human liver. Although some say that OATP1B1 and OATP1B3 are homologs of the murine protein [98, 103], their amino acid sequences are only 64 and 66% identical, respectively, to that of mouse oatp1b2. In addition, the mouse protein has a PDZ consensus binding site at its C-terminus, while these two human proteins do not (Table 21.1). However, they are thought to have overlap in some transported ligands [98, 105]. Interestingly, a large population-based study of patients taking simvastatin revealed that 85 subjects had developed signs of myopathy, a known side effect of this drug, and that this was associated with a polymorphism in the gene encoding OATP1B1 [106]. This study is in accord with several previous smaller studies that suggested that polymorphisms in OATP1B1 were associated with altered statin pharmacokinetics [107–111].

STUDIES OF Oatp1a1 AS A PROTOTYPICAL MEMBER OF THE Oatp FAMILY Membrane topology of oatp1a1 Analysis of oatp1a1 mRNA shows that it has an open reading frame of 2010 nucleotides, encoding a protein of 74 kDa. As is the case for all members of the oatp family, oatp1a1 is highly hydrophobic. The organization

of oatps within the plasma membrane has not been well defined. Although the oatps have been assumed to have 12 transmembrane domains, computer-based models have shown possible 12 as well as 10 transmembrane domain structures [112]. In a recent study, topology of oatp1a1 in the rat hepatocyte plasma membrane was assessed using an epitope-specific antibody designed to differentiate a 10 from a 12 transmembrane domain model [112]. The oatp1a1 sequence to which this antibody was prepared is present on the extracellular domain in the 10 transmembrane domain model, while it is on the cytosolic domain in the 12 transmembrane domain model. This antibody was used for immunofluorescence examination of rat hepatocytes with and without cell permeabilization. These studies showed that staining for oatp1a1 was seen only when cells were permeabilized. In the absence of permeabilization, oatp1a1 was not detectable, consistent with intracellular localization of the antigenic epitope, and the 12 transmembrane domain model. Additional insight into the membrane topology of oatp1a1 was obtained from studies of its N-linked glycosylation sites. Analysis on SDS-PAGE followed by immunoblot showed faster migration of oatp1a1 following incubation with the enzyme N-glycanase, indicating that it is a glycoprotein with at least one carbohydrate chain linked to an asparagine residue [113]. Computer analysis predicts four potential sites of N-glycosylation at amino acids at positions 62, 124, 135, and 492. Defining sites that undergo N-glycosylation provides important topological information, as glycosylation occurs only on extracellular domains of the protein [114]. Although these data show that oatp1a1 is N-glycosylated, they do not provide information regarding which of the four potential sites are utilized. To examine this issue, a series of mutagenized constructs were prepared in which the N-glycosylation consensus site was disrupted by mutation of the asparagine in the consensus sequence to a glutamine (N to Q). Constructs were expressed in HeLa cells and migration of oatp1a1 on immunoblot following SDS-PAGE was used to evaluate changes in its glycosylation state [112]. These studies showed that there was no effect on

314

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migration on SDS-PAGE of expressed oatp1a1 in which the asparagine at the first potential N-glycosylation site at amino acid 62 had been mutated to glutamine. Mutation of each of the three other potential N-glycosylation sites (asparagines 124, 135, and 492) resulted in faster migration of the expressed protein, consistent with loss of an N-linked carbohydrate chain from the protein. These studies thus indicate that of the four potential N-glycosylation sites on oatp1a1, three are utilized. These three sites (amino acids 124, 135, and 492) are on extracellular domains of the 12 transmembrane domain model, while the site that is not utilized (amino acid 62) is in a predicted transmembrane segment of this model, a position in which glycosylation would not occur.

Subcellular localization of Oatp1a1 Oatp1a1 is expressed on the basolateral (sinusoidal) plasma membrane of hepatocytes [113], on the apical (brush border) plasma membrane of the S3 segment of the proximal tubule [113], and on the apical (cerebrospinal fluid (CSF) side) plasma membrane of the choroid plexus epithelial cell [115, 116]. As seen in Table 21.1, seven of the oatp family members that have been described in rat, mouse, or human liver have PDZ consensus binding sites. PDZ consensus binding sites are defined by the sequence of a protein’s C-terminal four amino acids [117–119]. The crystal structures of known PDZ domains have been used to classify three types of PDZ consensus binding site [118]. The PDZ consensus sites that are present in the hepatic oatps are all of Class I, defined by the sequence X-S/T-X-
, where X is any amino acid and
is a hydrophobic amino acid [118]. There is still no way to predict which, if any, of the large number of PDZ proteins that have been described will bind a particular protein with a PDZ consensus binding site [118, 119]. Using a synthetic peptide corresponding to the C-terminal 16 amino acids of oatp1a1 for affinity isolation, interacting proteins from rat liver cytosol were purified. Protein mass fingerprinting identified PDZK1 as the major interacting protein [120]. PDZK1 and oatp1a1 coimmunoprecipitated from rat liver, indicating that they are bound to each other in vivo. Mouse and rat oatp1a1 are closely related to each other (82% amino acid identity) and have the same PDZ consensus binding sequence (KTKL). Studies performed in PDZK1 KO mice showed near normal expression of oatp1a1 protein. However, unlike wild-type mice, in which the transporter was localized to the basolateral plasma membrane, it was located predominantly in intracellular structures in PDZK1 KO mouse liver [120]. In agreement with these findings, plasma disappearance of the oatp1a1 ligand 35 S-BSP was reduced in the KO mice [120]. These studies strongly suggest that interaction of oatp1a1 with PDZK1 is essential for its proper subcellular localization and function. When studying function of transporters, it should be recognized that a protein must

be on the plasma membrane for it to be able to transport substances into a cell. A transporter’s subcellular distribution is thus an important potential regulatory factor that must be taken into account.

Studies of transport function of Oatp1a1 Initial studies characterized the transport function of oatp1a1 following its expression in Xenopus laevis oocytes [96, 121]. Subsequently, studies were performed in HeLa cells transiently co-transfected with vaccinia virus and an oatp1 expression plasmid [122–124]. These studies revealed a relatively broad range of substrates for oatp1-mediated transport, which included BSP, taurocholate, and estradiol 17β-D-glucuronide. Establishment of a stable HeLa cell line that expressed oatp1a1 under regulation of a metallothionein promoter [125] showed that oatp1a1-mediated transport was Na+ -independent, bidirectional, saturable, and of high affinity [125]. Studies in hepatocytes suggested that BSP uptake might be associated with HCO− 3 exchange [86]. This possibility was tested directly in the stably transfected cell system by examining the rates of taurocholate-dependent HCO− 3 efflux from alkali-loaded non-induced and induced cells that had been loaded with the pH-sensitive dye BCECF [126]. Addition of taurocholate to the outside of oatp1-expressing cells led to a rapid fall in intracellular pH, but had no effect on non-expressing cells. These studies show directly that oatp1a1 is an ion exchanger. Subsequent studies performed in Xenopus oocytes that had been microinjected with oatp1a1 cDNA showed that intracellular glutathione could also serve as an exchanged substrate [127]. As glutathione concentrations are high within hepatocytes, this could serve as an important driving force for organic anion uptake. The prostaglandin transporter (pgt), another member of the oatp family, has also been shown to be an anion exchanger [128].

Modulation of Oatp1a1 transport activity A number of physiological perturbations have been associated with altered oatp1a1 expression and function. In regenerating liver following two-thirds partial hepatectomy, influx of bilirubin falls by approximately 50% within six hours of surgery and returns to normal by four days [129]. Although bilirubin does not appear to be a substrate for oatp1a1, a similar reduction is seen for BSP uptake during liver regeneration [131]. Interestingly, hepatic oatp1a1 mRNA and protein levels show a similar pattern of expression during regeneration [132]. As regenerating liver in many ways recapitulates ontogeny of liver development, studies were also performed in

21: HEPATOCYTE BASOLATERAL MEMBRANE ORGANIC ANION TRANSPORTERS

the developing rat liver. Uptake of BSP in hepatocytes prepared from three-week-old animals was reduced by approximately 70% as compared to adults [116]. There was little oatp1a1 expression in liver until animals were one month of age, and this correlated with mRNA expression [116, 133]. Of note is the fact that oatp1a1 protein in the choroid plexus of one-day-old rats was essentially the same as in the adult, although examination of its subcellular distribution showed a major intracellular pool in contrast to an apical plasma membrane distribution in the adult choroid plexus [116]. The large differences in ontogeny of oatp1a1 expression seen in choroid plexus and liver suggest the presence of potent organ-specific transcription factors [134]. Oatp1a1 expression is also modulated following endotoxin administration and during cholestatic events. Eighteen hours following endotoxin administration to rats, uptake of indocyanine green by their isolated perfused livers is reduced by 40% as compared to controls [135]. Although oatp1a1 mRNA was unchanged, oatp1a1 protein expression was reduced by 50%. Following three to seven days of bile duct ligation, oatp1a1 mRNA and protein expression are reduced by approximately 50% [75]. Cholestasis due to administration of ethinyl estradiol for five days to rats was associated with an 80–90% reduction in expression of oatp1a1 mRNA and protein [136]. The time course over which reduced oatp1a1 protein expression takes place during cholestasis is relatively long, and the mechanism by which it occurs is thought to be related to alterations in expression of specific transcription factors [137–139]. A more rapid reduction in BSP transport was observed following exposure of hepatocytes to extracellular ATP [140]. Uptake was reduced by 80% within minutes and evidence was presented that this effect was mediated by a unique hepatocyte cell-surface purinergic receptor. Subsequent studies demonstrated that extracellular ATP reduces the Vmax for transport of BSP by rat hepatocytes without altering Km [141]. Similar reduced transport also occurs within minutes following incubation of hepatocytes with the phosphatase inhibitors okadaic acid or calyculin A [141]. When hepatocytes were preloaded for two hours with inorganic 32 P, subsequent short-term incubation in extracellular ATP resulted in serine phosphorylation of oatp1a1 with the appearance of a single major tryptic phosphopeptide. Transport of BSP by stably transfected HeLa cells is unaffected by incubation with extracellular ATP or phosphatase inhibitors, and phosphorylation of oatp1a1 does not occur in these cells, suggesting that they lack the kinase that mediates this effect. To establish the site of phosphorylation, oatp1a1 was purified from rat liver by immunoaffinity chromatography, subjected to trypsin digestion, and analyzed by mass spectrometry (MS) [142]. Except for predicted N-glycosylated peptides that would be difficult to detect by the technique that was used, 97% of oatp1a1 tryptic peptides were observed. A single tryptic phosphopeptide was found in the C-terminus (aa 626–647), existing in unphosphorylated,

315

singly phosphorylated, or doubly phosphorylated forms. Subsequent tandem MS/MS analysis revealed that phosphorylation at S634 accounted for all singly phosphorylated peptide, while phosphorylation at S634 and S635 accounted for all doubly phosphorylated peptide [142]. These findings identify the site of oatp1a1 phosphorylation and demonstrate that it is an ordered process, in which phosphorylation at S634 precedes that at S635. The mechanism by which phosphorylation results in loss of transport activity in hepatocytes remains to be established. Whether or not phosphorylation near the C-terminus regulates C-terminal oligomerization of oatp1a1 can be speculated upon but is as yet unknown.

SUMMARY A major function of the hepatocyte is uptake from the circulation of a large number of endogenous organic anionic compounds and xenobiotics. Several basolateral membrane transporters that can mediate these uptake processes have been described. These include ntcp and several members of the oatp family. Although a great deal has been learned about their transport function and expression, critical areas related to their cell biology and regulation of subcellular distribution remain to be elucidated. In order to fully understand their roles in health and disease, studies of these transporters directed toward structure–function relationships as well as factors required for cell-surface expression and activity will be of major importance.

ACKNOWLEDGMENTS The authors would like to acknowledge their support by NIH grants DK 23026, DK 41296, and DK41918.

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89. Min, A.D., Johansen, K.J., Campbell, C.G. and Wolkoff, A.W. (1991) Role of chloride and intracellular pH on the activity of the rat hepatocyte organic anion transporter. J Clin Invest , 87, 1496–502. 90. Stremmel, W., Gerber, M.A., Glezerov, V., Thung, S.N., Kochwa, S. and Berk, P.D. (1983) Physicochemical and immunohistological studies of a sulfobromophthalein- and bilirubin-binding protein from rat liver plasma membranes. J Clin Invest , 71, 1796–805. 91. Stremmel, W. and Berk, P.D. (1986) Hepatocellular uptake of sulfobromophthalein and bilirubin is selectively inhibited by an antibody to the liver plasma membrane sulfobromophthalein/bilirubin binding protein. J Clin Invest , 78, 822–26. 92. Mullock, B.M., Branch, W.J., van Schaik, M., Gilbert, L.K. and Luzio, J.P. (1989) Reconstitution of an endosome-lysosome interaction in a cell-free system. J Cell Pharmacol , 108, 2093–99. 93. Wolkoff, A.W. and Chung, C.T. (1980) Identification, purification, and partial characterization of an organic anion binding protein from rat liver cell plasma membrane. J Clin Invest , 65, 1152–61. 94. Goeser, T., Nakata, R., Braly, L.F., Sosiak, A., Campbell, C.G., Dermietzel, R., Novikoff, P.M., Stockert, R.J., Burk, R.D. and Wolkoff, A.W. (1990) The rat hepatocyte plasma membrane organic anion binding protein is immunologically related to the mitochondrial F1 adenosine triphosphatase beta-subunit. J Clin Invest , 86, 220–27. 95. Sottocasa, G.L., Baldini, G., Sandri, G., Lunazzi, G. and Tiribelli, C. (1982) Reconstitution in vitro of sulfobromophthalein transport by bilitranslocase. Biochim Biophys Acta, 685, 123–28. 96. Battiston, L., Passamonti, S., Macagno, A. and Sottocasa, G.L. (1998) The bilirubin-binding motif of bilitranslocase and its relation to conserved motifs in ancient biliproteins. Biochem Biophys Res Commun, 247, 687–92. 97. Battiston, L., Macagno, A., Passamonti, S., Micali, F. and Sottocasa, G.L. (1999) Specific sequence-directed anti-bilitranslocase antibodies as a tool to detect potentially bilirubin-binding proteins in different tissues of the rat. FEBS Lett , 453, 351–55. 98. Jacquemin, E., Hagenbuch, B., Stieger, B., Wolkoff, A.W. and Meier, P.J. (1991) Expression of the hepatocellular chloride-dependent sulfobromophthalein uptake system in Xenopus laevis oocytes. J Clin Invest , 88, 2146–49. 99. Jacquemin, E., Hagenbuch, B., Stieger, B., Wolkoff, A.W. and Meier, P.J. (1994) Expression cloning of a rat liver Na+-independent organic anion transporter. Proc Natl Acad Sci U S A, 91, 133–37. 100. Wolkoff, A.W. (1996) Hepatocellular sinusoidal membrane organic anion transport and transporters. Semin Liver Dis, 16, 121–27. 101. Hagenbuch, B. and Meier, P.J. (2003) The superfamily of organic anion transporting polypeptides. Biochim Biophys Acta, 1609, 1–18.

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Nuclear Receptors Regulate Bile Acid Synthesis Guorong Xu1,2 and Gerald Salen1 1

University of Medicine and Dentistry of New Jersey, New Jersey Medical School, NJ, USA 2 VA Medical Center, East Orange, NJ, USA

INTRODUCTION One of the most perplexing questions facing investigators interested in cholesterol homeostasis is the feedback regulation of bile acid synthesis. Bile acids represent the major catabolic product of cholesterol and their synthesis plays an important role in plasma cholesterol homeostasis. In the conversion of cholesterol to bile acids, the first reaction is the 7α-hydroxylation of cholesterol, such that every molecule of 7α-hydroxycholesterol is committed to bile acid synthesis. This reaction is catalyzed by the hepatic microsomal enzyme cholesterol 7α-hydroxylase (CYP7A1) and is considered rate-controlling for classic bile acid synthesis such that changes in CYP7A1 expression/activity mirror the direction of bile acid synthesis. Recently, it has been found that bile acids serve not only as physiological detergents to facilitate the digestion/absorption of fat and fat-soluble vitamins, but also as signaling molecules to activate nuclear receptors that regulate bile acid and cholesterol metabolism. Two important nuclear receptors, farnesoid X receptor (FXR) and liver X receptor (LXR) α have been found to function as the sensors for bile acid and cholesterol homeostasis, respectively, and to play a critical role in regulating CYP7A1 transcription. This review focuses on the ligand activation of the nuclear receptors and their effects, when activated, on the molecular regulation of CYP7A1 The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

expression/bile acid synthesis, in order to interpret the various physiological responses observed in animal studies.

DIFFERENT RESPONSES OF CYP7A1 TO DIETARY CHOLESTEROL IN RABBITS AND RATS Some humans are sensitive to dietary cholesterol, where marked plasma hypercholesterolemia develops when fed cholesterol, whereas others are relatively insensitive and plasma cholesterol levels barely increase when large amounts of cholesterol are eaten. However, it is not easy to predict these responses in patients [1, 2]. The opposite response to dietary cholesterol has been observed in other animal species. For example, rats fed a high-cholesterol diet do not develop hypercholesterolemia [3]. Responding to the increased dietary cholesterol load, rats upregulate CYP7A1 [3–6], the rate-controlling enzyme for the classic bile acid synthesis pathway [7], and divert the extra cholesterol to bile acid synthesis. The effect of cholesterol feeding in New Zealand white (NZW) rabbits, where dietary cholesterol downregulated CYP7A1 [8], was opposite to that seen in rats, so that

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cholesterol accumulated rapidly in the plasma compartment [9]. These observations suggest that a fundamental difference in regulation of CYP7A1 and bile acid synthesis between rabbits and rats contributes to their different susceptibility to develop plasma hypercholesterolemia. However, not all rabbits are sensitive to dietary cholesterol. Overturf et al. discovered a dietary cholesterol-resistant colony of NZW rabbits in which feeding 0.1% cholesterol for seven months did not produce hypercholesterolemia [10]. In these rabbits, fecal bile acid excretion was increased [11] and both activity and mRNA levels of CYP7A1 were higher than in normal NZW rabbits after feeding 0.1% cholesterol-enriched diet [12]. Thus, plasma cholesterol levels remained within the normal range after feeding cholesterol, presumably because the absorbed cholesterol was converted to more bile acids which were excreted in feces. African green monkeys fed cholesterol develop marked plasma hypercholesterolemia associated with inhibited CYP7A1 [13]. In addition, cholesterol-fed hamsters also manifest plasma hypercholesterolemia with suppressed CYP7A1 activity and mRNA levels [14, 15]. The information from different animal species suggests that plasma cholesterol concentrations are inversely related to hepatic CYP7A1 activity/mRNA, such that stimulation of CYP7A1 expression should be associated with reduced plasma cholesterol levels. This hypothesis is supported by findings in Watanabe heritable hyperlipidemic rabbits, where hepatic low-density lipoprotein (LDL) receptors are deficient. Stimulating CYP7A1 and classic bile acid synthesis by bile drainage reduced plasma cholesterol levels 40% in these rabbits [16]. Upregulation of CYP7A1 diverts significant amounts of cholesterol away from the plasma compartment to bile acid synthesis. In continuing studies to answer why cholesterol feeding in rabbits downregulates CYP7A1, it was discovered that the bile acid pool doubled in size and biliary bile acid outputs increased by 47% in NZW rabbits fed 2% cholesterol for 10 days [17]. In these cholesterol-fed rabbits, CYP7A1 activity decreased by 68%, while mitochondrial sterol 27-hydroxylase (CYP27) activity—reflecting the alternative bile acid synthesis pathway [18–20]—rose by 66%. The investigators hypothesized that CYP7A1 was feedback-regulated by the expanded circulating bile acid pool in cholesterol-fed rabbits. However, hepatic CYP27 was insensitive to changes in the bile acid pool and alternative bile acid synthesis was upregulated by increasing amounts of cholesterol taken in the liver via the hepatic LDL receptors. It was noticed that dietary cholesterol stimulated CYP27 activity in NZW rabbits but not in LDL receptor-deficient Watanabe rabbits [21]. Thus, LDL receptor seems essential for directing cholesterol to the alternative bile acid synthetic pathway and CYP27. Information about the regulation of sterol 27-hydroxylase and alternative bile acid synthesis is limited and controversial. Vlahcevic et al. [22] reported that in rats, malabsorption of bile acid produced by fed cholestyramine stimulated, and bile acid feeding inhibited, hepatic CYP27 activity.

Studies in rabbits from other investigators [23] agreed with the observations that CYP27 activity does not respond to cholic acid feeding nor bile acid malabsorption induced by cholestyramine treatment that strongly modulates CYP7A1 activity and mRNA levels. Then, the effect of dietary cholesterol on the regulation of classic (CYP7A1) and alternative (CYP27) bile acid synthesis in rabbits has been systematically studied [24]. Feeding 2% cholesterol for one day increased hepatic cholesterol and CYP7A1 activity with no significant change in the bile acid pool size or CYP27 activity. After three days of feeding cholesterol, the bile acid pool size increased significantly (83%), and further feeding of cholesterol produced increases in 10–20% increments, while CYP7A1 activity declined progressively to 60% below the baseline. In contrast, CYP27 activity rose after three days of cholesterol feeding and remained elevated with the continued increase of cholesterol intake. Bile drainage depleted the bile acid pool to stimulate the downregulated CYP7A1 activity. The more cholesterol deposited in the liver, the greater the upregulation of CYP7A1 activity in these rabbits. In contrast, bile drainage did not affect CYP27 activity. Thus, it was the enlarged bile acid pool, not cholesterol, that was responsible for the inhibition of CYP7A1. Increased hepatic cholesterol was not an inhibitor but rather a stimulator that initially induced CYP7A1 activity and resulted in greater upregulation of CYP7A1 after removal of the bile acid pool. CYP27 is insensitive to the bile acid flux, yet is upregulated by increasing hepatic cholesterol. It is proposed that activated CYP27 was responsible for the expansion of the bile acid pool size, because it continued actively making bile acids via the alternative bile acid synthesis pathway, regardless of the size of the expanded bile acid pool [24]. Activation of CYP27 was not simply compensation for the inhibition of classic bile acid synthesis/CYP7A1, but the result of continued stimulation by increased hepatic cholesterol levels. The next question considered was why the bile acid pool size enlarged in rabbits fed cholesterol, and whether the bile acid pool also changed size in cholesterol-fed rats. It has been found that the expression of the ileal apical sodium-dependent bile acid transporter (ASBT) protein did not change in rats, but rose 31% (p < 0.05) in rabbits after feeding cholesterol, and that the bile acid pool size did not expand in cholesterol-fed rats [25]. The investigators hypothesized that CYP7A1/classic bile acid synthesis was inhibited in cholesterol-fed rabbits because an enlarged bile acid pool developed from enhanced ileal bile acid reabsorption (transport) and alternative bile acid synthesis. In contrast, rat CYP7A1 was stimulated because the bile acid pool did not enlarge due to unchanged ileal ASBT expression. Therefore, although bile acid synthesis increased via different pathways in rats and rabbits, enhanced ileal bile acid transport was crucial for the expansion of the bile acid pool size, which exerted feedback regulation on CYP7A1.

22: NUCLEAR RECEPTORS REGULATE BILE ACID SYNTHESIS

THE NUCLEAR RECEPTOR LIVER X RECEPTOR (LXR) α Recently, LXRα, one of the members of the orphan nuclear receptor superfamily, has been found to be a positive regulator for CYP7A1 transcription [26–28]. LXRα is expressed most highly in the liver [29] and is activated by oxidized derivatives of cholesterol (oxysterols) [30]. In LXRα knockout (KO) mice, 2% cholesterol feeding did not upregulate CYP7A1, as occurred in wild-type mice (LXRα+/+) fed cholesterol [31]. These results demonstrate that CYP7A1 is a regulated target for LXRα and that cholesterol feeding that increases formation of oxysterols activates LXRα to induce CYP7A1 expression. In addition to mice, it is well documented that CYP7A1 is upregulated in rats fed cholesterol [3–6]. Chiang et al. reported that in the rat the CYP7A1 promoter was bound tightly to LXRα, and in the hamster the CYP7A1 promoter was bound loosely to LXRα, while the human CYP7A1 promoter had no LXR binding site at all [32]. It is presently unknown how the rabbit CYP7A1 promoter fits this scheme. It was asked if in rabbits, activation of LXRα induced by dietary cholesterol might inhibit CYP7A1 expression, as has been reported in human hepatocytes [33], and whether increased amounts of cholesterol would repress the rabbit but induce rat CYP7A1 transcription. To answer this question, Xu et al. cloned the rabbit CYP7A1 promoter and transfected the promoter into HepG2 cells (a cellular line of human hepatoma). The data demonstrated that not only does the rabbit CYP7A1 promoter contain a functional LXR binding site identical to that in the rat, but that LXRα/retionoid X receptor (RXR) stimulates rabbit promoter activity in a similar way to what is seen in those cells transfected with the rat promoter [34]. Furthermore, free cholesterol alone neither inhibited nor induced CYP7A1 promoter activity, but its oxidized products—oxysterols—stimulated the promoter activity in the HepG2 cells with either the transfected rabbit or the rat CYP7A1 promoter. Thus, theoretically, activation of LXRα should also upregulate CYP7A1 expression in the rabbit. So why in rabbits did cholesterol feeding result in repression of CYP7A1 expression and activity? These investigators believed that this occurred because the bile acid pool size was enlarged significantly in rabbits fed 2% cholesterol for more than three days [24].

THE NUCLEAR RECEPTOR FARNESOID X RECEPTOR (FXR) When we speak about the effect of an enlarged bile acid pool size on CYP7A1 expression, we need first to understand the role of another important nuclear receptor, FXR, in the regulation of CYP7A1 transcription and bile acid synthesis/homeostasis. FXR is expressed predominantly in the liver, kidney, intestine, and adrenals [35] and has recently been identified as a bile acid sensor and a

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negative regulator for CYP7A1 transcription [36–38]. The initial studies were carried out in CV-1 cells, and a variety of individual bile acids were tested. The hydrophobic free bile acids—chenodeoxycholic acid (CDCA), deoxycholic acid (DCA), and lithocholic acid—were found to be the most potent ligands for the activation of FXR. However, it should be pointed out that in mouse in vivo studies, cholic acid has been also demonstrated to be a potent ligand for FXR [39]. When bound to ligands (bile acids), FXR suppressed transcription of CYP7A1 and activated the gene-encoding ileal bile acid binding protein (IBABP) [37], which binds and transports bile acids within ileal enterocytes. However, the importance of IBABP in intestinal bile acid absorption has not yet been completely defined. Other target genes of FXR have been reported which are related to either bile acid synthesis or hepatic and intestinal bile acid transport. For example, activation of FXR downregulates cholesterol 12α-hydroxylase, CYP8B1, a key enzyme involved in the synthesis of the major primary bile acid, cholic acid [40]. FXR target genes also include: the hepatocanalicular bile acid transporter “bile salt export pump” (BSEP) [41], which is the major transporter responsible for hepatic bile acid excretion and generation of bile flow; the sodium-taurocholate cotransporting polypeptide (NTCP) [42], which is responsible for hepatic sinusoidal bile acid uptake from the portal blood [43]; and the heteromeric organic solute transporters, OSTα/OSTβ [44–46], which are basolateral bile acid transporters that transport bile acids from ileal enterocytes into the portal blood returning to the liver [47]. ASBT is the bile salt transporter in the terminal ileum responsible for the absorption (uptake) of bile acids from the intestinal lumen [48, 49]. Although there is no direct FXR binding site in the ASBT promoter, it has been reported that in the human [50], mouse [51], and rabbit [52], activation of FXR downregulates ASBT expression. In summary, bile acid-induced activation of FXR actually controls bile acid synthesis through these two key synthetic enzymes (CYP7A1 and CYP8B1), as well as the bile acid enterohepatic circulation via the bile acid transporters in and out of the liver and intestine. Thus, FXR plays the determining role in bile acid homeostasis in the body.

SHORT HETERODIMER PARTNER (SHP), LIVER RECEPTOR HOMOLOG-1 (LRH-1), AND HEPATOCYTE NUCLEAR FACTOR 4 (HNF4) Importantly, short heterodimer partner (SHP) [53] was found to be a direct target of FXR [54–56]. Chiang et al. found [57] that FXR/RXR did not directly bind to the bile acid response element (BARE) in the promoter

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THE LIVER: HETERODIMER PARTNER, LIVER RECEPTOR HOMOLOG-1, AND HEPATOCYTE NUCLEAR FACTOR 4

region of human CYP7A1 and suggested that bile acid-activated FXR inhibited CYP7A1 transcription by an indirect mechanism. Then two groups reported their very important findings [54, 56] that activated FXR enhanced SHP expression, which in turn formed a heterodimer with liver receptor homolog-1 (LRH-1) [58]. LRH-1 is a hepatic-specific transcription factor essential for CYP7A1 gene expression. It is also called fetoprotein transcription factor (FTF) in other species [59] and CYP7A1 promoter binding factor (CPF) in humans [60]. Because of the essential role of LRH-1 (FTF) in CYP7A1 transcription, these investigators proposed that the formation of heteromeric complex SHP/LRH-1 inactivates LRH-1 and results in inhibition of CYP7A1 transcription [54, 56]. This hypothesis of cascade reaction (bile acid–FXR–SHP–FTF–CYP7A1) may provide an explanation at the molecular level for the mechanism of repression of CYP7A1 indirectly by bile acid-activated FXR. However, it remains to be answered whether the proposed cascade via SHP/LRH-1 (or SHP/FTF) is the specific pathway for activated FXR to repress CYP7A1 transcription, as over-expressed SHP may also inhibit other receptors [58]. Recently, it was reported that in Shp–/–KO mice, feeding cholic acids could still downregulate CYP7A1 expression, and it was suggested that there are other SHP-independent pathways such as pregnane X receptor (PXR) or the c-Jun N-terminal kinase (JNK) for feedback regulation of CYP7A1 by bile acids [61, 62]. These findings demonstrated the existence of redundant pathways involving bile acids, which guarantee this very important feedback downregulation function in CYP7A1/bile acid synthesis in order to protect the liver from the toxicity of accumulation of bile acids. However, it does not yet rule out the crucial role of SHP in the feedback regulation of CYP7A1 transcription. More recently, it has been questioned whether FTF is always a positive transcriptional factor. The putative BARE-II contains a hepatocyte nuclear factor (HNF) 4α binding site, another major positive transcriptional factor for CYP7A1 expression, which overlaps with the FTF binding site [63, 64]. Feeding rats CDCA repressed CYP7A1 expression associated with induced FTF but not SHP mRNA expression [64]. The author proposed that FTF might compete with HNF4 for overlapping binding sites in the BARE-II. As HNF4 is essential for basal transcription of CYP7A1 [65], FTF also might be a suppressor of CYP7A1 expression. More surprisingly, it has been reported that in heterozygous KO LRH-1 mice, CYP7A1 and CYP8B1 mRNA levels were fiveto sevenfold higher than baseline value in the wild-type mice [66]. Adenovirus-mediated FTF overexpression in wild-type mice caused tenfold decrease of the expression in CYP7A1 and CYP8B1, with eightfold increase of SHP mRNA expression. Furthermore, the authors who proposed that feedback repression of CYP7A1 was via a cascade of bile acids–FXR–SHP–LRH-1 [54, 56] recently published another paper. In that paper

these investigators suggested that LRH-1 plays an important role in bile acid homeostasis but might not be essential for feedback regulation of CYP7A1 [67]. Using a special technique, they made a mouse model with only LRH-1 KO in either hepatocytes or the intestinal epithelium. Unexpectedly, they found that basal CYP7A1 expression was not decreased and the suppressive effect of activated FXR on CYP7A1 was still present in mice with LRH-1 KO in hepatocytes. Thus, another important question remains to be answered: what is the exact role of FTF in the regulation of CYP7A1? Indeed, much more work has to be done to define whether LRH-1 (FTF) is an enhancer or suppressor of CYP7A1 transcription and how FTF is involved in the regulation. Xu et al. observed the effect of FTF in the regulation of CYP7A1 in rabbits and rats. They demonstrated that there is a functional FTF binding site in the rabbit promoter similar to that in the rat. However, in HepG2 cells transfected with the rabbit CYP7A1 promoter, addition of increasing quantities of FTF did not further increase rabbit CYP7A1 promoter activity. The strong elevation of the promoter activity by the LXRα/RXR was nearly abolished when the FTF binding site in the rabbit CYP7A1 promoter was mutated [34]. This result agrees with others [54, 68] that FTF is a competent factor for the stimulation of CYP7A1 expression by LXRα. Xu et al. believe that FTF itself is not a stimulator but rather is a transcriptional factor for maintaining baseline level of the rabbit CYP7A1 promoter because: (i) mutation of FTF binding site abolished the promoter baseline activity; (ii) the amount of FTF protein naturally synthesized by HepG2 cells was sufficient to maintain rabbit CYP7A1 expression in culture but more FTF did not increase its activity; (iii) activity of the rabbit CYP7A1 promoter after its transfection into HEK 293 cells, which do not synthesize FTF, was barely detectable before but increased markedly after the cells were supplied with FTF; and (iv) the activation pattern in the two cell lines was similar, in that a baseline level of FTF (endogenous in HepG2 but exogenous in HEK) was needed to stimulate promoter activity but additional FTF led to no further increase in activity [34]. However, an excess amount of FTF did not further stimulate but in fact suppressed the promoter activity in HepG2 cells and HEK cells transfected with either the rabbit or rat CYP7A1 promoter. Furthermore, addition of a moderate amount of FTF with SHP enhanced the inhibitory effect of SHP, but not the stimulatory effect of activated LXRα. Thus, it is suggested that FTF may also have negative effects on the rabbit CYP7A1 promoter under other conditions. In previous in vivo animal studies, such as total bile drainage and feeding 1% cholic acid or 2% cholesterol in rabbits or rats, or feeding 2% cholesterol + 1% cholic acid in rats, the investigators did not find significant change of FTF mRNA levels in the liver tissues from these animals, though CYP7A1 expression in these tissues was increased or decreased, respectively. Thus, it is unlikely that CYP7A1 transcription is regulated by

22: NUCLEAR RECEPTORS REGULATE BILE ACID SYNTHESIS

means of significant changes in FTF expression under physiological conditions, but rather a steady level of FTF is required as FTF is one of the essential transcriptional factors needed for basal expression of CYP7A1. Similar to FTF, HNF4 is another major positive transcriptional factor crucial for CYP7A1 basal level transcription [65, 69]. It has been reported that SHP may directly interfere with HNF4 binding by repressing nuclear receptor-mediated transactivation through both competition with coactivators for binding to the nuclear receptor and its direct inhibitory effect on the nuclear receptor’s transcriptional activity [58]. As FTF and HNF4 binding sites overlap in BARE-II, it is proposed that increased expression of FTF suppresses CYP7A1 transcription by competing with HNF4 for binding to the promoter [63, 64]. In cholesterol-fed rabbits, no significant changes in FTF but a threefold increase in hepatic SHP mRNA were observed. Thus, at least in cholesterol-fed rabbits, the downregulation of CYP7A1 is not due to increased levels of FTF competing with HNF4 for binding to CYP7A1. However, it would be interesting to know whether SHP with FTF (SHP/FTF heterodimer) would also compete with HNF4 for binding to the BARE-II region, just as it does with LXRα for binding to the BARE-I (LXR binding site) [70].

THE ROLE OF NUCLEAR RECEPTORS IN THE REGULATION OF CYP7A1 The roles of hydrophobic DCA and hydrophilic ursocholic acid (UCA, a weak FXR ligand) were investigated in regulation of FXR in rabbits after total bile drainage for seven days where the bile acid pool had been completely depleted [71]. Replacing the bile acid flux with DCA alone restored activation of hepatic FXR, indicated by increased expression of its target genes, SHP and BSEP, and resuppressed CYP7A1 mRNA level and activity, which were elevated due to depletion of the bile acid pool. However, replacing with UCA did not activate FXR, as SHP mRNA and CYP7A1 mRNA/activity remained unchanged. The bile acid flux was composed of 82% UCA with only 17% CA and 0.6% DCA in the rabbits infused with UCA. The amounts of CA and DCA in the flux were apparently below the ligand-activating threshold for FXR. Therefore, a sufficient enterohepatic circulating flux of activating/high-affinity ligands, and not just the total size of the circulating pool, is required for activation of FXR. In other words, the composition of the bile acid flux through the liver plays a critical role in determining the nuclear activation state of FXR. This opinion has been supported by another report in Cyp8b1–/–mice [39]. In those mice, cholic acid was eliminated from the pool and approximately 85% of the bile acids in the pool were composed of non-activating FXR ligand bile acids

327

(muricholates and ursodeoxycholate). The bile acid pool increased in size, while bile acid synthesis via mainly Cyp7a1 did not decrease but rather increased significantly. Because of the significant increased proportion of non-activating ligands with the absence of FXR-activating ligand (cholate) in the bile acid pool, the feedback regulation of Cyp7a1 in the Cyp8b1–/–mice seemed “lost.” This in vivo study [39] demonstrated that cholate is the potent and important ligand for FXR activation in mice, which is different from the results observed in the in vitro cell studies [36–38]. There is as yet no method available to measure directly the activation status of FXR and LXRα in whole-animal models. To evaluate the regulatory role of nuclear receptors FXR and LXRα on CYP7A1 transcription, Xu et al. measured the changes in the expression of key target genes of FXR (SHP, BSEP, and CYP8B) or LXRα (cholesterol ester transfer protein (CETP) and ATP-binding cassette transporter ABC1 (ABCA1)) as markers to indicate the activation state of these nuclear receptors. It has been noted that SHP mRNA levels always mirrored the changes in FXR activation. However, although BSEP mRNA levels were regulated by activated FXR, other bile acid-dependent mechanisms also appear to be involved [71]. This FXR-independent regulation on BSEP has also been observed in mice [39]. The effects of dietary cholesterol on FXR and LXRα activation and their role in the regulation of CYP7A1 in NZW rabbits were also studied [72]. It was found that after one day of cholesterol feeding, hepatic concentrations of oxysterols, the activating ligands for LXRα, rose significantly and LXRα became activated, as evidenced by the rise in mRNA levels of its target genes, ABCA1 and CETP, in the liver. In contrast, mRNA levels of FXR target genes did not change, which indicated that FXR activation remained unchanged. In these rabbits fed cholesterol for one day, the bile acid pool—which contained 85–90% DCA, an activating ligand for FXR—did not increase, so that FXR was not activated. CYP7A1, the target gene of both FXR and LXRα, was upregulated in response to the enhanced activation of LXRα, secondary to the increase in its high-affinity oxysterol ligands. However, in rabbits fed cholesterol for 10 days, the bile acid (FXR ligand) pool size expanded significantly, leading to increased hepatic FXR activation, as indicated by the increased mRNA levels of its target genes, BSEP and SHP. Both activity and mRNA levels of CYP7A1 were decreased despite continued activation of LXRα, as evidenced by further increase in mRNA levels of its target genes, ABCA1 and CETP, in the liver. These results suggest that in rabbits, the inhibitory effect of FXR is dominant over the stimulatory effect of LXRα in regulation of CYP7A1 under these conditions. Further studies [73] showed that in rats, unlike in rabbits, feeding cholesterol did not expand the bile acid (ligand) pool for FXR but substantially increased LXRα ligands, oxysterols in the liver. FXR was not activated,

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as evidenced by unchanged mRNA levels of FXR target genes, SHP and BSEP, while LXRα was activated, with a significant increase in mRNA levels of LXRα target gene, ABCA1. As a result, CYP7A1 was upregulated by activated LXRα in cholesterol-fed rats. It is important to note that FXR was not activated in these cholesterol-fed rats, not only because the bile acid (ligand) pool size did not expand but because the proportion of DCA + CA + CDCA, the activating ligands for FXR, declined, whereas muricholic acids (none-activating ligands) rose in the bile acid pool. This observation was different from that reported by Gupta et al. [74]. These authors concluded that LXRα was the dominant regulator of CYP7A1 as (i) in cholesterol-fed rats, CYP7A1 was upregulated by LXRα, although at the same time FXR that was activated as the expression of FXR target gene, SHP was increased; and (ii) after adding FXR ligand CA to cholesterol diet, CYP7A1 mRNA levels were still increased 300% as compared to controls. However, Xu et al. believed that up-regulation of CYP7A1 in cholesterol-fed rats is not because rat CYP7A1 is only sensitive to activated LXRa and not to FXR activation but rather, because FXR is not activated. When FXR is activated, such as in rats fed CA with cholesterol, CYP7A1 is also suppressed, as seen in rabbits. This opinion has been further confirmed by recent in vitro studies in HepG2 cells where the rabbit and rat CYP7A1 promoters were transfected [34]. In the rabbit CYP7A1 promoter region, there is a functional LXR binding site identical to that in the rat promoter. Both rabbit and rat CYP7A1 promoter activity increased significantly in response to activated LXRα/RXR. Most importantly, this elevated promoter activity induced by LXRα/RXR in both rabbit and rat CYP7A1 promoters was decreased in a dose-dependent manner by adding SHP, which is increased by activation of FXR physiologically. Thus, the above studies in rabbits and rats suggest that the inhibitory effect of FXR activation is dominant over the stimulatory effect of LXR activation in the regulation of CYP7A1 transcription. When FXR and LXRα are simultaneously activated, the effect of FXR will override the effect of LXRα. The next question to be answered is how activation of FXR overrides the effect of LXRα activation. It has been identified that in addition to the FTF binding site located upstream of the rabbit CYP7A1 promoter (–129/–137), there is another FTF binding site (–54/–62) embedded within the LXR binding site [70]. The study demonstrated that FTF binds to the LXR binding site and competes with LXRα for binding to the CYP7A1 promoter. This observation provided a molecular basis for the involvement of SHP in the downregulation of CYP7A1 expression stimulated by LXRα/RXR, as SHP usually forms heterodimers with FTF. Adding SHP together with FTF in HepG2 cells transfected with the rabbit CYP7A1 promoter resulted in significantly lower promoter activity compared with SHP alone. The gel shift assay visibly demonstrated that although SHP itself did not directly

bind to the LXR binding site, adding SHP with FTF enhanced FTF binding to the LXR binding site and further diminished LXRα binding to its binding site in a dose-dependent manner. The above results suggest that increased amounts of SHP with FTF specifically act on the LXR binding site to block LXRα/RXR from binding, such that the positive effect of LXR activation is diminished. This effect of SHP plus FTF against LXRα activation is different from the possible direct effect of SHP in repressing LXRα’s transcriptional activity [75], or the indirect effect of SHP in inactivating FTF on the upstream of the promoter, as suggested in the feedback regulation of bile acid synthesis [54, 56]. Furthermore, the results from the chromatin immunoprecipitation (ChIP) assays demonstrated that SHP combined with FTF significantly reduced the amount of LXRα-bound DNA (LXR binding site in the CYP7A1 promoter) in HepG2 cells transfected with the rabbit CYP7A1 promoter (Figure 22.1a). The significant reduction of LXRα-bound DNA (CYP7A1 promoter) in ChIP assays confirmed that SHP combined Ctrl

FTF

SHP

F/S

Anti-LXRa IgG Input (a) Ctrl

Ch1d

Ch10d

Anti-LXRa IgG Input (b)

Figure 22.1 SHP combined with FTF prevents activated LXRα from binding to its natural binding site. LXRα-bound DNA (LXR binding site) determined by chromatin immunoprecipitation (ChIP) assays. Anti-LXRα: LXRα-bound chromatin precipitated by anti-LXRα antibody. IgG: the negative controls where the chromatin was precipitated using normal mouse IgG (non-specific antibody). Input was the positive controls, where no antibody was applied. The primer was designed to amplify the region around the LXR binding site in the rabbit CYP7A1 promoter by PCR. (a) In HepG 2 cells with activated endogenous LXRα/RXR and transfected rabbit CYP7A1 promoter. In each group, LXRα and RXR ligands, 22(R)-hydroxycholesterol and 9-cis-retinoic acid were added. FTF: FTF expression plasmid was transfected. SHP: SHP expression plasmid was transfected. F/S: both FTF and SHP expression plasmids were transfected. Only when SHP is added with FTF is the LXR-bound DNA (CYP7A1 promoter) almost undetectable. (b) In liver specimens from rabbits fed 2% cholesterol for one day (Ch 1d) and ten days (Ch 10d), or chew only (Ctrl). The LXR-bound DNA is reduced only in the liver from rabbits fed cholesterol for ten days. SHP expression increased in the liver of Ch 10d but not in those for Ch 1d

22: NUCLEAR RECEPTORS REGULATE BILE ACID SYNTHESIS

with FTF blocked LXRα binding to its binding site to diminish recruitment of LXRα to the CYP7A1 promoter. Most importantly, ChIP assays in liver specimens from the rabbit studies showed that LXRα-bound promoter DNA (LXR binding site) remained unchanged in rabbits fed 2% cholesterol for one day where SHP expression was not increased. The LXRα-bound DNA was decreased in rabbits fed cholesterol for ten days (Figure 22.1b) where FXR was activated and SHP mRNA/protein increased [70]. These data strongly support the hypothesis that 2% cholesterol diet feeding for 10 days results in an enlarged bile acid (FXR ligand) pool size, which in turn induces activation of FXR. The activated FXR stimulates SHP expression, which produces SHP/FTF heterodimers to block LXRα from binding to the CYP7A1 promoter. As a result of abolishing the stimulatory effect of LXRα, the inhibitory effect of FXR dominates such that CYP7A1 expression is repressed when FXR and LXRα are simultaneously activated. Xu et al. hypothesize that the induced expression of SHP by activation of FXR is likely to increase the formation of FTF/SHP heterodimers, which then occupy the LXR binding site via the embedded FTF binding element and diminish recruitment of LXRα [70].

ENTEROHEPATIC CIRCULATION AND REGULATION OF BILE ACID ABSORPTION IN THE ILEUM Figure 22.2 illustrates the enterohepatic circulation. The bile acids are synthesized in the liver and excreted into the bile by the hepatocanalicular bile acid transporter, BSEP [76, 77]. After entering the small intestine, 95% of the bile acids will be actively reabsorbed at the terminal ileum by an apical bile salt transporter, ASBT [48, 49]. Within the ileal enterocyte, bile acids are carried by IBABP [78] and finally transported into portal venous blood by the heterodimeric basolateral transporter OSTα/OSTβ [47]. The uptake of bile acids into the hepatocytes is mediated by another basolateral transporter, NTCP [43]. The bile acids are once again re-excreted into bile. This bile acid movement cycling within the liver and intestine is named enterohepatic circulation and the total amount of bile acids distributed within the circulation is defined as the bile acid pool. The enterohepatic circulation has been comprehensively reviewed elsewhere [79, 80]. It should be emphasized the four major transporters in charge of bile acids in and out of the liver or intestine, and IBABP, the vehicle for transporting bile acids within the enterocytes, all have their expression modulated by FXR, except for ASBT. Although it has been reported that activation of FXR in the ileum downregulates ASBT expression in the human [50], mouse [51], and rabbit [52], it is not clear whether ASBT is regulated by FXR in the rat. Regulation of ASBT

CYP7A1

329

Liver NTCP

BA BSEP

Duodenum

Portal vein OST a/b

Fecal BAs

ASBT ileum

IBABP

Figure 22.2 Schematic diagram of the enterohepatic circulation. NTCP and BSEP in the liver are the major nuclear receptors responsible for bile acid uptake and excretion, while ASBT and OSTα/OSTβ in the ileum are responsible for taking in and transporting out bile acids. Approximately 5% of the bile acids in the circulating bile acid pool will be lost in feces. Bile acids will be newly synthesized via CYP7A1 to compensate for the amount of bile acid lost in the feces

expression by intestinal bile acid flux has been studied at a number of laboratories in guinea pigs [81], rats [82–86], and mice [87]. However, whether ASBT expression is positively or negatively regulated by increasing bile acid flux in the intestinal lumen remains controversial. Observations in guinea pigs [81] and mice [87] showed that ASBT was negatively regulated by the intestinal bile acid flux, while in rats ASBT was positively regulated by bile acids [82–85]. However, Arrese et al. reported [86] that in rats no regulatory response to changes in the intestinal bile acid flux was observed. Furthermore, a physiologically functional LRH-1 transcriptional binding site was identified in the mouse ASBT promoter that was not present in the rat [51]. Thus, activation of ileal FXR downregulates ASBT expression via FXR–SHP–FTF cascade in mice but not in rats. The information about effect of dietary cholesterol on regulation of bile acid reabsorption is limited. It was suggested that in rats, cholesterol feeding might interfere with bile acid reabsorption in the intestine [88]. It has been reported that 1% cholesterol feeding induced CYP7A1 mRNA levels but suppressed ASBT gene expression in mice [87]. It has been shown that there is no difference between the rabbit and rat in response of CYP7A1 promoter to induced SHP/FXR activation [34] and that the opposite response of CYP7A1 to dietary cholesterol is due to the different bile acid pool size, which causes activation of FXR in rabbits but not in rats after cholesterol feeding [73]. It is also shown that ASBT mRNA/protein increased in rabbits but not in rats after cholesterol feeding [25]. Thus, enhanced ASBT expression in cholesterol-fed rabbits is likely the key factor that induces enlarged bile acid pool size and activation of FXR, and in turn results in suppression of CYP7A1 expression in rabbits fed cholesterol. In remains unknown why dietary cholesterol induces expression of

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ASBT in rabbits but not in rats. It has been reported that peroxisome proliferator-activated receptor (PPAR) upregulates human ASBT promoter activity [89], while thus far there has been no report suggesting a candidate PPAR binding site in the rat ASBT promoter. Furthermore, an FXR binding site has been identified in the human but not in the mouse PPARα promoter [90]. The above findings suggest that regulation of ASBT expression in rabbits is likely to be similar to that in humans but will likely be quite different from that in rodents. It has been confirmed that the rabbit ASBT promoter contains a functional PPAR binding site (data unpublished). It is expected that the rabbit’s PPAR promoter may also contain a functional FXR binding site such that PPAR expression would be induced in rabbits fed cholesterol for 10 days, because the FXR ligand DCA is greatly increased in the ileum of these rabbits. It is postulated that ASBT expression would be enhanced via bile acid–FXR–PPAR–ASBT cascade in these cholesterol-fed rabbits.

MEASUREMENT OF THE BILE ACID POOL Figure 22.3 shows the rabbit model with bile fistula to collect bile continuously for measurement of hepatic bile acid outputs and the bile acid pool size. After the operation for bile duct cannulation, rabbits are gently restrained in a rabbit restrainer by two chains (about 15 cm in length); one is connected to a ring around the animal’s neck and the other to a ring around the abdomen. These two rings are made of thick and soft gauze tissue such that the animal can tolerate the restraint. The other end of these two chains is fixed onto the side of the restrainer such that the animal can eat and drink freely and move within a limited distance but cannot turn around to reach the tubing system and the plastic bag for collecting bile. This plastic bag is taped to the elastic bandage, which is wrapped around to fix the gauze that covers the abdominal wound after the surgery. A silicon cannula for the intravenous infusion line is inserted into the femoral vein and the tip of the cannula is pushed to reach the level of inferior vena cava. The continuous infusion of Lactated Ringer’s solution with 5% glucose to compensate for the body fluid lost in bile is necessary to keep the animal body fluid in equilibrium at a physiological level. The amounts of replaced solution vary from animal to animal and depend on the bile outputs during different periods of time. Generally, each animal is given 4 ml kg−1 hour−1 in the first 24 hours after the operation and 2.5–3 ml kg−1 hour−1 for five to seven days afterwards. This model has been used to collect bile in Watanabe rabbits for as long as six weeks in order to observe the effect of stimulation of bile acid synthesis on reduction of plasma cholesterol levels in these LDL receptor-deficient rabbits. In rabbits, the bile acid pool size is calculated from measurements of

The chain

Bile cannula The bag for collecting bile IV line

Figure 22.3 The rabbit model with bile fistula. After the operation, rabbits are gently restrained in a restrainer

the total recovered DCA in the bile collected during bile drainage, divided by the percentage of DCA in the initial bile sample (approximately 85–95%) collected during the first 30 minutes after construction of the bile fistula. There are different opinions on whether or not cholesterol feeding would expand the circulating bile acid pool size in rats. An important point to consider about this issue is how to measure “the functional enterohepatic bile acid pool size” that circulates through the liver because only that pool (bile acid flux) reaches hepatic FXR and plays a determining role in activation of FXR and regulation of CYP7A1. Other investigators have reported increased bile acid pool sizes when cholesterol was fed to rats [91, 92] even though CYP7A1 was stimulated. In those experiments, the entire bile acid mass, including the functional bile acid pool which circulated through the liver plus the amount of bile acid which was not absorbed and was lost in the feces, was measured and included as the bile acid pool size. However, in cholesterol-fed rats, twice as many bile acids are excreted in the feces [25] and no longer circulate through the liver to affect FXR activation. Thus, in the previous reports [91, 92], the data for the bile acid pool size did not reflect the actual amount of bile acid returned through the liver that activated FXR. After bile fistula was established and the animals reached a steady state, the biliary bile acid secretion rate fell to a new low plateau, which represented basal hepatic bile acid synthesis equaling the amount of bile acid lost in the feces (fecal bile acid outputs). Xu et al. used the “washout” method originally reported by Eriksson [93] and later by Dowling et al. [94]. The “washout” bile acid pool is the total bile acid recovered in the bile collected from the beginning till the point when the low plateau has been reached.

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Biliary bile acid outputs (mg/h)

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piece of aluminum

15 cannula for bile drainage

10 Bottle for bile collection

Functional bile acidpool

5

Basal bile acid synthesis 0 0

2

4

6

8

10

12

Bile drainage time (hours) Figure 22.4 Measurement of the functional bile acid pool size in rats. After bile drainage, the biliary bile acid outputs decrease and reach a low plateau in about 10 hours. The low plateau represents the basal hepatic bile acid synthesis (the gray area), which equals the amount of bile acid lost in feces and is not part of the functional bile acid pool (white area) returning to the liver

The contribution from baseline bile acid synthesis (gray area in Figure 22.4) is calculated as the basal hepatic bile acid synthesis rate (mg hour−1 , at the low plateau) multiplied by the total hours spent collecting the washout bile acid pool. The amount of basal bile acid synthesis equals the bile acids lost in feces that will not return to the liver. Only those bile acids that circulate through the liver regulate hepatic FXR activation and CYP7A1 expression. This amount of bile acid is defined as the functional enterohepatic bile acid pool, which is calculated as the amount of the “washout” bile acid pool corrected by subtracting the contribution due to basal bile acid synthesis. Figure 22.5 shows the rat model with bile fistula. A small piece of light and soft aluminum was fixed onto the waist of the animal by elastic bandage such that the plastic bottle used to collect bile could be attached continuously. With this device, the rats were allowed to move around freely after the surgery but could not reach the bottle for bile collection, which was behind the metal sheet. The biliary tubing to drain the bile was exteriorized and its terminal end was inserted into the bottle through a tiny hole on the cap of the bottle. The lost body fluid was replaced by subcutaneous injection of Lactated Ringer’s solution.

THE EFFECT OF BILE ACID MALABSORPTION SC-435 is a competitive inhibitor of ASBT that produces ileal bile acid malabsorption [95]. It has been reported that after one week of treatment with SC-435, FXR was

Figure 22.5 The rat model with bile fistula. A small piece of light and soft aluminum was fixed onto the waist of the animal to facilitate the plastic bottle used to collect bile

inactivated, CYP7A1activity/expression induced, and plasma cholesterol decreased in rabbits [96] and mice [97]. However, in both rabbits and mice, ASBT mRNA levels unexpectedly increased after SC-435 treatment where the absorbed bile acid flux through the ileal enterocytes decreased substantially. It is possible that ASBT transcription rose to produce more protein to compensate for the reduced function of the ASBT protein that was competitively blocked with SC-435. Nevertheless, this finding suggested that the expression of ASBT would be upregulated when the bile acid flux through the ileal enterocytes was reduced. It remains unclear whether the sensor which triggers the pathway responsible for this regulation of ASBT expression is located on the surface of the brush border membrane or within the enterocyte. If the sensor is not on the surface then increasing amounts of bile acid in the lumen (feeding of bile acids) would have only limited effect on the sensor and ASBT expression. However, it is more likely that the sensor is located both on the surface and inside the enterocytes, in which case the expression of ASBT can be regulated coordinately. It was noted in the SC-435-treated rabbits [96] that although ileal bile acid absorption was significantly decreased, which resulted in diminished bile acids returning to the liver, biliary bile acid outputs and the bile acid pool sizes were similar to baseline levels. The unchanged biliary bile acid outputs and hepatic pool sizes were associated with increased bile acid synthesis to compensate for diminished bile acid return from the ileum and suggested that after treatment with SC-435, a new hepatic balance was reached. The reduced bile acids returning to the liver were compensated for by increased newly synthesized bile acids, such that biliary bile acid outputs from the liver and circulating pool size were maintained. That bile acid pool size was not changed due to compensatory increase in bile acid synthesis has also been observed in patients ingesting cholestyramine [98] and with small-intestine resection [99].

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NEW CHALLENGES For a long time it has been believed that the bile acid flux returning through the liver is the determining factor that controls activation of hepatic FXR, which in turn mediates CYP7A1 expression/bile acid synthesis. This opinion has been challenged recently by the finding from Kliewer’s group that fibroblast growth factor (FGF15) in the mouse or FGF19 in the human is the signal induced by bile acid-activated FXR in the ileum to suppress CYP7A1 expression and bile acid synthesis [100]. They suggested that FGF15/19 plays a role for intestinal FXR in the cross-talk between intestine and liver. More recently, they reported studies in mouse models with specific deletion of FXR gene in the liver (FxrL ) or intestine (FxrIE ) [101]. FXR agonist GW4064 did not induce SHP but repressed CYP7A1 expression in the liver of FxrL mice where hepatic FXR was knocked out. In FxrIE mice where FXR is competent in the liver but deficient in the intestine, GW4064 did not inhibit CYP7A1 expression, with no significant increase of SHP mRNA in the liver. The absence of significant repression of CYP7A1 in the intestinal FXR KO mice treated with FXR agonist GW4064 suggests that activation of FXR in the intestine but not in the liver regulates CYP7A1 expression and bile acid synthesis. The authors proposed that FGF15 induced by activation of intestinal FXR is responsible for signaling from intestine to liver in the regulation of CYP7A1. However, the following evidence supports the conventional opinion that increased expression of SHP by bile acid-induced activation of FXR in the liver also downregulates CYP7A1/bile acid synthesis without involvement of FGF15/19: (i) increased FXR activation in the liver induced SHP, which blocked the recruitment of LXRα to the CYP7A1 promoter in both in vitro [34, 70] and in vivo studies [39, 71, 73]; (ii) in HepG2 cells transfected with either rabbit or rat CYP7A1 promoter, increased amounts of SHP alone or with FTF suppressed CYP7A1 promoter activity [34, 70]; and (iii) Kliewer’s group also found [33] that in human primary hepatocytes, the activation of LXRα stimulated SHP expression, which suppressed the human CYP7A1 expression. Thus, induced expression of hepatic SHP with absence of FGF19 is able to suppress CYP7A1 transcription in human hepatocytes. It remains unclear why “bile acid sensor” FXR regulates its target CYP7A1 in the intestine via FGF15 instead of bile acids that will be returning to the liver anyway. The bile acid flux through the liver should be able to provide more accurate information to the hepatic FXR in order to regulate the hepatic bile acid input (NTCP) and output (BSEP). The current work from Dawson’s lab in Ostα–/–mice [102] provides further information on the role of FGF15 in the regulation of CYP7A1. Their study showed that the bile acid pool size was significantly decreased in the Ostα

KO mice, with an almost complete blockage (∼95%) of transileal taurocholate transport. Unlike in Asbt–/–mice, where Cyp7a1 and Cyp8b1 were upregulated, in these Ostα KO mice both Cyp7a1 and Cyp8b1 expression were downregulated. This in vivo study demonstrates that the heteromeric OSTs Ostα/Ostβ play an important role in bile acid homeostasis as well as intestinal bile acid transport. It seems that Ostα/Ostβ is not only a target of FXR, but is also able to inversely regulate FXR activation through some unknown pathway. As a result, Asbt, Shp, and Ibabp expression were repressed in Ostα–/–mice. The reduced expression of Cyp7a1/Cyp8b1, Ibabp, Asbt, and the decreased bile acid pool size all resulted from the “bile acid traffic jam” caused by Ostα deficiency. The body attempted to protect against the toxicity of overloaded bile acids by establishing a new point for homeostasis. One possible candidate of the regulatory protein would be FGF15. The distal segment of the small intestine showed an increase in FGF15 expression that should have suppressed CYP7A1 expression in the liver [100, 101]. One should note that in this study, FGF15 was not induced by FXR activation, since mRNA levels of Shp and Ibabp were both reduced in the ileum, indicating inactivation of FXR. Furthermore, the fecal bile acid excretion was similar in wild-type and Ostα–/–mice. This indicates that the bile acid synthesis was not decreased in Ostα–/–mice, since daily bile acid synthesis should equal the fecal bile acid excretion. Thus, the mechanism responsible for the induction of FGF15 expression and the effect of the elevated FGF15 on Cyp7a1/bile acid synthesis in these Ostα–/–mice remains to be ascertained. Nevertheless, this important work suggests that FGF15 is a signal from the ileum to downregulate the expression of CYP7A1/CYP8B1 when the ileal enterocyte is overloaded with bile acids; the Ostα–/–mouse is a good model to show that downregulation. However, the important role of the hepatic bile acid flux in the feedback regulation of CYP7A1 to protect hepatocytes against the toxicity of overloaded bile acids in the liver cannot be replaced by FGF15/19. In case of hepatic cholestasis or bile duct obstruction, hepatic bile acids accumulated with the activation of hepatic FXR will play the critical role in the downregulation of CYP7A1 expression, whereas ileal FXR activation/FGF19 probably will have little influence on this regulation. Evolution makes some regulatory pathways more complicated. Expression of certain important genes may be regulated by redundant pathways to ensure continuation of these genes’ proper function. The involvement of FGF15/19 in downregulation of CYP7A1 is probably to protect enterocytes from being overloaded with bile acids and to coordinate with—but not to replace—the role of the hepatic bile acid flux in the regulation of CYP7A1 transcription in the liver.

22: NUCLEAR RECEPTORS REGULATE BILE ACID SYNTHESIS

THE ROLE OF CELL SIGNALING IN THE REGULATION OF CYP7A1 There is some evidence demonstrating that activated cell signaling cascades may also play an important role in the regulation of CYP7A1. Stravitz et al. reported that bile acids activate different isoforms of protein kinase C (PKC) in a time- and concentration-dependent manner [103, 104]. Recently, the same group reported that bile acids activated the JNK/c-Jun cascade in primary rat hepatocytes, which rapidly downregulated CYP7A1gene transcription, and that SHP was also a direct target of activated c-Jun [105]. Therefore, the PKC pathway and FXR-mediated downregulation of CYP7A1 may interact with each other via a common receptor, SHP. It is proposed that bile acids induce inflammatory cytokines in hepatic macrophages, which in turn suppress CYP7A1 mRNA expression in hepatocytes [106]. It is also reported that bile acids repress CYP7A1 transcription by reducing transactivation activity of HNF4α via a mitogen-activated protein kinase (MAPK) pathway, and it is proposed that phosphorylation of HNF4α by JNK may reduce HNF4α ability to activate CYP7A1 transcription [107]. However, more recently, it was suggested that induction of the p38 kinase—one of the terminal kinases of the MAPK pathway—by insulin results in an increase in HNF4α protein and a concomitant induction of CYP7A1 expression [108]. The investigators proposed that the p38 pathway is directly involved in the insulin-mediated activation of CYP7A1 expression through phosphorylation of the nuclear receptor HNF4α. The cell-signaling pathways are important for the liver’s rapid response to overloaded bile acids in the hepatocytes.

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78. Nakahara, M., Furuya, N., Takagaki, K., Sugaya, T., Hirota, K., Fukamizu, A., Kanda, T., Fujii, H. and Sato, R. (2005) Ileal bile acid-binding protein, functionally associated with the farnesoid X receptor or the ileal bile acid transporter, regulates bile acid activity in the small intestine. J Biol Chem, 280, 42283–89. 79. Hofmann, A.F. (1989) Enterohepatic circulation of bile acids, in Handbook of Physiology. Section on the Gastrointestinal System (ed. S. Schultz), American Physiological Society, Bethesda, pp. 567–96. 80. Hofmann, A.F. (2009) The enterohepatic circulation of bile acids in mammals: form and functions. Front Biosci , 14, 2584–98. 81. Lillienau, J., Crombie, D.L., Munoz, J., LongmireCook, S.J., Hagey, L.R. and Hofmann, A.F. (1993) Negative feedback regulation of the ileal bile acid transport system in rodents. Gastroenterology, 104, 38–46. 82. Higgins, J.V., Paul, J.M., Dumaswala, R. and Heubi, J.E. (1994) Downregulation of taurocholate transport by ileal BBM and liver BLM in biliary-diverted rats. Am J Physiol , 267, G501–7. 83. Dumaswala, R., Berkowitz, D. and Heubi, J.E. (1996) Adaptive response of the enterohepatic circulation of bile acids to extrahepatic cholestasis. Hepatology, 23, 623–29. 84. Stravitz, R.T., Sanyal, A.J., Pandak, W.M., Vlahcevic, Z.R., Beets, J.W. and Dawson, P.A. (1997) Induction of sodium-dependent bile acid transporter messenger RNA, protein, and activity in rat ileum by cholic acid. Gastroenterology, 113, 1599–608. 85. Sauer, P., Stiehl, A., Fitscher, B.A., Riedel, H.D., Benz, C., Kloters-Plachky, P., Stengelin, S., Stremmel, W. and Kramer, W. (2000) Downregulation of ileal bile acid absorption in bile-duct-ligated rats. J Hepatol , 33, 2–8. 86. Arrese, M., Trauner, M., Sacchiero, R.J., Crossman, M.W. and Shneider, B.L. (1998) Neither intestinal sequestration of bile acids nor common bile duct ligation modulate the expression and function of the rat ileal bile acid transporter. Hepatology, 28, 1081–87. 87. Torchia, E.C., Cheema, S.K. and Agellon, L.B. (1996) Coordinate regulation of bile acid biosynthetic and recovery pathways. Biochem Biophys Res Commun, 225, 128–33. 88. Bj¨orkhem, I., Eggertson, G. and Anderson, U. (1991) On the mechanism of stimulation of cholesterol 7αhydroxylase by dietary cholesterol. Biochim Biophys Acta, 1085, 329–35. 89. Jung, D., Fried, M. and Kullak-Ublick, G.A. (2002) Human apical sodium-dependent bile salt transporter gene (SLC10A2) is regulated by the peroxisome proliferators-activated receptor-α. J Biol Chem, 277, 30559–66. 90. Torra, I.P., Claudel, T., Duval, C., Kosykh, V., Fruchart, J.C. and Staels, B. (2003) Bile acids induce the expression of the human peroxisome proliferators-activated receptor-α gene via activation of the farnesoid X receptor. Mol Endocrinol , 17, 259–72.

22: NUCLEAR RECEPTORS REGULATE BILE ACID SYNTHESIS

91. Smit, M.J., Kuipers, F., Vonk, R.J., Temmerman, A.M., J¨ackle, S. and Windler, E.E.T. (1993) Effects of dietary cholesterol on bile formation and hepatic processing of chylomicron remnant cholesterol in the rat. Hepatology, 17, 445–54. 92. Moundras, C., Behr, S.R., R´em´esy, C. and Demign´e, C. (1997) Fecal losses of sterols and bile acids induced by feeding rats guar gum are due to greater pool size and liver bile acid secretion. J Nutr, 127, 1068–76. 93. Eriksson, S. (1957) Biliary excretion of bile acids and cholesterol in bile fistula rats: bile acids and steroids. Proc Soc Exp Biol Med , 94, 578–82. 94. Mok, H.Y.I., Perry, P.M. and Dowling, R.H. (1974) The control of bile acid pool size: effect of jejunal resection and phenobarbitone on bile acid metabolism in the rat. Gut , 15, 247–53. 95. West, K.L., Ramjiganesh, T., Roy, S., Keller, B.T. and Fernandez, M.L. (2002) 1-[4-[4[(4R,5R)-3,3-Dibutyl7-(dimethylamino)-2,3,4,5-tetrahydro-4-hydroxy-1,1dioxido-1-benzothiepin-5-yl]phenoxy]butyl]-4-aza-1azoniabicyclo[2.2.2]octane methanesulfonate (SC-435), an ileal apical sodium-codependent bile acid transporter inhibitor alters hepatic cholesterol metabolism and lowers plasma low-density lipoprotein-cholesterol concentrations in guinea pigs. J Pharmacol Exp Ther, 303, 293–99. 96. Li, H., Xu, G., Shang, Q., Pan, L., Shefer, S., Batta, A.K., Bollineni, J., Tint, G.S., Keller, B.T. and Salen, G. (2004) Inhibition of ileal bile acid transport lowers plasma cholesterol levels by inactivating hepatic FXR and stimulating CYP7A1. Metabolism, 53, 927–32. 97. Bhat, B.G., Rapp, S.R., Beaudry, J.A., Napawan, N., Butteiger, D.N., Hall, K.A., Null, C.L., Luo, Y. and Keller, B.T. (2003) Inhibition of ileal bile acid transport and reduced atherosclerosis in apoE-/- mice by SC-435. J Lipid Res, 44, 1614–21. 98. Garbutt, J.T. and Kenney, T.J. (1972) Effect of cholestyramine on bile acid metabolism in normal man. J Clin Invest ., 51, 2781–89. 99. Poley, J.R. and Hofmann, A.F. (1976) Role of fat maldigestion in pathogenesis of steatorrhea in ileal resection. Fat digestion after two sequential test meals with and without cholestyramine. Gastroenterology, 71, 38–44. 100. Inagaki, T., Choi, M., Moschetta, A., Peng, L., Cummins, C.L., McDonald, J.G., Luo, G., Jones, S.A., Goodwin, B., Richardson, J.A., Gerard, R.D., Repa, J.J.,

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Mangelsdorf, D.J. and Kliewer, S.A. (2005) Fibroblast growth factor 15 functions as an enterohepatic signal to regulate bile acid synthesis. Cell Metab, 2, 217–25. Kim, I., Ahn, S., Inagaki, T., Choi, M., Ito, S., Guo, G.L., Kliewer, S.A. and Gonzalez, F.J. (2007) Differential regulation of bile acid homeostasis by the farnesoid X receptor in liver and intestine. J Lipid Res, 48, 2664–72. Rao, A., Haywood, J., Craddock, A.L., Belinsky, M.G., Kruh, G.D. and Dawson, P.A. (2008) The organic solute transporter α-β, Ostα -Ostβ, is essential for intestinal bile acid transport and homeostasis. Proc Natl Acad Sci U S A, 105, 3891–96. Stravitz, R.T., Vlahcevic, Z.R., Gurley, E.C. and Hylemons, P.B. (1995) Repression of cholesterol 7α-hydroxylase transcription by bile acids is mediated through protein kinase C in primary cultures of rat hepatocytes. J Lipid Res, 36, 1359–68. Stravitz, R.T., Rao, Y.P., Vlahcevic, Z.R., Gurley, E.C., Jarvis, W.D. and Hylemon, P.B. (1996) Hepatocellular protein kinase C activation by bile acids: implications for regulation of cholesterol 7α-hydroxylase. Am J Physiol , 271, G293–303. Gupta, S., Stravitz, R.T., Dent, P. and Hylemon, P.B. (2001) Down-regulation of cholesterol 7α-hydroxylase (CYP7A1) gene expression by bile acids in primary rat hepatocytes is mediated by the c-Jun N-terminal kinase pathway. J Biol Chem, 276, 15816–22. Miyake, J.H., Wang, S.L. and Davis, R.A. (2000) Bile acid induction of cytokine expression by macrophages correlates with repression of hepatic cholesterol 7α-hydroxylase. J Biol Chem, 275, 21805–8. De Fabiani, E., Mitro, N., Anzulovich, A.C., Pinelli, A., Galli, G. and Crestani, M. (2001) The negative effects of bile acids and tumor necrosis factor-α on the transcription of cholesterol 7α-hydroxylase gene (CYP7A1) converge to hepatic nuclear factor-4. A novel mechanism of feedback regulation of bile acid synthesis mediated by nuclear receptors. J Biol Chem, 276, 30708–16. Xu, Z., Tavares-Sanchez, O.L., Li, Q., Fernando, J., Rodriguez, C.M., Studer, E.J., Pandak, W.M., Hylemon, P.B. and Gil, G. (2007) Activation of bile acid biosynthesis by the p38 mitogen-activated protein kinase (MAPK): hepatocyte nuclear factor-4α phosphorylation by the p38 MAPK is required for cholesterol 7α-hydroxylase expression. J Biol Chem, 282, 24607–14.

23

The Function of the Canalicular Membrane in Bile Formation and Secretion Ronald P.J. Oude Elferink and Coen C. Paulusma AMC Liver Center, Academic Medical Center, Amsterdam, The Netherlands

THE MECHANISM OF BILE FORMATION Bile formation is an osmotic process; solutes are actively transported into the canaliculus by primary active transporters. The most important solutes driving bile formation are bile salts. These are pumped into primary bile at a concentration of about 20 mM. The plasma bile salt concentration is 1000-fold lower. Bile salts are anions, so an equal concentration of cations (Na+ ) needs to enter primary bile (mainly through the tight junctions). On top of this, there is a substantial flux of glutathione into bile (about 1–5 mM), which also needs to be accompanied by an equal flux of cations. Finally, the bile duct epithelial cells secrete bicarbonate. These solutes together create a hyperosmolarity of at least 35 mOsm, which attracts water through tight junctions and via aquaporins in the plasma membranes of hepatocytes and bile duct epithelial cells (aquaporins 0, 8, and 9) [1, 2] Various ATP-binding cassette (ABC) transporters in the canalicular membrane mediate the secretion of the main biliary solutes. The bile salt export pump (BSEP; encoded by the ABCB11 gene) pumps bile salts across the canalicular membrane. The ABC transporter MRP2 (encoded The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

by the ABCC2 gene) transports glutathione across the canalicular membrane. The mechanism of secretion of bicarbonate has not been fully elucidated yet, but this is thought to be mediated by the chloride/bicarbonate exchanger AE2 in the apical membrane of cholangiocytes [3]. Diseases associated with mutations in the various transporter genes involved in bile formation are discussed in Chapter 45. Figure 23.1 depicts the main transporter proteins important in bile formation, and their substrates. Bile formation serves two important functions: on the one hand it represents a major route for the elimination of drugs, toxins, and waste products; on the other hand bile formation ensures the secretion of bile salts, which are crucial for lipid emulsification and subsequent lipid absorption in the intestine.

TRANSPORT OF DRUGS, TOXINS, AND WASTE PRODUCTS INTO BILE Uptake of xenobiotics and waste products into the hepatocyte is mediated by a number of transporters, including members of the organic anion transporter protein (OATP)

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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OATPs

OATs

Bile salts Na +

OCTs

NTCP (SLC10A1)

MRP2 (ABCC2)

Drugs, toxins, and waste products

BCRP (ABCG2)

MDR1 Pgp (ABCB1)

BSEP (ABCB11) Bile salts

ABCG5/8 cholesterol Bile salts and lipids

FIC1 (ATP8B1) phosphatidylserine

MDR3 Pgp (ABCB4) phosphatidylcholine

Figure 23.1 Canalicular and basolateral transport proteins important for bile formation. The figure gives the name of each canalicular transporter, the trivial name of its gene product between brackets, the direction of transport, and the (putative) substrate

(SLC21A) [4], organic anion transporter (OAT), and organic cation transporter (OCT) (SLC22) [5] families. The driving force for this uptake has not been firmly established, but it has been postulated that various rat Oatps mediate uptake of compounds by countertransport against glutathione. It is also possible that these transporters do not mediate active transport but merely facilitated diffusion. The substrate specificity of these transporters is extremely broad and there is strong overlap between various members of the transporter families. They transport xenobiotics and their (conjugated) metabolites, but also endogenous compounds such as steroids and their metabolites, and other endogenous compounds such as opioid peptides [6], thyroid hormones [7], and possibly also bilirubin (although the latter is a controversial issue [8, 9]). The importance of the OATPs as a crucial step in drug elimination was illustrated by the recent finding of a strong association between a frequent polymorphism in OATP1B1 and simvastatin-induced myopathy [10]. Canalicular excretion of drugs, toxins, and waste products is mediated by at least three ABC transporters: MDR1 P-glycoprotein (Pgp) (encoded by ABCB1 ), MRP2 (encoded by ABCC2 ), and BCRP (encoded by ABCG2 ) [11]. MDR1 Pgp transports mainly neutral and cationic amphipaths, whereas MRP2 and BCRP have preference for anionic and neutral amphipaths and their conjugates. In addition, MRP2 transports glutathione (reduced and oxidized) and glutathione conjugates. An important endogenous substrate for MRP2 is conjugated (glucuronidated) bilirubin. Patients with the Dubin–Johnson syndrome have mutations in the ABCC2 gene and characteristically have a conjugated hyperbilirubinemia. Because

conjugated bilirubin (in contrast to unconjugated bilirubin) is not toxic, these patients do not suffer from clinical symptoms [12]. Knockout (KO) mice for these three ABC transporters do not display an endogenous phenotype, which might suggest that they do not have a crucial function in drug elimination. It has been demonstrated, however, that challenging these animals with certain drugs can induce major toxicity. Thus, oral doses of the antihelminthic drug ivermectin that were harmless to wild-type mice induced lethal neurotoxicity in Mdr1 Pgp-deficient mice [13]. It could be shown that the major function of Mdr1 Pgp is in the blood–brain barrier and to a lesser extent in the liver and intestine. A second example is the toxicity induced by pheophorbide in Abcg2 −/− mice [14]. This natural fermentation product of chlorophyll caused phototoxic lesions when administered to these mice, whereas it was completely harmless in wild-type animals. Pheophorbide is kept out of the body by Bcrp activity in both liver and intestine. For several of the canalicular transporters it is clear that their simultaneous expression in the hepatocyte and the enterocyte creates a highly efficient system of keeping unwanted compounds out of the systemic circulation [15]. This concept is also clearly illustrated by the phenotype of sitosterolemia patients and the corresponding mouse model, the Abcg5/8 −/− mouse. Abcg5/8 is an ABC transporter for plant sterols and cholesterol. The functional transporter is formed by heterodimerization of the two half-transporters Abcg5 and Abcg8 (see below). Plant sterol levels in plasma of humans and mice lacking this transporter are 10- to 300-fold increased [16]. Ingested plant sterols that diffuse into enterocytes are efficiently

23: THE FUNCTION OF THE CANALICULAR MEMBRANE IN BILE FORMATION AND SECRETION

pumped back into the gut lumen. As a consequence, only 5% of ingested plant sterols are absorbed (as opposed to more than 50% of ingested cholesterol) [17]. The small amount of plant sterol that does pass this gatekeeper system enters the liver via portal blood and is efficiently extruded into bile by the same transporter, preventing spillover of significant amounts of plant sterol into systemic blood. Clearly, absence of ABCG5/G8 in both tissues leads to a major increase in systemic plant sterol levels. This concept holds not only for the sterol transport by ABCG5/G8 but for all other drug transporters in the canalicular membrane. ABCC2 (MRP2), ABCG2 (BCRP), and ABCB1 (MDR1 Pgp) are all expressed in both hepatocytes and enterocytes [15]. ABCC2 is primarily expressed in the small intestine [18], while ABCG2 is expressed throughout the gut [19]. In the absence of these transporters there is enhanced absorption of drugs and toxins in the gut, leading to enhanced systemic concentrations of drugs and toxins [14, 20, 21].

BILIARY BILE SALT EXCRETION Bile salts are highly valuable compounds; they are synthesized from cholesterol in a complex enzymatic pathway (see Chapter 21). It is therefore not surprising that the body salvages bile salts very efficiently. After canalicular secretion via BSEP and delivery of bile to the intestine, more than 95% of secreted bile salts are reclaimed by uptake via the transporter apical sodium bile salt transporter (ASBT; encoded by SLC10A2 ) into enterocytes of the terminal ileum. The enterocytes subsequently secrete bile salts into portal blood via the transporter couple organic solute transporter (OST) α/β [22–24]. Hepatocytes filter the bile salts from portal blood by uptake via the sodium-dependent taurocholate transporter NTCP (SLC10A1 ), which completes the enterohepatic cycle of bile salts. As also discussed in Chapter 21, various types of bile salt exist with different physicochemical characteristics. Inherent to their function, however, is that all bile salts are detergents capable of dissolving (dietary) lipids. Consequently, bile salts are also able to solubilize phospholipids from the plasma membrane. Hence, bile salts are quite cytotoxic agents and the cells lining the biliary tree (i.e. hepatocytes and cholangiocytes) have developed several mechanisms of protection against this toxicity. In the last decade considerable progress has been made in elucidating these protective mechanisms, which will be discussed in more detail below.

BILIARY PHOSPHOLIPID EXCRETION The importance of canalicular lipid transport in the protection against bile salts has been elucidated mainly

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by unraveling several rare cholestatic diseases in which lipid transporter genes are mutated. This class of diseases is called progressive familial intrahepatic cholestasis (PFIC) type 1–3 and clinical aspects will be discussed in Chapter 45. Two forms of PFIC, type 1 and type 3, are caused by defects in the canalicular lipid translocators ATP8B1 and ABCB4, respectively. Apparently, lipid asymmetry of the canalicular membrane is essential for normal physiology. This is not very surprising because more than any other membrane in the body, the canalicular membrane has to withstand very high detergent concentrations. Bile salts, excreted via ABCB11 (BSEP), reach concentrations in the canaliculus well above the critical micellar concentration. This represents a condition which in principle leads to solubilization of the membranes, followed by immediate cell death. One mechanism of protection against this bile salt-mediated membrane solubilization is the excretion of phospholipid to form mixed micelles with bile salts. Addition of phospholipids to simple bile salt micelles reduces the capacity of these micelles to take up more phospholipid from the membrane. This organized phospholipid excretion is mediated by ABCB4 (MDR3 Pgp) in humans (and the orthologous Abcb4 (Mdr2 Pgp) in mice) [25, 26]. This protein mediates translocation of phosphatidylcholine (PC) from the inner to the outer leaflet of the canalicular membrane and hence is termed a floppase (Figure 23.2). The mechanism of the subsequent step, extraction from the membrane, is largely unknown. It has been known for many years that phospholipid excretion is driven by micelle-forming bile salts. Bile salt micelles may take up this translocated PC directly from the transporter Abcb4. Alternatively, the translocated PC may first be inserted into the outer leaflet of the canalicular membrane, from which it is subsequently extracted by bile salt micelles. It is also possible that both mechanisms of PC excretion occur simultaneously. Whichever mechanism is used for the extraction of phospholipids into bile, it has proven to be essential for the protection against bile salts. Mice with a disruption of the Abcb4 (Mdr2 ) gene (Abcb4 −/− mice) develop progressive liver disease [27, 28], while a more severe form of this disease occurs in patients with PFIC3 in which the ABCB4 (MDR3 ) gene is mutated [29–31]. The Abcb4 −/− mouse has proven to be a very instructive model for our understanding of the role of phospholipid excretion in protection of the liver. The toxic bile leads to a strong increase in hepatocyte turnover due to increased apoptosis. Abcb4 −/− mice also develop hepatocellular carcinoma after about eight months of life [27]. Not only hepatocytes but also cholangiocytes are particularly affected by the toxic phospholipid-free bile. Disruption of the tight junctions and basement membranes of the small bile ducts leads to bile leakage and a subsequent inflammatory response [32]. More recent reports suggest that mutations in ABCB4 are not only associated with the severe pediatric disease PFIC3, but also with adult

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floppases

flippase

ABCB4 MDR3

ABCG5 ABCG8

ATP8B1 FIC1

phosphatidylcholine

sterols (plant sterols + cholesterol)

phosphatidylserine

Figure 23.2 The different types of lipid translocator in the canalicular membrane of the hepatocyte.While ABCB4 (MDR3 P-glycoprotein) and ABCG5/G8 mediate outward translocation (“flopping”) of phosphatidylcholine (PC) and cholesterol, respectively, ATP8B1 (FIC1) mediates inward translocation (“flipping”) of phosphatidylserine

syndromes such as intrahepatic gallstones, cholangiopathy, cirrhosis, and intrahepatic cholestasis of pregnancy [33–36]. The relation between gallstone development and impaired phospholipid secretion is understandable since mixed micelles of bile salts with phospholipid are much more capable of solubilizing cholesterol than simple bile salt micelles.

BILIARY CHOLESTEROL EXCRETION Not only PC but also substantial amounts of cholesterol are secreted into bile in a bile salt-dependent fashion. This secretion depends on the function of the two ABC half-transporters, ABCG5 and ABCG8, which form an active heterodimer. Biliary cholesterol excretion in mice with disrupted Abcg5 and Abcg8 genes is reduced by about 80% [37, 38]. Similar observations have been made in mice with a disruption of only the Abcg8 gene [39, 40] or only the Abcg5 gene [41], indicating that an obligate heterodimer of Abcg5 and Abcg8 forms a functional ABC transporter for cholesterol excretion into bile [42]. These two half-transporters are expressed in both liver and intestine. In the intestine they primarily mediate the extrusion of plant sterols from enterocytes, but they may also limit cholesterol absorption. Patients with sitosterolemia, who have a mutation in either of the two half-transporter genes, do not only have increased plasma plant sterol levels but also suffer from diet-induced hypercholesterolemia [43]. In the liver the transporter couple appears to transport primarily cholesterol, because the plant sterol levels in liver are very low. Indeed, it has been demonstrated that certain polymorphisms, such as the D19H polymorphism in ABCG8, correlate with the formation of gallstones, suggesting that this polymorphism gives rise to enhanced activity [44]. The secretion of cholesterol appears not to play a role in the protection of membranes against bile salt toxicity: patients with sitosterolemia as well as Abcg5/g8 KO animals do not suffer from any form of liver damage [37, 39, 45].

Although actual translocation of cholesterol by the transporter has not been demonstrated, it may be assumed on the basis of sequence homology that the Abcg5/g8 protein complex represents a transporter. Hence, the minimum hypothesis must be that Abcg5/g8 translocates cholesterol from the inner to the outer leaflet of the membrane and represents another floppase (Figure 23.2). It can be questioned, however, whether the translocation of cholesterol from the inner to the outer leaflet is the essential step in this process. It is known that spontaneous flip-flop of cholesterol across the membrane is much faster than that of phospholipids [46], and this was in fact for a long time an argument to assume that no transporter is required for biliary cholesterol excretion. In line with this contention, biliary cholesterol excretion is not completely absent in Abcg5/g8 KO mice, but reduced to about 20% of wild-type levels [37–39], suggesting that in the absence of the transporter a significant amount of cholesterol still reaches the outer leaflet of the canalicular membrane and can be extracted from it. This is in stark contrast to the Abcb4 −/− mouse, which completely lacks phospholipids in bile even when highly hydrophobic bile salts are infused [47]. In the case of PC, translocation is required, and in the absence of Abcb4 there will be little if any PC in the outer leaflet. Nevertheless, the Abcg5/g8 transporter is apparently necessary for the bulk excretion of cholesterol, since in its absence cholesterol secretion is impaired. Two lines of reasoning should be considered here. First, the transporter might be important for the supply of cholesterol to the outer leaflet only under conditions of increased flux; that is, during high bile salt excretion rates, when cholesterol excretion should be high as well (floppase function). Second, it may be that simple extraction of cholesterol from the canalicular membrane by bile salts is not possible. It may be expected that cholesterol molecules are buried in the layer of sphingomyelin molecules, which are abundantly present in the outer leaflet of the canalicular membrane (see below) [48]. This increases the activation energy required for extraction to the extent that bile salts cannot extract cholesterol at body temperature. This possibility was already raised by Small

23: THE FUNCTION OF THE CANALICULAR MEMBRANE IN BILE FORMATION AND SECRETION

[49], who suggested that ABCG5/G8 might represent a “liftase,” an activity that reduces the activation energy, enabling bile salt-dependent extraction of cholesterol. The floppase and liftase models are certainly not mutually exclusive. Thus, in addition to translocating cholesterol from the inner to the outer leaflet of the membrane bilayer, the transporter may expose the cholesterol molecule from the bilayer sufficiently to allow transfer to acceptor bile salt micelles.

THE FLIPPASE FUNCTION OF ATP8B1 As discussed in Chapter 45, patients with PFIC1 have mutations in the ATP8B1 gene and suffer from chronic and progressive cholestasis. The question arises, what is the molecular function of ATP8B1 and how does the absence of this function cause cholestasis? ATP8B1 is a member of the type 4 subfamily of the P-type ATPase superfamily (P4 ATPase), harboring 14 members that are supposed to be phospholipid flippases [50]. ATP8B1 (formerly called FIC1) is localized in the canalicular membrane and was proposed to flip phosphatidylserine (PS) from the outer to the inner leaflet of this membrane [51]. We have recently demonstrated that ATP8B1, when expressed alone (in CHO cells), does not give rise to flippase activity in the plasma membrane because the protein is retained in the endoplasmic reticulum (ER). Work in yeast has demonstrated that P4 ATPases require interaction with chaperones, termed Cdc50 proteins, to be properly targeted to their cellular destination. In line with these observations, we found that ATP8B1 physically interacts with the human CDC50A and CDC50B proteins and that co-expression of ATP8B1 with either CDC50A or CDC50B leads to trafficking of the complex to the plasma membrane. Under this condition we could show that ATP8B1 is capable of flipping natural PS from the outer leaflet to the inner leaflet of the plasma membrane [52]. We have also demonstrated that bile salt-infused Atp8b1-deficient mice had significant amounts of PS in their bile (in contrast to wild-type mice, which have no PS in bile) [53]. Hence, it may be concluded that ATP8B1 is a PS flippase in the canalicular membrane (Figure 23.2) that contributes to maintaining a proper asymmetric distribution of phospholipids.

MEMBRANE ASYMMETRY CREATES RESISTANCE AGAINST BILE SALTS Phospholipid asymmetry is maintained in the plasma membranes of cells in all eukaryotes. In general, PC and sphingolipids are concentrated in the outer leaflet of the plasma membrane, whereas phosphatidylethanolamine

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(PE) and particularly PS are concentrated in the inner leaflet of the membrane. Most likely, cholesterol is more or less in equilibrium between the two membrane leaflets. This asymmetry serves many functions; a prominent example is the recognition of apoptotic cells by the exposure of PS on the outer leaflet of the membrane. The phospholipid asymmetry in the canalicular membrane may be particularly high and serve to protect the membrane against the detergent action of bile salt micelles. In vitro experiments have demonstrated that the combination of sphingolipids and cholesterol is the only way by which membranes can be rendered virtually detergent-insoluble [54]. Although direct proof that this mechanism is responsible for the detergent resistance of the canalicular membrane is lacking, the observed high sphingomyelin content of these membranes isolated from various species makes this hypothesis plausible [55–57]. Considerable information on detergent (in)solubility has come from studies on lipid rafts in biological membranes. Rafts are thought to play an important role in membrane protein and lipid trafficking, as well as cell signaling [48]. Lipid rafts contain specific membrane proteins such as glycosylphosphatidylinositol (GPI)-anchored proteins and have a high content of sphingolipids and cholesterol. At body temperature, membranes composed of glycerophospholipids (PC, PS, PE) are in the so-called “liquid disordered” phase (ld ), in which the lipids are rather loosely packed. The loose packing is mainly caused by the high content of unsaturated fatty acyl chains in glycerophospholipids. Such lipid bilayers are highly prone to intercalation of detergents and subsequent solubilization. However, the addition of sphingolipids and cholesterol to membranes induces a much more rigid membrane structure that is in the so-called “liquid ordered” phase (lo ). This phase represents tighter packing of lipids, mainly due to the long saturated fatty acyl chains of sphingomyelins, which combine quite well with the flat structure of the cholesterol molecule. The behavior of these constituting lipids leads to partial phase separation of the glycerophospholipids plus cholesterol on the one hand, and sphingomyelin and (a larger fraction of) cholesterol on the other hand, the first being in the ld phase and the second being in the lo phase [48, 58]. These phases coexist in the membrane and are most likely in a dynamic equilibrium. Membranes consisting of lipids forming an lo phase were found to be resistant to the detergent Triton X-100 [48]. By inference, rafts of similar composition in biological membranes are expected to have the same detergent resistance. The isolation of rafts as pre-existing structures from membranes by virtue of their detergent resistance is controversial [58]; it is unlikely that rafts are static structures that can be isolated by this highly perturbing method (i.e. Triton X100 extraction). Nevertheless, these experiments demonstrate that certain lipid compositions can render the membrane detergent-resistant and that this resistance depends on the extent to which the constituting lipids are in the lo phase.

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0,0

1,0

Canalicular membrane Basolateral membrane

PO PC of cti on

0,6

ol

Mo

0,4

ter les

ho fc

le fra

0,6

Lo

no

0,4

o cti

ra

f le

Mo

0,8

0,2

Ld + Lo

0,8

0,2 Ld + Lo + So

Ld

1,0 0,0

0,2

Ld + So 0,4

0,6

0,8

0,0 1,0

Mole fraction of P-SM

Figure 23.3 Phase diagram for the (co-)existence of the liquid disordered (ld ), the liquid ordered (lo ), and the solid ordered (so ) phases in membranes with various relative compositions of POPC, P-SM, and cholesterol. POPC, phosphatidylcholine (containing one palmitic and one oleic fatty acyl chain); P-SM, sphingomyelin with a palmitic acyl chain. The solid square represents the composition of the basolateral membrane and the solid circle the composition of the canalicular membrane as purified and analyzed by Nibbering et al. [60]. The arrow indicates the direction in which the canalicular membrane composition approximately changes if corrected for contaminating other membranes (mainly basolateral membranes and ER membranes). The broken arrow indicates the phospholipid composition of the canalicular outer membrane leaflet, taking into consideration that sphingomyelin is exclusively present in the outer leaflet. The upper (faint) part of the diagram represents relative cholesterol contents (>66%) at which cholesterol is not soluble in the membrane. This figure was taken and modified from [48, 59]

Using data obtained by different analytic methodologies, De Almeida et al. [59] constructed a phase diagram for the coexistence of the different phases in artificial membranes composed of various ratios of palmitoyl-oleoyl-phosphatidylcholine (POPC), palmitoylSM (P-SM), and cholesterol (Figure 23.3). Analyses of these membranes were performed at 23◦ C but extrapolated to a similar phase diagram at 37◦ C. This diagram reveals that at physiological membrane lipid compositions there is indeed coexistence of lo and ld phases. A “solid ordered” (so ) phase is recognized as well (also referred to as “gel phase”), which represents an even more rigid membrane structure, but this phase is not thought to occur in biomembranes. The diagram shows that the most important determinant of the lo phase is cholesterol. In membranes consisting of POPC and cholesterol, less than 50% cholesterol yields a membrane with coexisting lo and ld phases, while at 50% cholesterol the membrane is entirely in the lo phase. When increasing amounts of P-SM are added to the system, the required fraction of cholesterol to induce a complete lo phase drops from 50% (at 30% P-SM) to 35% (at 60% P-SM). The fact that the area of complete lo (phase) is largely above that of ld + lo demonstrates that the main determinant of the lo phase is cholesterol. Thus, upward movement in the diagram (which corresponds to increasing the cholesterol content) brings membranes closer to the area of entire lo phase. It must be emphasized that there is no sudden

change in the detergent resistance of membranes when the lines between the areas in the phase diagram are passed. Rather, there is a gradual increase in the fraction of the membrane that is in the lo phase with increasing cholesterol content, and this becomes complete when the line is passed. Extrapolation of these data to the canalicular membrane is highly speculative but provides important insights. Firstly, POPC and P-SM are quantitatively important constituents of the canalicular membrane [56, 60, 61], which means that in terms of bulk lipid, this phase diagram can be applied to this membrane. Evidently, the presence of proteins probably strongly influences the behavior of these phases, but this has not been studied. If we plot into this diagram the composition of hepatic basolateral (solid square) and canalicular (solid circle) membranes isolated from mice [60], it will be clear that the canalicular membrane is much nearer the border of an entire lo phase. Importantly, canalicular membrane preparations are not at all pure and the main contaminants are basolateral membranes and membranes from the ER. The basolateral membrane contains less SM and less cholesterol than canalicular membranes; ER membranes contain little if any cholesterol. Hence, the real canalicular membrane composition (as opposed to contaminated canalicular membranes as they are isolated) will be shifted more to the right (higher SM content) and more upward (higher cholesterol content) as indicated by the arrow, and thus

23: THE FUNCTION OF THE CANALICULAR MEMBRANE IN BILE FORMATION AND SECRETION

closer to the area of entire lo phase. This is even more the case if one considers that the main determinant of detergent resistance will be the outer leaflet, since this is the leaflet exposed to high bile salt concentrations. It is well known that SM is not equally distributed over the two leaflets but is almost exclusively present in the outer leaflet [62]. Hence, the relative SM content of the outer leaflet will be about twice as high (broken arrow). This means that the outer leaflet composition of the canalicular membrane may well be in or near the lo phase area of the diagram and thus be resistant toward detergent. Analysis of detergent resistance has mainly been performed with Triton X-100 and some other detergents. Unfortunately, elaborate studies of this kind have not been performed with bile salts. Some studies do, however, suggest that a similar behavior does occur with bile salts. Hofmann et al. [63] studied induction of liposome leakage by chenodeoxycholate (CDCA) and observed that addition of cholesterol to PC liposomes (1 : 1) caused a twofold reduction in leakage. Pure SM liposomes were in fact more sensitive to CDCA than pure PC liposomes. However, combination of the three lipids (SM/cholesterol/PC = 1 : 3 : 3) reduced the sensitivity by a factor 4.8 (compared to pure PC) or even a factor 35 (compared to pure SM) [63]. A similar behavior with respect to the induction of leakage in liposomes, composed of these three components, was observed by Moschetta et al. using taurocholate as the detergent [64]. These studies suggest a similar resistance of membranes containing SM and cholesterol toward bile salts to that observed with Triton X-100. The outer-leaflet composition of the canalicular membrane will be continuously (partly) dissipated by various processes, such as PC extraction, fusion of exocytic vesicles, and spontaneous flip-flop of other phospholipids. Exocytic vesicles may have a less asymmetric membrane composition than the canalicular membrane, and in addition the fusion process will lead to a certain extent of scrambling. The net effect of this process may be an increase of glycerophospholipids in the outer leaflet. As discussed above, ATP8B1 is localized in the canalicular membrane and functions as an (inward) flippase for PS. According to the arguments given above, ATP8B1 contributes to detergent resistance of the canalicular membrane by keeping the glycerolipid PS on the inside. In line with this, we observed that mice lacking Atp8b1 excrete much higher amounts of cholesterol as well as canalicular ectoenzymes into bile than wild-type mice [53]. The enhanced biliary cholesterol excretion in Atp8b1-deficient mice turned out to be independent of the cholesterol transporter Abcg5/g8, since Atp8b1-deficient mice with inactivated Abcg5/g8 also displayed enhanced cholesterol output [65]. The question arises, how does this phenotype relate to cholestasis? We have observed that the activity of Bsep in membrane preparations is critically dependent on the cholesterol content of these membranes. Plasma membranes isolated from normal mice were treated with

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increasing amounts of methyl-β-cyclodextrin, which selectively depleted these membranes of cholesterol. We observed that there was an inverse linear relationship between the cholesterol content of the membranes and the measured Bsep activity [66]. From these observations we hypothesize that in the absence of Atp8b1-mediated PS-flipping the canalicular membrane is less resistant toward bile salts; this leads to inappropriately high cholesterol extraction rates, which in turn may reduce the cholesterol content of the canalicular membrane. The latter will reduce the activity of Bsep and thus give rise to impaired bile salt transport and cholestasis.

CONCLUSION It will be clear that the phenotypical expression of mutations in canalicular transporter genes, both in patients and in animal models, has given us a tremendous insight into the function of the canalicular membrane. Flippases and floppases play a crucial role in the biliary lipid handling by mediating the actual transport required for excretion (ABCB4 and ABCG5/G8), but also by maintaining the membrane in a detergent-resistant state (ATP8B1) (Plate 23.1). The complete molecular mechanism of bile formation has not been elucidated yet; questions remain about the actual mechanism of lipid extraction by bile salts. It is at present especially unclear whether cholesterol and phospholipids are extracted from the membrane by bile salt micelles, or whether bile salt micelles accept the lipids directly from the transporters. Also, it remains unclear what the exact delivery mechanism of lipids to the canalicular membrane is. Is there direct supply to the canalicular membrane of vesicles containing lipids that are destined to be excreted into bile, or are biliary lipids obtained from the entire inner leaflet of the plasma membrane? Since inner-leaflet lipids can freely diffuse through tight junctions (unlike outer-leaflet lipids) [67], PC and cholesterol may laterally diffuse into the canalicular membrane inner leaflet from the entire plasma membrane. Finally, it will be important to dissect which regulatory and/or inhibitory mechanisms play a role in impaired biliary excretion during various states of cholestasis. Although many studies have been performed in animal models, it remains to be demonstrated whether similar mechanisms play a role in cholestatic patients.

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is independent of hepatocyte canalicular membrane lipid composition: a study in the diosgenin-fed rat model. J Hepatol , 35, 164–69. Coleman, R. and Rahman, K. (1992) Lipid flow in bile formation. Biochim Biophys Acta, 1125, 113–33. Van Meer, G. and Lisman, Q. (2002) Sphingolipid transport: rafts and translocators. J Biol Chem, 277, 25855–58. Hofmann, M., Schumann, C., Zimmer, G., Henzel, K., Locher, U. and Leuschner, U. (2001) LUV’s lipid composition modulates diffusion of bile acids. Chem Phys Lipids, 110, 165–71. Moschetta, A., Frederik, P.M., Portincasa, P., Vanberge-Henegouwen, G.P. and Van Erpecum, K.J. (2002) Incorporation of cholesterol in sphingomyelinphosphatidylcholine vesicles has profound effects on detergent-induced phase transitions. J Lipid Res, 43, 1046–53. Groen, A., Kunne, C., Jongsma, G., van den Oever, K., Mok, K.S., Petruzzelli, M., Vrins, C.L., Bull, L., Paulusma, C.C. and Oude Elferink, R.P. (2008) Abcg5/8 independent biliary cholesterol excretion in Atp8b1-deficient mice. Gastroenterology, 134, 2091–100. Paulusma, C.C., de Waart, D.R., Kunne, C., Mok, K.S. and Oude Elferink, R.P.J. (2009) Activity of the bile salt export pump (ABCB11) is critically dependent on canalicular membrane cholesterol content. J Biol Chem, 284, 9947–9954. Van Meer, G., Gumbiner, B. and Simons, K. (1986) The tight junction does not allow lipid molecules to diffuse from one epithelial cell to the next. Nature, 322, 639–41.

24

Apical Recycling of Canalicular ABC Transporters Yoshiyuki Wakabayashi1 and Irwin M. Arias2,3,4 1

Unit of Cellular Polarity, Cell Biology and Metabolism Program, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, MD, USA 2 Senior Scientist, National Institutes of Health, Bethesda, MD, USA 3 Emeritus Professor of Physiology, Tufts University School of Medicine, Boston, MA, USA 4 Visiting Professor of Medicine, Albert Einstein College of Medicine, Bronx, NY, USA

INTRODUCTION The bile canalicular membrane of the mammalian hepatocyte contains several primary active transporters which couple ATP hydrolysis to the transport of specific substrates into the bile canaliculus (BC) [1–4]. These transporters are members of the super family of ATP-binding cassette (ABC) transporters [5] and currently include BSEP (ABCB11) for bile acid, MDR3 (ABCB4) for phosphatidylcholine translocation, MDR1 (ABCB1) for organic cations, MRP2 (ABCC2) for non-bile acid organic anions, ABCG5/G8 for sterol, and BCRP (ABCG2) for organic anions and sulfate conjugates. Genetic studies have revealed that recessively inherited hepatobiliary phenotypes result from mutations in specific transporter genes: defects in ABCB11 produce PFIC type 2; defects in ABCB4 produce PFIC-3; defects in ABCC2 produce Dubin–Johnson syndrome; and defects in ABCG5/G8 produce sitosterolemia. Thus canalicular ABC transporters The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

play a key role in bile formation. The number of individual ABC transporters in the canalicular membrane is regulated by the physiological demand to secrete bile acids [6–8]. Insufficient numbers of ABC transporters or their malfunction in the canalicular membrane impair bile formation and can result in cholestasis. Regulation of the number of individual transporters in the apical membrane has been described in different cells. Examples include regulation of glucose transporters (GLUT-4) in adipose cells and muscle cells by insulin [9, 10]; sodium-d-glucose cotransporter (SGLT1) in enterocytes by extracellular D-glucose [11, 12]; H+ -K+ -ATPase in gastric parietal cells by cAMP [13]; aquaporin-2 water channel in MDCK kidney epithelial cells [14]; and cystic fibrosis transmembrane regulator (CFTR) channel in duodenal villous epithelial cells [15]. Each physiological response relies on different signaling molecules; however, all require recruitment from intracellular pools of the specific transporters to the apical membrane. Apical recycling of transporters is important

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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in physiological regulation by rapid post-translational altered concentration of active transporter in the plasma membrane. This chapter summarizes recent knowledge regarding apical recycling of canalicular ABC transporters, particularly ABCB11, in the hepatocyte.

TRAFFIC OF NEWLY SYNTHESIZED ABC TRANSPORTERS IN HEPATOCYTES Membrane targeting of newly synthesized canalicular single-transmembrane proteins (ectoenzymes dipeptidylpeptidase IV, aminopeptidase N and 5′ nucleotidase, and the canalicular cell adhesion molecule cCAM105 (also known as HA4)) has been studied in rat liver by in vivo metabolic pulse chase labeling. After biosynthesis, these canalicular proteins are transferred from the Golgi to the basolateral membrane and subsequently reach the BC only by transcytosis [16, 17]. Based on these results, it was proposed that all newly synthesized canalicular transmembrane proteins are targeted via this indirect route and that hepatocytes differ from other polarized epithelial cells in lacking a direct membrane trafficking pathway from the Golgi to the BC [18, 19]. An important observation was that the canalicular cell adhesion molecule cCAM105, but not canalicular ABC transporters, was readily detected immunologically in highly purified sinusoidal/basolateral membrane vesicles (SMVs) from rat liver [7, 20]. The presence of single-transmembrane proteins in SMVs can be explained by the fact that they are initially transferred to the basolateral membrane after biosynthesis and subsequently reach the apical pole by transcytosis. This scenario is in accordance with detectable steady-state levels of single-transmembrane proteins in SMVs. Furthermore, immunohistochemical analysis revealed restriction of ABC transporters to the BC and their absence from the sinusoidal/basolateral membranes. These observations suggested that canalicular ABC transporters do not undergo transcytosis after biosynthesis. The hypothesis of direct apical targeting of canalicular ABC transporters in rat hepatocytes was subsequently tested using metabolic pulse chase labeling [20]. Rats were metabolically labeled with 35 S -methionine, and the contents of newly synthesized cCAM105, ABCB1, ABCB4, and ABCB11 were serially determined in purified canalicular membrane vesicles (CMVs), SMVs, and Golgi membranes from rat liver by immunoprecipitation with specific antibodies. These studies confirmed the transcytotic pathway for apical targeting of newly synthesized cCAM105 (HA4). In contrast, at no time between passage through the Golgi and arrival at the BC were canalicular ABC transporters (ABCB1, ABCB4, and ABCB11) detected in SMVs,

indicating a direct Golgi-to-BC pathway for their membrane targeting [20]. Also, newly synthesized ABCB1, ABCB4, and ABCB11 were not initially transferred to the basolateral membrane; that is, their post-Golgi trafficking differed. After passage through the Golgi, ABCB1 and ABCB4 were rapidly delivered directly to the BC, whereas Golgi-to-BC trafficking of ABCB11 involved additional intermediate steps. At one hour after metabolic labeling, only the mature form of ABCB11 was detected in the homogenate, indicating that processing and passage through the Golgi were complete at this point. At this time point, ABCB11 was not detected in SMVs, the Golgi, or CMVs and therefore had not reached the cell surface, which occurred two hours after metabolic labeling. The most likely explanation is that ABCB11 is sequestered in an intracellular pool prior to delivery to the canalicular membrane. Intrahepatic sequestering of newly synthesized ABCB11 was demonstrated in a later study, which included a combined endosomal fraction in metabolic labeling experiments [8]. Membrane targeting pathways of newly synthesized canalicular proteins discovered by in vivo labeling studies are depicted in Figure 24.1. These studies promoted investigation of direct Golgi-to-BC trafficking of ABCB1-green fluorescent protein (GFP) in WIF-B cells, a polarized hepatocyte cell culture model. WIF-B cells, a hybrid of rat hepatoma cell and human fibroblast cell, have functional bile canaliculi, and are useful model for hepatocytes [21, 22]. Functional features of hepatocytes are also observed in WIF-B cells: basolateral-to-apical membrane transcytosis of canalicular ectoenzymes [23]; and secretion of fluorescent bile acids and substrates for ABCB1 [21] and ABCC2 [24].

Figure 24.1 Canalicular targeting of newly synthesized ABC transporter. ABCB1 traffics directly from the Golgi to the canalicular membrane without participation of post-Golgi compartments. ABCB11 traffics to the canalicular membrane through post-Golgi compartments. Post-Golgi trafficking of ABCB11 accelerates with administration of cAMP

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Live cell imaging revealed that ABCB1-GFP progressively moved from the Golgi to the canalicular plasma membrane within 30–60 minutes. This process was accelerated on incubation of cells with taurocholate (TC), and the entire process, including release from the Golgi, was prevented by preincubation with wortmannin, a PI-3 kinase inhibitor. At no time was basolateral membrane localization of ABCB1-GFP observed [25]. The mechanism and carriers involved in exocytic transport from the Golgi to the canalicular membrane were clarified using time-lapse live-cell-imaging techniques. These studies revealed that ABCB1-GFP move rapidly from the Golgi directly along straight or curvilinear paths and merge with the BC. Tubulovesicular movement of ABCB1-GFP was also observed between the BC and the pericanalicular region. Single, long tubules shrank, formed vesicles, and subsequently fused with the canalicular membrane. Other tubular structures extended from the canalicular membrane into the subapical region and retracted to the canalicular membrane. Time-lapse images reveal that tubular structures reach directly from the Golgi to the BC. Movement of ABCB1-GFP was not synchronous. Individual tubulovesicular structures frequently changed shape during translocation. Frequently, there was a brief delay following which tubular vesicles fused with the canalicular membrane. This event appeared distinct from movements of tubules in other directions (i.e. those presumably not fusing). Direct Golgi-to-bile canalicular trafficking of ABCB1-GFP in WIF-B9 cells is consistent with membrane targeting detected using C219 antibody (ABCB1, ABCB4) in rat metabolic labeling studies in vivo [20]. Furthermore, the movement of ABCB1-GFP from the Golgi to the canalicular membrane was tubulovesicular in appearance and intermittent (occurring every 5–20 minutes). A speed of 0.02–0.6 µm s−1 was slightly lower than values obtained from other studies, which ranged from 0.3 to 1 µm s−1 [26, 27]. The process closely resembles the movement of VSVG protein from ER to Golgi in non-polarized cells [26] and previously described Golgi-to-plasma-membrane trafficking involving large tubular-vesicular structures [26, 27] rather than discrete vesicles, which has been the conventional interpretation [28].

INTRAHEPATIC ABC TRANSPORTER POOLS Bile secretion increases in responses to the enterohepatic circulation of bile acids and the postprandial secretion of peptide hormones that increase cAMP production in hepatocytes [6, 7]. Intravenous administration of rat with cAMP or TC significantly enhances bile secretion, which results from increased amounts of ABC transporter in the bile canalicular membrane. Because the increase in canalicular ABC transporter amount requires an intact

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microtubule system and occurred within minutes of administration, it was proposed that intrahepatic pools provide additional transporters that can be rapidly recruited to the canalicular membrane. Previous morphological studies in rats rendered cholestatic by bile duct ligation [29] or administration of phalloidin [30] or lipopolysaccharide [31] also reveal that ABCC2 and ABCB11 traffic from the BC to intracellular sites, and ABCC2 [31] and ABCB11 [32] were observed by immunogold electron microscopy in undefined vesicular structures distinct from the bile canalicular membrane. ABC transporter trafficking from intracellular sites to the hepatocyte apical domain was induced by cAMP and TC, and used to confirm the existence and define the properties of potential intrahepatic ABC-transporter pools in rats in vivo [8]. Administration of cAMP or TC increased amounts of ABCB1, ABCB4, and ABCB11 in the bile canalicular membrane approximately threefold. These effects abated after six hours, and the bile canalicular content of ABCB1, ABCB4, and ABCB11 returned to basal levels. Pretreatment of rats with cycloheximide, which inhibits protein synthesis, did not prevent the increase in canalicular ABC transporter amounts upon administration of cAMP or TC. These data demonstrate that additional ABC transporters in the bile canalicular membrane do not result from enhanced transcription or translation and indicate recruitment from existing intracellular pools (Figure 24.2a). Using 35 S -methionine metabolic labeling, the overall half-life of ABCB1, ABCB4, and ABCB11 was five days in rat liver, suggesting that ABC transporters cycle between intracellular pools and the bile canalicular membrane prior to degradation. Previous studies [20] indicate that the effects of cAMP and TC on bile canalicular ABC transporter amounts are additive rather than alternative, which suggests the presence of at least two distinct intrahepatic pools of ABC transporters: one is mobilized to the canalicular membrane by cAMP (cAMP pool), and the other by TC (TC pool) (Figure 24.2a). The hypothesis of two distinct intrahepatic pools of ABC transporters was further supported by the observation that targeting of newly synthesized ABCB11 through intrahepatic sites to the bile canalicular membrane is accelerated by cAMP but not by TC [33] (Figure 24.1). After passage through Golgi, ABCB11 accumulates in an intrahepatic cAMP pool and later equilibrates with the TC pool. Whether equilibration of newly synthesized ABCB11 with the TC pool occurs from the bile canalicular membrane or the cAMP pool remains unclear. Newly synthesized ABCB1 and ABCB4 bypass intracellular pools on their journey to the BC [20, 25] (Figure 24.1). However, at steady-state levels, ABCB1 and ABCB4 are also mobilized to the bile canalicular membrane by cAMP and TC, suggesting that these ABC transporters also equilibrate with intrahepatic pools after reaching the bile canalicular membrane (Figure 24.2a). An interesting question is the distribution ratio of ABC transporters between the bile canalicular membrane and

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(a)

(b)

Figure 24.2 Apical recycling of ABC transporter. (a) Administration of cAMP or taurocholate (TC) increases the amount of ABCB1, ABCB4, and ABCB11 from intracellular pools. Because the effects of cAMP and taurocholate are additive, two distinct intracellular pools are postulated. (b) ABCB11 constitutively cycles between canalicular membrane and rab11a-positive endosomes, which causes perinuclear endosome to accumulate and tubular vesicular transport carrier to distribute cytosol. Transport carriers traffic along microtubules throughout the cell but only fuse with the canalicular membrane. ABCB11 trafficking to the canalicular membrane requires rab11a and myosin Vb; retrieval from the canalicular membrane requires HAX-1, cortactin, and EPS15 (clathrin-coated pit)

intrahepatic sites. For ABCC2, the canalicular : intracellular ratio was calculated to be 1 : 1 by quantitation of immunogold-stained ABCC2 in electron microscopy [31] in rat liver under basal conditions. Investigation of ABC transporters by immunoblots at steady-state levels revealed that the amounts of ABCB1, ABCB4, and ABCB11 significantly increased in the bile canalicular membrane after stimulation with cAMP or TC, whereas a change in ABC transporter amount in a combined endosomal fraction prepared from the same rat liver remained below detection limit [8]. This observation is in accordance with the presence of large intrahepatic pools. Under basal conditions, most ABCB1, ABCB4, and ABCB11 appears to reside in intrahepatic pools rather than in the bile canalicular membrane. However, the increase in the amount of ABC transporter in the canalicular membrane after stimulation with cAMP or TC is approximately threefold for each effector. Taking into account that cAMP and TC recruit transporters from different intracellular sources, the intrahepatic pool of ABC transporters is at least six times greater than the amount present in the bile canalicular membrane. Because this calculation presumes that all intracellular ABC transporters are translocated to the bile canalicular membrane upon stimulation, this number represents a lower limit. Thus the intrahepatic : canalicular ratio of ABCB1, ABCB4, and ABCB11 probably exceeds 6 : 1. In vivo biochemical studies suggested that at least two physiologically different ABC transporter intracellular pools exist in rat hepatocyte [34]. These studies promoted investigation of intracellular pools of canalicular ABC transporters. Using WIF-B9 cells, a hybrid cell line derived from a rat hepatoma and human fibroblast cell line, an intracellular pool of canalicular ABC transporters

was identified [35, 36]. Sucrose step gradient fractionation analysis revealed ABCB11 in plasma membrane and endosomal fractions. Confocal microscope imaging of yellow fluorescent protein-tagged ABCB11 (ABCB11-YFP) revealed localization in the bile canalicular membrane, in globular structures adjacent to the microtubule organizing center, and in tubular or vesicular transport carriers which distributed throughout the cytosol (Figure 24.2b). Quantification by confocal microscopy of the different pools of ABCB11 revealed that 40% of each is associated with the canalicular membrane region, 10% with cytoplasmic globular structures, and the remaining 50% in diffusely distributed tubulo-vesicular structures. Hepatocytes contain many different endosomal organelles. Co-immunostaining of specific organelle markers revealed that the major intracellular pool of ABCB11 and ABCC2 consists of rab11a small GTPase positive endosomes [35, 36]. Sucrose step gradient experiment demonstrated that ABCB11 segmentation pattern was similar to the distribution of rab11a, and distinct from other endosome and Golgi markers.

Constitutive canalicular cycling of ABC transporters The amount of ABC transporter in the canalicular membrane is physiologically regulated by the demand to secrete biliary components. The mechanisms and carriers involved in canalicular recycling were further determined using live cell imaging of ABCB11-YFP in polarized WIF-B9 cells [35]. Investigation of constitutive ABCB11 cycling from intracellular compartments and the canalicular membrane was carried out by quantitative confocal microscopy

24: APICAL RECYCLING OF CANALICULAR ABC TRANSPORTERS

combined with selective photobleaching. In the steady state, canalicular pool of ABCB11 was constant. After photobleaching of the canalicular membrane domain, extensive fluorescence recovery of the photobleached area occurred within 20 minutes, indicating that ABCB11 at the canalicular membrane constitutively exchanges with intracellular pools. The recovered pool of ABCB11-YFP did not result from new protein synthesis because pretreatment with cycloheximide did not prevent fluorescence recovery of the canalicular ABCB11 pool. Constitutive apical trafficking was abolished following microtubule disassembly by nocodazole or colchicine. Therefore, intact microtubule organization is required for intracellular trafficking of ABCB11 from intracellular sites. In addition, using combined photobleaching techniques (fluorescence loss in photobleaching, simultaneous photobleaching of two different pools), ABCB11 constitutively cycled between the canalicular membrane, globular structures adjacent to the microtubule-organizing center, and tubular or vesicular transport carriers which distributed in entire cytosol (Figure 24.2b). To investigate further the mechanisms involved in constitutive canalicular recycling of ABCB11, time-lapse imaging of ABCB11-YFP was performed. In the steady state, ABCB11 distributed in the canalicular membrane, globular structure, and tubular-vesicular transport carriers. ABCB11 containing tubular-vesicular cargos continually dissociated from the globular structures. After detaching, the carriers changed speed and direction, and moved at maximal rate of 0.8 µm s−1 to either canalicular or basolateral membranes. ABCB11-containing transport intermediates trafficked toward and along the basolateral membrane but never fused with these membranes. These results demonstrate that ABCB11-containing carriers traffic throughout the cell but specifically fuse with the canalicular membrane (Figure 21.b).

Mechanisms involved in canalicular cycling of ABC transporter PI3 kinase PI3 kinase is a prominent regulator of hepatic ABC transporters in vivo [7, 37]. PI3 kinases are ubiquitous lipid kinases that function as signal transducers downstream of cell-surface receptors and are essential for cell proliferation, adhesion, survival, and cytoskeletal rearrangement [38, 39]. Furthermore, PI3 kinase is required for vesicle trafficking in plant, yeast, and animal cells [40]. The products of PI3 kinase-catalyzed reactions serve as second messengers in many signal-transduction pathways. Studies of the function of PI3 kinase were facilitated by wortmannin, which, when used at the appropriate concentration, is a specific inhibitor for PI3 kinase in isolated systems and cells [41].

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Misra and coworkers [7] investigated the combined effects of wortmannin and TC on bile secretion in isolated perfused rat liver [42] and the amount of ABC transporter in the bile canalicular membrane. Different schedules of TC and wortmannin administration in perfused liver and comparison with ABC transporter amount in the bile canalicular membrane revealed two effects of wortmannin on TC-induced bile secretion. When compared with controls, TC perfusion increased bile secretion, which resulted from an increased amount of canalicular ABC transporter [6]. Administration of wortmannin before TC prevented increase in bile secretion and simultaneously prevented increase in the amount of canalicular ABC transporters. These observations indicate that active PI3 kinase is required for recruitment and vesicular trafficking of additional ABC transporters from intracellular pools to the bile canalicular membrane after stimulation by TC. These studies indicate that active PI3 kinase is required for TC-induced re-localization of ABC transporters in rat liver.

Rab11a Rab small GTPases, which regulate membrane trafficking, constitute a large family in mammalian cells [43]. Rab proteins are molecular switches which cycle between two nucleotide-bound states, a GDP-bound inactive state and a GTP-bound active state [44]. The GTP-bound active form of Rab protein is recruited to a specific membrane organelle, and regulates vesicle trafficking processes such as vesicle budding, motility, and fusion with specific membranes. Rab11a has been identified in the subapical region in many organs, including liver, intestine, kidney, prostate, and pancreas [45]. Rab11a regulates membrane protein apical recycling, such as IgA recycling in polarized MDCK cells [46], apical recruitment of H+-K+-ATPases in gastric parietal cell [47], and CFTR in human airway epithelial cells [48]. The role of rab11a in constitutive canalicular cycling of ABCB11 was investigated using polarized WIF-B9 cells with expression of rab11a GDP-locked form, which acts as a dominant negative. Time-lapse imaging revealed the presence of microtubule-dependent movement of ABCB11-containing endosomes throughout the cell. However, recovery of ABCB11-YFP after photobleaching the canalicular membrane domain was markedly reduced in rab11a-GDP locked form-expressing cells. These observations reveal that rab11a is required for apical recycling of ABCB11 in the canalicular membrane [36].

Myosin Vb: An actin-dependent motor protein The actin cytoskeleton is enriched around the bile canalicular domains [49, 50]. Administration of cytochalasin, which inhibits actin polymerization and disrupts

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cortical actin structure, inhibits bile secretion in rat liver and canalicular recycling of ABCB11 in WIF-B9 cells [35, 51]. These observations indicate that membrane trafficking to and from the canalicular membrane requires cortical actin structures. Myosin is an actin-based molecular motor and more than 20 distinct classes have been identified [52]. Class V myosin, a processive motor protein, contains three subtypes in mammalian cells. Class V myosin contains three different domains: a motor domain; IQ motifs with bound calmodulin or light chains; and a cargo-binding globular tail domain which is required to associate particular intracellular organelles [53]. Myosin Vb binds with rab11a and rab-FIP2, and is implicated in apical recycling of IgA receptor in polarized MDCK cells [54, 55]. To investigate the role of myosin Vb in canalicular targeting of ABC transporters, Myosin Vb tail domain, which lacks a motor domain and acts as dominant negative, was expressed in polarized WIF-B9 cells. Myosin Vb tail domain expression inhibited canalicular targeting of ABC transporters. Immunostaining data demonstrated that myosin Vb tail expression induced accumulation of rab11a positive endosome, which contains canalicular ABC transporters (ABCB1, ABCB11, ABCC2) and transcytotic membrane proteins. Immunostaining experiments demonstrate that endogenous myosin Vb localized with ABCB11 in intracellular pools. These observations indicate that myosin Vb is required for canalicular targeting of ABC transporters and may serve as a motor for delivery of endosomal cargo through the pericanalicular actin network [36] (Figure 24.2b).

HAX-1 Targeting and trafficking of integral membrane proteins depend on amino acid motif in the proteins, interactions with signaling and trafficking networks, and polypeptides that bind the membrane proteins. Several proteins have been identified that associate with ABC transporters and participate in their targeting and recycling. Interactions with PDZ domain proteins EBP50 and E3Karp [56, 57] are essential for apical targeting of CFTR [58]. ABCC2 retention in the apical membrane requires interaction with radixin [59] and a carboxyl-terminal moiety that binds PDZK1 [60, 61]. ABCB1, ABCB4, ABCB11 (MDR family) do not contain obvious PDZ-interacting motifs. Thus, trafficking of the MDR family is controlled by different subsets of proteins. Ortiz et al. identified HAX-1 as a binding partner for the MDR family [62]. HAX-1 is a small cytosolic protein (34 kDa) that interacts with a several different proteins, including polycystic kidney disease protein 2 and cortactin, an F-actin-binding protein [63]. Previously, the linker domains were thought to represent unstructured connecting loops that span the homologous halves of the MDR family. Yeast two-hybrid screening revealed

that HAX-1 specifically binds with linker domains of the MDR family. HAX-1 binding to the linker region of the MDR family was confirmed by HAX-1 pull-down assay using GST-fusion proteins containing ABCB4 and ABCB11 linker domains. Liver fractional studies revealed that HAX-1 is enriched in CMV and clathrin-coated vesicles. Co-immunoprecipitation studies using CMV demonstrated that HAX-1 binds to ABCB1, ABCB4, and ABCB11. Immunostaining analysis in polarized MDCK epithelial cells revealed that HAX-1 co-localized with ABCB11 at the apical membrane. To identify its role in ABCB11 trafficking, HAX-1 expression was ablated by RNAi. Immunoprecipitation studies revealed that the amount of ABCB11 in the apical membrane increased 71% in HAX-1-depleted polarized MDCK cells. Pulse-chase labeling revealed that newly synthesized ABCB11 directly trafficked from the Golgi to the apical membrane within one hour, and peaked in three to four hours. HAX-1 depletion did not affect ABCB11 translation, post-translational modification, or arrival in the apical membrane. However, apical membrane levels of newly synthesized ABCB11 were significantly higher in HAX-1-depleted cells at four (45%), six (72%), and nine hours (103%). These observations demonstrate that HAX-1 is required for ABCB11 internalization from the apical membrane (Figure 24.2b).

Clathrin-dependent endocytosis Clathrin-dependent endocytosis involves internalization of plasma membrane constituent. Immunostaining analysis and EM reveal clathrin-coated pit proximate to the canalicular membrane [64]. Administration of fluid phase maker and antibody against the external domain of single-transmembrane proteins demonstrates that endocytosed molecules co-localize with clathrin-coated pits at the canalicular domain. Direct interaction of canalicular membrane protein with clathrin has not been described. Identification of ABCB1, ABCB4, and ABCB11 binding partners provides new insight into mechanisms involved in canalicular endocytosis. As described above, HAX-1, a binding partner of ABCB1, ABCB4, and ABCB11, directly binds with cortactin, an F-actin-binding protein [62]. Cortactin has been implicated in clathrin-dependent endocytosis in non-polarized cells [65, 66]. Immunoprecipitation experiments using rat CMV fraction revealed that Hax-1 associates with cortactin. To determine whether cortactin participates in endocytosis of ABCB11, the dominant-negative form of cortactin, which lacks the carboxy-terminal SH3 domain, was expressed in polarized MDCK cells. Expression of dominant-negative cortactin increased the amount of ABCB11 (∼210%) in the apical membrane [62]. Liver fractionation studies revealed that ABCB1, ABCB4, and ABCB11 are enriched in clathrin-coated

24: APICAL RECYCLING OF CANALICULAR ABC TRANSPORTERS

vesicles. To determine whether the clathrin-dependent endocytosis machinery participates in ABCB11 internalization from the apical membrane, dominant-negative EPS15 was expressed in polarized MDCK epithelial cells. EPS15 specifically interacts with epsin and AP2 adaptor. A dominant-negative EPS15-EH21 mutant lacks the amino-terminal EH domains and inhibits clathrin-coated pit formation at the plasma membrane, as well as endocytosis [67]. Expression of dominant-negative EPS15 resulted in doubling the amount of ABCB11 in the apical membrane. These observations suggest that ABCB11 is internalized by a clathrin-dependent mechanism (Figure 24.2b).

rab11a

rab11a

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Apical membrane protein Basolateral membrane protein Transcytotic membrane protein

Myo Vb rab11a

rab11a

Polarization cue

BC

rab11a

Link between canalicular recycling system and bile canalicular formation Bile canalicular formation is essential for biliary secretion and is disrupted in cholestatic disorders. In response to cues provide by cell–cell and cell–substrate contacts, epithelial polarization is thought to require regulated coordination between sorting of plasma membrane proteins into post-Golgi vesicles and endosomal transport vesicles, delivery to specific plasma membrane sites, and redistribution into respective polarized domains. Membrane recycling machinery appears to be directly involved in the biogenesis and maintenance of membrane polarity in many organisms. For example, myosin Vb is required for membrane polarity in enterocytes [68]; lack of rab8a expression leads to mislocalization of apical membrane transporters into lysosome [69]; rab11a and myosin Vb are required for cellularization and cellular morphogenesis in developing Drosophila [70, 71]. Time-lapse imaging combined with photobleaching revealed that rab11a and myosin Vb are required for apical recycling of ABC transporter [36]. To investigate the consequences of impaired apical recycling for canalicular biogenesis, rab11a GDP-locked form, shRNAi against rab11a, and myosin Vb tail domain were expressed in non-polarized WIF-B9 cells [36]. Prior to formation of bile canaliculi, apical ABC transporters, which in polarized cells traffic by the direct route from Golgi membranes, are segregated from basolateral membrane proteins and restricted to rab11a- and myosin Vb-positive endosomes. Overexpression of rab11a GDP-locked form, shRNA against rab11a, and myosin Vb tail domain prevented canalicular formation. The impairement of rab11a and myosin Vb function caused apical ABC transporters to remain intracellularly co-localized with transcytotic membrane proteins that were also transported to the plasma membrane. These observations indicate that rab11a and myosin Vb are required for trafficking endosomes that contain apical ABC transporters to sites that initiate bile canalicular formation (Figure 24.3).

rab11a

Figure 24.3 Bile canalicular biogenesis. (a) Prior to formation of the bile canaliculus, canalicular ABC transporters localize in rab11a-positive endosome with transcytotic membrane proteins. Basolateral membrane proteins target to the plasma membrane with transcytotic membrane proteins. (b) Initiation of the polarization cue at the site of differentiation into the apical domain. (c) Delivery of canalicular membrane constituents to the site requires rab11a and myosin Vb

CONCLUSION The secretion of bile across the canalicular membrane is predominantly determined by the concentration of active bile salt excretory protein (ABCB11) in the canalicular membrane. However, the amount of active ABC transporter in the canalicular membrane is not static and involves complex dynamic interaction with regulated traffic and transporter activation. The demand to secrete bile varies with the nutritional, postprandial, and hormonal status of the individual, and the hepatocyte copes by cycling ABC transporters, particularly ABCB11, between intracellular and plasma membrane pools. The cycling mechanism is observed in different species and tissues and involves many other membrane transporters and pumps. It provides a physiological advantage in rapidly adapting to increased load of potentially deleterious cargo (i.e. bile acid) and confers a dynamic characteristic to the individual membrane transporter. It is analogous to an idling motor, which, when confronted with the need for increased performance, rapidly accelerates. Regulation of this dynamic mechanism is complex and slowly being resolved. Theoretically, it could be a major determinant in bile secretory failure as a result of viral infection, drugs, metabolic changes, hypoxia, or infection. To date, inheritable defects in ABC transporters related to cholestasis are restricted to primary sequence mutations. Trafficking and cycling defects have yet to be identified in liver. In the intestine, mutations in myosin Vb have resulted in microvillar inclusion body disease in which apical polarization is lost [68].

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Many other lipids and proteins undoubtedly participate in the ABC transporter recycling system [72, 73]. It is a challenge to identify the players involved in this concert of complex dynamic regulation of canalicular ABC transporters, and, of course, the conductor.

15.

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25

Cholangiocyte Functions in Health and Disease: The Ciliary Connection Anatoliy I. Masyuk1, Tatyana V. Masyuk1 and Nicholas F. LaRusso2 1 Mayo

Graduate School of Medicine, Mayo Clinic College of Medicine, Rochester, MN, USA 2 Gasteroenterology Research Unit, Mayo Graduate School of Medicine, Mayo Clinic College of Medicine, Rochester, MN, USA

INTRODUCTION Biliary epithelial cells (i.e. cholangiocytes) lining intrahepatic bile ducts, a complex three-dimensional network of conduits within the liver that provides delivery of bile to the gall bladder and the intestine, are capable of performing a wide variety of functions. Their major physiological function is ductal bile formation, which occurs via a series of secretory and absorptive processes contributing to the final composition of bile [1–12]. By releasing growth factors, peptides, nucleotides, proinflammatory and chemotactic cytokines, and other signaling molecules, cholangiocytes interact with nearby and downstream cells within the biliary tree, and with other liver cells, including hepatocytes, hepatic stellate, endothelial, mesenchymal, and inflammatory cells. Through these interactions, cholangiocytes are involved in regulation of ductal bile formation and in liver inflammatory processes, fibrogenesis, and angiogenesis [5, 10, 13–17]. Having a great capacity to proliferate, cholangiocytes are involved in repair of the biliary tree after injury, and in the development of liver diseases associated with cholangiocyte hyperproliferation (e.g. polycystic liver disease) [7, 10, The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

15, 17–30]. Thus, cholangiocytes possess extraordinary biological properties that are critically important for liver function in health and disease. Given that cholangiocyte biology and pathobiology have been discussed in a large number of excellent reviews [1, 2, 7–9, 17], in this chapter we have focused our attention mainly on cholangiocyte functions associated with primary cilia, which are currently recognized as sensory organelles that coordinate intracellular signaling and functional responses to extracellular stimuli [31–34]. We discuss the established ciliary connections to: (i) intracellular signaling in cholangiocytes, (ii) ductal bile formation, and (iii) cholangiocyte proliferation.

CHOLANGIOCYTE CILIA IN LIVER HEALTH A brief overview of cholangiocyte primary cilia Cholangiocytes lining the intrahepatic bile duct system are typical epithelial cells. They are flattened or cuboidal in

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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(a)

(b)

Figure 25.1 Cholangiocyte primary cilia. (a) A scanning electron microscopy image of primary cilia extending from the cholangiocyte apical plasma membrane into the bile duct lumen in rat liver—from Huang et al. [36], used with permission of The American Physiological Society (b) Transmission electron microscopy images of a longitudinal section and cross-section (inset) of a primary cilium in isolated rat intrahepatic bile duct. Reproduced from Masyuk, T.V., Huang, B.Q., Ward, C.J. et al. (2003) Defects in cholangiocyte fibrocystin expression and ciliary structure in the PCK rat. Gastroenterology, 125, 1303–10. Copyright  2003, Elsevier.

the small branches of the biliary tree, and are columnar in the large bile ducts [2, 35]. Cholangiocytes are polarized cells with well-defined basolateral and apical plasma membranes; the latter have a primary cilium, a solitary, long, tubular organelle extending into the bile duct lumen [19, 36] (Figure 25.1). The primary cilium arises from the mature mother centriole of the mother–daughter pair of centrioles and consists of a basal body and a membrane-bound, “9 + 0” axoneme that contains nine outer-doublet microtubules but lacks a central pair of microtubules and dynein arms [36]. Thus, primary cilia do not move spontaneously, in contrast to motile cilia, which have a “9 + 2” axoneme with nine outer-doublet microtubules, a central pair of microtubules, and dynein arms. Primary cilia in cholangiocytes were described in the 1960s and 1970s but their physiological and pathophysiological significance remained unclear until recently [31, 37, 38]. In rat liver, primary cilia range in length from 2 to 8 µm depending on cholangiocyte size and the diameter of the bile ducts; that is, in the large bile ducts the ciliary axonemes are three to four times longer than in the small bile ducts [36]. The correlation between the size of cholangiocytes and the length of cilia suggest that these organelles are likely functionally heterogenous along the biliary tree axis. The current generally accepted concept of the physiological significance of primary cilia is that these organelles perform sensory functions [32, 34]. Originally developed in the 1980s, this concept proposes that primary cilia in different cell types sense multiple mechano-, osmo-, and chemostimuli and transduce them into intracellular signaling and functional responses [39, 40]. Recent research provides multiple lines of evidence supporting this concept [41–44]. Studies on cholangiocyte cilia, which are

ideally positioned on the apical plasma membrane to detect changes in bile flow, composition, and osmolality, also suggest that in biliary epithelia, primary cilia function as mechano-, osmo-, and chemosensory organelles sensing and transducing extracellular biliary signals into intracellular signaling [31, 37, 38].

Cholangiocyte cilia and intracellular signaling In general, the Ca2+ and cAMP signaling pathways are key intracellular regulatory mechanisms through which extracellular stimuli affect ductal bile formation and other biliary epithelia functions. Importantly, sensory functions of cholangiocyte cilia are associated with both signaling pathways.

Calcium signaling Calcium signaling in cholangiocytes is initiated by two mechanisms: (i) release from intracellular Ca2+ stores in response to activation of G-protein-coupled receptors or receptor tyrosine kinases expressed on the cholangiocyte plasma membrane [45]; and (ii) influx of extracellular Ca2+ into the cell via Ca2+ channels localized to the plasma membrane [2]. Activation of certain cholangiocyte plasma-membrane receptors by hormones, neurotransmitters, growth factors, and nucleotides leads to an increase in the concentration of intracellular Ca2+ ; however, the patterns of [Ca2+ ]i signaling depend on the type of activated receptors and ligands [45, 46]. For example, Ca2+ signals induced

25: CHOLANGIOCYTE FUNCTIONS IN HEALTH AND DISEASE: THE CILIARY CONNECTION

by acetylcholine (ACh), which activates M3 muscarinic receptors localized to the cholangiocyte basolateral plasma membrane, begin as apical-to-basal Ca2+ waves [47]. In contrast, adenosine triphosphate (ATP), which activates both apically and basolaterally localized P2Y purinergic receptors, induces Ca2+ oscillations [47]. The Ca2+ -signaling response to ATP depends on ligand concentration: lower concentrations of ATP predominantly induce Ca2+ oscillations, while higher concentrations induce a single (either sustained or transient) increase in Ca2+ [45, 46]. The hormone/(neurotransmitter, growth factor, nucleotide)-induced increase in [Ca2+ ]i in cholangiocytes is associated with the production of inositol 1,4,5-trisphosphate (InsP3 or IP3), which is the endogenous ligand for the inositol 1,4,5-trisphosphate receptors (InsP3Rs) localized to the endoplasmic reticulum (ER), where intracellular Ca2+ is stored. InsP3Rs are tetrameric IP3-gated Ca2+ channels, activation of which results in Ca2+ release from the ER into the cytoplasm [46]. All three known isoforms of InsP3Rs (i.e. types I, II, and III) are expressed in cholangiocytes; however, InsP3R type III is the predominantly expressed isoform [45, 47]. The cholangiocyte apical plasma membrane expresses P2Y and P2X purinergic receptors, the activation of which by biliary nucleotides (i.e. ATP, ADP, UTP, and UDP) results in an intracellular Ca2+ signaling response [13, 14, 16, 48]. A family of P2Y purinergic receptors, which are G-protein-coupled receptors, consists of eight members (i.e. P2Y1 , P2Y2 , P2Y4 , P2Y6 , P2Y11 , P2Y12 , P2Y13 , and P2Y14 ) [49–51]. P2Y1 , P2Y2 , P2Y4 , and P2Y6 are associated with [Ca2+ ]i signaling, while P2Y11 , P2Y12 , P2Y13 , and P2Y14 are associated with cAMP signaling. P2Y11 is associated with both Ca2+ and cAMP signaling [49–51]. In rat, at least six P2Y receptors (i.e. P2Y1 , P2Y2 , P2Y4 , P2Y6 , P2Y12 , and P2Y13 ) are localized to the cholangiocyte apical plasma membrane [13]. However, there is no evidence that any of the P2Y purinergic receptors that are associated with Ca2+ signaling are localized to cholangiocyte cilia. In contrast, P2Y12 , which is associated with cAMP signaling (see below), is localized to these organelles [31]. A family of P2X purinergic receptors, which are ATP-gated, calcium-permeable, non-selective cation channels, consists of seven members (i.e. P2X1−7 ). They have two transmembrane domains, intracellularly oriented Cand N-termini, and an extracellular ATP-binding site [49, 50]. Normal rat cholangiocytes express four of them, P2X2 , P2X3 , P2X4 , and P2X6 [13, 48], however, there is no evidence that they are localized to cholangiocyte cilia. Nevertheless, in other cell types, P2X receptors are ciliary-associated proteins. It has been reported that one of the P2X receptors, called P2Xcilia , which is potentially a P2X4 receptor, is localized to motile cilia in airway epithelial cells in rabbits [52]. Thus, it is likely that P2Y and P2X receptors linked to [Ca2+ ]i signaling in cholangiocytes operate on their apical plasma membrane but

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not in cilia. However, cholangiocyte cilia are functionally connected to Ca2+ signaling in the cell via other mechanisms, which are associated with Ca2+ -entry channels, polycystin-2 (PC2), and a transient receptor potential vanilloid 4 (TRV4) channel. PC2 is a 968 amino acid protein with a molecular weight of 110 kDa. It has six transmembrane domains; both N- and C-termini are oriented intracellularly. Its carboxy terminal contains an EF-hand domain that has a calcium-binding site. PC2 acts as a non-selective cation channel permeable to Ca2+ , K+ , and Na+ [53]. Importantly, PC2 displays channel activity only in the presence of polycystin-1 (PC1), which is likely a mechanosensor. PC1 is a 4304 amino acid integral membrane glycoprotein with a molecular weight of ∼460 kDa. It has 11 transmembrane domains, a large extracellular N-terminus, and a short cytoplasmic C-terminus that is involved in intracellular signaling [54, 55]. These two proteins are thought to form a functional mechanosensory complex in primary cilia [56] (Plate 25.1). Both PC1 and PC2 are localized to cholangiocyte cilia and account for the mechanosensory functions of these organelles [37]. Cholangiocyte cilia respond to changes in bile flow rates by a transient increase in intracellular Ca2+ , which primarily depends on influx of extracellular Ca2+ into the cell via PC2 followed by Ca2+ release from intracellular Ca2+ stores [37]. The mechanisms of the flow-induced ciliary-mediated Ca2+ release from cholangiocyte intracellular stores remain uncertain. When intracellular InsP3-sensitive Ca2+ stores in rat cholangiocytes were depleted by thapsigargin, the flow-induced increase in Ca2+ was significantly reduced, suggesting a partial involvement of InsP3Rs in these mechanisms [37] (Plate 25.2). TRPV4 is another Ca2+ entry channel localized to cholangiocyte cilia. This channel is exquisitely sensitive to minute changes in osmolality of extracellular milieu, being activated by extracellular hypotonicity and inhibited by extracellular hypertonicity [57]. Activation of TRPV4 results in an increase in [Ca2+ ]i through influx of extracellular Ca2+ into the cell [58]. In epithelial cells lining the oviduct, TRPV4 is localized to motile cilia, where it may perform multiple roles linked to osmo-, chemo-, and mechanosensory ciliary functions [59, 60]. A homolog of TRPV4, a putative transient receptor potential (TRP)-like channel protein, OSMotic avoidance abnormal family member 9 (OSM 9) is expressed in primary cilia in C. elegans sensory neurons, where it functions as an osmosensor [61]. In rat and mouse cholangiocytes, TRPV4 is localized to primary cilia and is involved in cell responses to changes in bile osmolality [38]. Exposure of cholangiocytes to hypotonicity results in a sustained increase in intracellular Ca2+ , which depends on extracellular Ca2+ sources [38]. Taken together, these observations suggest that Ca2+ signaling in cholangiocytes associated with mechano- and osmosensory functions of primary cilia is provided by

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both mechanisms; that is, by entry of extracellular Ca2+ into the cell via Ca2+ channels, and by Ca2+ release from intracellular stores. These processes occur in a coordinated fashion given the fact that biliary osmolality not only induces the intracellular TRPV4-dependent Ca2+ increase but also induces the cholangiocyte TRPV4-dependent ATP release, which in turn may activate P2Y purinergic receptors linked via Gq protein to InsP3 formation, followed by Ca2+ release from intracellular stores [38].

cAMP signaling The major molecular components of the cAMP signaling cascade in any cell type, including cholangiocytes, are a receptor, adenylyl cyclase (AC), phosphodiesterase (PDE), the serine-threonine protein kinase A (PKA), the exchange proteins directly activated by cAMPs (EPACs), and A-kinase anchoring proteins (AKAPs). In electrically active cells, the major molecular components of cAMP signaling also include the cyclic-nucleotide-gated (CNG) ion channels [62, 63]. Typically, a receptor is a seven-transmembrane-domain protein, activation of which by specific ligands initiates a series of events including the interaction of a trimeric G protein with AC. There are ten different ACs; nine of them (AC1–AC9) are 12-transmembrane-domain proteins, and one is a soluble form of AC (sAC) [64]. The activities of membrane-bound ACs are regulated by different subunits of Gs, Gi, Go, and Gq proteins, resulting in an increase or a decrease of cAMP levels in the cell. The cAMP levels are also regulated by PDEs, which belong to at least 11 families of PDEs specific for cAMP. The activities of ACs and PDEs are regulated positively and negatively by other signaling mechanisms, which include but are not limited to Ca2+ signaling, G-protein-coupled signaling, and receptor tyrosine kinases signaling [62–64]. Rat cholangiocytes express 7 of 10 known ACs (i.e. AC4–AC9, and sAC), suggesting the existence of multiple mechanisms of the cAMP signaling pathway in biliary epithelia [37, 65]. Three ACs (i.e. AC4, AC6, and AC8) are localized to primary cilia [31]. Importantly, the activity of two of them, AC6 and AC8, depends on Ca2+ , suggesting an intersection between the cAMP and Ca2+ signaling cascades in cholangiocyte cilia. The main downstream targets of cAMP in cholangiocytes are PKA and EPAC [66, 67]. PKA, a heterotetramer, consists of two regulatory (R) and two catalytic (C) subunits, which in mammalian cells are represented by three C subunits (Cα, Cβ, and Cγ) and two types of R subunit. R subunits, numbered I and II, have an α and β subtype [67]. Rat cholangiocytes express all four PKA-R subtypes; that is, PKA-RIα, PKA-RIβ, PKA-RIIα, and PKA-RIIβ. Two of them, PKA-RIβ and PKA-RIIα, are localized to cholangiocyte cilia [68]. PKA is activated by the binding of cAMP to the R subunits, inducing their dissociation from the C subunits. The latter become active and

phosphorylate substrate proteins. In the absence of cAMP, C subunits are inhibited by reassociation with R subunits. EPAC proteins (i.e. EPAC1 and EPAC2, also known as RapGEF3 and RapGEF4, respectively) are other downstream targets of cAMP. They belong to a family of cAMP-regulated guanine nucleotide exchange factors controlling multiple cellular functions via PKA-independent mechanisms [67]. Both of these proteins are expressed in rat cholangiocytes; one of them, EPAC2, is localized to cilia [31]. Finally, the A-kinase anchoring protein AKAP150 is also localized to cholangiocyte cilia [31]. There are more than 30 AKAPs, representing a family of functionally related proteins that anchor PKAs to certain cellular compartments [69]. AKAP150 has unique abilities to bring together several components of the cAMP signaling cascade, such as AC, PKA, EPAC, and G-protein-coupled receptors [70] (Plate 25.3). Selective localization of ACs, PKA, EPAC, and AKAP150 to cilia suggests a subcellular compartmentalization of cAMP signaling in cholangiocytes, and that machinery involved in cAMP signaling in the ciliary axoneme differs from machinery providing a total cellular cAMP signaling response. The cAMP signaling cascade in cholangiocyte cilia plays an important role in their mechano- and chemosensory functions. Importantly, luminal fluid flow does not affect the basal level of cAMP in cholangiocytes. However, when the cellular concentration of cAMP is elevated by forskolin, which activates most isoforms of AC in the cell, luminal fluid flow completely inhibits the forskolin-induced cAMP increase in cholangiocytes via ciliary-associated mechanisms [37]. This observation suggests that the major function of the ciliary cAMP-signaling machinery is inhibition of the cAMP signaling pathway in cholangiocytes where this pathway was initially activated by other stimuli. The flow-induced ciliary-mediated cAMP decrease in forskolin-stimulated cholangiocytes depends on two ciliary-associated proteins, AC6 and PC2 [37]. AC6 is a Ca2+ -inhibitable isoform that is usually colocalized on the cellular plasma membrane with Ca2+ -entry channels [64]. In cholangiocyte cilia, the channel is likely PC2, given that inhibition of its expression by specific siRNAs abolished the flow-induced decrease in forskolin-stimulated cAMP levels [37]. Importantly, the forskolin-induced cAMP increase in cholangiocytes is inhibited via ciliary-associated mechanisms not only by mechanical stimuli (i.e. fluid flow) but also by biliary nucleotides [31]. As mentioned above, one of the P2Y purinergic receptors, P2Y12 linked to ACs via Gi, is localized to cholangiocyte cilia. Its activation by ADP, the most potent endogenous ligand, resulted in inhibition of ACs and a decrease in cAMP levels; that is, it causes effects similar to the flow-induced ciliary-mediated inhibition of cAMP signaling [31]. Biliary ADP is a product of degradation of ATP, which is released in bile by both hepatocytes and cholangiocytes [71]. In response to different stimuli, cholangiocytes may release ATP via the

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apically located cystic fibrosis transmembrane conductance regulator (CFTR) chloride channel, or at least with the involvement of this channel [5, 30]. Thus, recent observations suggest that cholangiocyte cilia possess mechanisms linked to both the intracellular Ca2+ and cAMP signaling pathways. Ca2+ signaling in cholangiocytes is activated by extracellular stimuli such as bile flow and bile osmolality, while cAMP signaling is inhibited by bile flow and biliary nucleotides. The latter are released by cholangiocytes in response to different stimuli, including changes in bile osmolality [14, 16, 29, 30, 38]. The physiological implications of the flow- and osmolality-induced ciliary-mediated increase in [Ca2+ ]i and the flow- and nucleotide-induced decrease in cAMP in cholangiocytes remain unclear. The fact that PC1, PC2, TRPV4, P2Y12 , ACs, and other components of the cAMP signaling cascade are localized to cholangiocyte cilia suggests the existence of previously unknown ciliary-associated mechanisms that may provide regulation of ductal bile secretion by different stimuli originating in the lumen of intrahepatic bile ducts—that is, via the apical pole of cholangiocytes.

Cholangiocyte cilia and ductal bile formation Bile is an aqueous solution of organic and inorganic compounds initially secreted by hepatocytes (primary bile) and subsequently modified by cholangiocytes (ductal bile) through secretory and absorptive processes. Cholangiocytes modify the fluidity and alkalinity of primary bile by secreting ions, primarily Cl− and HCO− 3 , and by absorbing bile salts, glucose, and amino acids, followed by passive movement of water into or out of the bile duct lumen depending on osmotic gradients [2–4, 6–8, 12,]. Ductal bile formation is a regulated process and is affected by many extracellular regulatory stimuli, such as hormones, neurotransmitters, peptides, and nucleotides, via both the intracellular cAMP and Ca2+ signaling pathways [2, 8, 11]. The cAMP signaling pathway is activated by secretin and other hormones, followed by CFTR-dependent chloride transport into the lumen of intrahepatic bile ducts and chloride-bicarbonate exchange via the apically located AE2/SLC4A2 anion exchanger [8]. The cAMP/CFTR pathway is thought to be of primary importance in maintaining ductal bile formation [5]. The secretin-stimulated ductal bile secretion is terminated by somatostatin, which inhibits cAMP signaling in cholangiocytes [2]. The [Ca2+ ]i signaling pathway is involved in regulation of ductal bile formation by neurotransmitters and nucleotides via activation of apically located Ca2+ -dependent chloride channels, followed by chloride–bicarbonate exchange via the apically located AE2/SLC4A2 anion exchanger [5, 14, 16, 48].

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Thus, hormones (with the exception of somatostatin), neurotransmitters, and nucleotides stimulate ductal bile secretion via specific receptors localized primarily to the cholangiocyte basolateral plasma membrane. Biliary nucleotides stimulate ductal bile secretion via apically located P2Y and P2X purinergic receptors [13, 14, 48]. It is likely that cholangiocyte cilia function as apically located sensory organelles that monitor changes in bile flow, bile osmolality, and concentrations of biliary nucleotides within intrahepatic bile ducts, and adjust cholangiocyte functional responses (i.e. ductal bile secretion) to such changes. Physiologically, bile flow in intrahepatic bile ducts is pulsatile. Thus, minute-to-minute changes in bile flow alter the mechanical forces to which cholangiocyte cilia are exposed and may affect cholangiocyte secretory and absorptive processes. Functionally, luminal bile flow via activation of [Ca2+ ]i signaling in cholangiocytes may terminate both global and local cAMP signaling (i.e. turn off regulation) initially activated by secretin and other regulatory molecules (i.e. turn on regulation), thus providing a coordinated regulation of ductal bile secretion and fast functional adjustments of the intrahepatic ductal system to physiological needs. Changes in bile osmolality also contribute to regulation of ductal bile formation. Bile is considered isotonic; however, transient changes in ductal bile tonicity as a result of secretory and absorptive functions of cholangiocytes affect (i.e. activate or inhibit) a ciliary osmosensory protein, TRPV4. Its activation or inhibition in turn induces or inhibits ion and water transport by cholangiocytes, restoring the isotonicity of ductal bile. This observation is supported by experiments demonstrating that a retrograde infusion of 4-alpha-phorbol 12, 13-didecanoate (4αPDD), an agonist to TRPV4, into rat intrahepatic bile ducts in vivo resulted in stimulated ductal bile secretion that was abolished by Gd3+ , a TRP channel inhibitor [38]. Finally, changes in bile tonicity induce ATP release by cholangiocytes, which in turn activates apically located P2Y and P2X receptors and coordinates ductal bile secretion via the intracellular Ca2+ and cAMP signaling pathways. Biliary nucleotides stimulate ductal bile secretion via apically located P2Y and P2X purinergic receptors linked to intracellular Ca2+ signaling [13, 14, 16, 48], but may inhibit this process via the ciliary associated P2Y12 purinergic receptor if the secretory activity of cholangiocytes was already activated by other stimuli [31].

CHOLANGIOCYTE CILIA IN LIVER DISEASES Cholangiopathies associated with primary cilia (Cholangiociliopathies) Cholangiocytes are the target of a large group of acquired and inherited liver diseases that are collectively called

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Table 25.1 Cholangiociliopathies Disease

Gene

Chromosome

Protein

Hepatic pathology

ADPKD

PKD1 PKD2 PKHD1

16p13.3–p13.12 4q21–q23 6p21.1

Polycystin 1 Polycystin 2 Fibrocystin

Cysts

PRKCSH SEC63 INV/ NPHP2 NPHP3 NPHP4 MKS1

19p13.2–2p13.1 6q21–q23 9q22–q31 3q21–q22 1p36 17q21–q24

Hepatocystin Sec63 Inversin or nephrocystin 2 Nephrocystin 3 Nephrocystin 4 MKS1

MKS2 MKS3

11q13 8q21.13–q22.1

— Meckelin

ARPKD ADPLD NPHP2 NPHP3 NPHP4 MKS

“cholangiopathies” [7, 10]. A number of cholangiopathies, which we propose to call “cholangiociliopathies” [22, 31], are caused by mutations in genes encoding ciliary-associated proteins, which account for the structural and functional characteristics of cholangiocyte primary cilia. The cholangiociliopathies are characterized by liver cysts and/or hepatic fibrosis, and include: (i) autosomal dominant polycystic kidney disease (ADPKD), caused by mutations in either PKD1 or PKD2 , genes encoding PC1 and PC2, respectively; (ii) autosomal recessive PKD (ARPKD), caused by mutations in PKHD1 , a gene that encodes FC; (iii) nephronophthisis (NPHP), caused by mutations in six genes (NPHP1 –6 ) that encode nephrocystin 1–6; (iv) Bardet–Biedl syndrome (BBS), caused by mutations in 12 BBS genes that encode BBS1–12; (v) Meckel–Gruber syndrome (MKS), caused by mutations in three genes (MKS1 –3 ) that encode MKS1–3; and (vi) potentially other liver diseases (Table 25.1). ADPKD is characterized by massive, progressive liver cysts, while ARPKD and MKS are associated with biliary dysgenesis/cystogenesis and hepatic fibrosis [22, 72]. In contrast, NPHP and BBS are characterized mainly by congenital hepatic fibrosis [22].

Pathogenesis of the cholangiociliopathies The mechanisms involved in the pathogenesis of cholangiociliopathies in general and polycystic liver diseases in particular are obscure. However, recent developments suggest that cholangiocyte hyperproliferation and abnormal secretory/absorptive functions play major roles in hepatic cyst growth and expansion [22].

Cholangiocyte proliferation Cholangiocytes in healthy adult liver are mitotically dormant, but in cholangiociliopathies they hyperproliferate.

Cysts, congenital hepatic fibrosis, biliary dysgenesis Cysts Congenital hepatic fibrosis Ductal plate malformation, fibrosis, biliary dysgenesis

For example, in the PCK rat, a well-characterized animal model of ARPKD, bile ducts are markedly distorted, displaying multiple saccular dilatations and many hepatic cysts of different sizes (Figure 25.2) [20]. Overgrowth of liver cysts is associated with an increased proliferation of cystic cholangiocytes compared to normal cholangiocytes of age-matched normal rats [21, 73]. The ability to proliferate is preserved in a spontaneously immortalized cholangiocyte cell line derived from the PCK rat (PCK-CCL) that retains the in vivo phenotype of cystic cholangiocytes. The rate of PCK-CCL proliferation is significantly higher and the doubling time twice as fast as in a cell line derived from normal rats (NRC) [74]. Accelerated cell proliferation is associated with alterations in the cell cycle, which is regulated, in particular, by a family of dual-specificity phosphatases, called Cdc25. One of the Cdc25 phosphatases, Cdc25A, is principally responsible for G1–S and G2–M transitions [75, 76]. In polycystic liver, the expression of Cdc25A is increased, correlating with cholangiocyte hyperproliferation and hepatic cyst formation. The link between Cdc25A and hepatic cystogenesis is provided by microRNAs (miRNAs), which are small non-coding RNAs that post-transcriptionally inhibit target mRNA transcripts via sequence-specific base-paring [77].

miRNAs in cholangiocyte hyperproliferation Biochemical and genetic studies suggest that miRNAs play important roles in a variety of cellular processes, including cell proliferation [77]. In particular, the miR-17 cluster and miR-21 are known to promote cell proliferation, while miR-15a, miR-16, and the family of let-7 miRNAs suppress this process [78, 79]. miRNAs are transcribed as long primary precursor transcripts (pri-miRNAs) that are cleaved into pre-miRNAs (∼70-nucleotide precursor hairpins) by the RNase III-like enzyme, Drosha. The pre-miRNAs are exported to the cytoplasm by Exportin-5 and cleaved to short (about 22

25: CHOLANGIOCYTE FUNCTIONS IN HEALTH AND DISEASE: THE CILIARY CONNECTION

(a)

(b)

(c)

(e)

(f)

365

BT

(d)

Figure 25.2 Liver manifestation of ARPKD—an example of cholangiociliopathies. Light microscopy images of normal (a) and PCK (b) rat liver tissues stained with hematoxylin and eosin show healthy liver (a) and liver with large numerous cysts (b). Light microscopy image of PCK rat liver tissue stained with picrosirius red shows the presence of hepatic fibrosis around hepatic cysts (c). Scanning electron microscopy images of normal (d) and PCK (e,f) rat liver tissues show healthy liver with a conventional biliary triad (BT) consisting of the portal vein, hepatic artery, and intrahepatic bile duct (d), and unhealthy liver in which cysts replace most of the liver parenchyma (e,f). Epithelial cells lining cysts are cholangiocytes in origin; they contain primary cilia, which are functionally abnormal due to mutation in Pkhd1 encoding fibrocystin. From Masyuk, Masyuk & LaRusso. Cholangiocyte primary cilia in liver health and disease. Dev Dynamics 2008; 237:2007–2012, reproduced with permission from John Wiley & Sons Inc.

nucleotides), partially double-stranded RNA, in which one strand is the mature miRNA. The mature miRNA then binds to specific messenger RNA targets and mediates either translation repression or direct mRNA cleavage [80]. In polycystic liver, the expression of mir-15a is significantly reduced, while its target, the cell-cycle regulator Cdc25A, is overexpressed. These interrelated changes promote cholangiocyte hyperproliferation and hepatic cyst formation. This conclusion is supported by the observation that in PCK-CCL transfected with pre-miR-15a, levels of Cdc25A expression, G1-S cell-cycle transition, and cholangiocyte proliferation are all decreased [81].

Intracellular signaling in hyperproliferative cholangiocytes The connections between the sensory functions of cholangiocyte cilia, their structure, intracellular signaling pathways, and hepatic cyst development have recently been established.

cAMP signaling Increased cAMP levels in cholangiocytes represent a common feature of the cholangiociliopathies. cAMP increases

the rate of cholangiocyte proliferation in polycystic liver and affects downstream targets, such as EPAC and PKA, in proliferating cells. Both EPAC isoforms (EPAC1 and EPAC2) and one of the PKA subtypes (PKA-RIβ) are overexpressed in polycystic liver, accelerating cholangiocyte proliferation via the MEK/ERK1/2 signaling cascade [68]. The importance of cAMP signaling in the development of hepatic cysts is supported by the observation that treatment of PCK rats (an animal model of ARPKD) with octreotide, a synthetic analog of somatostatin, decreases cAMP levels in cholangiocytes, suppressing hepatic cyst growth, and reducing hepatic fibrosis [21].

Ca2+ signaling In contrast to intracellular cAMP levels, which are elevated in cholangiocytes of ARPKD liver, the concentration of [Ca2+ ]i in proliferating cholangiocytes is low. The mechanisms that cause the [Ca2+ ]i decrease in proliferative cholangiocytes remain unknown. Restoration of [Ca2+ ]i in cholangiocytes of polycystic liver inhibited both basal and PKA-regulated cholangiocyte proliferation via the PI3K/AKT pathway but had no effect on EPAC-regulated cholangiocyte proliferation in PCK rats,

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suggesting a differential interaction between [Ca2+ ]i signaling and the two downstream effectors of the cAMP signaling cascade [68]. Thus, cholangiocyte hyperproliferation in ARPKD liver is associated with abnormalities in two intracellular signaling pathways: the cAMP pathway, which is upregulated, and the intracellular Ca2+ pathway, which is downregulated.

Hedgehog signaling Hedgehog (Hh) signaling is currently recognized as one of the key regulatory pathways involved in embryonic development, maintenance of adult tissues, and carcinogenesis. The term “Hedgehog signaling” originates from an intercellular signaling molecule called Hedgehog, which is a 19 kDa dually lipid modified protein. In mammals, three different genes encode Hh: sonic hedgehog (Shh), desert hedgehog (Dhh), and Indian hedgehog (Ihh) [82]. The Hh signaling pathway is activated by an Hh protein, which interacts with the Hh receptor, “Patched” (Ptch). In an inactive form (i.e. in the absence of Hh), Ptch inhibits another receptor called “Smoothened” (Smo), a downstream protein in the pathway. Activation of Ptch liberates Smo, which in turn activates transcription of Hh target genes via transcription factors that belong to a glioblastoma (Gli) family. Importantly, in embryonic cells, components of Hh signaling are localized to primary cilia. How primary cilia are involved in the Hh signaling in adult mature organs and tissues remains obscure [82]. There is not much activity of Hh signaling in the healthy adult liver. However, this signaling pathway is activated when the liver is damaged or is developing diseases such as hepato- and cholangiocarcinomas [83, 84]. It is likely that Hh signaling is involved in cholangiocyte hyperproliferation in polycystic liver diseases. Our initial observations suggest that components of Hh signaling normally absent in healthy cholangiocytes are expressed in cystic cholangiocytes in ADPKD, and some of them (i.e. Ptch) are localized to cholangiocyte cilia.

Cholangiocyte secretion and absorption in hepatic cyst formation In polycystic liver, defects in ciliary structure and function induce cholangiocyte hyperproliferation and alter their secretory and/or absorptive functions, resulting in production of cystic fluid. In 3D culture, microdissected single PCK cysts expand more rapidly than normal bile ducts, both spontaneously and in response to secretin. Also, while AQP1, CFTR, and AE2, the functionally related proteins involved in cholangiocyte fluid secretion, are localized preferentially to the apical membrane in normal cholangiocytes, a striking overexpression of these three proteins is found at the

basolateral membrane of PCK cholangiocytes [85]. PCK cysts secrete more fluid into the cystic lumen than normal bile ducts when exposed to hypo-osmolality, suggesting that hepatic cystogenesis in PCK rats may be the result of alterations in expression and topographic location of AQP1, CFTR, and AE2 proteins [85]. cAMP-stimulated CFTR-dependent Cl− secretion has also been reported to contribute to fluid accumulation in hepatic cysts of ADPKD [86, 87]. Thus, elevated cAMP levels in PCK cholangiocytes may induce a permanent activation of apical CFTR, leading to continuous Cl− efflux into the bile duct lumen, which in turn is exchanged for HCO− 3 through AE2, thus accounting for ion-driven water transport via AQP1. Basolaterally overexpressed AQP1, CFTR, and AE2 may facilitate basolateral-to-apical ion/water movement, accounting for increased luminal fluid accumulation. Thus, elevated expression and altered topography of AQP1, CFTR, and AE2 are associated with hepatic cyst expansion in ARPKD.

CONCLUSION Cholangiocyte cilia functioning as mechano-, osmo-, and chemosensory organelles play important roles in liver health and disease. In liver health, primary cilia are involved in regulation of ductal bile formation; when the structures and/or functions of these organelles are affected by mutations in genes encoding specific proteins, the cholangiociliopathies result. Understanding cholangiocyte function controlled by extracellular stimuli via ciliary-associated mechanisms in normal and pathological conditions should benefit the biology of epithelial cells in general and clinical hepatology in particular. Indeed, clinical trails based on recent developments in physiology and pathophysiology of cholangiocyte primary cilia are underway to assess novel therapeutic approaches in humans.

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SECTION C : SINUSOIDAL CELLS

26

The Hepatic Sinusoidal Endothelial Cell: Morphology, Function, and Pathobiology Laurie D. DeLeve Division of Gastrointestinal and Liver Diseases, University of Southern California, Keck School of Medicine, Los Angeles, CA, USA

INTRODUCTION It was long a point of controversy whether the endothelial cells of the microcirculation and the Kupffer cells were of the same cell type or different. Recognition of the hepatic sinusoidal endothelial cell (SEC) as a unique cell type distinct from the Kupffer cell did not occur until Eddie Wisse combined perfusion fixation with electron microscopy [1, 2]. The next major advance in the field was the development of the first method to isolate a pure population of SECs [3, 4] that could then be studied in primary culture. This development permitted characterization of SEC functions. One of the remaining barriers to SEC research is the expense of the equipment needed for the original method of isolating SECs; that is, a centrifuge optimized for elutriation and an elutriator rotor. This has spawned a number of other methods for isolating SECs, some of which have never been properly validated [5]. As a result, there are many conflicting observations about SECs in the literature, based on studies using different methods to isolate SECs [6]. Three of the most commonly used methods for SEC isolation are briefly described at the end of this chapter. “Endothelial biomedicine” has come into its own, as evidenced most recently by two multi-author textbooks The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

[7, 8] and a recurring Gordon Conference on “Endothelial cell phenotypes in health and disease.” Electron microscopy, cell culture, and proteomic and genomic studies have changed our perspective on endothelial cells. These cells are no longer viewed as simple cells whose only function is to line the vascular spaces. The endothelium is now viewed as a physiologically important organ with complex functions and with tremendous heterogeneity, sometimes even within a single vascular loop. The SEC is one of the many fascinating members of this family. Although our studies of the SEC are still in their infancy, the literature is already too vast to cover all the important aspects of the SEC. This chapter reviews the normal SEC morphology and function, as well as some of the diseases that have been recognized to date as involving the SEC.

MORPHOLOGY SEC morphology SECs have a morphological phenotype that is unique among mammalian endothelial cells (Figure 26.1).

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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space of Disse sinusoid

Figure 26.1 Scanning electron micrograph of sinusoidal endothelial cells within the hepatic sinusoid. The image demonstrates a hepatic sinusoid lined by sinusoidal endothelial cells with fenestrae grouped in sieve plates. Hepatocyte villi can be seen in the space of Disse. Courtesy of Margaret McCuskey and Robert S. McCuskey, University of Arizona

Endotheliums of the microcirculation can be categorized as continuous or discontinuous (Table 26.1). Continuous endothelial cells have continuous cytoplasm and the only areas of fusion between the luminal and abluminal plasma membranes are at the cell junctions. Discontinuous endotheliums have gaps within the cell or between adjacent cells. Fenestrated endothelial cells are a subset of discontinuous endotheliums that have pores within them which connect the luminal and abluminal surfaces of the cell. Fenestrated endothelial cells have pores that are either closed by a diaphragm or are fully open. Endothelial fenestrae are arranged in clusters, which were termed “sieve plates” when they were first described in the hepatic sinusoid [1]. Finally, continuous and discontinuous endothelial cells may or may not have an underlying organized basement membrane. SECs and endothelial cells of the renal glomerulus are the only endothelial cells in mammals with open (i.e. non-diaphragmed) fenestrae, but SECs lack an organized basement membrane and renal glomerular

endothelial cells have an organized basement membrane. Thus the liver microcirculation has the most porous of all endothelial barriers. The endothelial cells of the liver, bone marrow, and spleen are all called sinusoidal endothelial cells. “Sinusoidal” is said to describe the more tortuous course of the capillaries and the presence of a discontinuous endothelium. However, the morphological phenotype of these three SECs is very different. Splenic endothelial cells have continuous cytoplasm; the cells themselves are rod-shaped and run parallel to the longitudinal axis of the capillary, whereas the basement membranes form ring-like structures around the capillaries [9–11]. The bone marrow has diaphragmed, fenestrated endothelial cells [12] and a discontinuous, irregular basement membrane [13]. Both the bone marrow and the spleen have inter-endothelial cell slits that allow red cells to migrate through the sinusoids. In this chapter, “SEC” will refer to cells from the hepatic sinusoid. The dimension of SEC fenestrae depends on the method of visualization. By transmission electron microscopy the diameter of fenestrae ranges from 150 to 175 nm, and by scanning electron microscopy from 105 to 110 nm [14]; the smaller size on scanning electron microscopy has been attributed to shrinking during preparation of the sample. The number of fenestrae, porosity (number of fenestrae per µm2 ), number of fenestrae per sieve plate, and size of sieve plates are greater in the centrilobular region than in the periportal region [15, 16].

Regulation of normal SEC phenotype The unique SEC phenotype is maintained by its proximity to other neighboring liver cells [17], but this paracrine effect requires downstream autocrine regulation. Paracrine secretion of vascular endothelial growth factor (VEGF) by adjacent hepatocytes and hepatic stellate cells (HSCs) stimulates SEC production of nitric oxide (NO) by

Table 26.1 Endothelial cell phenotypes Continuous endothelium

Discontinuous, not fenestrated

Lung

Spleen

Heart Brain Muscle Lymph nodes

Fenestrated Diaphragmed fenestrae

Open fenestrae

Peritubular capillary ascending vasa recta Intestinal villi Pancreas Adrenal cortex Endocrine glands Choriocapillaries of brain and eye Bone marrow

SEC Glomerulus

26: THE HEPATIC SINUSOIDAL ENDOTHELIAL CELL: MORPHOLOGY, FUNCTION, AND PATHOBIOLOGY

endogenous nitric oxide synthase (eNOS) [18]. Exogenous VEGF alone mimics the effect of co-culture with either hepatocytes or HSCs. Inhibition of eNOS completely blocks the effect of VEGF on SEC phenotype, indicating that autocrine production of NO is essential to the VEGF effect [18]. However, VEGF contributes to fenestration of various types of endothelium [19]. Application of exogenous VEGF induces fenestrae in vitro in non-fenestrated endothelial cells [20]. High constitutive expression of VEGF by adjacent epithelial cells is characteristic of various organs with fenestrated endothelium, including the glomerulus and the choroid plexus [19]. Given that other fenestrated endothelial cells are exposed to high VEGF levels but do not have a phenotype combining open fenestrae and lack of a basement membrane, it is likely that micro-environmental factors other than just high VEGF levels contribute to the induction and maintenance of this unique SEC phenotype.

Sinusoidal morphology The normal liver is comprised of cords of hepatocytes that are two hepatocytes thick (Figure 26.2). Bile flows within the canaliculus that lies between the apical surfaces of the hepatocyte. The sinusoidal surface of the hepatocyte is bordered by the space of Disse. The space of Disse is a virtual space composed of a loosely organized extracellular matrix, hepatocyte villi, and hepatic stellate cells. The hepatic stellate cells are vitamin A-storing, contractile pericytes with dendritic-type cytoplasmic processes which are in contact with hepatocytes and other cytoplasmic processes that encircle SECs and regulate the caliber of the sinusoid. The tissue macrophage of the liver, the Kupffer cell, resides on the luminal

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surface of the SEC. Other non-parenchymal cells include pit cells, the liver-associated natural killer cells that adhere to the luminal side of the SEC, and dendritic cells, which are antigen-presenting cells and which may reside in the lumen, the space of Disse, or within the parenchyma [21–23].

FUNCTION Functional consequences of sinusoidal morphology Oxygen delivery Blood flow to the liver is a combination of arterial and venous blood. Thirty percent of blood flow to the liver is well-oxygenated blood provided by the hepatic artery, whereas 70% of the blood supply is poorly oxygenated blood derived from the portal vein. Normal adult arterial pO2 is 90–100 mmHg, whereas the reported value for portal vein pO2 is 45–55 mmHg [24]. Consequently, tissue oxygenation of the liver is lower than that for other tissues. Reported values depend on the method used, but one mouse study found liver tissue pO2 to be 14 mmHg (i.e. 14 torr or 2%), whereas normal tissue pO2 is 21 mmHg (i.e. 3%). Fick’s law of diffusion states that oxygen delivery is inversely correlated with distance. The thin SEC cytoplasm, open fenestrae, and lack of an organized basement membrane reduce the distance from the sinusoid to the hepatocyte and thereby optimize oxygen deliver. One may speculate that the open structure of the liver microcirculation is a tissue response to relative hypoxia. As will be discussed in Section “Capillarization and pseudocapillarization”, in various forms of Canaliculus

Hepatocytes Stellate cell Kupffer cell Sinusoidal endothelial cell

Sieve plate of SECs

sinusoid

Sinusoidal endothelial cell space of Disse

Figure 26.2 Cartoon of the hepatic sinusoid. The image depicts the hepatic sinusoid lined by sinusoidal endothelial cells, with Kupffer cells on the luminal side and the space of Disse on the contraluminal side. Stellate cells reside in the space of Disse and make contact with both sinusoidal endothelial cells and hepatocytes

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liver disease these adaptations of the SEC are lost and the hepatocyte becomes hypoxic.

Sieve function Dietary triglycerides, cholesterol, phospholipids, and apolipoproteins B48, AIV, and AI are formed into chylomicrons by the enterocyte and secreted into the mesenteric lymphatics [25]. Chylomicrons travel through the lymphatic system and enter the systemic circulation via the thoracic duct. In the circulation, chylomicrons are metabolized by endothelial lipoprotein lipase to chylomicron remnants. Chylomicron remnants smaller than 100 nm pass through SEC fenestrae into the space of Disse, as first noted by Wisse when he described SEC morphology [1]. The critical role of the liver in chylomicron remnant clearance was first demonstrated in studies with functional hepatectomy [26]. Subsequent electron microscopic studies demonstrated that the size of SEC fenestrae did indeed determine the size of chylomicron remnants cleared by the liver [27]. Similarly, chemically-induced defenestration leads to a decrease in chylomicron remnant clearance and an accompanying increase in plasma lipids [28], more definitively demonstrating the importance of SEC fenestration in lipid clearance. The porous nature of the hepatic microcirculation also enhances drug clearance by the liver. In tissues with continuous endothelium, only unbound drug—that is, drug that is not bound to serum proteins—leaves the circulation. In the liver, both unbound and bound drug enter the space of Disse. As unbound drug is taken up by hepatocytes from the space of Disse, the free and bound fractions within the space of Disse can re-equilibrate, allowing more unbound drug to be taken up. Thus the fraction of drug cleared during one pass through the liver can exceed the unbound fraction.

SEC functions Scavenger function of the SEC SECs and Kupffer cells together form the liver portion of the reticuloendothelial system. SECs contribute significantly to innate immunity by clearing waste macromolecules and antigens, whereas Kupffer cells phagocytose larger particulate matter and insoluble waste. Smedsrod et al. have outlined several characteristics that favor the contribution of SECs and Kupffer cells to the reticuloendothelial system [29], which will be outlined here. SECs are the first contact point for gut-derived macromolecules and antigens that enter through the portal vein. SECs form a large surface area and sinusoidal flow is slow and intermittent, particularly in the periportal region, which increases exposure of SECs to macromolecules and antigens. SECs have numerous

coated pits that have a cationic surface charge, which aids endocytosis of negatively charged molecules [30]. There are three distinct endocytosis receptors that rapidly take up ligands by receptor-mediated endocytosis: hyaluronan receptor/scavenger receptor, mannose receptor/collagen, α-chain receptor, and Fc gamma receptor. Through these receptors, SECs take up large amounts of colloidal and macromolecular waste, such as breakdown products of extracellular matrix, cellular constituents such as lysosomal enzymes, IgG-antigen immune complexes, CpG-oligonucleotides (present in bacterial DNA), advanced glycation end products, acetylated and oxidized low-density lipoprotein, and modified blood proteins [29, 31–36]. Specific activity of several lysosomal enzymes in SECs is as high as or higher than in Kupffer cells and much higher than in hepatocytes, so that endocytosed macromolecules can be degraded [37]. Experimental studies using viral vectors in recent years have taught us that the liver plays a significant role in viral clearance. The assumption has been that this is due to clearance of viral particles by Kupffer cells. A recent review by Elvevold et al. [6] noted that viral particles form colloidal particles that are usually 95%, and cell yields are high: around 10–12 million cells per mouse liver or 80–120 million cells per liver from a 200 g rat. The disadvantages are that the equipment is very expensive and that optimal separation requires a large number of cells at the time of elutriation.

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Density gradient centrifugation plus selective adherence The second method separates out the SECs using Percoll density gradient centrifugation followed by selective adherence and has been described in detail for both the rat [157] and the mouse [158] (original description [159], detailed description for rat [160]). Selective adherence is based on the principle that Kupffer cells will attach firmly in a culture dish after 15–20 minutes; non-adherent SECs are then collected and plated. Kupffer cell adherence is improved when the non-parenchymal mixture obtained after density gradient centrifugation is plated on glutaraldehyde-fixed albumin [161]. Cells isolated by this method have fenestrae organized in sieve plates and are endocytic, cell purity is 95%, viability is close to 100%, and yields are around 10–15 million cells per mouse liver or 150–200 million cells per rat liver. The advantages of this method are that it is rapid and inexpensive; the disadvantage is that it requires selective adherence to provide highly purified cells.

Variable first step plus immunomagnetic separation The third method of isolation uses immunomagnetic isolation; there are multiple variations to this method. After collagenase perfusion, the liver slurry may first undergo a density gradient step. The cell suspension is then incubated with magnetic beads with an attached antibody. After an incubation period, cells that have antibody attached can be isolated using a magnet. The major advantage of this approach is that it is relatively rapid and inexpensive. There are however several potential disadvantages. The major concern is that there are no known antigens specific to SECs, so that immunomagnetic separation may yield a mixed population of cells. Antigenic targets that have been used for immunomagnetic separation include CD31 (not present on the SEC surface [18, 46]), SE-1 (identity unpublished), Fc gamma receptor III/II (present on dendritic cells and macrophages), CD105 (endoglin, present on activated stellate cells and macrophages), CD146 (melanoma cell adhesion molecule (MelCAM or MCAM), present on hepatic stellate cells and other mesenchymal cells), and stabilin-2 (present on monocyte-derived macrophages). Most papers have confirmed that the population isolated by immunomagnetic separation is about the right size, takes up diL-labeled acetylated low-density lipoprotein (diL-Ac-LDL), and forms a cobblestone-type pattern in culture, but few of the investigators or commercial vendors of SECs using these methods have ever validated the identity of the population isolated using SEM. Viability and yields of the isolated population have rarely been provided in papers using these methods. Immunomagnetic

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separation from rat liver may yield as little as half a million to two million cells (compared to 100–200 million by the other two methods listed above), which raises concerns about isolation of sub-populations. An attempt to validate immunomagnetic separation based on CD31 demonstrated that the cells isolated looked like SECs on light microscopy, but did not have the scanning electron microscopic features of SECs [5]. The population isolated had a low number of fenestrae; yields were 640 000 cells per rat liver or 0.5% of the yield obtained by elutriation [5]. In our laboratory, purity of the isolated population is determined by uptake of diL-Ac-LDL, which identifies both SECs and Kupffer cells. The percentage of contaminating small Kupffer cells is determined by a peroxidase stain. Contaminating hepatocytes can be readily identified on light microscopy. In the opinion of this author, the next step forward in this field will occur when reviewers consistently demand validation of methods used for isolation of SECs. Criteria for methods of isolation might include documentation of fenestrae in sieve plates visualized by scanning electron microscopy, purity of the isolated cell population, and the number of cells isolated per liver. The latter criterion relates to the concern that a method that only isolates a tiny fraction of a cell population may be retrieving a subset of SECs rather than a representative population, or may even be isolating a completely different population of endothelial cells.

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cells: role of actin disassembly and ICAM-1. Liver Transpl , 9, 1286–94. Topp, S.A., Upadhya, G.A. and Strasberg, S.M. (2004) Cold preservation of isolated sinusoidal endothelial cells in MMP 9 knockout mice: effect on morphology and platelet adhesion. Liver Transpl , 10, 1041–48. Sindram, D., Porte, R.J., Hoffman, M.R., Bentley, R.C. and Clavien, P.A. (2000) Platelets induce sinusoidal endothelial cell apoptosis upon reperfusion of the cold ischemic rat liver. Gastroenterologia, 118, 183–91. Sindram, D., Porte, R.J., Hoffman, M.R., Bentley, R.C. and Clavien, P.A. (2001) Synergism between platelets and leukocytes in inducing endothelial cell apoptosis in the cold ischemic rat liver: a Kupffer cell-mediated injury. FASEB J , 15, 1230–32. Cottart, C.H., Do, L., Blanc, M.C., Vaubourdolle, M., Descamps, G., Durand, D., Galen, F.X. and Clot, J.P. (1999) Hepatoprotective effect of endogenous nitric oxide during ischemia-reperfusion in the rat. Hepatology, 29, 809–13. Morisue, A., Wakabayashi, G., Shimazu, M., Tanabe, M., Mukai, M., Matsumoto, K., Kawachi, S., Yoshida, M., Yamamoto, S. and Kitajima, M. (2003) The role of nitric oxide after a short period of liver ischemia-reperfusion. J Surg Res, 109, 101–9. Serracino-Inglott, F., Virlos, I.T., Habib, N.A., Williamson, R.C. and Mathie, R.T. (2003) Differential nitric oxide synthase expression during hepatic ischemia-reperfusion. Am J Surg, 185, 589–95. Oishi, A., Inagaki, M. and Tanaka, N. (1997) Correlation between nitric oxide production and preservation injury of sinusoidal endothelial cells during cold ischemia. Transplant Proc, 29, 1338–39. Ohmori, H., Dhar, D.K., Nakashima, Y., Hashimoto, M., Masumura, S. and Nagasue, N. (1998) Beneficial effects of FK409, a novel nitric oxide donor, on reperfusion injury of rat liver. Transplantation, 66, 579–85. Peralta, C., Rull, R., Rimola, A., Deulofeu, R., Rosello-Catafau, J., Gelpi, E. and Rodes, J. (2001) Endogenous nitric oxide and exogenous nitric oxide supplementation in hepatic ischemia-reperfusion injury in the rat. Transplantation, 71, 529–36. Geller, D.A., Chia, S.H., Takahashi, Y., Yagnik, G.P., Tsoulfas, G. and Murase, N. (2001) Protective role of the L-arginine-nitric oxide synthase pathway on preservation injury after rat liver transplantation. J Parenter Enteral Nutr, 25, 142–47. Man, K., Fan, S.T., Lo, C.M., Liu, C.L., Fung, P.C., Liang, T.B., Lee, T.K., Tsui, S.H., Ng, I.O., Zhang, Z.W. and Wong, J. (2003) Graft injury in relation to graft size in right lobe live donor liver transplantation: a study of hepatic sinusoidal injury in correlation with portal hemodynamics and intragraft gene expression. Ann Surg, 237, 256–64. Man, K., Lo, C.M., Ng, I.O., Wong, Y.C., Qin, L.F., Fan, S.T. and Wong, J. (2001) Liver transplantation in rats using small-for-size grafts: a study of hemodynamic and morphological changes. Arch Facial Plast Surg, 136, 280–85.

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135. Leong, S.S., Cazen, R.A., Yu, G.S., LeFevre, L. and Carson, J.W. (1992) Abdominal visceral peliosis associated with bacillary angiomatosis. Ultrastructural evidence of endothelial destruction by bacilli. Arch Pathol Lab Med , 116, 866–71. 136. Goerdt, S. and Sorg, C. (1992) Endothelial heterogeneity and the acquired immunodeficiency syndrome: a paradigm for the pathogenesis of vascular disorders. Clin Oral Investig, 70, 89–98. 137. Scoazec, J.Y., Marche, C., Girard, P.M., Houtmann, J., Durand-Schneider, A.M., Saimot, A.G., Benhamou, J.P. and Feldmann, G. (1988) Peliosis hepatis and sinusoidal dilation during infection by the human immunodeficiency virus (HIV). An ultrastructural study. Am J Pathol , 131, 38–47. 138. Wanless, I.R. (1990) Micronodular transformation (nodular regenerative hyperplasia) of the liver: a report of 64 cases among 2,500 autopsies and a new classification of benign hepatocellular nodules. Hepatology, 11, 787–97. 139. Hillaire, S., Bonte, E., Denninger, M.H., Casadevall, N., Cadranel, J.F., Lebrec, D., Valla, D. and Degott, C. (2002) Idiopathic non-cirrhotic intrahepatic portal hypertension in the West: a re-evaluation in 28 patients. Gut , 51, 275–80. 140. Ibarrola, C. and Colina, F. (2003) Clinicopathological features of nine cases of non-cirrhotic portal hypertension: current definitions and criteria are inadequate. Histopathology, 42, 251–64. 141. Nakanuma, Y., Hoso, M., Sasaki, M., Terada, T., Katayanagi, K., Nonomura, A., Kurumaya, H., Harada, A. and Obata, H. (1996) Histopathology of the liver in non-cirrhotic portal hypertension of unknown aetiology. Histopathology, 28, 195–204. 142. Shedlofsky, S., Koehler, R.E., DeSchryver-Kecskemeti, K. and Alpers, D.H. (1980) Noncirrhotic nodular transformation of the liver with portal hypertension: clinical, angiographic, and pathological correlation. Gastroenterology, 79, 938–43. 143. Wanless, I.R., Godwin, T.A., Allen, F. and Feder, A. (1980) Nodular regenerative hyperplasia of the liver in hematologic disorders: a possible response to obliterative portal venopathy. A morphometric study of nine cases with an hypothesis on the pathogenesis. Medicine, 59, 367–79. 144. Shimamatsu, K. and Wanless, I.R. (1997) Role of ischemia in causing apoptosis, atrophy, and nodular hyperplasia in human liver. Hepatology, 26, 343–50. 145. Bioulac Sage, P., Dubuisson, L., Bedin, C., Gonzalez, P., de Tinguy Moreaud, E., Garcin, H. and Balabaud, C. (1992) Nodular regenerative hyperplasia in the rat induced by a selenium-enriched diet: study of a model. Hepatology, 16, 418–25. 146. Croquelois, A., Blindenbacher, A., Terracciano, L., Wang, X., Langer, I., Radtke, F., Heim M.H. (2005) Inducible inactivation of Notch1 causes nodular regenerative hyperplasia in mice. Hepatology, 41, 487–496.

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147. Maione, D., Di Carlo, E., Li, W., Musiani, P., Modesti, A., Peters, M., Rose-John, S., Della Rocca, C., Tripodi, M., Lazzaro, D., Taub, R., Savino, R. and Ciliberto, G. (1998) Coexpression of IL-6 and soluble IL-6R causes nodular regenerative hyperplasia and adenomas of the liver. EMBO J , 17, 5588–97. 148. McEntee, M.F., Wright, K.N., Wanless, I., DeVovo, R., Schneider, J.F. and Shull, R. (1998) Noncirrhotic portal hypertension and nodular regenerative hyperplasia of the liver in dogs with mucopolysaccharidosis type I. Hepatology, 28, 385–90. 149. Snover, D.C., Weisdorf, S., Bloomer, J., McGlave, P. and Weisdorf, D. (1989) Nodular regenerative hyperplasia of the liver following bone marrow transplantation. Hepatology, 9, 443–48. 150. Haboubi, N.Y., Ali, H.H., Whitwell, H.L. and Ackrill, P. (1988) Role of endothelial cell injury in the spectrum of azathioprine-induced liver disease after renal transplant: light microscopy and ultrastructural observations. Am J Proctol Gastroenterol Colon Rectal Surg, 83, 256–61. 151. Mion, F., Napoleon, B., Berger, F., Chevallier, M., Bonvoisin, S. and Descos, L. (1991) Azathioprine induced liver disease: nodular regenerative hyperplasia of the liver and perivenous fibrosis in a patient treated for multiple sclerosis. Gut , 32, 715–17. 152. Russmann, S., Zimmermann, A., Krahenbuhl, S., Kern, B. and Reichen, J. (2001) Veno-occlusive disease, nodular regenerative hyperplasia and hepatocellular carcinoma after azathioprine treatment in a patient with ulcerative colitis. Eur J Gastroenterol Hepatol , 13, 287–90. 153. Vora, A., Mitchell, C.D., Lennard, L., Eden, T.O.B., Kinsey, S.E., Lilleyman, J. and Richards, S.M. (2006) Toxicity and efficacy of 6-thioguanine versus 6-mercaptopurine in childhood lymphoblastic leukaemia: a randomised trial [see comment]. Lancet , 368, 1339–48. 154. Dubinsky, M.C., Vasiliauskas, E.A., Singh, H., Abreu, M.T., Papadakis, K.A., Tran, T., Martin, P., Vierling, J.M., Geller, S.A., Targan, S.R. and Poordad, F.F. (2003) 6-thioguanine can cause serious liver injury in inflammatory bowel disease patients. Gastroenterology, 125, 298–303. 155. Arotcarena, R., Cales, V., Berthelemy, P., Parent, Y., Malet, M., Etcharry, F., Ferrari, S. and Pariente, A. (2006) Severe sinusoidal lesions: a serious and overlooked complication of oxaliplatin–containing chemotherapy? Gastroenterol Clin Biol , 30, 1313–16. 156. Steensma, A., Beamand, J.A., Walters, D.G., Price, R.J. and Lake, B.G. (1994) Metabolism of coumarin and 7-ethoxycoumarin by rat, mouse, guinea pig, cynomolgus monkey and human precision-cut liver slices. Xenobiotica, 24, 893–907. 157. Smedsrød, B. and Pertoft, H. (1985) Preparation of pure hepatocytes and reticuloendothelial cells in high yield from a single rat liver by means of Percoll centrifugation and selective adherence. J Leukocyte Biol , 38, 213–30.

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158. Hansen, B., Arteta, B. and Smedsrød, B. (2002) The physiological scavenger receptor function of hepatic sinusoidal endothelial and Kupffer cells is independent of scavenger receptor class A type I and II. Mol Cell Biochem, 240, 1–8. 159. Eriksson, S., Fraser, J.R.E., Laurent, T.C., Pertoft, H. and Smedsrød, B. (1983) Endothelial cells are a site of uptake and degradation of hyaluronic acid in the liver. Exp Cell Res, 144, 223–28.

160. Pertoft, H. and Smedsrød, B. (1987) Separation and characterization of liver cells, in Cell Separation: Methods and Selected Applications (eds T.G.I. Pretlow and T.P. Pretlow), Academic Press, pp. 1–24. 161. Laakso, T. and Smedsrød, B. (1987) Cellular distribution in rat liver of intravenously administered polyacryl starch and chondroitin sulfate microparticles. Int J Pharm, 36, 253–62.

27

Fenestrations in the Liver Sinusoidal Endothelial Cell Victoria C. Cogger and David G. Le Couteur Centre for Education and Research on Ageing, University of Sydney and Concord RG Hospital, Sydney, Australia

INTRODUCTION

HISTORICAL BACKGROUND

The liver sinusoidal endothelial cell (LSEC) occupies a strategic position in the liver, separating blood in the sinusoid from the extracellular space of Disse and surrounding sheets of hepatocytes [1]. It has long been recognized that LSECs have a role facilitating, and perhaps regulating, the bi-directional transfer of substrates between the blood and the liver parenchyma, forming a blood–hepatocyte barrier [2–4]. LSECs have an unusual morphology compared to other capillaries, which underpins their physiological role in substrate transfer. The cytoplasmic extensions of LSECs are very thin and perforated with pores called fenestrations that are true discontinuities in the endothelium [5]. The regulation and normal function of these fenestrations, and their changes in disease and aging, support the concept that fenestrations have widespread implications for hepatic metabolism and subsequent systemic exposure to many substrates and toxins [3]. Recently, the other roles of LSECs in endocytosis [6] and the immune response [7, 8] have been identified. Any understanding of the LSEC must recognize that it is not a conventional endothelial cell, but a unique hybrid with features of macrophages, antigen presenting cells, and lymphoid endothelial cells [9].

In the late nineteenth century, it was deduced on the basis of injection of various dyes into the liver blood vessels that there is free communication via small channels between the hepatic capillaries and perivascular lymphatics [10, 11]. Fraser and Fraser injected frog liver vessels and concluded that “blood serum is brought into much closer relation to epithelial cells than has been before understood” as a result of passages that “are in direct communication with the blood vessels though of course far too small to transmit the corpuscles” [12]. Herring and Simpson performed a light microscopic examination of the cat liver and noted the “sinusoidal character of the blood-vessels and the incomplete nature of their endothelial lining” [10]. Although this is suggestive of fenestrations, it should be noted that the size of most fenestrations (50–200 nm) falls below that detectable using light microscopy. They injected “melted and filtered hogs lard” into the portal vein (PV) of a dog liver and found uptake of fat globules by the liver cells, demonstrating the passage of fat particles across the sinusoidal endothelium [10]. In the mid 1950s, electron microscopy was first used to detect pores or fenestrations in the sinusoidal endothelium that allowed direct contact between liver cells and blood [13–17]. Wisse subsequently

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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(a)

(b)

Kupffer cell Hepatic sinusoid

Space of Disse Lipoprotein Agranular reticulum

Golgi complex

(c)

Figure 27.1 Early transmission electron micrographs of the rat liver sinusoid demonstrating fenestrations in the LSEC. (a) [15]. J, fenestrations; G, endothelial cell; S, space of Disse; L, sinusoidal lumen. Reprinted from Bennett et al. and used with permission from The American Physiological Society. (b) [1]. →, fenestrations; end, endothelial cell; DS, space of Disse; L, sinusoidal lumen reproduced from Wisse (1970), Copyright  1970, Elsevier. (c) By 1968 the concept of fenestrations had entered texts [17]. Reproduced from Bloom and Fawcet (1968), Copyright  1968, Elsevier. The diagram shows fenestrations and lipoproteins in the space of Disse, but also the incorrect historical assumption that Kupffer cells also line the sinusoid

established the structure of fenestrations and identified their arrangement in sieve plates (SPs), in a publication that has become the foundation for future studies on fenestrations [1] (Figure 27.1). The demonstration by Fraser of the role of fenestrations in regulating the passage of lipoproteins according to size (“the liver sieve”) had implications for the development of atherosclerosis [2, 18]. Arias [19–21] and Oda [22, 23] recognized that fenestrations are dynamic structures that can be altered by numerous endogenous and exogenous agents, foreseeing the physiological role and therapeutic potential of regulating fenestration dimensions.

MORPHOLOGY OF SINUSOIDAL VESSELS AND THEIR FENESTRATIONS AND SIEVE PLATES Hepatic sinusoids connect afferent portal triads to exiting central hepatic venules, and supply the hepatocytes which

lie in sheets of single cells between the sinusoids [2, 24, 25]. At the periportal region (zone 1), the sinusoids form a reticulated network of vessels that become more linear and parallel as they enter the pericentral region (zone 3). The sinusoids have an average diameter of approximately 5–10 µm and occupy between 10 and 30% of the total liver volume [26, 27]. The fractal dimension of the sinusoidal vessels (a measure of complexity) exceeds two, also indicating the space-filling characteristic of the sinusoids [27]. This degree of vascularity facilitates the exchange of substrates between blood and the liver and provides an extensive endothelial surface area for interactions with circulating immune cells and various colloid and soluble macromolecular waste products. LSECs, which represent 2.5% of total liver volume and 15–20% of all liver cells [19, 28], are highly specialized endothelial cells that line the wall of the hepatic sinusoid. The cytoplasmic extensions of LSECs are very thin and perforated with fenestrations, which are circular and oval pores approximately 50–200 nm in diameter. Fenestrations are complete gaps in the endothelium, lacking

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10 µm

50 µm (a)

391

(b)

1 µm

1 µm (c)

1 µm (d)

5 µm

(e)

1 µm

(f)

Figure 27.2 Electron micrographs (EMs) of LSECs and fenestrations. (a) Scanning EM of vascular cast showing portal vein (PV) branching into sinusoids (S). (b) Scanning EM of sinusoids (S) showing intercalated plates of hepatocytes (H). Fenestrations (→) are apparent in the endothelial wall. (c) Scanning EM of sinusoidal endothelial lumen showing fenestrations clustered into sieve plates (SP). (d) Transmission EM showing sinusoidal lumen, LSEC (E), stellate cell process (SC), and extravascular space of Disse containing microvilli. (e) Scanning EM of isolated LSEC showing fenestrations distributed in the cytoplasm away from nucleus (Nuc). (f) Scanning EM of isolated LSEC showing fenestrations clustered in sieve plates and a network of fenestrations resembling a vesiculo-vacuolar organelle (VVO). Magnification bars are shown

either a diaphragm or underlying basal lamina. There are approximately 3–20 fenestrations per µm2 of endothelial surface, and between 2 and 20% of the surface of the LSEC is covered by fenestrations. Some but not all studies have suggested that there is a zonal gradient, with slightly smaller fenestrations in the periportal sinusoids

and greater porosity in the pericentral sinusoids [2, 5, 25, 26, 29–31] (Figure 27.2). The diameter of fenestrations has a Gaussian distribution, skewed to the right by the presence of some larger pores [5, 26]. Attempts have been made to classify fenestrations on the basis of their diameter [32]. Very small

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non-perforating pores are often called pits, and larger pores (greater than 200–300 nm diameter) are usually called gaps [5]. Gaps represent large deficits in the liver endothelium, probably caused by loss or fusion of all the fenestrations in the liver SPs [5, 32, 33]. Gaps seem to occur as a result of cellular injury, technical issues related to high-pressure perfusion [5, 26, 34], hypoxia [35], fixation, or pathological states [36]. The absence of large chylomicrons in the space of Disse has been proposed as evidence that gaps do not normally occur in vivo [5]. Sometimes dumbbell-shaped fenestrations are seen, presumably occurring as a result of fusion of two adjacent fenestrations. Fenestrations are also found in meshlike structures reminiscent of vesiculo-vacuolar organelles (VVO) [37] and pored domes [38]. Fenestrations have been detected using a variety of methodologies (transmission electron microscopy, scanning electron microscopy, electron tomography, freeze–fracture microscopy, cryo-electron microscopy, and atomic force microscopy [2, 5, 33, 39]) and have been observed in numerous vertebrate species (human, rat, mouse, guinea pig, sheep, goat, rabbit, fowl, monkey, baboon, bat, kitten, dog, turtle, gold fish [29, 36, 38, 40–48]), where they have similar appearances, including SP formation (Table 27.1). Even so, the exact size and morphology of fenestrations are difficult to measure [39]. On scanning electron microscopy, it is problematical to differentiate smaller fenestrations from caveolae and other membrane pits and this distinction can only be made by demonstrating the presence or absence of complete discontinuity of

the cytoplasm on transmission electron microscopy. Furthermore, scanning electron microscopy underestimates the size of fenestrations. There is shrinkage artifact of approximately one-third induced by critical point drying utilized in scanning electron microscopic preparations, and gold coating used in scanning electron microscopy will also reduce the apparent diameter of fenestrations by about 10–20 nm [39, 49]. Thus the diameter of fenestrations was 108 nm on scanning electron microscopy versus 161 nm on transmission electron microscopy in the rat [50]. Likewise in dogs, the values were 118 and 177 nm, respectively [48]. Recently, atomic force microscopy of wet fixed cells indicated that fenestrations in living LSECs have a diameter of 223 nm and a depth of 183 nm [39]. Fenestrations are either scattered individually across the endothelial surface or clustered into groups of tens to hundreds called liver sieve plates. Between 60 and 75% of fenestrations are found within SPs in rats [30]. SPs are particularly apparent in healthy young liver endothelial cells and are decreased with actin disruptors such as cytochalasin B [58]. In isolated LSECs, there are usually tens of SPs present in the cytoplasmic extension of a single cell, representing many hundreds or even thousands of fenestrations per cell [59, 60]. Fenestrations may not be single and independent pores but instead interconnected labyrinthine structures, which have been seen on transmission electron microscopy of thicker cell sections and isolated cells [39, 61]. Immunogold electron microscopic studies have shown the presence of caveolin-1, endothelial nitric oxide synthase [62], and calcium- and calcium-magnesium-ATPase

Table 27.1 Fenestrations have been reported in all species studied, and a very wide range of species. Some of the many reports of different species are given in the table in order to show that fenestrations are widespread, and quite similar, in animals and humans Species Rat (zone 1) Rat (zone 3) Rat (zone 1) Rat (zone 3) Rat Rat Mouse Rabbit Rabbit Chicken Chicken Rainbow trout Gold fish Dog Sheep Baboon Baboon Baboon Human (zone 1) Human (zone 3) Human (zone 1) Human (zone 3)

Porosity (area %)

Diameter (nm)

Frequency (per µm2 )

Citation

9.6 28.5 6.0 ± 0.2 7.9 ± 0.3 4.1 ± 2.3 12.0 ± 2.1 4.1 ± 2.2 5.2 ± 0.9 4.0 ± 1.5 3.6 ± 1.6 2.2 ± 0.6

73 ± 0.13 94 ± 0.11 111 ± 1 105 ± 0.2 73 ± 1 110 ± 7 74 ± 4 60 ± 5 69 ± 8 99 ± 15 90 ± 18 123 50–200 118 ± 2 60 ± 2 50 ± 1 58 ± 1 82 — — 170 ± 12 160 ± 10

5.7 ± 0.1 10.2 ± 0.01 9.1 ± 0.3 13.3 ± 0.5 2.7 ± 1.1 12.4 ± 3.6

[30] [30] [26] [26] [51] [52] [41] [53] [52] [53] [52] [54] [46] [48] [36] [55] [43] [56] [29] [29] [57] [57]

6.7 2.6 ± 0.2 4.2 ± 0.5 1.8 7.6 9.1 3.4 ± 0.2 4.0 ± 0.4

17.3 ± 3.8 12.7 ± 2.5 3.9 ± 0.9 2.9 ± 0.3 7.2 12.1 ± 0.8 9.4 ± 0.9 3.3 19 23.5 9.8 ± 1.8 11.2 ± 2.6

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[63, 64] in the walls of the fenestrations. Caveolin-1 is connected to the actin cytoskeleton and its presence in fenestrations suggests a commonality between them and caveolae. However, research into the cellular biology of fenestrations has been limited by the ephemeral nature of fenestrations in isolated LSECs, the obvious inability to isolate fenestrations from endothelial cells, and the requirement for fixation and electron microscopy to identify fenestrations.

PHYSIOLOGICAL ROLES OF FENESTRATIONS The fenestrated endothelium acts as a filter and hence was termed “the liver sieve” [1, 18, 65]. Fenestrations permit the passage of a wide range of substrates (plasma and substrates within plasma, plasma proteins including albumin, smaller lipoproteins, colloidal particles, and polystyrene microspheres) into the extravascular space of Disse, although the proportion of each substrate that enters the space of Disse via fenestrations remains unknown [3]. Blood cells could massage fluid through the fenestrations by virtue of the fact that their diameter is greater than that of a typical sinusoid [5]. The narrowness of hepatic endothelial cells and the lack of basal lamina and collagen in the space of Disse ensure that any permeability barriers to the diffusion of substrates between blood and hepatocytes are also minimized [3]. Extensive investigations of substrate transfer by Goresky using a distributed physiological model and Roberts using the dispersion model have established that under normal conditions there is no barrier to the transfer of soluble substrates, albumin, and albumin-bound substrates at the LSEC—instead, transfer into the space of Disse is flow-limited [66–72]. Of particular clinical importance is the role of fenestrations in the hepatic metabolism of lipoproteins [2, 73]. The first stage in the metabolism of lipoproteins is the production of chylomicrons. Chylomicrons are triglyceride-rich, spherical lipoproteins formed in the intestine from dietary lipids. They are large particles with diameters of 100–1000 nm that are unable to pass through the fenestrations of the hepatic sinusoidal endothelium because of their size. Besides, most chylomicrons bypass the liver via the thoracic duct. Chylomicrons are metabolized to chylomicron remnants by lipoprotein lipase present on the endothelium of systemic capillaries. Chylomicron remnants are smaller particles (30–80 nm) that have acquired apoE. Remnants pass through the LSEC and are sequestered within the space of Disse for receptor-mediated uptake into hepatocytes [2, 74]. There is accumulating evidence for the role of fenestrations in regulating lipoprotein disposition in the liver. There is a close correlation between the diameter of fenestrations and chylomicron remnants in the space of Disse and chylomicrons larger than fenestrations have not been seen in the extracellular space [65]. Membranes

393

isolated from hepatocytes cannot differentiate between chylomicrons and chylomicron remnants, whereas in vivo, livers selectively take up chylomicron remnants [75]. Electron microscopy has demonstrated that large chylomicrons are only found in the sinusoidal blood whereas smaller chylomicron remnants are also observed in the space of Disse. There is differential trapping by the liver of radiolabeled chylomicrons of different sizes such that smaller particles are trapped to a greater extent than those larger than 100 nm [18]. Similar results indicating exclusion on the basis of size have been reported for large and small liposomes [76] and colloidal gold particles of different diameters [77]. Defenestration will lead to impaired clearance of chylomicron remnants after meals, and because remnants are still relatively rich in triglycerides, this is manifested as postprandial hypertriglyceridemia [2, 74]—a condition first described in 1641 as Tulp syndrome [78]. Recently it has been shown that conditions associated with reduced fenestrations cause impaired lipoprotein uptake and hypertriglyceridemia: old age [79]; vascular endothelial growth factor receptor (VEGFR) knockout mice [80]; and treatment with the remarkable defenestrating surfactant, poloxamer 407 [81]. Conversely, the increase in fenestrations and gaps that occurs following partial hepatectomy [82] and some toxic LSEC injury might contribute to increased fat uptake and steatosis. Both diameter and frequency of fenestrations will determine diffusive and convective transfer across the LSEC, whereas permselectivity is determined only by fenestration diameter [4]. It is possible to quantify the effects of changes in the liver endothelium and fenestrations on the transfer of substrates such as lipoproteins by application of the engineering principles related to membrane filtration, specifically ultrafiltration. In ultrafiltration, the volume flux (J ) is described by the Hagen–Poiseuille formula: 2 J = f R8ηlP where f is the porosity of the membrane, R the diameter of the pores, P the pressure gradient across the membrane, η the viscosity, and l the thickness of the membrane. The membrane permeability, Q, for a substance of molecular weight M is given by the equation [83]: Q = K. fl M 0.76 where K is a constant, f the porosity, and l the thickness of the membrane. In defenestrating conditions such as liver disease and aging, there is typically a reduction of porosity of 30–50%, a reduction of fenestration diameter of 10%, and an increase in endothelial thickness of 50%. Changes of this magnitude will be associated with a reduction in flux and membrane permeability by two-thirds or more. The reduction in the diameter of fenestrations will also influence the size of particles that are able to transfer across the endothelium. It can be seen from Figure 27.3 that alteration of the diameter of the fenestrations between 25 and 150 nm has a dramatic effect on the transfer of lipoproteins depending upon their diameter [41]. This is likely to be of particular relevance to chylomicron remnants, because their diameters range from 30 to 80 nm.

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Sieving coefficient

0.8

Fenestration diameter 25 nm 50 nm 75 nm 100 nm 150 nm

0.6

0.4

0.2

0.0 0

20

40

60 80 100 Lipoprotein diameter (nm)

120

140

Figure 27.3 The effect of fenestration diameter on the sieving coefficient (the fraction of particles able to pass through ultrafiltration pores) according to lipoprotein diameter [41]

Viral particles have also been postulated to enter the liver parenchyma via fenestrations, although membrane proteins are also important for the uptake of hepatotrophic viruses [84]. Fenestrations may be important in gene therapy for the transfecting virus to gain access to the liver cells, and interestingly most of the strategies that improve uptake of gene therapy vectors such as partial hepatectomy, liver ischemia, and cyclophosphamide cause increased porosity and/or gap formation in the liver sinusoidal endothelium [85]. For example, the human adenoviral vector is 93 nm with protruding fibers of 30 nm, thus was less able to infect rabbits with fenestral diameter of 103 nm than C57BL/6 mice with fenestral diameter of 141 nm. An increase in transfection rates was associated with ischemia-induced increase in fenestration diameter [86]. Fenestrations permit interactions between cells in the sinusoidal lumen and the space of Disse. This is especially important for the immune role of the liver. LSECs express many antigens important for interactions with leukocytes and lymphocytes and have a possible role in antigen presentation [87, 88]. The liver appears to be a key site for the development of immunotolerance by inducing apoptosis in lymphocytes [89]. Using a murine transgenic model of autoimmune hepatitis, it was shown that na¨ıve T-cells interact directly with hepatocytes through fenestrations (trans-endothelial hepatocyte lymphocyte interactions, TEHLIs), and this appears to be the first step in the development of immunotolerance [88, 90]. It was also found that activated lymphocytes and other leukocytes accessed the liver tissue in hepatitis via passage through the fenestrations, and conversely that loss of fenestrations almost fully abolished any hepatitis [90]. It is likely that

TEHLIs are the equivalent of the lymphocytes podosomes, which are projections that initiate transendothelial diapedesis in other circulatory systems [91]. It has also been suggested that contraction of the fenestrations might increase vascular resistance, thereby influencing hepatic blood flow and pressure [24, 25, 92]. Accordingly, an endothelin (ETA -R) antagonist was found to dilate fenestrations and cause a reduction in portal perfusion pressure of about 2.5 cmH2 O [93]. Finally, fenestrations are involved in the formation of hepatic lymph [3, 4]. The space of Disse is continuous with lymphatic vessels found in the portal triads [94–96]. This morphology suggests that plasma flows through the fenestrations and upstream along the space of Disse, finally emptying into the lymphatic vessels around the PV [3, 4, 94]. A hydrodynamic analysis of flow in the hepatic sinusoids, specifically application of Bernouilli’s law, is also consistent with the concept of retrograde plasma flow along the space of Disse [97]. The hepatic lymph has unusually high levels of plasma proteins [94] and the filtration coefficient is about five times greater than in other capillaries [4], indicating that there is no significant colloid osmotic pressure across the LSEC. This probably reflects the fact that, unlike parenchymal tissue elsewhere, the hepatocytes produce albumin, which is released into the extravascular space.

REGULATION OF FENESTRATIONS Fenestrations are dynamic structures that change in frequency and diameter in response to numerous stimuli

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395

Table 27.2 Physiological factors involved with the regulation of fenestrations in LSECs. In all studies, the quantification was performed using scanning electron microscopy and was performed in isolated LSECs, unless otherwise indicated. In some cases the values have been recalculated to provide comparable units (∼) between studies Porosity (area %) Treatment

Control

Vascular endothelial growth factor 10 ng ml−1 ∼1.3 100 ng ml−1 ∼1.3 10 ng ml−1 — 100 ng ml−1 — Actin (actin disruptors) Cytochalasin B 5 g ml−1 Cytochalasin B 10 µg ml−1 Cytochalasin B 10 µg ml−1 (perfused mouse liver) Cytochalasin B 10 µg ml−1 Cytochalasin B 10 µg ml−1 Latrunculin A 0.1 µg ml−1 Misakinolide 25 nM Swinholide A 25 nM Dihydrohalichondramide 100 nM Vasoactive agents Serotonin (rat in vivo) Serotonin Serotonin Noradrenaline (rat in vivo) Endothelin 1 Endothelin 1 Prostaglandin E1 ETA -R antagonist (BQ123) Phorbol myristate acetate Pantethine (rabbit)

Diameter (nm)

Frequency (per µm2 )

Treatment

Control

Treatment

Control

Treatment

Citation

∼2.8 ∼3.3 — —

∼167 ∼167 — —

∼176 ∼183 — —

∼0.6 ∼0.6 2.4 ± 0.7 2.4 ± 1.7

∼1.1 ∼1.3 3.4 ± 1.2 4.1 ± 0.7

[59] [59] [105] [105]

114 ± 37 130 ± 41 99 ± 18

157 ± 67 129 ± 16 103 ± 20

— 5.6 ± 0.6 14.0 ± 5.0

— 22.0 ± 2.0 23.0 ± 2.0

[58] [106] [106]

— — 7.7 ± 4.0 23.0 ± 3.2 10.0 ± 4.0 21.5 ± 3.2 — ∼11.2 ∼11.2 ∼12.3 ∼12.3 ∼10.6

— ∼24.8 ∼22.2 ∼21.2 ∼17.8 ∼12.8

— 211 ± 69 211 ± 69 215 ± 2 215 ± 2 212 ± 82

— 205 ± 67 201 ± 80 195 ± 1 158 ± 1 165 ± 60

5.5 3.2 3.2 3.4 3.4 3.0

1.4 1.0 1.0 0.2 0.2 0.2

9.5 ± 1.3 ∼7.5 ± 1.5 ∼7 ± 2 7.1 ± 0.3 9.1 ± 0.3 6.0 ± 0.2

[49] [107] [107] [108] [108] [109]

— — — — — — — — — 3.2 ± 1.7

— — — — — — — — — 5.8 ± 2.3

103 ± 1 164 ± 15 220 ± 66 103 ± 1 123 ± 35 161 ± 29 161 ± 29 133 ± 12 228 ± 49 61 ± 9

87 ± 1 127 ± 16 215 ± 72 94 ± 1 46 ± 21 101 ± 29 227 ± 40 338 ± 14 229 ± 52 74 ± 16

— — — — — — — 9.3 ± 0.7 2.2 ± 0.5 8.4 ± 3.7

— — — — — — — 8.1 ± 0.6 1.5 ± 0.3 10.3 ± 3.7

[110] [20] [111] [110] [112] [64] [64] [93] [111] [113]

in vitro. In vivo it is likely that fenestrations open and close in response to various stimuli such as inflammation, dietary fat load, and/or circulating vasoactive cytokines and hormones [19]. Local paracrine and autocrine factors presumably establish and maintain porosity at a level presumptively required for health. There are several issues that confound the interpretation of studies of regulatory factors. LSECs isolated from rat livers have been the major model for studying fenestration biology. This is dependent on the methodology, with some methods failing to generate well-fenestrated cells [60, 98, 99]. Isolated LSECs are only viable for one to two days and there is a dramatic change in fenestrations during this period [20, 98, 100–102]. Maintenance of fenestrations in isolated LSECs requires VEGF [59, 98–100] and extracellular matrix derived from the liver [102, 103]. It is likely that fenestrations are regulated in vivo by a variety of paracrine and circulating factors as well as the extracellular matrix, and of course these are absent in isolated cell studies [99, 100]. Another important issue relates to the measurement of the fenestrations and porosity. The formation of large gaps induced by non-specific cellular injury will appear to increase the average diameter of

± ± ± ± ± ±

the fenestrations, yet the mechanisms and implications of gap formation may be quite unrelated to those involved in the regulation of fenestrations. Porosity is calculated from the percentage of the surface area of the cytoplasm that is perforated by fenestrations. These measurements may be quite subjective and have not been standardized. Even so, thousands of fenestrations are measured in many studies and this no doubt increases confidence in the reported values. Accepting such caveats, there are many reports of agents that influence fenestrations [104] but current focus has been on the roles of the actin cytoskeleton and vascular endothelial growth factor (Table 27.2).

Calcium–calmodulin and the actomyosin cytoskeleton In endothelial cells, calcium that enters the cell from the extracellular space binds calmodulin, which leads to the activation of myosin light chain kinase (MLCK). Activated MLCK phosphorylates the myosin light chain, which allows crossbridges to form with actin and cell retraction to occur. Activation of the Rho pathway, for

396

THE LIVER: REGULATION OF FENESTRATIONS

example by ligands acting on the G-protein-coupled receptor, also increases phosphorylation of the myosin light chain [114]. Actin retraction in capillaries and post-capillary venules opens up the gaps between adjacent cells and enhances vascular leakiness. Liver SPs and fenestrations themselves are supported by the actin cytoskeleton [39, 58, 107–109, 115–117]. A variety of actin-based structures involved with the maintenance of fenestrations have been identified, such as the fenestrae-associated cytoskeleton ring, SP-associated cytoskeleton, fenestrae forming center, and defenestration-associated center. It has been proposed that these are involved in the maintenance of fenestrations [61, 108, 109, 118]. Recently it has been shown that filamentous actin is absent in the cytoplasm separating fenestrations in the SPs [39]. The diameter of the fenestration-associated cytoskeleton ring is somewhat larger than a fenestration (270 ± 58 nm) [117]. Agents that disrupt actin, such as cytochalasin B, misokinolide, and latrunculin, increase the number of fenestrations, usually in the order of twofold [103, 106–109, 119] (Table 27.2). This is associated with a marked reduction in SPs [58], and in most but not all studies the size of the fenestrations is not increased. On the other hand, microtubule inhibitors do not influence fenestrations [106]. Fenestrations probably form as a result of fusion of opposing plasma membranes [106, 107, 119, 120] by a process of membrane or pore fusion [121]. In other cell types, this process generates membrane pores with diameters of 100–500 nm [122] and is highly sensitive to actin-modifying agents [123]. The actin cytoskeleton prevents the development of protein-free patches in membranes and the subsequent contact of the protein-free membrane domains that are required for membrane fusion to be initiated (“actin barrier”). Actin is then involved in the subsequent development and stabilization

Calcium

Calmodulin

Rho

Impaired completion of membrane fusion

Myosin light chain kinase

Myosin light chain phosphorylation

G-proteincoupled receptor

of fusion pores [121, 123]. Thus there is a biphasic dose-dependent response to actin-modifying agents, with low doses stimulating fusion and high doses inhibiting fusion [123]. It has been proposed that actin depolymerization is initially required to allow membranes to dock, whereas the final membrane fusion process requires the re-establishment of an actin network [123]. The time for the opening of a fusion pore in other cells is less than 20 minutes [122], which provides some estimate of the rate of fenestration formation. In LSECs, actin reorganization might allow more cell-membrane fusion to be initiated by removing the intervening actin barrier, leading to increased fenestrations. This is consistent with the observation that actin disruptors cause increased fenestrations and decreased SPs. On the other hand, a different level of actin reorganization will impede the subsequent completion of membrane fusion, causing defenestration. The effects may also be time-dependent and many studies showing increased fenestration were measured over longer periods than those showing reduced fenestrations. The roles of various fusion proteins (e.g. SNAREs, Rabs) [121] and other pathways involved in the regulation of membrane fusion [121, 123] have not been investigated with respect to fenestration formation. However, a complete peristomal ring of sterols, considered to contribute to membrane fusion, has been detected lining the rim of fenestrations, but not around gaps [120]. A possible mechanism for regulating fenestrations, derived from Arias et al. [19], is shown in Figure 27.4. The key role of calcium in regulating fenestrations through effects on the cytoskeleton was reported by Gatmaitan et al. [20]. Several agents were identified that reduced the diameter of fenestrations by about 20% in rat LSECs. All were associated with an increase of intracellular calcium by two- to threefold. Agents that reduced fenestration diameter included serotonin, metoclopramide, propranolol, indomethacin, and calcium ionophore, while

Myosin light chain phosphatase

Actin

Reduced barrier to initiation of membrane fusion

Figure 27.4 Some of the pathways involved in the regulation of fenestrations [19]. Actin depolymerization and reorganization may act via a process of membrane fusion on both the number and diameter of fenestrations

27: FENESTRATIONS IN THE LIVER SINUSOIDAL ENDOTHELIAL CELL

agents with no activity included verapamil, diltiazem, nifedipine, ketanserin, imipramine, mianserin, pertussis toxin, and dexamethasone. Calcium channel blockers (diltiazem, verapamil, and nifedipine) and the calcium chelator EGTA reversed the effect of serotonin. In addition, the increase of calcium induced by serotonin was linked with phosphorylation of myosin light chain and reduced levels of cAMP [20]. In another study, serotonin reduced the diameter of fenestrations by 20%, associated with an increase in the thickness of the fenestrae-associated actin ring of 6 nm, confirming the interaction between serotonin, actin, and fenestrations [49]. Members of the Rho-like GTPase family also regulate the actin cytoskeleton in endothelial cells and are critical for membrane fusion [123]. Inhibition of the Rho pathway by C3-transferase caused reduction of myosin light chain phosphorylation, loss and retraction of actin filaments, and increased porosity and the formation of large gaps. Activating Rho with lysophosphatidic acid increased myosin light chain phosphorylation and actin filaments and led to defenestration [61]. Other factors that influence fenestrations, presumably via actions on actin, are endothelin 1 and nitric oxide. Endothelin 1 increased intracellular calcium and decreased fenestration diameter, whereas prostaglandin decreased intracellular calcium and increased fenestration diameter [64]. In another study, endothelin 1 decreased the diameter of fenestrations from 123 to 46 nm, an effect abolished by blockade of the ETB -R, and only partially abolished by ETA -R antagonism [112]. Antagonism of the ETA -R caused a marked increase in fenestration diameter, associated with gap formation [93]. Nitric oxide is involved in the maintenance of fenestrations. Caveolin-1 and endothelial nitric oxide co-locate in the cell membranes lining fenestrations and caveolin-1 is attached to actin [124]. Activation of calmodulin by increased levels of intracellular calcium releases endothelial nitric oxide synthase from caveolin-1, thereby increasing production of nitric oxide [62]. Importantly, it has been shown that the effects of VEGF on the phenotype of LSECs requires autocrine production of nitric oxide [99].

Vascular endothelial growth factor In endothelial cells, VEGF activates cell division, angiogenesis, and vascular permeability. VEGF increases intracellular calcium, phosphorylates myosin light chain, and causes retraction of the cytoskeleton [125, 126]. VEGF generates fenestrations and caveolae in a number of different endothelial cells, including tumor [127], renal [128], and adrenal [129] endothelial cells. In the liver, hepatocytes produce VEGF, which acts on liver endothelial cells via the receptors: VEGFR-1 (Flt-1) and VEGFR-2 (KDR/Flk-1), of which VEGFR-2 is the most important

397

[59, 130, 131]. VEGF is expressed more highly in the pericentral regions reflecting hypoxia, which is the primary stimulus for VEGF production, whereas VEGFR-2 is found along the entire sinusoidal endothelium [132]. In isolated liver endothelial cells, VEGF increases porosity about twofold, mostly through its effects on the number of fenestrations [59, 105] (Table 27.2). VEGF converts punctate caveolin-1 staining to aggregates of staining, the majority of which are located at the periphery of the LSECs. VEGF did not change total caveolin-1 protein expression. Indeed, caveolin-1 labeling might eventually appear reduced by the redistribution of caveolin-1 onto the markedly increased number of fenestrations [62]. Systemic VEGF exposure generated by VEGFexpressing CHO cells implanted into nude mice stimulated mitosis and proliferation of liver endothelial cells and led to increased complexity and branching of sinusoids [130]. Conversely, transgenic inhibition of VEGF receptors altered the hepatic endothelium of early postnatal mice, including loss of endothelial lining in many sinusoids [133], and was associated with defenestration and hyperlipidemia [80]. VEGF is considered to be the major cytokine involved in the regulation of fenestrations [128]. There is a mechanistic paradox with respect to the relationship between phosphorylation of myosin light chain and the subsequent effects on actin and fenestrations. VEGF increases intracellular calcium and phosphorylates myosin light chain in endothelial cells [125, 126], and this is associated with increased fenestrations in sinusoidal and other endothelial cells [59, 105, 128]. On the other hand, serotonin, calcium ionophore, and activation of the Rho pathway also phosphorylate myosin light chain, yet this leads to reduced fenestration diameter [21, 61].

PATHOPHYSIOLOGY OF FENESTRATIONS There are numerous reports of diseases and pathological processes that influence fenestrations, including: primary liver disease (cirrhosis [134, 135], fibrosis [136], steatosis [137], hepatitis [90, 138], hepatic vascular diseases and the sinusoidal obstruction syndrome [139], vena caval obstruction [140]), liver toxins (acetaminophen [141–144], oxidants [145, 146], bacterial toxins [35, 147, 148]), systemic disease (diabetes mellitus [55, 149–152]), and other liver processes (aging [3, 51, 153–155], partial hepatectomy [82], hypoxia [35], high pressure [34], ischemia reperfusion, and transplantation [156–159]). These changes have not usually been diagnostic [160, 161] but the overall trends are that: (i) acute toxic injury and acute medical conditions are associated with loss of endothelial integrity characterized by gap formation; and (ii) subacute and chronic conditions have been associated with defenestration and reduced porosity. Three important conditions are described below.

398

THE LIVER: PATHOPHYSIOLOGY OF FENESTRATIONS

Table 27.3 The effects of ethanol on fenestrations Porosity (area %) Experimental procedure Rat in vivo 33% Rat in vivo 40% Rat LSECs 20 g l−1 Rat in vivo 5% (zone 1) Rat in vivo 5% (zone 3) Rat in vivo 2 g kg−1 (zone 1) Rat LSECs 400 mg dl−1 Rat in vivo 38 mM (zone 1) Rat in vivo 38 mM (zone 3) Baboon in vivo 50%

Control

Treatment

— 12.2 — 4.3 5.6 — — — — 1.8

— 11.7 — 4.1 5.3 — — — — 1.5

Diameter (nm) Control 116 120 208 105 93 108 ± 172 ± 133 ± 109 ± 82

Aging and pseudocapillarization Old age is associated with thickening and defenestration of the LSEC, sporadic deposition of collagen and basal lamina in the extracellular space of Disse, and increased numbers of fat-engorged, non-activated stellate cells [41–43, 51, 154, 155, 162, 163]. There also are several immunohistochemical age-related changes, including endothelial upregulation of von Willebrand factor [42, 51, 154], VEGFR-2 [132], and ICAM-1 [155], and reduced expression of caveolin-1 [162]. These changes occur in the absence of light microscopic evidence of liver disease and have been termed age-related pseudocapillarization. Given that old age is the major risk factor for most diseases including liver disease, studies of the effects of disease on the liver should take this age-related change into account. Pseudocapillarization is reversed by caloric restriction, associated with reduced caveolin-1 expression [162], and not associated with stellate cell activation [154]. These findings indicate that pseudocapillarization is a primary aging change, rather than undiagnosed or sub-clinical liver disease. In aged rats the loss of fenestrations leads to impaired transfer of lipoproteins with average diameter of approximately 50 nm from the sinusoid into the space of Disse [79, 164]. This provides a novel mechanism and therapeutic target for impaired chylomicron remnant clearance and post-prandial hyperlipidemia associated with old age [3, 73, 153].

4 7 7 4

Frequency (per µm2 )

Treatment

Control

Treatment

Citation

140 162 251 112 98 122 ± 5 169 ± 8 81 ± 4 69 ± 6 116

— 15.1 — 9.3 13.7 — — 9.3 ± 11.8 11.3 ± 1.7 3.3

— 7.9 — 7.9 11.2 — — 5.5 ± 1.4 8.2 ± 1.3 1.4

[180] [181] [181] [173] [173] [182] [182] [172] [172] [56]

rats, porosity reduced from approximately 10 to 1% [169, 170]. In thioacetamide-induced cirrhosis in rats, there was a reduction in porosity (2.5–0.4% zone 1) and diameter (108–99 nm), and loss of SPs [134]. The defenestration seen in vivo was maintained when LSECs were isolated, indicating that changes are intrinsic to LSECs and not related to the in vivo microenvironment [134]. Defenestration and loss of SPs were seen in carbon tetrachloride-induced cirrhosis in rats [135] and porcine serum-induced rat liver fibrosis [171]. Because of the association between alcohol consumption and cirrhosis, there have been several studies of the acute and subacute effects on fenestrations (Table 27.3). Results are inconsistent, but overall ethanol causes dilatation of fenestrations of about 20%, associated with a reduction of about one-third in the number of fenestrations. Ethanol has an effect on the sinusoidal endothelial cytoskeleton [117], although the in vivo effects may also be secondary to associated steatosis, fibrosis, hepatitis [168], or altered blood flow [172]. It has been postulated that defenestration of endothelial cells prevents the uptake of vitamin A, leading to activation of stellate cells [169, 173]. In addition, it has been suggested that hyperlipidemia associated with cirrhosis is secondary to defenestration [167, 170]. Impaired transfer of numerous substrates (albumin [174], lidocaine [175], propranolol [176–178], prazosin, labetolol, diltiazem [178], and IgM [179]) across the capillarized sinusoidal endothelium has been documented in cirrhotic livers [3].

Cirrhosis and capillarization

Sinusoidal obstruction syndrome

The term “capillarization” was first used in 1963 by Schnaffer and Popper to describe the ultrastructural changes seen in the sinusoidal endothelium in cirrhosis, including a thickened endothelium with underlying basement membrane and loss of fenestrations [165]. These findings in human cirrhosis and alcoholic liver disease have been frequently observed [138, 166–168]. In animal models of cirrhosis, similar changes have been seen. In dimethylnitrosamine-induced cirrhosis in

The sinusoidal obstruction syndrome is the prototypic disease of LSECs and probably the only recognized primary disease of the LSEC. The two main causes of this syndrome are dietary pyrrolizidine alkaloids and chemo-irradiation, especially associated with bone marrow transplantation [183, 184]. The major experimental model is induced by monocrotaline [185], a pyrrolizidine alkaloid that is activated by cytochrome P450 and acts as an actin disruptor [139, 183]. After administration

27: FENESTRATIONS IN THE LIVER SINUSOIDAL ENDOTHELIAL CELL

of monocrotaline, there is gap formation, endothelial swelling, and defenestration. This is followed by extensive sinusoidal endothelial cell injury, associated with dissection into the space of Disse and eventually embolization of the sinusoidal endothelial fragments [186]. Centrilobular necrosis ensues as a result of impaired perfusion and clinically is associated with jaundice, hepatomegaly, and ascites, and a high mortality. Matrix metalloproteinases (MMPs), particularly MMP-9 and MMP-2, are key mediators of the syndrome, but appear to influence the dehiscence of LSECs rather than fenestrations [183, 187]. On the other hand, the decrease in nitric oxide levels that occurs early [183, 188] could directly influence fenestrations. Of great significance is the observation that strategies that maintain the integrity of the LSEC, such as MMP inhibition and preservation of nitric oxide levels, totally abrogate any hepatocellular injury [183]. The loss of LSEC morphology appears to be an initiating step for some hepatotoxic agents.

6.

7.

8.

9.

10.

11.

FUTURE THERAPEUTIC OPPORTUNITIES It has now been well established that fenestrations can be regulated with a variety of pharmacological agents [19, 104]. Changes in fenestration morphology appear to have systemic implications, particularly for lipoprotein metabolism [53], clearance of medications [3], and immunity [8], as well as hepatoprotective effects [183]. Thus the modulation of fenestrations, for example to treat dyslipidemia in aging [153, 189] and diabetes mellitus [55], has become a novel target for future pharmaceutical development.

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147. Cheluvappa, R., Jamieson, H.A., Hilmer, S.N., Muller, M. and Le Couteur, D.G. (2007) The effect of Pseudomonas aeruginosa virulence factor, pyocyanin, on the liver sinusoidal endothelial cell. J Gastroenterol Hepatol , 22, 1350–51. 148. Dobbs, B.R., Rogers, G.W., Xing, H.-Y. and Fraser, R. (1994) Endotoxin-induced defenestration of the hepatic sinusoidal endothelium: a factor in the pathogenesis of cirrhosis? Liver, 14, 230–33. 149. Jamieson, H., Dobbs, B.R., Day, W.A., Rogers, G.W.T. and Fraser, R. (2001) The liver sieve in diabetes: are the ultrastructural changes similar to those seen in alcoholism? in Cells of the Hepatic Sinusoid (eds E. Wisse, D.L. Knook, R. De Zanger and M.J.P. Arthur), The Kupffer Cell Foundation, Leiden, pp. 123–24. 150. Harrison, S.A., Brunt, E.M., Goodman, Z.D. and Di Bisceglie, A.M. (2006) Diabetic hepatosclerosis: diabetic microangiopathy of the liver. Arch Pathol Lab Med , 130, 27–32. 151. Latry, P., Bioulac-Sage, P., Echinard, E., Gin, H., Boussarie, L., Grimaud, J.A. and Balabaud, C. (1987) Perisinusoidal fibrosis and basement membrane-like material in the livers of diabetic patients. Hum Pathol , 18, 775–80. 152. Bernuau, D., Guillot, R., Durand, A.M., Raoux, N., Gabreau, T., Passa, P. and Feldmann, G. (1982) Ultrastructural aspects of the liver perisinusoidal space in diabetic patients with and without microangiopathy. Diabetes, 31, 1061–67. 153. Le Couteur, D.G., Fraser, R., Cogger, V.C. and McLean, A.J. (2002) Hepatic pseudocapillarisation and atherosclerosis in ageing. Lancet , 359, 1612–15. 154. Le Couteur, D.G., Warren, A., Cogger, V.C., Smedsrod, B., Sorensen, K., de Cabo, R., Fraser, R. et al. (2008) Old age and the hepatic sinusoid. Anat Rec, 291 (6), 672–83. 155. Ito, Y., Sorensen, K.K., Bethea, N.W., Svistounov, D., McCuskey, M.K., Smedsrod, B.H. and McCuskey, R.S. (2007) Age-related changes in the hepatic microcirculation of mice. Exp Gerontol , 48, 789–97. 156. Enomoto, K., Nishikawa, Y., Omori, Y., Tokairin, T., Yoshida, M., Ohi, N., Nishimura, T. et al. (2004) Cell biology and pathology of liver sinusoidal endothelial cells. Med Electron Microsc, 37, 208–15. 157. Huet, P.M., Nagaoka, M.R., Desbiens, G., Tarrab, E., Brault, A., Bralet, M.P. and Bilodeau, M. (2004) Sinusoidal endothelial cell and hepatocyte death following cold ischemia-warm reperfusion of the rat liver. Hepatology, 39, 1110–19. 158. Morgan, G.R., Sanabria, J.R., Clavien, P.A., Phillips, M.J., Edwards, C., Harvey, P.R. and Strasberg, S.M. (1991) Correlation of donor nutritional status with sinusoidal lining cell viability and liver function in the rat. Transplantation, 51, 1176–83. 159. Takeda, Y., Arii, S., Kaido, T., Niwano, M., Moriga, T., Mori, A., Hanaki, K. et al. (1999) Morphologic alteration of hepatocytes and sinusoidal endothelial cells in rat fatty liver during cold preservation and the protective effect of hepatocyte growth factor. Transplantation, 67, 820–28.

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173. Tanikawa, K., Noguchi, K. and Sata, M. (1991) Ultrastructural features of Kupffer cells and sinusoidal endothelial cells in chronic ethanol-fed rats, in Cells of the Hepatic Sinusoid , Vol. 3 (eds E. Wisse, D.L. Knook and R.S. McCuskey), The Kupffer Cell Foundation, Leiden, pp. 445–48. 174. Huet, P.M., Goresky, C.A., Villeneuve, J.P., Marleau, D. and Lough, J.O. (1982) Assessment of liver microcirculation in human cirrhosis. J Clin Invest , 70, 1234–44. 175. Varin, F. and Huet, P.M. (1985) Hepatic microcirculation in the perfused cirrhotic rat liver. J Clin Invest , 76, 1904–12. 176. Fenvyes, D., Gariepy, L. and Villeneuve, J.P. (1993) Clearance by the liver in cirrhosis: I. Relationship between propranolol metabolism in vitro and its extraction by the perfused liver in the rat. Hepatology, 17, 301–6. 177. Gariepy, L., Fenyves, D., Kassissia, I. and Villeneuve, J.P. (1993) Clearance by the liver in cirrhosis. II. Characterization of propranolol uptake with the multiple-indicator dilution technique. Hepatology, 18, 823–31. 178. Hung, D.Y., Chang, P., Cheung, K., McWhinney, B., Masci, P.P., Weiss, M. and Roberts, M.S. (2002) Cationic drug pharmacokinetics in diseased livers determined by fibrosis index, hepatic protein content, microsomal activity, and nature of drug. J Pharmacol Exp Ther, 301, 1079–87. 179. Henriksen, J.H. (1983) Permselectivity of the liver blood-lymph (ascitic fluid) barrier to macromolecules in decompensated cirrhosis: relation to calculated pore size. Clin Physiol , 3, 163–71. 180. Fraser, R., Bowler, L.M. and Day, W.A. (1980) Damage of rat liver sinusoidal endothelium by ethanol. Pathology, 12, 371–76. 181. Charels, K., de Zanger, R.B., Van Bossuyt, H., Van Der Smissen, P. and Wisse, E. (1986) Influence of acute alcohol administration on endothelial fenestrae of rat livers: an in vivo and in vitro scanning electron microscopic study, in Cells of the Hepatic Sinusoid, Vol. 1 (eds A. Kirn, D.L. Knook and E. Wisse), The Kupffer Cell Foundation, Rijswijk, pp. 497–502. 182. Mori, T., Okanoue, T., Sawa, Y., Itoh, Y., Kanaoka, H., Hori, N., Enjyo, R. et al. (1991) Effect of ethanol on the sinusoidal endothelial fenestration of rat liver—in vivo and in vitro study, in Cells of the Hepatic Sinusoid, Vol. 3 (eds E. Wisse, D.L. Knook and R.S. McCuskey), The Kupffer Cell Foundation, Leiden, pp. 469–71. 183. Deleve, L.D. (2007) Hepatic microvasculature in liver injury. Semin Liver Dis, 27, 390–400. 184. DeLeve, L.D., Shulman, H.M. and McDonald, G.B. (2002) Toxic injury to hepatic sinusoids: sinusoidal obstruction syndrome (veno-occlusive disease). Semin Liver Dis, 22, 27–42. 185. DeLeve, L.D., McCuskey, R.S., Wang, X., Hu, L., McCuskey, M.K., Epstein, R.B. and Kanel, G.C. (1999) Characterization of a reproducible rat model of hepatic veno-occlusive disease. Hepatology, 29, 1779–91.

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186. DeLeve, L.D., Ito, Y., Bethea, N.W., McCuskey, M.K., Wang, X. and McCuskey, R.S. (2003) Embolization by sinusoidal lining cells obstructs the microcirculation in rat sinusoidal obstruction syndrome. Am J Physiol , 284, 1045–52. 187. Deleve, L.D., Wang, X., Tsai, J., Kanel, G., Strasberg, S. and Tokes, Z.A. (2003) Sinusoidal obstruction syndrome (veno-occlusive disease) in the rat is prevented by matrix metalloproteinase inhibition. Gastroenterology, 125, 882–90.

188. DeLeve, L.D., Wang, X., Kanel, G.C., Ito, Y., Bethea, N.W., McCuskey, M.K., Tokes, Z.A. et al. (2003) Decreased hepatic nitric oxide production contributes to the development of rat sinusoidal obstruction syndrome. Hepatology, 38, 900–8. 189. Le Couteur, D.G., Cogger, V.C., McCuskey, R.S., De Cabo, R., Smedsrod, B., Sorensen, K.K. and Warren, A. et al. (2007) Age-related changes in the liver sinusoidal endothelium: a mechanism for dyslipidemia. Ann N Y Acad Sci , 1114, 79–87.

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Hepatic Stellate Cells Marcos Rojkind and Karina Reyes-Gordillo Department of Biochemistry and Molecular Biology, The George Washington University Medical Center, Washington, DC, USA

INTRODUCTION The liver, like any other tissue/organ, is a small ecosystem in which all its cellular elements and extracellular matrix components play key roles in generating its structure and function [1]. The interaction of the different cell types produces the correct geometry for the formation of cords, and from these cell–cell interactions the normal quality and quantity of connective tissue components will be deposited. Each of the different liver cell types produces various molecules (metabolites, cytokines, and growth factors) that control the proliferation and regulate the function of other cells within the liver. Accordingly, any alteration in the components present in a unit, whether it is the extracellular matrix or a cellular element, could generate two distinct signal cascades. The first one induces regeneration of the lost cells and modulate the production of normal extracellular matrix. This results in complete restoration of normal liver architecture and function. The second signal cascade, which could be independent of or occur simultaneously with the regeneration signals, will result in transdifferentiation of hepatic stellate cells (HSCs) into myofibroblasts (Mybs). This will result in the production and deposition of excess extracellular matrix components present in scar tissue, distortion of the liver architecture, and alterations in liver function. The predominance of either one of the signals will depend on several factors, namely the nature, intensity, and chronicity of the injury. In general, acute and extensive lesions that do not produce liver failure will induce liver regeneration. However, less intense and chronic injury will result in fibrosis and cirrhosis. The multiple cytokines and growth

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

factors involved in liver regeneration and/or scarring are produced by the various cell types within the liver as well as by the inflammatory cells that arrive from the circulation after injury [2–5]. They can also be derived from the extracellular matrix, which acts as a reservoir for multiple cytokines and growth factors, and these could be released when matrix remodeling and/or degradation take place [6–11]. Therefore, they may play a key role in the repair process. The release of these cytokines and growth factors could represent the earliest form of response to injury and could play a key role in the activation of other liver cell types. In vivo studies to produce liver fibrosis are physiologically relevant. However, the alterations induced are quite complex and therefore it is difficult to determine the key roles that each liver cell type and/or inflammatory cell plays in the process. On the other hand, studies performed with isolated cells lack the normal cell–cell and cell–matrix interactions and therefore may not always be physiologically relevant. Nonetheless, studies in vitro with isolated HSCs, the main collagen-producing cells of the liver, have significantly advanced our knowledge of the pathophysiology of liver fibrosis [5, 12–20]. Moreover, these studies have provided us with multiple novel tools to treat and revert liver fibrosis [19–44]. While the old antifibrogenic drugs were directed to enhance the degradation of collagen and/or prevent its secretion [45–49], the new tools are directed toward key steps in the activation of HSCs and their capacity to produce scar collagen [19–44]. Due to the relevance of liver fibrosis as one of the main causes of death worldwide and the significant advances made in the understanding of HSCs physiology, the

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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number of publications on HSCs has increased dramatically and multiple reviews have been published covering the various aspects of HSC physiology and pathobiology [4, 5, 12–20, 50, 52]. Therefore, in this chapter we will attempt to summarize important aspects of the physiology of HSC and the molecular mechanisms involved in their transformation into Mybs.

WHAT ARE HSCs? HSCs are liver sinusoidal cells localized to the space of Disse. They have a relatively small body, loaded with vitamin A-containing fat droplets and very long projections. They account for 5–8% of total liver cells. They were first described in 1876 by Carl von Kupffer (Sternzellen or star-shaped cells). Subsequently, they were rediscovered and renamed “fat-storing cells” by Toshio Ito in the early 1950s, and were further characterized and established as perisinusoidal cells containing vitamin A by Kenjiro Wake (for reviews see [4, 5, 13–15, 50, 52]). HSCs are in close contact with liver endothelial cells and are embedded in the loose extracellular matrix of the Space of Disse (Figure 28.1). This space contains several extracel-

lular matrix components, including basement membrane macromolecules such as collagen type IV and laminin, but it is devoid of a continuous basement membrane [53–55]. HSCs express connexin 43 [56–59], which is localized to the tips of the projections, where it forms functional gap junctions with other HSCs as well as with hepatocytes (Figure 28.2). The capacity of HSCs to store vitamin A and triglycerides is what enables them to have the characteristic low density that has been used for their isolation after density gradient centrifugation [60–62]. In general, HSCs regulate retinoid metabolism and retinoids regulate HSC phenotype [63–70]. Isolation procedures based on density gradient centrifugation can yield over 50 million cells with purities greater that 95% when older normal rats are used [50, 52, 60–62]. However, the yield and purity vary greatly when rats from different models of fibrosis are used. As described below, HSCs obtained from fibrotic livers are heterogeneous and contain HSCs with vitamin A and Mybs devoid of fat droplets. Therefore, more than a single band will be obtained on a gradient and Mybs will sediment in regions where Kupffer and endothelial cells sediment. Additional methods for isolation of the cells include culture explants, centrifugal elutriation, and cell sorting using specific cellular markers [5, 71].

BC

Hepatocytes

Space of Disse

HSC

KC

Fenestrae

KC

Sinusoid

Space of Disse HSC

Hepatocytes

Quiescent HSC

Myofibroblast

Figure 28.1 Cartoon illustrating the organization of the liver sinusoid. It corresponds to a longitudinal section of a sinusoid. It shows Kupffer cells within the lumen of the sinusoid, endothelial cells with fenestrae, and HSCs that are located beneath the endothelial cells within the space of Disse. HSCs contain the characteristic lipid droplets with triglycerides and vitamin A. The hepatocytes are polarized and show the microvilli within their sinusoidal domain and the formation of bile canaliculi. When there is liver injury, HSCs transform into Mybs and lose their fat droplets (bottom of the figure). Modified from [72]

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Figure 28.2 Confocal microscope images of co-cultures of hepatocytes and HSCs showing the localization of tubulin in cytoplasmic microtubules. Projection of 31 serial optical sections of the entire depth of HSCs (range of 12–18 µm measured at cell center) and of hepatocytes (range of 20–31 µm measured at cell center). The spatial relations of HSCs (I) and hepatocytes (H) are evident. Note the long tubulin-positive projections of HSCs (range of 8–14 µm in depth and 5–9 µm in length) extending to the hepatocyte and contacting a small region of the hepatocyte surface (arrows). Reprinted from Am J Pathol 1995, 146, 1508–20, with permission from the American Society for Investigative Pathology

WHAT IS THE FUNCTION OF HSCs? The function of quiescent HSCs is yet to be fully determined. In the normal liver we call them “quiescent” HSCs to hide our ignorance regarding their multiple functions in addition to their role in vitamin A metabolism [5, 50, 52]. They are the main storage site of vitamin A, and only when a large excess of vitamin A is administered to animals are stellate cells present in other tissues loaded with this auto-fluorescent vitamin [70]. HSCs express several proteins involved in binding and transporting retinols, and store vitamin A largely as retinyl palmitate [63–70]. It is of interest to mention that vitamin A plays a key role in HSC physiology, although the molecular mechanisms involved remain to be fully elucidated [59–66]. HSC transdifferentiation into Mybs is accompanied by a significant decrease of retinol-binding proteins and vitamin A stores [5, 50, 52]. However, whether this is a cause or a consequence of the transdifferentiation process remains to be determined. While vitamin A ameliorates liver fibrosis [73], excess vitamin A induces liver fibrosis and cirrhosis. Whether this is associated with excess vitamin A alone or is dependent on the type of retinoids ingested is yet to be established [74–78].

An additional function of HSCs is the regulation of blood flow [15, 79, 80]. In their transdifferentiated stage, they express actins and myosin present in muscle and non-muscle cells [81–83] (Table 28.1) and have a significant contractile capacity [81–83]. Multiple factors, including transforming growth factor-β (TGF-β) [84], nitric oxide (NO) [85], endothelin-1 [86–88], angiotensin II [89], and lysophosphatidic acid [90] among others, induce HSC/Myb contraction, Rho kinases being key players in their contraction [91–93]. A less understood function of HSCs is that of regulating growth and metabolic activities of other cells. This can occur either by direct cell–cell interaction or by the release of cytokines and growth factors that will act via autocrine/paracrine loops. HSCs produce several growth factors, including epidermal growth factor [115], platelet-derived growth factor (PDGF) [116, 117], hepatocyte growth factor (HGF) [118–124], fibroblast growth factors [120, 125, 126], connective tissue growth factor (CTGF) [127–133], and insulin growth factor [134–136] among others [5]. All these factors modulate the function and proliferation of many liver cell types, including HSCs and hepatocytes. Thus, they could be targets for therapeutic intervention. Inhibition and/or abolition of CTGF, HGF or PDGF expression prevent HSC transdifferentiation and ameliorates liver fibrosis [137–145]. On the other hand,

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Table 28.1 Genes of various cellular origins expressed by HSC Genes of neural origin expressed by HSC Alpha B-crystallin Brain-derived nerve growth factor (BDNF) Connexin 43 Glial fibrillary acidic protein (GFAP) N-cadherin Neurabin 1 and 2 Nestin Neural cell adhesion molecule (N-CAM) Neurotrophin-3 (NT-3) Neurotrophin-4/5 (NT-4/5) Nerve growth factor (NGF) Synaptophysin (SYP) Genes of epithelial origin expressed by HSC Albumin Cytokeratins 8 (Ck-8) Cytokeratins 18 (Ck-18) Cytokeratins 19 (Ck-19) E-cadherin Hnf-4α Genes of muscle origin expressed by HSC A-actinin A-smooth muscle actin (α-SMA) Desmin (type III intermediate filament) Sarcomeric myosin heavy chain Synemyn Vimentin Myo D Myogenin Genes of oval cell origin expressed by HSC Alpha-fetoprotein Muscle pyruvate kinase (MPK)

overexpression of PDGF A or C in mice induces liver fibrosis [146, 147]. It is interesting to note that the expression of insulin-like growth factor I by HSCs ameliorates fibrosis and enhances regeneration [136]. HSCs also produce several cytokines, such as TGF-β, a major fibrogenic cytokine [136, 148], and IL-6 [56, 149–151], a cytokine whose production is increased during the acute phase response and further enhances the fibrogenic capacity of Mybs [152, 153]. HSCs also express IL-10, a cytokine with anti-inflammatory potential [154–158]. It has been shown that IL-10 modulates neutrophil infiltration and hepatocyte proliferation after chronic injury with CCl4 [159]. Expression of IL-6 and IL-10, both of which modulate inflammation, could play a key role in the immunological functions of HSCs (see below). As expected from an ecological system, all the elements of the liver system are important and therefore regulate each other’s functions. Thus, other liver cell types regulate activation and proliferation of HSCs at the same time that HSCs modulate the functions of the other cell types. Growth factors produced by HSCs could play a role in sustaining hepatocyte differentiation and/or inducing epithelial/mesenchymal transition in order to either enhance

[94, 95] [96] [50] [97] [98] MR, unpublished data [99] [100, 101] [96, 102] [96] [96] [103, 104] [105] [106] [106] [105, 106] [107] [105] [108] [109, 110] [110, 111] [112, 113] [114] [104, 105] [108] [108] [105] [105]

regeneration or recruit cells for collagen production during healing. Several examples of these regulatory activities are described below. It has recently been demonstrated that fibroblast growth factor (FGF) enhances differentiation of cells from adipose tissue into hepatocyte-like cells [160]. Hedgehog (Hh) ligands produced by HSCs enhance the proliferation and differentiation of bone marrow-derived stem cells into hepatocyte-like cells [161]. HSCs express HGF, a growth factor that induces hepatocyte proliferation and ameliorates liver cirrhosis when administered to cirrhotic animals [138–143]. Fenestrated but not capillarized liver endothelial cells prevent the transdifferentiation of HSCs via production of vascular endothelial growth factor (VEGF) and NO [162]. HSCs also secrete angiopoietin II, which stimulates angiogenesis [163]. Mybs stimulated with PDGF release exosome-enriched microparticles that contain active Hh ligands and regulate angiogenic responses in liver sinusoidal endothelial cells [164]. These multiple autocrine and paracrine loops are necessary to maintain liver homeostasis and each cell type contributes to this process. When homeostasis is disturbed, such as it occurs after liver injury or activation of Kupffer cells, the change in the

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balance of the various loops results in liver regeneration or in transdifferentiation of HSCs and liver fibrosis [2–5]. Recently, an additional function has been assigned to HSCs, namely immunoregulation. HSCs are phagocytic and have the capacity to present antigens [164–168]. They also produce monocyte chemoattractant protein 1 (CMP-1) [169–171], which as its name indicates is a potent chemoattractant for monocytes and macrophages. Although levels of expression of CMP-1 are low, nonetheless cytokines such as tumor necrosis factor (TNF) and or interleukin 1 upregulate its expression [171]. Therefore, the activation of HSCs could result in increased expression of CMP-1 and the recruitment of inflammatory cells, which will in turn produce additional cytokines and growth factors that will help in sustaining the inflammatory process and the deposition of extracellular matrix components, including collagen [2–5]. Accordingly, transdifferentiation of HSCs and their response to inflammatory cytokines, as well as their capacity to produce hydrogen peroxide after treatment with TGF-α or acetaldehyde (ethanol’s first metabolite), could play an important role in amplifying inflammation and fibrosis [172–176]. HSCs express toll-like receptors [177] that are involved in the binding of endotoxin [171, 172]; they produce neutrophil chemoattractants [178–180] and numerous molecules that are involved in antigen presentation, or modulate their interaction and/or response to inflammatory cells (for an excellent review see [168]). Some of these immunoregulatory functions are altered as the HSCs age, when they express cytokines and receptors that enhance the function of natural killer (NK) cells and thus their elimination [181].

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express multiple genes present in epithelial cells (Table 28.1) [105–107], a strong indication that some HSCs may undergo mesenchymal/epithelial transition [105, 106]. When cultured in serum-free media they express cytokeratin 19 and E-cadherin, transcription factors found in hepatocytes; that is, hepatocyte nuclear factor (HNF) 4α, or markers present in oval cells such as muscle pyruvate kinase (MPK) and in hepatoblasts such as α-fetoprotein [105]. HSCs also express genes present in muscle cells, including sarcomeric myosin, MyoD, and myogenin among others [108–113] (Table 28.1), and these myosins play key roles in their contractile capacity. HSCs contract with endothelin 1 and this growth factor induces the expression of sarcomeric myosin [113]. Altogether, these findings suggest the possibility that HSCs are derived from a common precursor cell that gives rise to all the different cell types within the liver. Recently, it has been suggested that HSCs represent a pool of precursor cells that undergoes a transition to mesenchymal cells prior to acquiring an epithelial phenotype [163]. However, additional work is required to confirm this provocative suggestion. It has been suggested that bone marrow-derived cells (fibrocytes) may contribute to the pool of activated Mybs in the fibrotic liver [4, 5, 15, 114, 182, 183]. However, although the liver contains some bone marrow-derived Mybs, these do not account for the large number of cells involved in fibrogenesis for which are localized around fibrous septa and portal tracts. Further studies are needed to determine their quantitative contribution in normal and fibrotic livers.

WHAT IS THE ORIGIN OF HSCs?

HSC TRANSDIFFERENTIATION (ACTIVATION)

Current evidence suggests that HSCs may be derived from the endoderm [50, 52]. Around the fourth week of human fetal development, a portion of the ventral endoderm (the hepatic diverticulum) traverses the septum transversum and comes in close contact with the heart mesenchyme. Therefore, it is possible that not only do these tissues provide signals for the bud to differentiate into a liver, but the migrating cells present in the cardiac mesoderm or septum transversum could invade the bud and eventually differentiate into HSCs. HSCs are clearly visualized in the space of Disse at the end of the first trimester of human embryo development. At this stage, cells are quite rudimentary, they contain few fat droplets, and their cytoplasmic extensions are small. Full maturation and development of the adult rat HSC morphology is observed several weeks after birth [50, 52]. Overall, HSCs have unique features that are present in pluripotential cells, and under different experimental conditions they switch phenotype. HSCs express multiple genes present in cells of neural origin [94–104] (Table 28.1). However, they also have the ability to

One unique feature of HSCs is their capacity to reprogram gene expression during liver injury, when they are detached from their extracellular matrix, and/or when they are removed from their microenvironment. During injury and inflammation, they differentiate into Mybs, lose their vitamin A stores, and acquire the capacity to contract and produce type I collagen found in scar tissue [4, 5, 17, 184, 185]. Mybs develop a very contractile cytoskeleton that allows them to control sinusoidal blood flow and thus play a critical role in the development and sustenance of portal hypertension [5, 16, 186]. Once HSCs are removed from their environment and placed in culture, they differentiate into Myb-like cells by a process that resembles that observed in vivo. However, there are significant differences between the patterns of gene expression in vitro and in vivo [187]. The transformation of HSCs into Mybs is referred to as activation or transdifferentiation and involves the formation of reactive oxygen species, namely hydrogen peroxide [188, 189]. Two distinct stages have been described during HSC differentiation, namely initiation and perpetuation. The first

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stage is characterized by the expression of a variety of new receptors and significant changes in transcriptional factors that regulate the adipogenic phenotype of quiescent HSCs (see Table 28.2). The second stage corresponds to the interactions of cytokines and growth factors that are produced either by HSCs (autocrine loops) or by cells in their surroundings (paracrine loops) with the corresponding receptors on HSCs. These growth factors and cytokines subsequently induce cell migration, proliferation, and upregulation of the expression of matrix components found in scar tissue [4, 5, 17]. Transdifferentiation of HSCs is accompanied by multiple changes in the expression of genes that regulate cell migration, proliferation, collagen production, and wound contraction [4, 5, 17]. Some of these genes are illustrated in Tables 28.1 and 28.2. Of the multiple genes expressed by HSCs, those of α-smooth muscle actin (α-SMA) and PDGF-β receptor are considered hallmarks of transdifferentiation [5, 17]. Of these two markers, the latter has been used as a target for antifibrogenic therapy [144, 145]. Antioxidants, such as N-acetyl-cysteine (NAC), have antifibrogenic potential. They prevent activation of HSCs by inducing the degradation of PDGF-β receptors via cathepsins secreted by the cells [234]. Although there has been significant advancement in our understanding of the role of PDGF in HSC migration and proliferation, the role of the various PDGFs and their receptors needs to be further investigated. Transgenic mice that overexpress PDGF-C develop liver fibrosis and cancer [146]. On the other hand, HSCs express PDGF-A, -B, -C, and –D, and the latter appears to have a more significant fibrogenic effect on cultured HSCs and Mybs. In contrast with the fibrogenic effects of PDGF-C in vivo, this growth factor is not strongly fibrogenic in vitro and does not induce a significant mitogenic effect on HSCs and Mybs [117]. It has been shown that lysophosphatidic acid is involved in HSC attachment to the extracellular matrix and regulates cell migration and gel contraction. These effects are mediated by the activation of Rho kinase, as well as the phosphorylation and activation of myosin light-chain kinase [90–93, 235]. Several years ago, it was demonstrated that adenosine ameliorates liver fibrosis [236]. Although the molecular mechanisms whereby adenosine exerts its antifibrogenic effect remain to be elucidated, adenosine inhibits Ca++ signaling and chemotaxis in HSCs [237]. In addition, adenosine inhibits Rho kinase and the contraction of HSCs, and eliminates formation of stress fibers [238]. Recent work from several laboratories has shown that the adenosine A(2A) receptor and the adenosine monophosphate-activated protein kinase (AMPK) play key roles in the transdifferentiation of HSCs [239–241]. The occupancy of the A(2A) receptor upregulates collagen gene expression by complex mechanisms involving ERK 1/2 or p38 protein kinase signaling pathways [241].

Additional genes that reflect and/or affect the status of transdifferentiation of HSCs are leptin and adiponectin, two adipokines expressed in adipocytes [242, 243]. Leptin and its receptor are expressed in HSCs during transdifferentiation and leptin is elevated in cirrhotic livers [244–249]. This adipokine induces the expression of tissue inhibitor of metalloproteinase 1 (TIMP-1) and enhances collagen gene expression by cultured HSCs [247–250]. While the former effect is mediated by several kinases, including JAK, MAPK, and PI3-AKT [247], the latter effect appears to be mediated mainly by the PI3K–AKT pathway and ERK [248]. The inhibition of leptin upregulation of TIMP-1 results in apoptosis of HSCs. This could be taken as an additional indication that TIMP-1 acts as a survival factor for HSCs, as previously suggested [26, 27]. In addition, leptin induces HSC proliferation by a mechanism dependent on the upregulation of the PDGF-β receptor [249]. Interestingly, leptin downregulates the expression of matrix metalloproteinase 1 (MMP-1) [250], a further indication of the reciprocal modulation of interstitial MMPs and collagens occurring in HSCs and other cell types [251]. Adiponectin is an antiangiogenic and antiatherogenic agent [252] that inhibits cell proliferation by binding in its polymeric form to several growth factors, such as PDGF-BB, BFGF, and heparin-binding growth factor [243, 253, 254]. Adiponectin inhibits HSC proliferation by a mechanism dependent on AMPK. TGF-β is the fibrogenic cytokine involved in most types of wound-healing situation [13, 148]. It is upregulated in differentiated HSCs and plays an important role in type I collagen gene upregulation in HSCs by a mechanism dependent on the activation of members of the Smad family [255–259]. It is produced by HSCs, Kupffer cells, and other inflammatory cells, and via paracrine or autocrine loops it induces activation of HSCs and upregulates the expression of the type I collagen genes [13, 148]. In addition to stimulating the activation of Smads, TGF-β acts via a second messenger, namely H2 O2 [195]. While H2 O2 reproduces some of the TGF-β-mediated effects, catalase, a scavenger for H2 O2 , prevents them. Similarly, acetaldehyde induces the expression of type I collagen genes by TGF-β-dependent and -independent mechanisms, Moreover, H2 O2 is also a mediator of the fibrogenic effects of this ethanol metabolite [196, 205, 260]. Thus oxidative stress plays a key role in the transdifferentiation of HSCs and in their increased production of scar tissue components [188, 189]. However, the source of the reactive oxygen species needs additional studies. Although HSCs express functional nicotinamide adenine dinucleotide phosphate-oxidase (NADPH oxidase) [261–266], mitochondria are an additional source of H2 O2 , specially associated with the alterations induced by acetaldehyde/alcohol [196, 205, 260]. In the cirrhotic liver there are alterations in bile acid metabolism and therefore their putative fibrogenic actions have been investigated. These are also mediated by

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induction of the transdifferentiation of portal fibroblasts [258] and HSCs. In the former, the effects are mediated by TGF-β [267–269] via the production of thrombospondin 1, a matrix component that plays an important role in the activation of the latent cytokine [269].

THE ADIPOGENIC PHENOTYPE Quiescent HSCs express multiple transcription factors whose activity is upregulated during transdifferentiation (see Table 28.2) [18, 82, 151, 184, 190–232, 270–272]. However, of the transcription factors, those involved in preserving the adipocyte phenotype are the ones necessary to sustain quiescence [231, 232, 273]. They include, among others, the peroxisome proliferator-activated receptor gamma (PPARγ) and members of the CCAATenhanced binding proteins (C/EBPs) family (see Table 28.2). These quiescent cells express low levels of collagens type I and III and high levels of collagen type IV [274]. However, the adipogenic transcription factors are downregulated during differentiation of HSCs into Mybs and there is a switch in collagen gene expression [274, 275]. Moreover, treatment of Mybs with an adipocyte differentiation cocktail containing 0.5 mM isobutylmethylxanthine, 1 µM dexamethasone, and 1 µM insulin restores the adipogenic phenotype and reverts

Mybs to a quiescent phenotype [195]. The expression of PPARγ is a key element in the adipogenic phenotype. PPARγ inhibits α1(I) collagen promoter activity [276]. Moreover, the inhibition of PPARγ by hydrogen peroxide generated in acetaldehyde-treated HSCs [277] could account for one possible mechanism of the fibrogenic actions of this ethanol metabolite [196, 205]. Thus, multiple studies have been performed to investigate the antifibrogenic potential of agonists of this receptor. It has been shown that SC-236, a selective COX2 inhibitor, reduces liver fibrosis by a mechanism involving PPARγ activation [278]. In a rat model of chronic cholestasis, thiazolidone, a PPARγ agonist, reduced bile duct proliferation and fibrosis [279]. PPARγ agonist inhibited liver fibrosis in thioacetamide-treated animals. This effect was additive to that of agonists of the retinoic acid receptor [280]. Similarly, the interaction of agonists of the farnesoid X receptor (FXR) with PPARγ may be responsible for the antifibrogenic effects of FXR [281]. Recent studies regarding the antifibrogenic properties of curcumin have revealed that its main effect is on PPARγ, through several distinct mechanisms. It inhibits the expression of CTGF [282], induces apoptosis [283], disrupts TGF-β signaling [284], and interrupts PDGF-dependent signaling [285, 286]. Overall, PPARγ attenuates oxidative stress, a key element in the fibrogenic cascade [287].

Table 28.2 Transcription factors associated with HSC activation Activator protein (AP-1) c-Fos c-jun Fra 1 JunB JunD CCAAT/enhancer binding protein (C/EBP) c/EBPα c/EBPβ c/EBPδ c-myb and other E-box-binding transcription factors Kruppel-like transcriptional factors Sp1 BTEB1 KLF5 KLF6 ZNF267 Myogenic transcriptions factors MyoD Myf5 Myogenin MEF-2 Nuclear factor κB (NFκB) Peroxisome proliferator activated receptor-γ (PPARγ)

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[151, 184, 190–194]

[151, 184, 193, 195–199]

[151, 184, 193, 200–204] [151, 184, 193, 205–216]

[113, 151, 154, 204]

[151, 193, 217–226] [151, 193, 227–233]

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THE WINGLESS (WNT) PATHWAY AND HSC TRANSDIFFERENTIATION Wnts represent a family of small-molecular-weight proteins containing lipid moieties that stimulate the nuclear translocation of β-catenin and modulate cell differentiation, inflammation, and scarring (Figure 28.3a and 28.3b). Numerous reviews covering different aspects of the Wnt pathway, its receptors and signaling mechanisms have recently been published [288–293]. HSCs express several components of the Wnt pathway, including various Wnts, the receptor Frizzled (FZ), the co-receptor lipoprotein like receptor protein 6(LPR6), the inhibitor Dickkopf-1, and several transcription factors involved in Wnt signaling (see Table 28.3) [294–296]. The Wnt pathway is involved in transdifferentiation of quiescent HSCs into Mybs [296, 297]. Although the molecular mechanisms involved in Wnt-dependent transdifferentiation remains to be elucidated, it has been suggested that they involve the downregulation of PPARγ [297]. Based on these findings, an additional therapeutic approach to prevent liver fibrogenesis has been proposed [297].

THE HEDGEHOG (HH) PATHWAY AND HSC TRANSDIFFERENTIATION The Hh pathway is another important pathway in cell differentiation and development [299] (Figure 28.4a and 28.4b). In addition to Wnts, HSCs express several components of the Hh pathway, including Sonic Hh (SHh), Indian Hh, patch, smoothened (SMO), the transcription factors Gli1, Gli2, and Gli 3, and the inhibitor Hip [300–306]. It has been shown that the Hh pathway plays a key role in survival and transdifferentiation of HSCs, is involved in epithelial–mesenchymal transitions, and plays an important role in tissue repair [300–306]. In addition, it is the key player in cell–cell communication via the release of exosomes containing functional members of the Hh pathway [163]. Under normal conditions, the repressor Hip prevents signaling (see Figure 28.4a). However, when ligands bind to the receptor, expression of Hip decreases significantly and the expression of SHh is upregulated, resulting in the activation of Hh target genes (Figure 28.4b) [301]. When HSCs are cultured with neutralizing antibodies to SHh they undergo apoptosis. The addition of PDGF-BB to induce Myb proliferation is accompanied by SHh expression through a mechanism dependent on the activation of AKT. The proliferative action of PDGF-BB can be blocked by the addition of neutralizing antibodies to SHh. These findings suggest that an autocrine SHh loop plays a key role in the differentiation of HSCs to Mybs and involves

the AKT survival pathway [303]. Based on these and other findings it has been suggested that an autocrine SHh loop plays an important role in the survival of HSCs [163, 303]. In experiments performed after bile duct ligation, Hh appeared to play an important role in epithelial/mesenchymal interactions [303].

THE ROLE OF THE PHYSICAL NATURE OF THE EXTRACELLULAR MATRIX ON HSC TRANSDIFFERENTIATION The physical nature of the extracellular matrix, and not only its composition, plays an important role in HSC transdifferentiation. Pressure loading increases the expression of α-SMA by a mechanism dependent on the activation or ERK and Jun kinases; however, it does not downregulate the expression of type IV collagen [307–310]. It has also been suggested that the fibrogenic cytokine TGF-β, via activation of Smad 3, regulates stiffness of the matrix, and this in turn plays a key role in transdifferentiation of HSCs (for a review see [310]). Similarly, differentiation of portal fibroblasts is also modulated by mechanical tension and TGF-β [309]. In addition to changes in the organization of the extracellular matrix, alterations in the expression of receptors to extracellular matrix components, such as galectin-3, a putative laminin receptor, occur during activation of HSCs [311]. It is important to mention that stiffness of the matrix can also prevent its degradation by MMPs, due in part to the formation of cross-links [308]. Indeed, physical hindrance associated with increased formation of cross-links delays and/or prevents collagen degradation [312–316]. This is in agreement with previous studies showing that the half-lives of liver collagens I and III are decreased by 50% in CCl4 -cirrhotic liver, and that the collagens undergoing degradation are mainly those recently synthesized and least cross-linked [316].

THE ROLE OF LYMPHOCYTES IN HSC TRANSDIFFERENTIATION HSCs are phagocytic [317–319], have the capacity to present antigens [166, 320], control lymphocyte infiltration [321], and bind lymphocytes [322]. Therefore, the role of inflammatory cells obtained from patients with hepatitis C virus (HCV) on the transdifferentiation of HSCs has been investigated. The analysis of liver samples from patients with HCV revealed that cells CD4/CD8 and NK were in close proximity to transdifferentiated, α-SMA-positive HSCs localized to the

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1

415

FZ LRP6 DKK

Dvl CK1

2

GSK3

APC

PO4

Axin

3

-cat

4

PROTEASOME (a) 1 Wnt 3a LRP6 3

2

FZ Dvl PO4 APC

DKK

GSK3

CK1

Axin

-cat 3

4 Cyclin D1 c-Myc

-cat-TCF-LEF Nucleus

(b)

Figure 28.3 (a) Illustrates the components of the Wnt pathway. There are two basic components. (i) The membrane complex with the receptor Fz and the co-receptor LRP6. This complex includes Dickkopf (DKK), the natural inhibitor of this pathway, which is bound to the co-receptor, thus preventing the interaction of LRP6 with Fz and inhibiting\downstream signaling via Dvl. (ii) The β-catenin destruction complex contains multiple components that facilitate the phosphorylation of β-catenin by CK1 and GSK3β, tagging it for proteasomal degradation. (b) shows that in the presence of Wnt ligands, DKK is dislodged, LRP6 dimerizes with FZ, and Dvl is phosphorylated. The β-catenin degradation complex is inhibited and β-catenin is translocated to the nucleus, where it forms transcriptional complexes with LEF and TCF, enhancing the transcription of key genes involved in cell proliferation and HSC transdifferentiation

periportal area [322]. Some of these inflammatory cells were not only in close proximity to the HSCs, but were actually engulfed by them. When HSCs were cultured in the presence of CD8+ cells obtained from patients with HCV, they were activated. However, NK cells, known to eliminate activated HSCs, prevented activation [318]. Interestingly, phagocytosis of lymphocytes was prevented

by blocking the intracellular adhesion molecule I-CAM, as well as integrin αV [318]. It has been further shown that members of the Rho GTPase family of kinases, which are known to be important in HSC migration [90], are also involved in phagocytosis [318]. Leptin also induces phagocytosis of apoptotic bodies by HSCs via the Rho kinase [319].

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THE LIVER: IS TRANSDIFFERENTIATION A REVERSIBLE PROCESS?

Table 28.3 Members of the Wnt pathway expressed by HSCa B-catenin Dickkopf-1 (Dkk) Frizzle receptor 1 (Fz1) Frizzle receptor 2 (Fz2) Frizzle receptor 3 (Fz3) Frizzle receptor 4 (Fz4) Frizzle receptor 6 (Fz6) Frizzle receptor 7 (Fz7) Frizzle receptor 8 (Fz8) Frizzle receptor 9 (Fz9) Lipoprotein-receptor-related protein 5 (LRP)5 Lipoprotein-receptor-related protein 6 (LRP)6 Receptor-like tyrosine kinase (Ryk) T-cell factor (TCF) T-cell factor/lymphocyte enhancer factor (TCF/LEF) Wnt 2, 2b Wnt 3 Wnt 4 Wnt 5a, 5b Wnt 6 Wnt 7a Wnt 9a Wnt 9b Wnt 10a, 10b

Hedgehog Pathway Shh

Ihh

Hip

Ptc

Pathway OFF

Smo

Gli1

Nucleus Gli1

(a)

Hedgehog Pathway Shh

Ihh

Hip

Pathway OFF

a Many of these Wnt components were detected by microarrays

performed on quiescent and activated HSCs [294–298].

THE ROLE OF HYPOXIA IN HSC TRANSDIFFERENTIATION When HSCs are cultured in the presence of low concentrations of O2 , they express hypoxia inducible factor 1 (HIF-1) [323–325]. The expression and/or degradation of this factor are associated with the hydroxylation of critical proline residues by a proline hydroxylase [230–232]. Although this enzyme is different from that involved in the hydroxylation of proline residues in collagen, nonetheless it has the same requirements, including O2 [326]. In the presence of oxygen the hydroxylation of proline residues induces the proteasomal degradation of HIF-1α. However, in anoxia, proline residues are not hydroxylated and HIF is translocated to the nucleus, where it induces the expression of multiple genes, including VEGF [323–328]. Using microarrays, it was shown that an HSC line cultured under anoxic conditions upregulated the expression of HIF-1α, VEGF, α-SMA, and activated HSCs via a TGF-β1-mediated signal transduction pathway [324].

IS TRANSDIFFERENTIATION A REVERSIBLE PROCESS? One of the important questions pertaining to the transdifferentiation of HSCs is whether this is a reversible process or a “suicidal” command ultimately leading to cell death. Unfortunately, the results obtained so far are controversial

Ptc

Smo

Gli1

Nucleus Gli1

(b)

Figure 28.4 Cartoons illustrating the basic components of the hedgehog pathway. As shown in (a), the inhibitor Hip interacts with Hh ligands and prevents hedgehog signaling via Smo. Accordingly, there is no formation of transcriptional complexes with Gli1. However, as illustrated in (b), during liver injury Hip is downregulated and the Hh ligands can now bind to the receptor Pct, allowing Smo to signal downstream, enhancing nuclear translocation of Gli and activating the formation of transcriptional complexes that regulate transcription of genes involved in development as well as epithelial/mesenchymal transitions. Kind gift of Drs. Choi, S.S., Sicklick, J.K., and Diehl, A.M., Division of Gastroenterology, Duke University Medical Center, Durham, NC

and additional studies are required to determine whether the phenotype is indeed completely reversible. The evidence for a “suicidal” command is derived from in vivo studies showing that Mybs present in the various animal models of liver cirrhosis disappear by apoptosis once the fibrogenic stimulus has been discontinued [329–335]. In addition, there is new evidence indicating that Mybs show signs of senescence and that they exit the cell cycle, switch

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from a fibrogenic to an inflammatory phenotype, and are eliminated by NK cells [38]. These cells decrease the production of extracellular matrix components and upregulate the production of metalloproteinases involved in their degradation. Indeed, in animal models of liver fibrosis induced in mice that lack genes casually associated with aging, the number of α-SMA-positive cells is increased, fibrosis is exaggerated, and when the HSCs of these animals are placed in culture, they do not show markers of senescence and continue to actively proliferate [38]. Altogether, these findings suggest that transdifferentiation is a “suicidal” command that destines HSCs to proliferate, migrate, differentiate, and die, either by apoptosis or by destruction by NK cells. Indeed, a form of antifibrogenic therapy applied to animal models of fibrosis is the use of drugs that induce apoptosis of Mybs and enhance the resolution of liver fibrosis [331, 335]. Also, it would be interesting to investigate whether targeting drugs that induce senescence to Mybs would increase their destruction by NK cells and enhance the resolution of fibrosis. In favor of the reversibility of the Myb phenotype, there are studies in vitro showing that, under particular circumstances, HSCs can revert phenotype. HSCs cultured on laminin-containing gels do not transdifferentiate [336]. These observations are of interest as they suggest that while HSCs are quiescent in the space of Disse, the loose connective tissue, mainly laminin and other basement membrane components, may play a role in sustaining their quiescence [53, 54, 336]. On the other hand, as soon as there are changes in matrix composition after injury, these may stimulate the transdifferentiation of HSCs [55, 337]. An additional procedure to revert their phenotype in culture is to add to the cells an adipogenic mixture that favors the differentiation of 3T3L1 cells into adipocytes [231].

HETEROGENEITY OF HSCs HSCs are quite heterogeneous regarding the storage of vitamin A, the expression of cytokeratins, intermediate filaments, and gene expression patterns [50]. In addition to the portal Mybs that have different properties from HSCs [338, 339], HSCs localized to different areas of the hepatic lobule express one or more markers present in Mybs. However, not all the cells express the same markers at a given time. While some cells express α-SMA and desmin, others express only α-SMA. Similarly, some cells may express α-SMA and GAFP, while others express either one of the two markers. Likewise, the contents of vitamin A-containing fat droplets vary in HSCs from different zones of the liver [50]. Work from our laboratory reveled that HSC clones derived from a cell line generated from a single cirrhotic liver were heterogeneous regarding proliferation, expression of cytokines and growth factors, production of collagen and TGF-β, and their response to cytokines and growth factors [56]. The heterogeneity of HSCs may

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recognize two distinct origins. The first could be a real heterogeneity of the cells, which will depend on the zone of the liver they are derived from. Based on the known heterogeneity of hepatocytes [340–345], it is conceivable that the microenvironment for the HSCs differs in the different zones of the hepatic acinus and that this leads HSCs to express distinct phenotypes. Alternatively, and based on our current knowledge of epithelial–mesenchymal transitions, it is possible that the heterogeneity of HSCs may be due to differences in the state of differentiation of HSCs undergoing epithelial–mesenchymal transition [105, 106]. Finally, we cannot rule out completely the possibility that the heterogeneity of HSCs is due to the contribution of different cells populating the liver, including stem cells, fibrocytes, and epithelial–mesenchymal transitions [17].

FIBROGENESIS AND FIBROLYSIS Liver fibrosis results from the imbalance of extracellular matrix synthesis and degradation. The fibrogenic process includes the synthesis and deposition of multiple extracellular matrix components, including collagens, fibronectin, and proteoglycans [4, 5, 17, 346–354]. Although collagens I and III are the main components of liver scar tissue [354], there is excess deposition of type I, the collagen type that can form thick bundles and get highly cross-linked with intra- and intercellular cross-links. This post-translational modification, which is carried out by lysyl oxidase, makes collagens more resistant to degradation, and therefore this collagen is longer lasting and more difficult to degrade [355–358]. Nonetheless, liver fibrosis is a reversible process both in humans and in different animal models [49, 359–362]. However, similar to other chronic diseases, there is a point where the process is not reversible anymore, and this may be associated with the change in ratios of type I/III collagen, the distortion of the liver architecture, and the formation of shunts that prevent or impede the access of MMPs to the scar tissue. Investigations pertaining to the half-life of liver collagen in rats treated with CCl4 to produce cirrhosis that were labeled in vivo with radioactive proline revealed that collagen is active metabolically. Indeed, in cirrhotic animals the half-lives of collagens type I and III are shortened by 50% as compared to controls. While in control animals the half-life of type I collagen was 30 days, in the cirrhotic animals, after discontinuation of CCl4 , it was 15 days. However, only newly synthesized, labeled collagen, or that in the external layer of the collagen bundles, is degraded [316]. These findings suggest a possible steric hindrance due to formation of inter- and intramolecular cross-links. Thus, it seems that MMPs degrade the most superficial fibrils of thick collagen bundles, which are less cross-linked than those closer to the center of the fiber. The process of degradation of extracellular matrix components recognizes two antagonistic processes, namely the

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activity of the various MMPS and the expression of their various inhibitors (TIMPs) [363–371]. It has been shown that after liver injury the expression of TIMPs increases in parallel with the increase in collagen gene expression. Moreover, there is a reciprocal modulation of collagen and MMPs (MMP-13) [251] and therefore, while the expression of MMPs decreases, that of TIMP and collagen increases. However, this explanation is too simple to account for the numerous events in which both enzymes and inhibitors are involved. For example, MMP-13 appears to play a key role in activating MMP-9 during HSC transdifferentiation [372] and an additional MMP, namely MMP-14, which is a collagenase present in the plasma membrane, is also involved. This is an interesting observation that may be related to increased migration and proliferation of HSCs during transdifferentiation. It is interesting to mention that a similar mechanism has recently been described for the migration of keratinocytes [373]. The functions of TIMPs are being evaluated beyond their role as inhibitors of MMPS. There is strong evidence to suggest that TIMP-1 is a survival factor for HSCs [26, 27] and that collagen reabsorption from cirrhotic livers and elimination of HSCs is associated with a decrease in TIMP-1 expression. Thus, additional work would be required to clarify the role of TIMPs and MMPs in fibrogenesis and fibrolysis of the fibrotic livers.

CONCLUSION A significant number of discoveries over the past 30 years have paved the way for a change to the belief that cirrhosis is an irreversible process and, as such, is not susceptible to treatment. The initial demonstration that cirrhosis was a reversible process was observed in patients with hemochromatosis. Experimental evidence was obtained with drugs that modified the synthesis, deposition, or secretion of collagen. However, with the isolation of HSCs and the investigations pertaining to the molecular mechanisms involved in their trans-activation, more specific antifibrogenic approaches have been and will continue to be developed. These include the use and development of drugs that can specifically inhibit key events involved in the trans-activation of HSCs and/or induce their apoptosis. Conceivably, these approaches will prevent the migration and proliferation of HSCs, enhance their elimination, and therefore block the production of type I collagen produced when HSCs have been transformed into Mybs. Another important aspect of therapy is the delivery of the drugs to the correct sites. This has been accomplished by the delivery of drugs using vitamin A-containing liposomes or through the use of viral vectors that have liver specificity. The new studies on the plasticity of HSCs, the presence of epithelial mesenchymal transitions, and hepatic stem cells open a new avenue for the treatment of liver fibrosis and the enhancement of liver-cell regeneration. Thus, the future is bright as physicians have accepted that the

disease is reversible and therefore susceptible to treatment. However, additional strategies need to be developed to upregulate the production and/or enhance the activity of MMPs in order to accelerate the removal of collagen and thus facilitate the regeneration of the liver parenchyma. Although the future is challenging, it is nonetheless quite bright.

ACKNOWLEDGMENT This work was supported with NIH grants RO1-AA10541 and RO1-AA-09231 (M. Rojkind).

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29

Hepatic Fibrosis Ram´on Bataller1 and David A. Brenner2 1 Liver

Unit, Institut de Malalties Digestives i Metab`oliques, Hospital Cl´ınic, IDIBAPS, Barcelona, Spain 2 UCSD School of Medicine, University of California, San Diego, CA, USA

INTRODUCTION Liver fibrosis is the excessive accumulation of extracellular matrix (ECM) proteins, including collagen, that occurs in most types of chronic liver disease [1]. The main causes of liver fibrosis in developed countries include chronic hepatitis C virus (HCV) infection, alcohol abuse, and non-alcoholic steatohepatitis (NASH). The accumulation of ECM proteins distorts the hepatic architecture by forming a fibrous scar and the subsequent development of nodules of regenerating hepatocytes defines cirrhosis [2]. Cirrhosis produces hepatocellular dysfunction and increased intrahepatic resistance to blood flow, resulting in hepatic insufficiency and portal hypertension, respectively. Hepatic fibrosis is considered a wound-healing response to chronic liver injury [3]. Our understanding of liver fibrosis was limited until the 1980s, when hepatic stellate cells (HSCs) were identified as the main collagen-producing cells in the liver [4] (Figure 29.1). The molecular mechanisms leading to HSC activation and increased collagen synthesis in liver fibrosis have been elucidated using cultured HSCs and experimental models of chronic liver injury in rodents [5–7]. Besides HSCs, portal myofibroblasts (Mybs) and cells of bone marrow origin have recently been shown to have fibrogenic potential [8]. The demonstration that even advanced liver fibrosis is reversible has greatly stimulated researchers to identify antifibrotic therapies [9]. However, the most effective therapy to treat hepatic fibrosis to date is still to remove the causative agent [10, 11]. A number of drugs are able to reduce the accumulation of scar tissue in experimental models of chronic liver injury, although evidence-based therapies are not yet available [12]. The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

This chapter reviews our current understanding of the pathogenesis of hepatic fibrosis and highlights those elements of the process for which intervention has been attempted. The etiology and cellular and molecular mechanisms of liver fibrosis will be detailed. The basis for pathophysiologically-based therapies will be also detailed.

CAUSES OF HEPATIC FIBROSIS Many types of chronic liver injury can cause fibrosis (Table 29.1). The main causes of fibrosis in developed countries are HCV infection and alcoholic liver disease (ALD), which account for more than half of cases [1]. Other major causes are hepatitis B virus (HBV) infection, autoimmune hepatitis (AIH), chronic cholestasis (primary biliary cirrhosis (PBC) and primary sclerosing cholangitis (PSC)), and genetic metabolic diseases (hemochromatosis and Wilson disease). Due to the current epidemics of obesity, NASH is increasingly recognized as a major cause of fibrosis, yet its actual prevalence is unknown. Chronic hepatitis viral infections are a common cause of liver fibrosis. More than 170 million people worldwide are chronically infected by HCV. The natural history of patients with chronic HCV is characterized by a slow progression of liver fibrosis [13]. Following infection, cirrhosis may develop after an average of 20–30 years. In some patients, the rate of fibrosis progression is much faster, and cirrhosis develops after 10–15 years, whereas in others the rate of progression is negligible. Several factors are associated with fibrosis progression rate: duration of infection, older age, male sex, alcohol or cannabis consumption, and

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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THE LIVER: CAUSES OF HEPATIC FIBROSIS

αSMA positive

procollagen α1 positive

(a)

(b)

Figure 29.1 Identification of activated hepatic stellate cells in fibrotic livers. (a) Demonstration of smooth α-muscle actin immunoreactivity in a liver biopsy of a patient with HCV-induced liver fibrosis. Magnification 400×. (b) Identification of activated HSCs expressing green fluorescent protein (GFP)-procollagen α1(I) in transgenic mice with bile duct ligation. Magnification 400× Table 29.1 Main causes of liver fibrosis Chronic viral diseases Hepatitis B Hepatitis C Hepatitis D Alcohol abuse Autoimmune diseases Autoimmune hepatitis Primary biliary cirrhosis Primary sclerosing cholangitis Graft versus host disease Hepatotoxic agents Drugs: methotrexate, α-methyldopa, amiodarone, isoniazid, others Vitamin A intoxication Metabolic diseases Non-alcoholic steatohepatitis Wilson disease Hemochromatosis α1-antitrypsin deficiency Glycogen storage disease Fructosemia Galactosemia Lipid abnormalities (e.g. Gaucher disease) Tyrosinemia Vascular diseases Chronic right-sided heart failure Budd–Chiari syndrome Veno-occlusive disease Inferior vena cava thrombosis Miscellaneous Secondary biliary cirrhosis Cryptogenetic Congenital hepatic fibrosis

HIV co-infection. Metabolic disorders such as obesity and diabetes are emerging as independent co-factors of fibrogenesis [14]. The host genome has a major effect on the rate of fibrosis with chronic HCV infection. HBV infection is the single most common cause of fibrosis globally, although the prevalence rate is influenced by geographic

region [15]. The natural history of liver fibrosis due to chronic HBV infection can vary dramatically between individuals [16]. Some individuals with chronic HBV will have clinically insignificant or minimal liver disease and never develop complications. Others will have clinically apparent chronic hepatitis that can progress to fibrosis and cirrhosis. Patients with chronic HBV and “replicative” infection defined by the presence of detectable HBeAg have a generally worse prognosis and a greater chance of developing advanced fibrosis and/or cirrhosis than those without HBeAg. Liver fibrosis progresses rapidly to cirrhosis in patients with dual infection with HBV and hepatitis delta virus [17]. Co-infection of HIV with hepatitis viruses is a common cause of liver fibrosis [18]. Importantly, advanced liver fibrosis and cirrhosis are the main causes of mortality among HIV-infected individuals in developed countries, in which highly active antiretroviral therapy is widely available. Both HBV and HCV are more common in HIV-infected individuals than in the general population as a result of shared risk factors for viral acquisition. Co-infection with HIV results in greater likelihood of chronicity and enhanced viral replication in the setting of both HBV and HCV infections. Alcoholic liver disease (ALD) is one of the commonest causes of liver fibrosis worldwide [19]. Both genetic and environmental factors are associated with fibrosis progression in these patients [20]. Patients developing alcoholic hepatitis are at high risk of progressing to advanced fibrosis and cirrhosis [21]. This condition is characterized by evidence of hepatocyte necrosis and apoptosis, leukocyte infiltration, and perisinusoidal/pericellular fibrosis. The only effective maneuver leading to fibrosis regression in these patients is alcohol abstinence. Until now, the development of specific antifibrotic treatments has been hampered by the lack of detailed knowledge of the disease mechanisms [22]. This situation may now be changing, with improved insight into pathogenesis obtained from

29: HEPATIC FIBROSIS

studies that focus on the roles of inflammation, endotoxin, and immunity [23]. Among causes of metabolic origin, non-alcoholic fatty liver disease (NAFLD) has recently emerged as a major cause of liver fibrosis [24]. NAFLD encompasses fatty liver alone as well progressive disease in a subset of patients who develop hepatocellular injury, inflammatory changes, and progressive fibrosis (NASH). In the general adult population, NAFLD prevalence ranges between 20 and 30%, while the prevalence of NASH is 1–5%. The presence of necroinflammation has been associated with a significant risk of progression to advanced fibrosis and eventually to liver cirrhosis and hepatocellular carcinoma [25]. At age >45 years, obesity and diabetes have also been associated with an increased risk of liver fibrosis and progression to cirrhosis. Hereditary hemochromatosis is the most common genetic cause of liver fibrosis [26]. Factors such as moderate alcohol consumption can accelerate the progression of fibrosis [27]. Wilson disease is an autosomal recessive disorder of copper metabolism that may result in liver fibrosis [28]. Resultant liver damage leads to steatosis, inflammation, and progressive fibrosis. Wilson disease should be suspected in a patient with liver fibrosis of unknown origin and neurosychiatric symptoms. Alpha-1-antitrypsin deficiency is the most common genetic cause of liver fibrosis in children [29]. In adults, alpha-1-antitrypsin deficiency may cause severe liver injury and is a common finding in patients with hepatic fibrosis of unknown origin. Heterozygocity for the Pi*Z allele may also predispose to liver fibrosis in patients with ongoing liver injury (HCV infection and/or NASH). Other inherited causes of fibrosis include glycogen storing diseases, cytokeratin mutations, galactosemia, and congenital tyrosinosis. Even diabetes mellitus itself has been associated with the development of fibrosis. Several autoimmune liver diseases may result in liver fibrosis. PBC is a common cause of liver fibrosis in women [30]. Affected patients are typically middle-aged women with abnormal serum concentrations of alkaline phosphatase. Identification of PBC is important, because effective treatment with ursodeoxycholic acid has been shown to halt fibrosis progression and improve survival without need for liver transplantation [31]. PSC is a chronic cholestatic liver disease characterized by strictures of the biliary tree. Small-duct PSC is a distinct clinical entity associated with a benign course [32]. Patients eventually show progressive fibrosis and cirrhosis. AIH is characterized histologically by interface hepatitis, and by the presence of non-organ and liver-specific autoantibodies [33]. In severe cases, which are characterized by repeated relapses, fibrosis often develops and its progression is attenuated and even reversed if effective immunosuppressant therapy is initiated. Other diseases can also cause liver fibrosis. Cardiac fibrosis includes a spectrum of hepatic derangements that occur in the setting of right-sided heart failure.

435

Prolonged biliary obstruction (secondary biliary fibrosis) results in liver fibrosis [9]. It can result from common bile-duct stones, bile-duct or head-of-the-pancreas carcinoma, biliary-tract infections or strictures, and, in children, biliary atresia. This latter condition is the main cause of advanced liver fibrosis requiring liver transplantation among children. Granulomatous diseases can cause liver fibrosis, including sarcoidosis and infections (mycobacterial infections and schistosomiasis). Hepatotoxic drugs can also cause liver fibrosis [34]. Most cases resolve after withdrawal of the drug, but in undiagnosed cases the liver disease can progress to cirrhosis. Drug-induced liver disease can mimic AIH or veno-occlusive disorders. Herbal and traditional medicines, which are currently ingested by many people, should always be considered in any patient with liver fibrosis of unknown origin. Venous outflow obstruction (e.g. Budd–Chiari syndrome) can result in severe liver fibrosis and cirrhosis [35]. Most cases are due to an underlying hypercoagulable disorder from myeloproliferative disorders or abnormalities in the coagulation cascade. Fibrosis progresses rapidly to severe portal hypertension and liver failure in some patients with Budd–Chiari syndrome. Hepatic veno-occlusive disease is a common complication of bone marrow transplantation [36]. Perivenular fibrosis develops, which can result in jaundice and portal hypertension. The diagnosis of cryptogenic fibrosis is made when all known causes of liver disease are ruled out. Therefore, medical history, consumption of alcohol and/or drugs, virological tests, tests for metabolic diseases, tests for autoimmune diseases, and careful imaging analysis of the liver should be performed. Importantly, recent data suggest that many patients with cryptogenic fibrosis and cirrhosis show features of the metabolic syndrome, including obesity, diabetes, and dyslipemia. Therefore, NASH may be the underlying cause of cases of cryptogenic cirrhosis [37].

CELLULAR BASIS The cellular mechanisms of liver fibrosis have been uncovered during the past two decades [3]. The key finding was the identification of activated HSCs as the major source of ECM in the fibrotic liver [7]. Activation of HSCs is currently considered the critical common step in liver fibrogenesis. HSCs from normal liver represent about 15% of the total liver cell population and have a “quiescent” phenotype (rich in vitamin A fat droplets). When freshly isolated and cultured, quiescent HSCs show a low proliferative rate and very modest expression of fibrogenic genes, little cytokine secretion, and lack of contractile properties [38, 39]. Therefore, the main function of these quiescent HSCs is the storage and metabolism of vitamin A [40]. However, following liver injury of any etiology, HSCs undergo a process termed “activation,” which represents a transition into proliferative, fibrogenic, pro-inflammatory, and contractile Mybs [41] (Figure 29.2). The initiation

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THE LIVER: CELLULAR BASIS

Apoptosis

Autocrine cytokines Quiescent HSC Retinoid droplets

Neural markers

Activated HSC

Smooth muscle a-actin

Proliferation

Cytokine receptors Proliferative cytokines (PDGF)

Fibrogenic cytokines (TGFb)

Fibrogenic stimulus Type IV collagenase Initiation Phase

Myofibroblast

Perpetuation Phase

Fibrogenesis Matrix induced activation R.

Ri

pp

e

Figure 29.2 Activation process of quiescent hepatic stellate cells into myofibroblastic fibrogenic cells. TGF, transforming growth factor; PDGF, platelet-derived growth factor

phase of this process is probably caused by paracrine stimuli from injured neighboring cells, which include hepatocytes, Kupffer cells, sinusoidal endothelial cells, platelets, and infiltrating inflammatory cells [42, 43]. Some cells (i.e. hepatocytes, Kupffer cells) promote HSC activation by producing lipid peroxides, leading to oxidative stress [44–46]. A number of cytokines released by damaged neighboring cells also activate HSCs. These include transforming growth factor β (TGFβ) 1, platelet-derived growth factor (PDGF), and endothelin (ET) 1 [6, 47, 48]. Activated HSCs perpetuate their own activation by several autocrine loops, including the secretion of TGFβ1 and upregulation of its receptors [42]. Based on its importance, several therapeutic options currently under evaluation are aimed at inhibiting HSC activation (i.e. antioxidants, antagonists to TGFβ1 activity). One important feature of activated HSCs is their proliferative phenotype (Figure 29.3). Following liver injury there is a marked accumulation of α-smooth-muscle actin positive cells at the sites of active liver fibrogenesis [5, 49]. Growth factors for HSCs and their intracellular signaling pathways have been largely characterized in culture [50]. The most powerful growth factor for HSCs is PDGF, but stimulation of other tyrosine-kinase receptors by epidermal growth factor (EGF), fibroblast growth factor (FGF), or insulin-like growth factor (IGF) also increases HSC proliferation [6]. Moreover, substances acting through G-protein-coupled receptors, such as angiotensin II, promote HSC proliferation [51]. Resistance to apoptosis of activated HSCs is a key event in liver fibrogenesis [52]. Soluble survival factors are likely to be important to HSC survival. During liver

injury, hepatocytes and HSCs express IGF type 1, a potent prosurvival factor. In addition, overexpression of TGFβ1 in the injured liver may promote stellate cell survival. The separate regulation of proliferation and survival in HSCs provides a further control on HSC numbers. Moreover, adipokines such as leptin and factors regulating collagen degradation such as tissue inhibitor of metalloproteinases type 1 (TIMP-1) may also act as survival factors for activated HSCs [53, 54]. The abnormal ECM in the fibrous scar also provides key survival signals [55]. There is a striking correlation between the degradation of the collagen-I-rich matrix during recovery from liver fibrosis, and apoptosis of activated HSCs [56]. A decreasing concentration of soluble survival factors, the degradation of specific matrix components, or separation of cells from matrix contact, either separately or in combination, may precipitate an apoptotic response in HSCs. Apoptosis of activated HSCs plays a critical role in the spontaneous recovery from experimental fibrosis in different experimental models (carbon tetrachloride (CCl4 ) administration, bile duct ligation) [57]. Conversely, resistance to apoptosis and the consequent prolonged survival of activated HSCs may contribute to the progression of hepatic fibrosis [58]. Migration of HSCs contributes to their accumulation at sites of liver injury. Following activation, cultured HSCs migrate in response to several stimuli, including growth factors (PDGF), vasoactive substances (angiotensin II, ET-1), and chemokines (monocyte chemotactic protein (MCP) 1) [59, 60]. Although the importance of migration of HSCs in vivo has not been determined, substances that

29: HEPATIC FIBROSIS

ROS NADPH

Retinoid Loss Prolifera

437

id

H p

Chemotaxis

Degradation

Figure 29.3 Biological properties of activated HSCs that participate in liver fibrogenesis. HGF, hepatocyte growth factor; ET-1, endothelin-1; PDGF, platelet-derived growth factor; MCP-1, monocyte chemotactic factor type 1; MMP, metalloproteinase

inhibit migration of HSCs could theoretically reduce their accumulation in the injured liver. The interaction of HSCs with the ECM is of major importance in the profibrogenic behavior of activated HSCs. Proteins including type IV collagen, fibronectin, proteoglycans, and urokinase-type plasminogen activator are released by sinusoidal endothelial cells and accumulate around the hepatic sinusoids [61]. These factors contribute to the activation of resident HSCs by the stimulation of latent cytokines such as TGFβ [62]. Of particular interest is a “fetal” isoform of fibronectin, [EIIIA]Fn, which is de novo synthesized by the damaged liver and contributes to the activation of resident HSCs [63]. Once HSCs are activated, they secrete fibrillar collagens and other ECM proteins, which also accumulate in the space of Disse. This altered microenvironment can amplify the fibrogenic activity of HSCs by different mechanisms. First, fibrillar collagens can bind to discoidin domain receptors (DDRs) and activate intracellular signaling pathways [55]. Second, the altered ECM can serve as a reservoir for a number of growth factors (PDGF, TGFβ, FGF) and matrix metalloproteinases (MMPs), which are released and reach neighboring cells including HSCs [64]. This implies that proteolysis of the ECM during liver inflammation can initiate reparative processes by the released growth factors and MMPs. Third, HSCs express a number of integrins, heterodimeric transmembrane proteins whose ligands are matrix proteins, which transduce the extracellular signals from the ECM into the cells [65]. Several activities of HSCs can be regulated by integrins, including cell proliferation, contraction, migration, and collagen synthesis [66]. Moreover, integrins can also activate latent TGFβ, thus amplifying the fibrogenenic action of this key cytokine.

HSCs also play an active role in hepatic inflammation [67]. Activated HSCs migrate in response to cytokines released by monocytes, as well as secreting a number of proinflammatory cytokines and chemokines [68]. The mechanisms involved in cytokine secretion by HSCs include activation of the transcription factor nuclear factor κB (NFκB) [69]. Moreover, activated HSCs express a number of chemokine receptors, which activate several intracellular pathways, stimulating the inflammatory action of these cells [70]. In addition, HSCs can take up and process antigens and, under stimulation with cytokines, express the cell machinery required for antigen presentation and thereby modulate the growth of lymphocytes [71]. Another potential fibrogenic mechanism is the capability of HSCs to perform phagocytosis of apoptotic bodies and lymphocytes [72]. Therefore, the use of substances with combined antifibrotic and anti-inflammatory effect should be considered in antifibrotic therapy. Promoting angiogenesis is another biological property of HSCs that may influence fibrogenesis. Following activation, HSCs secrete a number of angiogenic factors, such as vascular endothelial growth factor (VEGF) and angiopoietin-1 [73, 74]. Hypoxia and fibrogenic cytokines such as angiotensin II and leptin stimulate the pro-angiogenic behavior of HSCs [75]. Importantly, agents that inhibit angiogenesis attenuate liver fibrosis in vivo, suggesting an important role for angiogenesis during liver fibrogenesis [76]. Hepatic cell types other than HSCs also have fibrogenic potential (Figure 29.4). Mybs derived from small portal vessels proliferate around biliary tracts in cholestasis-induced liver fibrosis to initiate collagen deposition [77]. HSCs and portal Mybs differ in specific

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THE LIVER: CYTOKINES INVOLVED IN HEPATIC FIBROSIS

CD45CD34Colα1(I)+ α-SMA+ Desmin+ FSP-1+ GFAP+ Synamin+

CD45Albumin+/CK19+ FSP1+ Colα1(I)+ α-SMA+

CD45+ Colα1(I)+ CD11b+ MHC II+ ICAM I+ α-SMA-

CD45-/+ CD34Colα1(I)+ α-SMA-/+

by releasing ROSs and cytokines [84, 85]. Recently, a role for macrophages in hepatic wound healing has been proposed [86]. Macrophage depletion during progressive fibrosis produces reduced scarring and fewer Mybs. Macrophage deletion during recovery, by contrast, leads to a failure of matrix degradation. Endothelial sinusoidal cells can also influence the activation of HSCs. In fact, capillarization of the hepatic sinusoids precedes hepatic fibrosis, and capillarization of sinusoidal endothelial cells is permissive for activated HSCs [87]. Finally, changes in the composition of the ECM can directly stimulate fibrogenesis. Fibrillar collagens can bind and stimulate HSCs via DDR2 and integrin alphavbeta6. Moreover, the altered ECM can serve as a reservoir for growth factors and MMPs [64].

Activated myofibroblast

Figure 29.4 Different sources of myofibroblastic cells with fibrogenic potential in livers with chronic damage. SMA, smooth muscle actin; GFAP, glial fibrillary acidic protein; Col, collagen; ICAM, intercellular adhesion molecule; FSP, fibroblast-specific protein

cell markers and response to apoptotic stimuli [8]. Culture of CD34 + CD38- hematopoietic stem cells with various growth factors has been reported to generate HSCs and Mybs of bone marrow origin that infiltrate human livers undergoing tissue remodeling [78]. These data suggest that cells of bone marrow origin can be a source of fibrogenic cells in the injured liver. Other potential sources of fibrogenic cells are circulating fibrocytes [79]. Finally, epithelial-to-mesenchymal transition, a TGFβ1-driven mechanism that participates in renal fibrosis, may also occur during chronic liver injury [80]. The role of this mechanism is uncertain, since epithelial-to-mesenchymal transition does not seem to be involved in experimentally-induced liver fibrosis. A complex interplay among different hepatic cell types takes place during hepatic fibrogenesis [81]. Hepatocytes are targets for most hepatotoxic agents, including hepatitis viruses, alcohol metabolites, and bile acids. Damaged hepatocytes release reactive oxygen species (ROSs) and fibrogenic mediators and induce the infiltration by inflammatory cells [82]. Apoptosis of damaged hepatocytes stimulates the fibrogenic actions of liver Mybs [52]. Inflammatory cells, either lymphocytes, macrophage, or polymorphonuclear cells, activate HSCs. Activated HSCs secrete inflammatory chemokines, express cell adhesion molecules, and modulate the activation of lymphocytes [67]. Therefore, a vicious circle in which inflammatory and fibrogenic cells stimulate each other is likely to occur. Fibrosis is influenced by different T-helper (Th) subsets. The Th2 response is associated with more active fibrogenesis [83]. Kupffer cells are resident macrophages that play a major role in liver inflammation

CYTOKINES INVOLVED IN HEPATIC FIBROSIS Cytokines include chemokines, interleukins (ILs), interferons (IFNs), growth factors, angiogenic factors, vasoactive substances, soluble receptors, and soluble proteases. Unregulated cytokine synthesis and release coordinate the hepatic response to injury and participates in the initiation, progression, and maintenance of fibrosis. Inflammatory cytokines in the hepatic wound-healing response are derived from Kupffer cells, hepatocytes, HSCs, natural killer (NK) cells, and lymphocytes including CD4+ Th [1]. Th cells can differentiate into Th1 and Th2 subsets, a classification that is based on the pattern of cytokines produced. In general, Th1 cells produce cytokines that promote cell-mediated immunity (IFNγ, tumor necrosis factor (TNF)α, and IL-2) and protect against hepatic fibrosis, while Th2 cells promote humoral immunity (IL-4, IL-5, IL-6, and IL-13) and induce fibrosis, as evidenced by a study using two mice strains with different polarities of Th cells and different susceptibilities to liver fibrosis [83]. TNFα participates in the activation process of HSCs and in cholestasis-induced liver fibrosis [88]. In contrast, IL-10 exerts net antifibrogenic effects in the liver, both in animals and in humans [89]. IFNα has been shown to exert a direct antifibrotic effect and is used as antiviral therapy in patients with chronic HCV [90]. IFNγ inhibits HSC proliferation and procollagen mRNA expression in vitro and reduces liver fibrosis in rodents [91]. Among cysteine-cysteine (CC) chemokines, MCP-1 is a profibrogenic chemokine that is markedly overexpressed in the injured liver [92]. MCP-1 induces chemotaxis of HSCs and participates in experimentally induced fibrosis in rats [59]. CCL5 (RANTES) is upregulated in livers from HCV patients and in experimental models of liver fibrosis. RANTES induces HSC proliferation but not increased ECM production [68]. Several studies have demonstrated that cysteine-X-cysteine (CXC) chemokines are also involved in liver fibrosis. For instance, serum IL-8 is increased in patients with ALD,

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NASH, and PBC, and GROα expression is upregulated in livers from patients with alcoholic hepatitis [21]. Several growth factors play a key role in liver fibrogenesis by promoting activation and accumulation of HSCs and stimulating collagen synthesis [93]. PDGF and transforming TGFβ are the most important mediators due to their effects on HSC proliferation and ECM protein production, respectively. PDGF-B is the most potent mitogenic factor for HSCs, acting through PDGFRβ [6]. All isoforms of PDGF and PDGFR are upregulated in injured livers and correlate with the degree of inflammation and fibrosis [94]. Moreover, inhibition of PDGF-B attenuates experimental liver fibrogenesis [95]. TGFβ1 is a key mediator of liver fibrogenesis. In the injured liver, TGFβ1 is upregulated [47] and favors the transition of resident HSCs into Myb-like cells, stimulating synthesis of ECM proteins and inhibiting their degradation. Strategies aimed at disrupting TGFβ1 synthesis and/or signaling pathways markedly decreased fibrosis in experimental models [63, 96]. A newly discovered regulator of TGFβ activity is bone morphogenic protein and activin membrane-bound inhibitor (BAMBI), which is a TGFβ pseudoreceptor that inhibits TGFβ signaling by preventing the formation of receptor complexes [97]. Downregulation of BAMBI is a mechanism of fibrogenesis induced by lipopolysaccharide (LPS) through toll-like receptor (TLR)4. Established angiogenic growth factors such as VEGF and FGF play a role not only in angiogenesis but also in chronic wound-healing conditions. VEGF and its receptors (VEGFR-1 and VEGFR-2) are upregulated in chronic liver injury and promote fibrogenic effects in HSCs by stimulating cell proliferation, collagen production, and migration [98]. A number of vasoactive substances are locally produced in the injured liver and act on HSCs through an autocrine and/or paracrine manner. Vasodilator substances (e.g. nitric oxide (NO), prostaglandin E2, atrial natriuretic peptide, adrenomedullin, and relaxin) exert antifibrotic effects, while vasoconstrictors (e.g. ET-1, norepinephrine (NE), angiotensin II, and thrombin) have opposite effects [99]. In livers with advanced fibrosis, there is a predominance of vasoconstrictors over vasodilators, favoring fibrosis. Interestingly, advanced fibrosis is associated with endothelial dysfunction and decreased NO production, which may contribute to disease progression [100]. Prostaglandin E2 is a vasodilatory molecule synthesized by virtually all liver cells that inhibits HSC proliferation and TGFβ1-mediated collagen synthesis and attenuates fibrosis in vivo [101]. Drugs that deliver either NO or PGE2 have been proposed for the treatment of patients with liver fibrosis. Among vasoconstrictors, angiotensin II is the most widely studied. Accumulating evidence indicates that angiotensin II, the main effector of the renin-angiotensin system (RAS), is a true cytokine that plays a major role in liver fibrosis [102] (Figure 29.5). A complete intrahepatic RAS is expressed in chronically damaged livers, and angiotensin II induces an array of

439

fibrogenic actions in HSCs, including increased collagen synthesis and secretion of inflammatory mediators [51]. These effects are mediated by NADPH oxidase-generated ROSs and are prevented by angiotensin type 1 receptor blockers [60]. Importantly, inhibition of the RAS markedly attenuates experimentally induced liver fibrosis in rodents [103]; the effect on humans is being investigated in clinical trials. Thrombin is produced by activated HSCs and regulates cell migration, proliferation, and fibrogenesis [104]. Both thrombin and PAR-1, its main receptor, are overexpressed in fibrotic livers. Moreover, antagonism of thrombin attenuates liver fibrosis in animal models [105]. ET is another important vasoconstrictor implicated in liver fibrosis. ET and its receptors are upregulated in the fibrotic liver [106]. In the early phase of activation, HSCs have a majority of ETA receptors, which stimulate an increase in intracellular free calcium in HSCs coupled with cell contraction and proliferation. This is linked to stimulation of fibrogenesis. In later stages, ETB receptors become more abundant and their stimulation promotes an antiproliferative effect. The use of ETA/ETB receptor blockers has yielded conflicting results [107, 108], possibly due to the different relative activities toward each of the two receptors. NE is a catecholamine with a dual role as a neurotransmitter and a hormone. There is evidence indicating that NE stimulates liver fibrogenesis. Activated HSCs are capable of secreting mature NE, which induces pro-inflammatory and fibrogenic effects. Moreover, α1 adrenoreceptors are upregulated in livers with advanced fibrosis and their blockade attenuates the development of liver fibrosis in rats with chronic liver injury [109, 110]. Adipokines are biologically active peptides mainly secreted by adipose tissue. Adipokines include leptin, resistin, visfatin, and adiponectin. Circulating adipokines secreted by excessive fat accumulation may regulate hepatic fibrosis in diseases such as NASH [111]. Moreover, several adipokines are locally synthesized in the liver and may regulate fibrogenesis in an autocrine/paracrine manner. Leptin is secreted by activated HSCs and stimulates cell proliferation, secretion of chemokines, and collagen synthesis. Moreover, leptin is required for fibrosis development [112]. Serum adiponectin levels are decreased in obese individuals. Adiponectin markedly inhibits liver fibrogenesis in vitro and in vivo [113]. Therefore, obesity may induce hepatic fibrosis by increasing leptin and decreasing adiponectin levels. Resistin is upregulated in ALD and exerts pro-inflammatory effects on HSCs, suggesting a role in liver fibrogenesis [114]. Finally, cannabinoid s ( CBs) display a wide variety of peripheral functions, including regulation of wound-healing response to injury. The endogenous CB system has been implicated in liver fibrosis. Both CB1 and CB2 receptors and endocannabinoids are upregulated in chronic liver diseases. Pharmacological and/or genetic inactivation of CB1 reduces fibrosis in different models of chronic liver injury [115]. In contrast, activation of

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THE LIVER: SIGNALING PATHWAYS INVOLVED IN HEPATIC FIBROSIS

Source of Ang II

Locally generated Ang II

Systemic Ang II

Increased Ang II

AT1 receptors

Receptors in HSC

Signaling pathways

Gene transcription

Cell functions

NADPH oxidase

PKC

MAP kinases PI3K/AKT

Calcium channels IP3 receptors

NFκB AP-1

Intracellular calcium

Procollagen α1(I) PAI-1 TIMP-1 MCP-1/RANTES

INCREASED COLLAGEN SYNTHESIS DECREASED COLLAGEN DEGRADATION INFLAMMATION GROWTH/MIGRATION

Clinical consequences

LIVER FIBROSIS

CELL CONTRACTION

PORTAL HYPERTENSION

Figure 29.5 Mechanisms of the pathogenic effect of the renin-angiotensin system in the liver. Increased angiotensin II (Ang II) binds to angiotensin receptors type 1 (AT1) located in activated hepatic stellate cells (HSCs). AT1 receptors activate a non-phagocytic NADPH oxidase to generate ROSs which stimulate redox-sensitive intracellular pathways. Increased gene transcription leads to mitogenic, fibrogenic, and inflammatory properties, promoting fibrogenesis. Ang II increases intracellular calcium and induces cell contraction, increasing intrahepatic vascular resistance and participating in the pathogenesis of portal hypertension. AP-1, activating protein type-1; MAPK, mitogen-activated protein kinase; MCP-1, monocyte chemotactic protein type 1; NFκB, nuclear factor κB; PI3k, phosphoinositol 3 kinase; PAI-1, plasminogen activator inhibitor type 1; PKC, protein kinase C; TIMP-1, tissue inhibitor of metalloproteinases type 1

CB2 receptors attenuates liver injury, inflammation, and oxidative stress, and CB2 knockout mice exposed to CCl4 show enhanced liver fibrosis [116]. Globally, CBs may worsen liver injury since daily cannabis use exacerbates liver fibrosis in patients with chronic HCV [117].

SIGNALING PATHWAYS INVOLVED IN HEPATIC FIBROSIS A variety of intracellular pathways are involved in the pathogenesis of liver fibrosis (Figure 29.6). There is a complex cross-talk between them, which determines the global effect on liver fibrosis [102].

The focal adhesion kinase ( FAK ) phosphoinositol-3phosphate kinase ( PI3K )–Akt signaling pathway mediates a variety of profibrogenic actions in HSCs, including proliferation, chemotaxis, and transcription of profibrogenic genes [118]. This pathway may be activated by growth factors that trigger tyrosine-kinase activity (PDGF, VEGF) or activation of cytokine receptors (MCP-1), but also by other signals including integrins, stimulators of G-protein-coupled receptors (angiotensin II, thrombin), and adipokines (leptin) [60]. Activated Akt induces mTOR activity and signals through mTOR increase the phosphorylation of p70S6 kinase, which phosphorylates a ribosomal subunit and 4E-BP1, leading to upregulation of protein synthesis and stimulation of cell growth signals. Rapamycin, which inhibits mTOR activity, attenuates liver fibrosis, possibly by decreasing growth of HSCs [119].

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TGFβ1 ECM RII

FA

K

RI SMAD2

PDGF

AKT Ras

PDGF

P SMAD3

PI3-K Rac

Raf

mTOR

SMAD2

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PKA MKK

MEK

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4E-BP1 SMAD2

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ATP Fatty acids PPAR

RIP

A TR

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FADD

Wnt

TN

F

TNF-R1

DD

TRAF

MyD88 D 8

Caspases/Apoptosis

IRAK

β-catenin

T TLR4 CD14 4 LBP LPS

Figure 29.6 Signaling pathways involved in the fibrogenic properties of the hepatic stellate cell. Following binding of cytokines such as TGFβ1, PDGF, or TNFα to their cell-surface receptors, activation of several intracellular signaling pathways occurs. PDGF stimulation can induce activation of mitogen-activated protein kinase (MAPK) signaling as well as the phosphatidylinositol 3-kinase-Akt-p70 S6 kinase (PI3K-Akt-p70S6K) signaling pathway. The matrix-associated focal adhesion kinase (FAK) also stimulates the PI3K-Akt-p70S6K signaling pathway. TGFβ1 stimulates transcription of profibrogenic genes by activating the SMAD signaling pathway. Fatty acids and other agonists activate peroxisome proliferator-activated receptors (PPAR) to regulate gene expression. The Wnt/β-catenin pathway is also involved in transcriptional regulation. The bacterial product lipopolysaccharide (LPS) bind to TLR4 and stimulates IL-1 receptor-associated kinase (IRAK) to induce fibrogenic signals. TNFα binds to the protein TRADD to activate c-Jun N-terminal kinase (JNK) and NFκB. Finally, agonists such as PGE2 induce cAMP production and protein kinase A (PKA) activation to inhibit MAPK signaling

Members of the mitogen-activated protein kinase ( MAPK ) family, including extracellular-regulated kinase (ERK), c-Jun N-terminal kinase (JNK), and p38 MAPK are activated by several growth factors and vasoactive peptides and subsequently translocated to the nucleus where they phosphorylate several transcription factors, resulting in cellular responses. In HSCs, ERK regulates cell proliferation, secretion of chemokines, cell migration, and collagen synthesis. This pathway is activated by peptides that induce proliferation (PDGF, thrombin, angiotensin II, VEGF, and leptin) as well as by chemokines [70, 120]. Importantly, ERK activation is induced in vivo in rats with chronic liver injury and after chronic elevation of angiotensin II [121]. In HSCs, JNK

or stress-activated protein kinase (SAPK) is activated in response to cellular stress, bacterial products, FasL, oxidative stress, vasoactive substances (angiotensin II), adipokines (leptin), chemokines (RANTES), and growth factors (TGFβ1 and PDGF) [102]. JNK is a profibrogenic pathway in HSCs, modulating cell growth and secretion of inflammatory cytokines [122]. The p38 pathway seems to have an antiproliferative role in HSCs since blocking p38 activity increases cellular proliferation [123]. The Smad pathway plays a major role in liver fibrosis by signalling TGFβ1 in activated HSCs. TGFβ1dependent Smad signaling may also mediate other fibrogenic factors such as hypoxia [124]. TGFβ1 binds to its type II receptor, which induces the dimerization with

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THE LIVER: SIGNALING PATHWAYS INVOLVED IN HEPATIC FIBROSIS

its type I receptor. Inhibitory Smad proteins mediate the effect of IFNγ on TGFβ signaling in the liver. Interestingly, Smad signalling participates in TGFβ1-dependent mesenchymal-to-epithelial transition in cultured hepatocytes, a mechanism that may produce Mybs in liver fibrogenesis [125]. In vivo, inhibition of Smad signaling suppresses collagen gene expression and hepatic fibrosis in mice [126]. Nuclear factor-κB (NF κB) is a major downstream effector of pro-inflammatory cytokines. Other peptides such as angiotensin II and leptin also activate NFκB signalling [73]. Following HSC activation, NFκB becomes persistently activated and many NFκB-responsive genes (e.g. IL-6, intercellular adhesion molecule (ICAM) 1, etc.) are constitutively expressed [127]. NFκB plays a pivotal role in the inflammatory effects of TNFα and other mediators on HSCs. Its activity is not required for proliferation or activation, but protects activated HSCs against apoptosis by TNFα and other agonists [128]. Nuclear receptors directly bind to DNA and regulate the expression of adjacent genes. Several nuclear receptors have been described in HSCs. The pregnane X receptor (PXR) is a nuclear receptor that seems to exert an antifibrotic role. PXR activators inhibit the proliferation, transdifferentiation, and expression of TGFβ1 in HSCs [129]. In addition, treatment with a PXR activator markedly reduces the degree of liver fibrosis in animal models [130]. Peroxisome proliferator-activated receptors (PPARs) regulate HSCs’ biological actions and are potential targets for antifibrotic therapy [104]. There are three isoforms encoded by three different genes, PPARα, PPARβ, and PPARγ. PPARs mainly regulate metabolic functions in the liver, but also inflammation and fibrogenesis. Following activation of HSCs, expression of PPARγ diminishes, and the expression of PPARβ increases. PPARγ activation inhibits the pro-inflammatory and profibrogenic actions in HSCs and attenuates liver fibrosis in vivo [131], while PPARβ seems to exert opposite effects. Most importantly, PPARγ ligands (e.g. thiazolidinediones) are currently being tested to treat liver fibrosis in the context of NASH [25]. The Wnt/ β-catenin pathway is crucial in normal development, including embryogenesis. This pathway also signals cytokines and promotes inflammation. Recently, it has been implicated in hepatic fibrogenesis [132]. Wnt is an extracellular secreted glycoprotein that binds to its cell-surface receptor, Frizzled (Fz), and induces specific downstream events. In the liver there is evidence indicating that Wnt signalling has a profibrogenic role [133]. In cultured activated HSCs, mRNA for Wnt genes and co-receptors are increased, and protect cells from apoptosis [132]. Moreover, Wnt activity is enhanced in liver cirrhosis. These observations suggest that Wnt signaling promotes hepatic fibrosis by enhancing HSC activation and survival. Toll-like receptors (TLRs) are pattern-recognition receptors that recognize pathogen-associated molecular patterns

and signal through adaptor molecules. Recent studies suggest a role for intracellular pathways driven by TLRs in liver inflammation [134]. In particular, TLR4 and TLR9 are implicated in liver fibrogenesis and LPS signaling [135, 136]. Mice lacking a functional TLR4 have reduced liver fibrosis compared to wild-type mice. The mechanism by which TLR4 promotes liver fibrogenesis has recently been uncovered. TLR4 activation in HSCs reduces BAMBI expression, which is a TGFβ pseudoreceptor, thus TGFβ signaling is enhanced in HSCs by TLR4 activation [97]. The intracellular domain of TLR is similar to that of IL-1 receptor, thus they share intracellular pathways. Stimulated toll/IL-1 receptors activate MyD88, and then the receptor recruits interleukin-1 receptor-associated kinase (IRAK), which becomes activated [137]. This leads to the phosphorylation of the TNF receptor-associated factor (TRAF), which in turn activates pro-inflammatory transcription factors (AP-1 and NFκB). Several pathways implicated in cell death mediate cytokine signalling in activated HSCs. The receptors for TNF belong to a superfamily that includes several transmembrane molecules which bind cytokines and other molecules. TNFα is a critical activator of the pro-inflammatory role of HSCs [138]. TNFα activates JNK in both quiescent and activated HSCs. TNFα also activates ERK1/2 and p38 MAPK, which regulates collagen synthesis in HSCs. Another receptor on HSCs, CD40, interacts with its ligand to amplify the inflammatory behavior of HSCs through TNFR-associated factor 2- and IKappaB kinase (IKK)2-dependent pathways [139]. Cell death is mediated by the interaction of tumor necrosis factor receptor 1-associated death domain protein (TRADD) with Fas-associated protein with dead domain (FADD), which stimulates caspases leading to apoptotic cell death. Fas (CD95) is also expressed in quiescent HSCs and drives proliferation and resistance to apoptosis [140]. Another ligand, tumor necrosis factor-related apoptosis-inducing ligand (TRAIL) binds to TRAIL-R2 in activated HSCs to induce apoptosis [141]. Janus kinases ( JAK s) are intracellular signaling kinases that can bind to several receptors. Signaltransducer-and-activator-of-transcription (STAT) proteins possessing SH2 domain are recruited to the receptors and are phosphorylated at tyrosine residues by JAKs. Activated STAT dimers translocate the cell nucleus and activate transcription of their target genes. In HSCs, this pathway is stimulated by a variety of cytokines and mediators, including INFγ and leptin [142]. Activation of STAT1 plays an important role in liver injury, inflammation, and inhibition of liver regeneration. Mice lacking STAT1 exhibit accelerated liver fibrosis by inhibition of HSC proliferation, suppression of PDGF expression, and inhibition of TGFβ/Smad3 signaling [143]. Adenosine monophosphate-activated protein kinase (AMPK ) is a fuel-sensing enzyme capable of regulating cellular metabolism in response to different stimuli. Once activated, AMPK switches on catabolic pathways,

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leading to adenosine triphosphate (ATP) generation, and inactivates ATP-consuming processes not essential for short-term survival. Adenosine monophosphate-activated protein inhibits cell proliferation, migration, chemokine secretion, and collagen production in HSCs [144]. Interestingly, adiponectin activates AMPK, and this pathway mediates the antifibrogenic effect of adiponectin in HSCs [145].

FIBROSIS REMODELING AND RESOLUTION In contrast to the traditional view that cirrhosis is an irreversible disease, recent evidence indicates that even advanced fibrosis is reversible [146]. In experimentally-induced fibrosis, cessation of liver injury results in fibrosis regression [147]. In humans, resolution of liver fibrosis can occur after successful treatment of the underlying liver disease. This observation has been described in patients with iron and copper overload, alcohol-induced liver injury, chronic hepatitis C, B, and D, hemochromatosis, secondary biliary cirrhosis, NASH, and AIH [1]. The time taken to achieve significant regression may be years and varies depending on the underlying cause and the severity of the liver disease. Chronic HCV infection is the most extensively studied condition, and successful therapy (IFNα plus ribavirin) with viral clearance results in fibrosis improvement. Importantly, nearly half of patients with cirrhosis reverse to a significant degree [148]. Whether this beneficial effect is associated with improvements in long-term clinical outcome, including decreased portal hypertension, is unknown. Increased collagenolytic activity is a major mechanism of fibrosis resolution [149]. Fibrillar collagens (I and III) are degraded by interstitial MMPs (MMP-1, -8, and -13 in humans and MMP-13 in rodents). During fibrosis resolution, MMP activity increases due to a rapid decrease in the expression of the inhibitor TIMP-1. Partial degradation of fibrillar collagen occurs, and the altered interaction between activated HSCs and ECM favors apoptosis of the activated HSCs [147]. Removal of activated HSCs by apoptosis precedes fibrosis resolution. Stimulation of death receptors in activated HSCs and a decrease in survival factors, including TIMP-1, can precipitate HSC apoptosis [57]. HSC senescence is an alternative mechanism for terminating fibrogenesis. Senescent cells, mainly derived from HSCs, accumulate in livers with advanced fibrosis. Interestingly, senescent activated HSCs exhibit gene expression profile consistent with cell-cycle exit, reduced secretion of ECM components, enhanced secretion of ECM-degrading enzymes, and enhanced immune surveillance. NK cells probably kill senescent activated HSCs, thereby facilitating the resolution of fibrosis. Activated HSCs are the main hepatic cell type implicated in collagen degradation in fibrotic liver [150],

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while recruited monocytes also express collagenases. Following cell activation, HSCs express virtually all the key components required for matrix degradation. In particular, they are a key source of MMP-2, as well as stromelysin/MMP-3. Importantly, through the activation of TIMP-1 and -2, activated HSCs also inhibit the activity of interstitial collagenases, which degrade fibrillar collagen, thus favoring the accumulation of ECM [151]. This hypothesis is supported by studies in experimental liver fibrosis and in human liver disease, in which TIMP-1 expression is markedly upregulated in cirrhotic compared with normal liver, whereas expression of collagenases remains unchanged [152]. Moreover, transgenic mice overexpressing TIMP-1 rapidly develop liver fibrosis following injury [153]. Administration of TIMP-1 antibodies to rats with established fibrosis partially removes collagen from the liver, suggesting that this approach may be useful in patients with chronic liver diseases [154].

PATHOPHYSIOLOGY-ORIENTED ANTIFIBROTIC THERAPIES There is no gold-standard treatment for liver fibrosis. Experimental studies have revealed targets to prevent fibrosis progression in rodents [11] (Table 29.2). However, the efficacy of most treatments has not been tested in humans. This is due to the need to perform serial liver biopsies to accurately assess changes in liver fibrosis and the requirement of long follow-up studies. The development of reliable non-invasive markers of liver fibrosis will positively impact the design of clinical trials. The ideal antifibrotic therapy would be the one that is liver-specific, well tolerated when administered for prolonged periods of time, and effective in attenuating excessive collagen deposition without affecting normal ECM synthesis. The removal of the causative agent is the most effective maneuver in the treatment of liver fibrosis. This strategy is effective in most etiologies of chronic liver diseases [9, 148, 155–157]. For patients with cirrhosis and clinical complications, liver transplantation is currently the only curative approach [158, 159]. In randomized clinical trials using IFNα plus ribavirin in patients with chronic HCV, the persistent clearance of viral infection is associated not only with the resolution of hepatic inflammation but also with an improvement in liver fibrosis [148]. Moreover, long-term treatment with IFNα is commonly associated with improvement of liver fibrosis, including in patients with cirrhosis [160]. These findings raise the possibility that IFNα has an intrinsic antifibrotic effect independent of its antiviral effect. This hypothesis is supported by experimental models in which IFNα inhibits the fibrogenic action of HSCs and prevents the development of liver fibrosis [91]. However, when chronic HCV patients who failed to have a sustained viral response were treated with chronic IFNα therapy (hepatitis C antiviral long-term

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Table 29.2 Potential antifibrotic drugs to treat liver fibrosis Agent

Antifibrotic effects in hepatic stellate cells

Antifibrotic effect in experimental fibrosis

Angiotensin inhibitors Colchicine Corticosteroids

Consistent positive data Not tested Limited data

Consistent positive data Limited data Limited data

Endothelin inhibitors Interferon-α Interleukin-10

Not tested Consistent positive data Limited data

Limited data Consistent positive data Consistent positive data

Pentoxifylline Phosphatidylcholine

Consistent positive data Limited data

Consistent positive data Consistent positive data

PPAR antagonists S-adenosyl-methionine

Consistent data Not tested

Consistent positive data Not tested

Sho-saiko-to (asiatic herb)

Consistent positive data

Consistent positive data

TGFβ inhibitors Tocopherol CB1 antagonists

Consistent positive data Consistent positive data Consistent positive data

Consistent positive data Limited data Consistent positive results

treatment against cirrhosis (HALT-C) trial), there was no improvement in fibrosis. Because inflammation precedes and promotes the progression of liver fibrosis, anti-inflammatory drugs have been proposed. Corticosteroids are only indicated for the treatment of hepatic fibrosis in patients with AIH and acute alcoholic hepatitis [155]. Inhibition of the accumulation of activated HSCs by either inhibiting their activation/proliferation or promoting their apoptosis is another strategy. Antioxidants such as vitamin E, silymarin, phosphatidylcholine, and S-adenosyl-L-methionine inhibit HSC activation, protect hepatocytes from undergoing apoptosis, and attenuate experimental liver fibrosis [161–163]. Antioxidants may exert beneficial effects in patients with alcohol-induced liver disease and NASH [164], but the results need to be confirmed. Disrupting TGFβ1 synthesis and/or signaling pathways prevents scar formation in experimental liver fibrosis [165]. This strategy includes inhibition of key members of the TGFβ superfamily, such as bone morphogenic protein 7 [166]. Moreover, administration of growth factors (e.g. insulin growth factor, hepatocyte growth factor, and cardiotrophin) or their delivery by gene therapy attenuates experimental liver fibrosis [167, 168]. Inhibition of growth factors involved in liver fibrogenesis (i.e. PDGF-BB) also attenuates liver fibrosis in rats [169]. However, these approaches have not been tested in humans and may favor cancer development. Substances that inhibit key signal transduction pathways involved in liver fibrogenesis also have potential to treat liver fibrosis [10]. These include pentoxifylline (phosphodiesterase inhibitor), amiloride (Na+ /H+ pump inhibitor), and S-farnesylthiosalicylic acid (RAS antagonist). Stimulation of pathways with antifibrogenic actions

Antifibrotic effect in humans Retrospective study Discrepant results Effective in autoimmune hepatitis Not tested Effective in chronic hepatitis C Isolated reports in chronic hepatitis C Not tested Not proven in alcohol-induced fibrosis Isolated reports in NASH Effective in alcohol-induced fibrosis Isolated reports in chronic hepatitis C Not proven Isolated reports in NASH Epidemiological data

(e.g. STAT1) can also reduce collagen deposition [143]. Ligands of PPARα and/or PPARγ such as thiazolindiones exert beneficial effects in experimental liver fibrosis and in patients with NASH [170, 171]. Drugs that interfere with AKT/PI3k pathways, such as rapamycin, inhibit fibrosis in rats [119]. The usefulness of rapamycin in humans deserves further investigation. Pharmacological manipulation of nuclear factors can also modulate liver fibrosis in vivo. For example, stimulation of pregnate receptors inhibits liver fibrosis in rats [129, 172]. Promoting apoptosis of activated HSCs is an alternative approach to treating liver fibrosis. Substances such as gliotoxin induce apoptosis of cultured cells and remove activated HSCs from fibrotic livers [173–175]. These effects can be also achieved by inhibiting antiapoptotic factors in HSCs such as IKKs or by using proteasome inhibitors [176, 177]. Another substance that causes death of HSCs is anandamide [178]. Anandamide is an endogenous CB, a family that has been shown to modulate hepatic fibrogenesis both in vitro and in vivo. Endocannabinoids bind to CB1 and CB2 receptors, which exert powerful pro- and antifibrogenic effects, respectively [115, 116]. The CB1 antagonist ramaban inhibits experimental liver fibrosis. The inhibition of the renin-angiotensin system is probably the most promising strategy for the treatment of liver fibrosis [12]. There is overwhelming experimental data indicating that pharmacological and/or genetic inhibition of this system attenuates liver fibrogenesis in rodents. Renin-angiotensin inhibitors are widely used as antifibrotic agents in patients with chronic renal and cardiac diseases and appear to be safe when administered for prolonged periods of time [179]. Little information is available on liver fibrosis. Transplanted patients receiving

29: HEPATIC FIBROSIS

RAS inhibitors as antihypertensive therapy show less fibrosis progression than patients receiving other types of drugs [180]. Moreover, AT1 receptor blockers reduce liver fibrosis in patients with NASH [181]. Large prospective clinical trials should be performed to confirm this promising data. Other vasoactive substances are potential targets for treatment of liver fibrosis. The blockade of ET-1 type A receptors and the administration of vasodilators (prostaglandin E2 and NO-donors) exert antifibrotic activity in rodents, yet the effects in humans are unknown [182]. Cyclo-oxygenase 2 (COX-2), a key enzyme in the generation of vasodilatory prostaglandins, is a novel target for the treatment of liver fibrosis. COX-2 inhibition has yielded contradictory results, probably due to differences of specificity between the inhibitors used. In particular, COX-2 inhibitors with intrinsic PPARγ activity have net antifibrotic effect [183]. Finally, the mineralocorticoid receptor antagonist spironolactone also suppresses liver fibrogenesis in rodents [184]. Different herbal compounds, many of them traditional medicines used in Asian countries to treat liver diseases, have been demonstrated to have antifibrotic effects [10]. They include sho-saiko-to, glycyrrhizin, and savia miltiorhiza. An alternative approach is to inhibit collagen production and/or promote its degradation [185]. Inhibitors of prolyl-4 hydoxylase and halofuginone prevent the development of experimental liver cirrhosis by inhibiting collagen synthesis. MMP-8 and urokinase-type plasminogen activator stimulate collagen degradation in vivo [186]. Antibodies that block the tissue inhibitors of metalloproteinases such as TIMP-1 reverse established liver fibrosis, suggesting that they may be useful for patients with advanced liver diseases [154]. The efficacy of these drugs in humans is unknown and they may result in undesirable side effects. Limitations of the current antifibrotic approaches are that antifibrotic drugs are not efficiently taken up by activated HSCs and may produce unwanted side-effects. Cell-specific delivery to HSCs could provide a solution to these problems. Promising preliminary results have recently been obtained using different carriers (e.g. cyclic peptides coupled to human serum albumin (HSA)-recognizing collagen type VI receptor and/or PDGF receptor, vitamin A bearing liposomes) [187]. Drugs such as pentoxifylline and micophenolic acid are efficiently released into activated HSCs and reduce liver fibrogenesis [188, 189]. Interestingly, this approach could also be useful for treating portal hypertension, a condition that involves chronic contraction of activated HSCs.

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(2006) Selective targeting of pentoxifylline to hepatic stellate cells using a novel platinum-based linker technology. J Control Release, 111, 193–203. 189. Greupink, R., Bakker, H.I., Reker-Smit, C., van Loenen-Weemaes, A.M., Kok, R.J., Meijer, D.K., Beljaars, L. et al. (2005) Studies on the targeted delivery of the antifibrogenic compound mycophenolic acid to the hepatic stellate cell. J Hepatol , 43, 884–92.

30

Matrix Giuliano Ramadori and Jozsef Dudas Department of Internal Medicine, Section of Gastroenterology and Endocrinology, Georg-August-University Goettingen, Goettingen, Germany

DEFINITION AND BACKGROUND The extracellular matrix (ECM) provides cells with positional information and a mechanical scaffold for adhesion and migration. The ECM includes the interstitial matrix and the basement membrane. The interstitial matrix is present between various cells (i.e. in the intercellular spaces). Gels of polysaccharides and fibrous proteins fill the interstitial space and act as a compression buffer against the stress placed on the ECM. Basement membranes are sheet-like depositions of ECM on which various epithelial cells rest [1]. The ECM consists of peptidic components such as the collagen and non-collagen proteins, and of non-peptidic components such as the glycosaminoglycans and hyaluronanes. Several peptides and proteins are bound specifically by the ECM, such as certain growth factors/cytokines, matrix metalloproteinases (MMPs), and processing enzymes such as tissue transglutaminase and procollagen propeptidases. Partially via defined oligopeptide sequences or structural domains, the ECM transfers specific signals to cells that act in concert with growth factors/cytokines. These signals either confer stress activation, with a resultant fibrogenic response, or stress relaxation, with a fibrolytic response. Alternatively, ECM-derived peptides can modulate angiogenesis, or growth factor and MMP availability and activity [2].

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

ECM SYNTHESIS AND REMODELING DURING EMBRYONIC LIVER DEVELOPMENT Our knowledge of the molecular mechanisms of ECM synthesis in liver development is still limited. Recently Sugiyama et al. provided novel data in this regard [3]. Primitive sinusoidal structures are well developed in the fetal mouse liver, and have a typical vessel structure accompanied by ECM deposition and hepatic stellate cells (HSCs) in their surroundings, as seen in the adult hepatic sinusoids. During liver development, these constituent cells intimately interact with one another, which leads to mature liver lobule development. Several growth factors or cytokines, including fibroblast growth factors (FGFs), oncostatin M, and tumor necrosis factor (TNF) α, and components of the ECM, are known to act on these interactions [3]. Types I and III collagen were deposited mainly in mesenchymal cell regions of aggregates consisting of hepatoblasts and non-parenchymal cells as fine fibrils or fibers (Table 30.1). Such deposition was not seen in most aggregates that lacked hepatoblasts, or in small aggregates irrespective of the presence of hepatoblasts. Nidogen and type IV collagen immunoreactions started in aggregates containing at least hepatoblasts and desmin-positive cells;

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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THE LIVER: ECM SYNTHESIS AND REMODELING DURING EMBRYONIC LIVER DEVELOPMENT

Table 30.1 Liver ECM during liver development, in normal liver and after liver injury

During embryonic liver development In normal liver

ECM of the subendothelial space of Disse

ECM of the portal area

Nidogen and type IV collagen deposited by desmin-positive mesenchymal cells just around hepatoblasts Interstitial and a basement membrane-like ECM composed of fibrillar collagens type I, III, and V, microfibrillar collagen VI, the small proteoglycan decorin, FN, tenascin-C, laminin, nidogen, heparan sulfate proteoglycans, and others

Types I and III collagen deposited mainly in mesenchymal regions of cell aggregates

Liver inflammation

No significant change

Liver fibrosis

Formation of an incomplete subendothelial basement membrane, creating diffusion barriers between hepatocytes and the liver sinusoid (capillarization of sinusoids)

they were mainly deposited in mesenchymal cells just around hepatoblasts or hepatocytes (Table 30.1). ECM deposition preferentially occurred in aggregates of both hepatoblasts and mesenchymal cells, suggesting that their cellular interactions were pivotal for ECM deposition. As collagen and nidogen immunoreactions were seen in the non-parenchymal regions, their production appeared to take place in non-parenchymal cells. Because small aggregates tended not to deposit ECM components even though they contained mesenchymal cells and hepatoblasts, ECM deposition might depend on cell population sizes. These cellular interactions (hepatoblast–mesenchymal cell interactions and community effects) might take place during liver development, and determine ECM synthesis [3]. Fibronectin (FN) and type I collagen are reported to be involved in the differentiation of the bile duct epithelium in vitro. On the other hand, laminin and type IV collagen are broadly expressed in the hepatic connective tissue, including the portal vein. Laminin and type IV collagen could enhance the ability of the type I collagen gel to induce the bile duct epithelium. Laminin and type IV collagen are additional players regulating bile duct differentiation of the hepatic epithelium in the embryonic liver. Culture of the hepatic epithelium on Matrigel led to their differentiation into hepatocytes [4]. Interestingly, the matrix-remodeling enzymes are also expressed during embryonic liver development and play an important role in the liver lobule formation. MMP-1 was found from the sixth gestational week (GW) of human embryonic development onward in hepatocytes, and later also in outer limiting plate hepatocytes, early bile ducts, and periportal mesenchymal cells. In the sixth GW, MMP-2 was found only in the microvascular endothelium. In the seventh GW, MMP-2 was also detected in hepatocytes. From the ninth GW onward, MMP-2 was detectable in all hepatocytes and erythropoietic, endothelial, and

Dense, interstitial ECM is largely confined to the portal area. Basement membranes underlie the vascular structures, but not the bile canaculi or small bile ducts. The interstitial ECM contains more of the fibrillar collagens and associated molecules, and less of the basement membrane components Synthesis of provisional matrix early after induction of a liver injury Up to 10-fold increase of ECM that comprises several types of collagen, structural glycoproteins, sulfated proteoglycans (glycosaminoglycans), and hyaluronan

periportal mesenchymal cells. MMP-7 was present in the sixth GW in some hepatocytes and endothelial cells, but from the seventh GW onward, only in hematopoietic cells. MMP-13 was found exclusively in hematopoietic cells. These results indicate that the synthesis of MMP-1, MMP-2, MMP-7, and MMP-13 during human liver development occurs from the sixth GW [5]. Recently a regulatory insight was given into the gene regulation of MMP-2 in hepatoblasts [5]. MMP-2 was upregulated in prospero-like homeobox (Prox)1-transfected met-murine hepatocytes (MMH), which represent hepatoblastic cells expressing hepatocyte and bile duct markers. These cells originate from a mouse embryo at the 14th gestational day. MMH cells do not express Prox1, which is a transcription factor with two highly conserved domains, a homeobox, and a prospero domain [6]. Artificial (ectopic) expression of Prox1 in these cells after transfection induced approximately 26-fold upregulation in the microarrays and 16-fold upregulation in quantitative reverse-transcription polymerase chain reaction (qRT-PCR) of MMP-2. As further evidence, zymography showed that MMP-2 was secreted by Prox1-transfected MMH cells, and it was functionally active [6]. As a comparison, in mouse embryos, Prox1-positive hepatoblasts expressed MMP-2. Those hepatoblasts located in the periphery of the liver, if interacting with the mesenchyme, had the highest MMP-2 expression [6]. MMP-2 expression in human hepatoblasts was mentioned before. Additionally, the transverse septum also displayed high levels of MMP-2. Prox1 is already expressed in endodermal cells well before the formation of the liver anlage [7]. Prox1 remains a hepatocyte differentiation marker in adult hepatocytes, and also after liver injury or during liver regeneration [7]. The new experimental proof that Prox1 induces MMP-2 expression in hepatoblasts might be of major importance for early liver morphogenesis.

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Interestingly, Prox1 upregulates not only MMP-2 but also type IV α3 procollagen. This places Prox1 in the center of ECM turnover, making it important for both liver development and maintenance [6].

ECM OF THE NORMAL ADULT LIVER In the normal liver the ECM has a positive function. It facilitates the exchange of fluids and their flow through the tissue, and enables elasticity and rigidity at the same time. The normal adult liver is soft and elastic (Figure 30.1a). Nevertheless, it provides the required resistance for the tissue. These functions (softness, elasticity, and resistance) are provided by the dynamic interaction between liver cells and matrix. In adult livers, the matrix chemistry is dominated by FN, a high ratio of type I to type III collagen, highly sulfated proteoglycans, and a low representation of laminin (Figure 30.2a), hyaluronans (HAs), and type IV collagen [8]. In the normal liver, ECM components are detected in the subendothelial space of Disse, in the portal area, at the wall of the central veins, and in the liver capsule (Figures 30.1b and 30.2a). The subendothelial space of Disse separates the epithelium (hepatocytes) from the sinusoidal endothelium. In normal liver this space contains both an interstitial and a basement membrane-like ECM of low density. This perisinusoidal matrix is composed of fibrillar collagens type I, III, and V, microfibrillar collagen VI, basement membrane collagens IV and XVIII, and traces of collagen XIV and of the small proteoglycan decorin, FN, tenascin-C, laminin, nidogen, heparan sulfate proteoglycans (HSPGs), and others (Table 30.1). This low-density ECM is critical for maintaining the differentiated functions of resident liver cells (i.e. hepatocytes, stellate cells, sinusoidal endothelium, and Kupffer cells; Figures 30.1b, 30.2a, and 30.3). In contrast, the dense, interstitial ECM is largely confined to the portal area, the central veins, and the liver

(a)

455

capsule (Table 30.1) [2]. Basement membranes underlie the vascular structures, but not the bile canaculi or small bile ducts (Figure 30.2a). The larger bile ducts in the region of the hepatic hilus are underlined by basal membrane [9]. The interstitial ECM contains more of the fibrillar collagens and associated molecules and less of the basement membrane components. The ECM components are distributed heterogeneously in the liver acinus. At the periportal zone more laminin, collagen type III and IV, chondroitin sulfate proteoglycan, and HA can be found, while in the pericentral zone more FN, type I collagen, heparin proteoglycan, and dermatan sulfate proteoglycan can be recognized [8].

ECM DURING LIVER INFLAMMATION The ECM remodeling is a quick response to liver damage, which precedes and later parallels the reparative progress, and is part of tissue healing (Figure 30.3a–d). In short-term liver damage a “clotting-like process” occurs, containing fibrin/FN, fibrinogen-like protein (FGL) 1, von Willebrand factor, and gelsolin, constituting a “provisional matrix” which affects the attraction and proliferation of inflammatory and matrix-producing cells [10]. In this relation, there seems to be a putative pacemaker role of FN. It is deposited soon after liver injury in necrotic areas (Table 30.1). In contrast to normal liver, mononuclear phagocytes of acutely injured livers synthesize and secrete FN in abundant amounts [10]. Fibrinogen (FBG) is closely linked to FN during clotting processes; FBG transcripts are localized in cells of the non-necrotic areas. In vitro studies showed FBG de novo synthesis restricted to hepatocytes. Hepatic FBG is upregulated during an acute phase response (APR), and might be induced by glucocorticoids and interleukin (IL)-6 [11]. FBG is a principal factor in the maintenance of hemostasis, and during disruption of homeostasis FBG is upregulated as part of the systemic APR [11].

(b)

Figure 30.1 (a) Laparoscopic image of the normal liver showing the elasticity of the liver touched by the biopsy-needle. (b) Goldner trichrome staining: detection of collagen fibers in the periportal area of the normal human liver. The liver is soft but resistant at the same time

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(a)

(b)

Figure 30.2 Human liver sections stained with an antibody against laminin. (a) Normal human liver and (b) in human liver cirrhosis. In normal liver (a) basement membranes underlie the vascular structures and the liver capsule but not the bile canaculi or small bile ducts. In normal liver the subendothelial space contains both an interstitial and a basement membrane-like ECM of low density. In human liver cirrhosis (b) induction of subendothelial laminin leads to the formation of an incomplete subendothelial basement membrane, creating additional diffusion barriers between hepatocytes and the liver sinusoid at the border of the fibrotic septum (capillarization of sinusoids). Contrastaining of nuclei with DAPI

After short-term liver damage, the synthesis of provisional matrix components may be related to the APR or to the induction of an inflammatory process (Table 30.1, Figure 30.3c, d). For example, FGL1 is a hepatocyte-derived protein that is upregulated in regenerating rodent livers following partial hepatectomy (PH). Following induction of acute inflammation in rats by subcutaneous injection of turpentine oil, serum FGL1 levels are also enhanced. The enhancement of FGL1 levels in vitro by IL-6 and its induction after turpentine oil injection suggest that it is an acute-phase reactant [12]. A further component of the provisional matrix, gelsolin, a 90 kDa protein, was suggested to be involved in cell motility, to inhibit apoptosis, and to have a protective role for tissue. In human fulminant hepatic failure, positivity in the necrotic areas was detected. Gelsolin has an important protecting effect in clinical situations where actin is released from injured cells, as happens under conditions of the crash syndrome, shock, and fulminant hepatic failure. Under these conditions, a decreased gelsolin plasma level has been detected. An enhanced expression of gelsolin under conditions of liver damage may exert a protective role; the inflammatory cells and the hepatocytes are responsible for the enhanced gelsolin expression. This finding was confirmed in the rat liver injury model of CCl4 treatment, where either a direct action of CCl4 on liver cells, cytokines produced in the CCl4 -treated liver, or recruited inflammatory mononuclear phagocytes may be responsible for the induction of the “defense protein” gelsolin in acute and chronic human and rat liver injury [13]. The findings related to the synthesis of provisional matrix early after induction of a liver injury reveal that

matrix remodeling is strongly related to active inflammation (Figure 30.3c, d). The ECM may also be directly involved in the action of the damaging agents. For example, recently a new leptospiral protein that exhibits ECM-binding properties was described: Lsa21 (leptospiral surface adhesin, 21 kDa). Attachment of Lsa21 to laminin, collagen IV, and plasma FN was found to be specific and dose-dependent. A binding of this leptospiral protein in the liver was reported [14]. As a further step, the liver matrix is also involved in the recruitment of inflammatory cells such as neutrophil granulocytes. In fact, matrix components such as gelsolin may also be synthesized by inflammatory cells. As a further example, HA is disproportionately expressed in the liver under both basal and inflammatory conditions. Spinning-disk intravital microscopy revealed that constitutive HA expression was restricted to liver sinusoids. Experimental blocking of CD44–HA interactions reduced neutrophil adhesion in the sinusoids of endotoxemic mice, with no effect on rolling or adhesion in postsinusoidal venules. Neutrophil but not endothelial CD44 was required for adhesion in sinusoids, yet neutrophil CD44 avidity for HA did not increase significantly in endotoxemia. Instead, activation of CD44–HA engagement via qualitative modification of HA was demonstrated by a dramatic induction of serum-derived HA-associated protein in sinusoids in response to lipopolysaccharide (LPS). LPS-induced hepatic injury was significantly reduced by blocking CD44–HA interactions. Administration of anti-CD44 antibody four hours after LPS rapidly detached adherent neutrophils in sinusoids and improved sinusoidal perfusion in endotoxemic mice [15].

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(a)

(b)

(c)

(d)

Figure 30.3 (a,b) Goldner trichrome stainings of human liver in a case of subacute liver failure. The images show histological distribution of fibrillar collagen around the intact hepatocytes, which will restore the normal parenchyma after recovery. The ECM keeps the tissue together and accompanies regeneration. (c,d) Inflammatory infiltrate in a case of chronic HCV in a transplanted liver. Connective tissue fibers underlie the inflammatory cells which occupy space previously occupied by normal hepatocytes

As a further example, recent data reveal that osteopontin (OPN) has an important role in cell signaling that controls inflammation. OPN is a highly-modified integrin-binding ECM glycophosphoprotein produced by cells of the immune system, epithelial tissue, smooth muscle cells, osteoblasts, and tumor cells. It interacts with the integrin receptors expressed on inflammatory cells through its arginine-glycine-aspartate (RGD) and non-RGD motifs, by which it promotes migration and adhesion of cells. In the liver, it has been reported that hepatic Kupffer cells secrete OPN, by which they facilitate macrophage infiltration into necrotic areas following carbon tetrachloride liver toxicity [16]. In addition to full-length OPN, the thrombin-cleaved osteopontin (cOPN) product was also present in the liver of mice following Concanavalin A (Con A) injection. Infiltrating leukocytes purified from the liver after Con A injection migrated toward thrombin cOPN more efficiently than to the full-length form of OPN. Neutrophils were the predominant leukocyte population that responded to the cleaved OPN. After Con A-induced activation, natural killer T-cells (NKT-cells) secrete OPN, which is cleaved by thrombin in the liver. The interaction of NKT-cells and thrombin cOPN through its receptors further activates

the NKT-cells, which likely produces macrophage inflammatory protein (MIP) 2, a known chemotactic factor for neutrophils. Thrombin cOPN also interacts with its receptors, α9 and α4 integrins, on neutrophils so that the neutrophils become activated and contribute to additional liver damage [17]. Further data indicate that OPN is chemotactic for macrophages; in addition, activated macrophages themselves produce abundant OPN. In the liver, expression of OPN was highest in Kupffer cells, macrophages, and HSCs. Because OPN stimulates cell migration, it may act as a chemotactic factor in the recruitment of macrophages to sites of liver injury. Higher neutrophilic inflammation, necrosis, and liver injury correlated well with the levels of both intrahepatic and circulating levels of OPN [18] in male Sprague–Dawley rats fed EtOH-containing Lieber–DeCarli liquid diet for six weeks, followed by LPS injection. OPN has been identified as an important cytokine whose expression is increased early in the course of the disease in an experimental dietary rodent model of non-alcoholic steatohepatitis (NASH) [16]. Transgenic mice were created that express OPN in hepatocytes under the control of the serum amyloid

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P-component promoter [19]. These mice developed mononuclear cell infiltration starting at 12 weeks of age, and becoming more extensive with aging. This infiltrate was primarily cytotoxic T-cells. When these mice were treated with CCl4 , the extent of necrosis and cellular infiltration was reduced compared with control mice, suggesting that OPN overexpression in hepatocytes may play a protective role [20]. Studies using OPN-deficient mice have further clarified the role of OPN in liver injury. Granuloma formation in the liver was induced by β-glucan administration to both wild-type and OPN-deficient mice [21]. In wild-type mice, β-glucan administration induced OPN expression in the liver at both the RNA and protein levels; the kinetics of the changes in OPN expression paralleled those of granuloma formation, and OPN expression was detected within the granulomas. OPN−/− mice had significantly reduced granuloma numbers and size two weeks after β-glucan administration. Although there was no difference in the histological appearance of the granulomas between the two genotypes, there was a twofold reduction in macrophage, CD4+ T-cell, and dendritic cell accumulation in the livers of OPN−/− mice as compared with wild-type animals. Liver damage, as assessed by alanin-aminotransferase (ALT) and aspartat-aminotransferase (AST) in serum, was twofold worse in OPN-deficient mice than in wild-type controls, measured one day after CCL4 treatment, and the area of necrosis was increased two and three days after treatment.

ECM OF THE LIVER FIBROSIS The finely-tuned ecosystem of liver ECM is pushed toward increased deposition when repeated damage followed by tissue loss takes place and regeneration is not possible (Figure 30.3a,b). This process has to be considered as part of the defense apparatus of the organ. In fact, it has the task of maintaining the organ function (Figure 30.3a,b). Furthermore, matrix synthesis aims at defensive action by producing anti-apoptotic proteins such as gelsolin. The key cell in this defense function is the myofibroblast (Myb), which when activated serves as the primary collagen-producing cell. Mybs are generated from a variety of sources. The processes of Myb activation in the liver are discussed in other chapters of this book. Fibrosis of the liver is now characterized by an up to 10-fold increase of ECM that comprises several types of collagens, structural glycoproteins, sulfated proteoglycans (glycosaminoglycans), and HA; by a histological redistribution with preferred initial matrix deposition in the perivenular zone 3 of the acinus along the subendothelial space of Disse leading to the formation of an incomplete subendothelial basement membrane creating additional diffusion barriers between hepatocytes and the liver sinusoid (capillarization of sinusoids [22] (Table 30.1); by changes to the ECM profile; and by changes to the

fine structure of collagens (e.g. degree of hydroxylation of proline and lysine), glycoproteins (variations of the carbohydrate structure), and proteoglycans (Figure 30.3c,d). Over time, the subendothelial matrix composition changes from one comprised of type IV collagen, HSPG, and laminin (the classic constituents of a basal lamina) to one rich in fibril-forming collagens, particularly types I and III ([23]; Figure 30.3b). These progressive changes in ECM composition as fibrosis accumulates instigate several positive-feedback pathways, which further amplify fibrosis. First, dynamic changes in membrane receptors, in particular integrins, sense altered matrix signals that provoke stellate cell activation and migration through focal adhesion disassembly [24]. Matrix-provoked signals also engage membrane-bound guanosine triphosphate (GTP)-binding proteins, in particular Rho67 and Rac68, which transduce signals to the actin cytoskeleton that promote migration and contraction. Second, activation of cellular matrix metalloproteases leads to release of growth factors from matrix-bound reservoirs in the extracellular space that may stimulate cellular growth and fibrogenesis [2]. Third, the enhanced density of ECM leads to increasing matrix stiffness, which is a significant stimulus to stellate cell activation, at least in part through integrin signaling [25]. It is important to claim that the ECM deposition is a substantial part of the healing process, which occurs in coordination with the hepatocellular regeneration (Figure 30.3a–d). The synthesis of ECM facilitates the abrogation of tissue damage, and provides positive induction for repair (regeneration) processes (Figure 30.3a,b).

NEWLY DESCRIBED COMPONENTS OF THE LIVER ECM Recently, novel ECM components were described as hepatocyte-derived FBG-related-protein-1 (HFREP-1 or hepassocin), agrin [12, 26, 27], matrilin-2 [28, 29], reelin [30, 31], and OPN [16]; new findings are also available regarding fibulins. Hepassocin, a new member of the fibrin matrix of a plasma clot, was described recently, analyzed by 2D gel electrophoresis. A relatively abundant spot was identified as HFREP-1. HFREP-1 in plasma almost completely bound to the fibrin matrix during clot formation. Several purified FBG preparations proved to be contaminated with HFREP-1. It is concluded that HFREP-1, a protein with liver cell growth regulatory properties, occurs in plasma and strongly associates with fibrin and possibly FBG [32]. Also recently, reelin was suggested to be not only a component of the liver ECM, but a plasma protein as well. Reelin is an ECM protein secreted by a variety of

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cell types in both embryonic and adult tissues, including the liver. The HSCs are the major source of reelin in the injured liver [30]. Increased liver reelin in bile duct-ligated rats leads to a pronounced 165% increase in the plasma levels. The data provide evidence that a fraction of plasma reelin is synthesized in the liver. In human subjects suffering liver cirrhosis, the level of the 180 kDa reelin fragment was also increased by 140% in the plasma. Reelin fragments in the plasma of cirrhotic patients are differentially glycosylated compared to non-diseased control subjects [31]. OPN is a newly-discovered matrix constituent, which might be included in the provisional matrix, and later also in the ECM of the fibrotic liver. OPN was discussed in detail before [16]. Fibulins belong to an emerging family of ECM proteins characterized by tandem arrays of calcium binding epidermal growth factor (EGF)-like domains and a unique C-terminal structure [33]. To date, fibulin-1 and -2 are characterized in most detail [33] and have been shown to display distinct yet overlapping molecular interactions and expression patterns, whereas fibulin-3–6 are less defined [34]. Fibulin-1 and -2 are localized in basement membranes, elastic fibers, and other connective tissue structures. Fibulin-2-positive MFs have been detected in the portal field, vessel walls, and hepatic parenchymas of the normal liver, and their number was increased in the septal regions during liver fibrogenesis [35]. Fibroblasts are regarded as the typical cellular source for biosynthesis, in which fibulin-2 is deposited with FN into a fibrillar matrix [36], but little is known concerning the hepatic source of fibulin-1. After liver damage both, fibulin-1 and -2 were found to participate in hepatic tissue repair, but interestingly each showed a distinct individual expression. Following an acute liver damage comprising tissue necrosis, inflammation, and regenerative events, fibulin-1 and -2 showed different expression patterns. Fibulin-1-specific mRNAs increased early after CCl4 administration in the rat, and peaked at 48 hours. Interestingly, similar changes were not observed for fibulin-2, whose specific mRNA levels and immunohistochemical expression remained unmodified during the entire kinetics of acute tissue damage and repair, up to the 96th hour. In particular, the participation of hepatocytes in the synthesis of fibulin-1, which is also a circulating plasma protein, should be considered. Moreover, fibulin-1 may be involved in the provisional matrix formation, similarly to FN. In addition, fully activated HSCs significantly contribute to the fibulin-1 synthesis in the liver. At the same time, activated smooth muscle actin (SMA)-positive HSCs of the necrotic pericentral area remain negative for fibulin-2 after an acute damage, whereas other molecules selectively expressed by activated HSC, such as neural cell adhesion molecule and glial fibrillary acidic protein, are reported to appear at immunohistochemistry in the same condition [37].

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The relative increases of the two fibulin-1 variants C and D were not identical during acute and chronic hepatic tissue repair. After an acute liver injury the increase of the two isoforms was rather similar, whereas in chronic liver damage a prevalence of the isoform D was manifest. The two variants differ for their binding affinity to FN and laminin and, most of all, to nidogen [38], and their ratio may therefore be a mechanism contributing to the fate of hepatic tissue repair. Recently, functions of further fibulins were reported. The expression of fibulin-5 and fibrillin-1 is essential for elastogenesis. Immunohistochemical labeling of fibulin-5 was observed in the major portal vein branches and the distribution corresponded to that of elastic fibers in the vessel walls, while the peripheral portal tracts totally lacked fibulin-5 in spite of the presence of dense elastic fibers [39]. Fibrillin-1 was detected in the connective tissue of the hilar region and peripheral portal tracts in idiopathic portal hypertension, chronic viral hepatitis, liver cirrhosis, and normal livers, with the expression varying greatly among cases. Fibulin-5, rather than fibrillin-1, was expressed in the major portal vein branches of idiopathic portal hypertension livers related to phlebosclerosis, leading to an increase in presinusoidal vascular resistance and portal hypertension [39]. Agrin is a multidomain HSPG with different modules homologous to domains found in basal membrane proteins. Its functions include binding of various growth factors and cytokines in the extracellular milieu outside the basement membranes. Agrin may play an important role in cellular growth, differentiation, adhesion, and motility in pathophysiological conditions such as tumor cell growth, invasion, and metastasis. Only small amounts of agrin were detected in the normal liver. The presence of agrin in normal and cirrhotic liver as well as in hepatocellular carcinoma (HCC) has recently been reported by T´atrai et al. [26]. Agrin may play a role in the vascular and ductular proliferation characteristic of cirrhosis and may promote tumor progression by supporting stromal cell growth and neoangiogenesis in HCC. Agrin, like other HSPGs, is able to bind a variety of growth factors, such as basic fibroblast growth factor (bFGF), vascular endothelial growth factor (VEGF), and hepatocyte growth factor (HGF). The presence of agrin in the basement membrane enables a local enrichment in these factors [27]. Agrin may also contribute to the presentation of these growth factors to their receptors and thereby promote the corresponding signaling cascades, eventually resulting in downstream effects such as tumor cell survival, proliferation, and migration. These functions also interfere with the growth of hepatic tumors as cholangiocellular carcinoma (CCC). As long as CCC cells possess an intact basement membrane, as observable in well-differentiated tumors, they may derive benefit from the growth factor-presenting property of agrin, using it as a supplier of survival ligands.

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Similarly, the recently described matrilin protein family is part of the ECM. Matrilins have been found to play roles in cell growth and tissue remodeling, and their possible involvement in carcinogenesis has been raised. Matrilin-2 protein expression was detected in normal liver around bile ducts and portal blood vessels, while sinusoids were negative by immunohistochemistry. Cirrhotic surrounding tissue showed intensive matrilin-2 staining along the sinusoids. Tumorous neovasculature was found strongly positive by immunohistochemistry. These data indicate that matrilin-2 is a novel basement membrane component in the liver that is synthesized during sinusoidal “capillarization” in cirrhosis and in HCC [28, 29]. During establishment of liver fibrosis, the number of endothelial fenestrae of the involved sinusoids initially decreases. Then the basement membrane components collagen type IV, FN, and laminin increase and form a true basement membrane-like structure within the space of Disse. These changes of the sinusoids are called “capillarization” because the altered structure of the sinusoids resembles that of capillaries [40]. In the acetyl-aminofluorene-partial hepatectomy (AAF/PH) model of the rat, where a sequential intragastric AAF treatment is combined with PH, the oval cells but not the hepatocytes produced matrilin-2 mRNA. Increase in protein level in the AAF/PH regenerating liver model was demonstrated [29]. The fibrosis process is not only associated with qualitative changes highlighted by the novel expression of ECM components, but also with quantitative changes. There are efforts to use the deposition of ECM components as prognostic markers. Some of the new contributions to this aim are discussed below. Serological markers have been proposed for monitoring hepatic fibrosis in chronic liver disease. Among fibrosis markers, type III procollagen (PIIIP) and hyaluronic acid have been studied in these patients. A prospective cross-sectional study was carried out with hepatitis C virus (HCV)-positive blood donors. Liver function tests were not associated with tissular fibrosis, elevated PIIIP was correlated with 66.7% chance of fibrosis, and hyaluronic acid, when elevated, gave a 33.3% chance of fibrosis. Authors suggest that PIIIP should be useful to assess fibrosis in patients with chronic HCV [41]. In contrast, a further study analyzing 72 patients with histologically verified NASH revealed that type IV collagen 7s domain, but not hyaluronic acid, was significantly elevated in patients with advanced fibrosis after adjustment for age, sex, platelet count, prothrombin time, aspartate aminotransferase/alanine aminotransferase ratio, body mass index, and presence of underlying type 2 diabetes mellitus, all of which have previously been reported as useful predictors of advanced fibrosis in patients with NASH [42]. These studies showed that the use of ECM components as serological markers for monitoring hepatic fibrosis requires correct statistical analysis and may be different in NASH and in chronic HCV.

MATRIX REMODELING The metabolism of ECM is based on the balance between activities of MMPs and those of tissue inhibitor metalloproteinases (TIMPs) [43]. MMPs other than MMP-1, MMP-8, and MMP-13 cannot degrade type I collagen, which is very stable, and a net deposition of type I collagen has been observed in progressive liver fibrosis [44]. MMP and TIMP coding transcripts were detectable in all liver cell types by reverse transcription-polymerase chain reaction; however, the cellular expression levels were markedly different as assessed by Northern blot analysis. Gelatinase-B was predominantly expressed in Kupffer cells, gelatinase-A in HSCs and rat liver Mybs, and stromelysins-1 and -2 as well as collagenase in HSCs. Membrane type-1 MMP (MMP-14) was found in significant amounts in all liver cells. TIMP-1-coding mRNAs were present mainly in HSCs and rat liver Mybs, TIMP-2 additionally in Kupffer cells, while TIMP-3 expression was detectable only in hepatocytes. During in vitro activation of HSCs, MMP expression was mostly downregulated, while TIMP expression was enhanced, thereby providing an explanation for matrix accumulation co-localized with these cells during chronic liver injury. In general, TNFα stimulated both MMP and TIMP expression of HSCs, while TGFβ1 induced TIMP expression only [45]. Following a single toxic liver injury, MMPs and TIMPs were induced in a distinct time frame, in that expression of most MMPs was induced during the early phase of liver injury, was maximal during the inflammatory reaction, and was diminished in the recovery phase. In contrast, TIMP and MMP-2 steady-state mRNA levels remained constant in the early phase, were strongly induced during tissue inflammation, and remained increased until the recovery phase. Interestingly, hepatic TNFα expression paralleled the MMP induction profile, while the increase of TGFβ1 expression mapped to the increase of TIMPs. Chronic liver injury was accompanied by an increase in the steady-state mRNA levels of MMP-2 and TIMPs, while other MMPs remained more or less unchanged or were diminished. PH was followed by a dramatic increase of MMP-14 and to a lesser extent also of TIMP-1 expression; other MMPs and TIMPs were not significantly induced. Liver injury is accompanied by profound changes in hepatic MMP/TIMP expression, the latter being critically dependent on the type of injury [46]. In recent studies, this knowledge was extended by novel findings of an unequivocal role for scar-associated macrophages (SAMs) in the spontaneous resolution of liver fibrosis. It was demonstrated that SAMs may be the source of MMP-13 (collagenase 3), which is considered to be the primary interstitial collagenase in rodents. mmp13 gene expression was restricted to regions of fibrosis that were rich in SAMs. Both MMP-13 mRNA and protein co-localized to large phagocytes within and directly apposed to hepatic scars [47].

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A further source of MMP-expressing cells involved in matrix remodeling is the bone marrow (BM). Mice were treated with repeated carbon tetrachloride injections after hematopoietic reconstitution with enhanced green fluorescent protein (EGFP)-expressing BM cells. A large number of EGFP+ cells were observed in liver tissue at peak fibrosis, which decreased during the recovery from liver fibrosis. Some of them, as well as EGFP− liver resident cells, expressed MMP-13 and MMP-9. Whereas MMP-13 was transiently expressed mainly in the cells clustering in the periportal areas, MMP-9 expression and enzymatic activity were detected over the resolution process in several different kinds of cell located in the portal areas and along the fibrous septa. Autologous BM cells contribute to the spontaneous regression of liver fibrosis, and their therapeutic derivation could be a new treatment strategy for intractable liver fibrosis [48]. As a further source of matrix remodeling enzymes, the hepatocyte itself was suggested. Authors clearly detected the mRNAs of four MMPs (MMP-2, MMP-3, MMP-10, and MMP-13) and of two TIMPs (TIMP-1 and TIMP-2) in hepatocytes from both normal and cirrhotic rats. They also demonstrated MMP-2, MMP-3, and MMP-13, and TIMP-1 and TIMP-2 proteins in the same hepatocyte preparations by immunostaining [49]. Delivery of interstitial collagenases such as MMP-1 in the liver could be an attractive strategy to treat advanced hepatic fibrosis, from the point of view that the imbalance between too few interstitial collagenases and too many of their inhibitors is the main obstacle to resolution of fibrosis. Remodeling of hepatic ECM by delivered interstitial collagenases also facilitates the disappearance of activated HSCs and promotes the proliferation of hepatocytes [50]. Relaxin is a recently-described, naturally-occurring matrix-remodeling enzyme which may have a therapeutic potential. Since its discovery as a reproductive hormone 80 years ago, relaxin has been implicated in a number of pregnancy-related functions involving ECM turnover and collagen degradation. It is now becoming evident that relaxin’s ability to reduce matrix synthesis and increase ECM degradation has important implications in several non-reproductive organs, including the heart, lung, kidney, liver, and skin. The identification of relaxin and RXFP1 (relaxin family peptide receptor-1) mRNA and/or binding sites in cells or vessels of these non-reproductive tissues has confirmed them as targets for relaxin binding and activity. Recent studies on Rln1 and Rxfp1 gene-knockout mice have established relaxin as an important naturally-occurring and protective moderator of collagen turnover, leading to improved organ structure and function. Furthermore, through its ability to regulate the ECM and in particular collagen at multiple levels, relaxin has emerged as a potent antifibrotic therapy, with rapid-occurring efficacy. It not only prevents fibrogenesis, but also reduces established scarring (fibrosis), which is a

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leading cause of organ failure and affects several tissues regardless of etiology [51].

MATURATION OF LIVER ECM, MATRIX STIFFNESS Mechanical forces may be particularly important in the liver, where a number of common pathological conditions including fibrosis and non-alcoholic fatty liver disease result in significant mechanical changes at whole-organ, regional, and cellular levels. Cells within tissues contact and sense the mechanical stiffness of both the ECM and other cells. The stiffness of a matrix is determined by both its chemical makeup and its organization. The combination of various collagens with other matrix proteins and proteoglycans typically seen in tissues would be expected to alter mechanical stiffness. Cell-directed organization of collagen as well as FN matrices changes the mechanical properties of the ECM and may contribute significantly to stiffness when compared with the passive mechanical properties of the matrix alone [25, 52]. Increases in liver stiffness precede fibrosis and potentially Myb activation. It may also be the consequence of the deposition of clot-forming provisional matrix containing FBG, hepassocin, and FN. Liver stiffness appears to result from matrix cross-linking and possibly other unknown variables in addition to matrix quantity. Increased liver stiffness may play an important role in initiating the early stages of fibrosis [53]. Post-translational modifications of ECM components also regulate stiffness. For collagen, these changes can include non-enzymatic glycation as well as cross-linking by enzymes such as tissue transglutaminase, the lysyl oxidases, and the lysyl hydroxylases [25, 52–54]. All three families of cross-linking enzymes contribute to collagen cross-linking in liver cirrhosis and may thereby alter the stiffness of the cirrhotic liver out of proportion to increases in the quantity of collagen. The lysyl oxidases are upregulated early in liver injury. Lysyl oxidase staining increased after bile-duct ligation, when it was located in areas surrounding the Myb cells. After bile-duct ligation, ECM deposition and lysyl oxidase expression occur very early in portal connective tissue surrounding proliferating ductules, and precede activation of Mybs; that is, SMA expression. In addition, these data are compatible with the suggestion that in the bile-duct ligation model, Myb differentiation represents an adaptive response to modification of the ECM environment [54]. A further recently-described matrix-modifying enzyme is ADAMTS2. ADAMTS2 belongs to the “ADAM metallopeptidase with thrombospondin type 1 motif” (ADAMTS) family. Its primary function is to process collagen type I, II, III, and V precursors into mature molecules by excising the aminopropeptide. This process allows the correct assembly of collagen molecules

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into fibrils and fibers, which confers to connective tissues their architectural structure and mechanical resistance. The extent of fibrosis is reduced in ADAMTS2-deficient (TS2−/−) mice in comparison with their wild-type littermates [55]. In addition to the above-mentioned maturation modification of the liver ECM, mainly by collagen cross-linking, proteoglycans and hyaluronic acid also regulate matrix hydration and therefore resistance to compression. Proteoglycan expression increases dramatically during liver fibrosis and could, like collagen cross-linking, increase liver stiffness even in the absence of increased collagen [56]. The altered mechanical properties of the ECM affect the cells getting into contact with it. Integrins, the heterodimeric receptors for the ECM, are the primary cellular mechanosensors for adhesion-dependent mechanical forces. There is a push–pull relationship between the matrix, which exerts force on the cell via integrins, and the cellular cytoskeleton, which in turn resists this force. Focal adhesions, the multiprotein complexes linking integrins to the actin cytoskeleton, are central to the process. Rho GTPases are crucial downstream mediators, as is the contractile actin-myosin cytoskeleton [57, 58]. Interestingly, endogenous or heterologous Thy-1 expression was found to promote focal adhesion and stress fiber formation, characteristic of increased Rho GTPase activity, and to inhibit migration. Thy-1 expression resulted in a net increase in active Rho, and increased stress fibers and focal adhesions [59, 60]. Recently, Thy-1 expression was demonstrated in liver Mybs, and the lack of Thy-1 expression was proved in HSCs [61–64]. These data indicate a crucial difference in mechanosensing and stress management of HSCs and liver Mybs. HSCs cultured on soft matrices mechanically similar to normal liver remain quiescent, whereas cells cultured on stiff matrices similar to cirrhotic liver undergo activation manifested by induction of SMA expression. Portal fibroblasts in vitro adopt a fibrogenic phenotype only when cultured in a mechanically stiff environment similar to a cirrhotic liver. Portal fibroblasts, increasingly recognized to be key mediators of biliary fibrosis, require both the soluble factor transforming growth factor-beta and a stiff matrix for differentiation in culture. Similarly, HSCs lose their lipid droplets and start to express SMA (characteristic of Mybs) as the stiffness of their underlying support increases [25]. Sinusoidal endothelial cells maintain their fenestrations when cultured on human amnion basement membrane but not on Matrigel or individual matrix components. The role of mechanical stiffness in the function of other non-parenchymal cells of the liver, including Kupffer cells, dendritic cells, and other cells of the immune system, has not been investigated; similarly, the role of mechanics in regulating the differentiation of BM-derived stem cells is unknown [25].

ECM REMODELING HAS A MAJOR IMPACT ON LIVER REGENERATION In contrast to the fact that hepatic fibrosis is characterized by excessive deposition of ECM, altered liver architecture, and impaired hepatocyte proliferation, the fibrotic liver can still regenerate after PH. Liver collagen content was reduced to 75% after PH in cirrhotic rats when compared with sham-operated cirrhotic rats. The regenerating fibrotic liver oxidized actively free proline and had diminished transcripts for alpha-1 (I) collagen mRNA, resulting in decreased collagen synthesis. PH also increased collagenase activity, accounted for by higher amounts of pro-MMP-9, MMP-2, and MMP-13, which largely coincided with a lower expression of TIMP-1 and TIMP-2. Therefore, an early decreased collagen synthesis, mild ECM degradation, and active liver regeneration were followed by higher collagenolysis and limited deposition of ECM, probably associated with an increased mitochondrial activity. Stimulation of liver regeneration through PH contributes to restore the balance in ECM synthesis and degradation, leading to ECM remodelling, and it may reach an almost complete resolution of liver fibrosis [65]. These data also support the central role of hepatocytes in the regulation of liver matrix synthesis and remodeling, not only during liver lobule development, but also during normal adult liver function and in liver injury. Alternatively, in advanced cirrhosis, hepatic regeneration can also occur via an intrinsic facultative stem cell compartment located within the terminal branches of the biliary tree. Hepatic progenitor cells (called “oval cells” in rat and mice) are found in close anatomical approximation to Mybs and macrophages, within their stem cell niche. Stem cell fate may be determined by constituents of the ECM, such as collagen I and laminin, as well as by growth factors secreted by the supporting mesenchymal cells. The r/r collagen transgenic mouse expresses mutated collagen I resistant to collagenase degradation, has an exaggerated fibrotic phenotype to CCl4 liver injury, and shows impairment of collagenolysis post-injury. Chronic liver injury and liver fibrosis were induced by eight weeks of repeated intraperitoneal CCl4 administration in r/r mice and wild-type C57B6 controls. As expected, r/r mice developed significantly more liver fibrosis than wild-type controls, and showed persistence of collagen into recovery. Surprisingly, r/r mice had a markedly attenuated oval cell response throughout the recovery phase. There was no significant difference in parenchymal regeneration via proliferation of mature hepatocytes. Persistence of collagen I and a paucity of laminin deposition after hepatic injury are associated with an inability to mount an oval cell response. Matrix remodeling appears critical

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to parenchymal regeneration via hepatic stem cells, and laminin–progenitor cell interactions may be an important feature of this [66].

IMPACT OF MATRIX ON HEPATOCYTES IN CULTURE Accumulating evidence demonstrates the differential effect of matrix components on cultured hepatocytes. Basal-level enzyme activities and enzyme inducibility were compared for rat hepatocytes that were cultured with three different ECM configurations: single-layer (SL) collagen type I, SL Matrigel, and collagen/Matrigel (C/M) sandwich. Overall, C/M sandwich and SL Matrigel plates were both superior to SL collagen type I plates in maintaining enzyme activities and inducibility, and C/M sandwich plates had higher induced activity for CYP3A enzymes than SL Matrigel plates [67]. When isolated mature hepatocytes are cultured on type I collagen-coated dishes, the cells appear as a flattened monolayer and express low levels of liver function-specific mRNA and proteins. In dramatic contrast, when hepatocytes are cultured on a model basement membrane Engelbreth–Holm–Swarm (EHS) gel, hepatocytes retain their normal polarity and structure, and the products of liver-specific genes continue to be secreted for prolonged periods of culture [68, 69]. Cell–matrix interaction influences the determination of the differentiated phenotype of hepatocytes in cell culture, and maintains liver-specific functions for long-term culture, which effects have been associated with upregulation of liver-enriched transcription factors, including hepatocyte nuclear factor (HNF). Upregulation of liver-specific genes induced by ECM is mediated via upregulation of HNF-4α and HNF-1 induced by ECM. A collagen gel matrix increased the levels of HNF-3α in the hepatocyte-derived cell line H2.35, but not those of HNF-3β and -3γ, responsible for the transcription of liver-specific genes. ECM regulates HNF-4 and tissue-specific gene expression in fetal hepatocytes as well as adult hepatocytes [70, 71]. The ECM may influence hepatocyte differentiation via the assembly of endocytic components. A remarkable level of phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2 ) phosphatase mRNA production was observed in hepatocytes cultured on EHS gel, resulting in the decline of the actin-regulatory molecule PI(4,5)P2 and the promotion of the actin depolymerization in the cells. These changes in cytoskeletal filament assembly that occur following cell attachment to the ECM and/or alterations in the pattern of actin-regulatory molecules, including PI(4,5)P2 , cofilin, and LIM kinase, are likely to determine and maintain the differentiated phenotype of hepatocytes by regulating the liver-specific transcription factor HNF-4 [72].

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Further data revealed that ECM functions mechanistically to augment the differentiation character of primary human hepatocytes in culture by mediating a reduction in cellular stress response signaling and by enhancing gap-junctional cell–cell communication [5].

ECM OF HEPATOCELLULAR AND CHOLANGIOCELLULAR CARCINOMA (HCC AND CCC) Several hepatic tumors develop on the basis of liver cirrhosis. On special changes of the ECM of liver tumors, only scattered reports are available. The balance between the proteolytic activity of MMP-2 and TIMP-2 was analyzed in more studies. Gianelli et al. found that MMP expression was similar among primary nodule tissues of patients with and without metastasis of hepatic tumors. Serum and tissue levels of MMP-2 were not statistically different between patients with and without metastasis, but MMP-2 was concentrated at the invasive edge of the metastatic tissue. On the contrary, serum and tissue levels of TIMP-2 were significantly higher in HCC patients without metastasis than in those with. This situation was not only observed in the primary HCC tissues, but also in the metastatic nodules. These results correlated with the clinical outcome, because more than 90% of the patients with high levels of TIMP-2 were still alive after two years, whereas less than 30% with low levels of TIMP-2 had survived. Authors found a strict correlation between tissue and serum levels of TIMP-2, this suggesting that an MMP-2/TIMP-2 imbalance, and in particular TIMP-2 levels, could represent an important prognostic factor in patients with HCC [73]. In an attempt to seek tumor-associated proteins of HCC, OPN was identified as highly overexpressed, and overexpression correlated with tumor grade, tumor stage, and early recurrence. OPN was identified as one of the genes whose expression was most highly upregulated in metastatic as compared with non-metastatic tumors. OPN identification by immunohistochemistry in hepatitis B virus (HBV)-positive HCC was strongly positively correlated with portal vein and lymph node invasion and negatively correlated with worse disease-free and overall survival [16]. Recent findings underline an important role of ECM in the development of CCC. Fibrotic changes in the matrix microenvironment, typified by increased type I and III collagens and fibroblast recruitment, were shown to stimulate biliary epithelium hyperplasia, with subsequent progression to malignant intrahepatic CCC only in mice harboring a p53 mutant allele. These murine CCCs bear histological and genetic features of human intrahepatic CCC, including dense peritumoral fibrosis, increased inducible nitric oxide

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synthase, nitrotyrosine, and cyclooxygenase-2 expression, c-Met activation, cErbB2 overexpression, downregulation of membrane-associated E-cadherin, and p53 codon 248 mutation. Thus, p53 deficiency, chronic bile duct injury/proliferation, and the fibrotic matrix microenvironment cooperate to induce intrahepatic CCC, highlighting the key role of the ECM microenvironment in this common liver cancer [74]. Hepatoma cells and HCC cells are also capable of the synthesis of some matrix components, and modifying the action of matrix-degrading enzymes. Carr et al. [75] recently provided evidences for this issue. Prothrombin inhibited DNA synthesis in hepatocytes cultured on FN, but not on collagen matrix. At the same time, hepatoma cell lines were not inhibited. This difference was based on the matrix produced by hepatoma cells. Prothrombin decreased FN but not collagen amounts in cases where hepatocytes were present, but in cases where hepatoma cells were used, not. Prothrombin also caused changes in cell shape and actin depolymerization [75].

CONCLUSION The liver ECM plays an important role in the elasticity and rigidity of the organ. It is a dynamic system adjusted to the cells, and provides appropriate interactions with the cells within the tissue. It facilitates communication between the liver cells and with the other organs. The liver ECM suffers qualitative and quantitative changes during acute phase, inflammation, fibrosis, cirrhosis, regeneration, and tumor development. In the liver, in response to injury, a rapid clot formation occurs, providing a provisional matrix. The inflammatory reaction also drives the tracking pathways of later fibrosis. Inflammatory processes are closely related to matrix deposition. The ECM plays a positive role even in advanced diseases, and it contributes to the tissue repair. Recent findings revealed novel data regarding the dynamic communication between liver cells and matrix components.

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56. Griffith, L.G. and Swartz, M.A. (2006) Capturing complex 3D tissue physiology in vitro. Nat Rev Mol Cell Biol , 7, 211–24. 57. Bershadsky, A., Kozlov, M. and Geiger, B. (2006) Adhesion-mediated mechanosensitivity: a time to experiment, and a time to theorize. Curr Opin Cell Biol , 18, 472–81. 58. Ingber, D.E. (2006) Cellular mechanotransduction: putting all the pieces together again. FASEB J , 20, 811–27. 59. Barker, T.H., Grenett, H.E., MacEwen, M.W., Tilden, S.G., Fuller, G.M., Settleman, J., Woods, A., Murphy-Ullrich, J. and Hagood, J.S. (2004) Thy-1 regulates fibroblast focal adhesions, cytoskeletal organization and migration through modulation of p190 RhoGAP and Rho GTPase activity. Exp Cell Res, 295, 488–96. 60. Rege, T.A., Pallero, M.A., Gomez, C., Grenett, H.E., Murphy-Ullrich, J.E. and Hagood, J.S. (2006) Thy-1, via its GPI anchor, modulates Src family kinase and focal adhesion kinase phosphorylation and subcellular localization, and fibroblast migration, in response to thrombospondin-1/hep I. Exp Cell Res, 312, 3752–67. 61. Dudas, J., Mansuroglu, T., Batusic, D., Saile, B. and Ramadori, G. (2007) Thy-1 is an in vivo and in vitro marker of liver myofibroblasts. Cell Tissue Res, 329, 503–14. 62. Mansuroglu, T., Dudas, J., Saile, B., Batusic, D. and Ramadori, G. (2006) Thy-1 expression in human liver cirrhosis and in rat models of liver damage and regeneration. J Hepatol , 44, S77. 63. Dezso, K., Jelnes, P., L´aszl´o, V., Baghy, K., B¨od¨or, C., Paku, S., Tygstrup, N., Bisgaard, H.C. and Nagy, P. (2007) Thy-1 is expressed in hepatic myofibroblasts and not oval cells in stem cell-mediated liver regeneration. Am J Pathol , 171, 1529–37. 64. Mueller, L., Goumas, F.A., Affeldt, M., Sandtner, S., Gehling, U.M., Brilloff, S., Walter, J., Karnatz, N., Lamszus, K., Rogiers, X. and Broering, D.C. (2007) Stromal fibroblasts in colorectal liver metastases originate from resident fibroblasts and generate an inflammatory microenvironment. Am J Pathol , 171, 1608–18. 65. Su´arez-Cuenca, J.A., Chagoya de S´anchez, V., ArandaFraustro, A., S´anchez-Sevilla, L., Mart´ınez-P´erez, L. and Hern´andez-Mu˜noz, R. (2008) Partial hepatectomyinduced regeneration accelerates reversion of liver fibrosis involving participation of hepatic stellate cells. Exp Biol Med (Maywood), 233 (7), 827–39. 66. Kallis, Y.N., Bird, T.G., Iredale, J.P. and Forbes, S.J. (2008) Extra-cellular matrix remodelling is critical to the oval cell response after chronic liver injury. J Hepatol , 48 (2), S195. 67. Mingoia, R.T., Nabb, D.L., Yang, C.H. and Han, X. (2007) Primary culture of rat hepatocytes in 96-well plates: effects of extracellular matrix configuration on cytochrome P450 enzyme activity and inducibility, and its application in in vitro cytotoxicity screening. Toxicol in Vitro, 21, 165–73. 68. Nagaki, M., Shidoji, Y., Yamada, Y. et al. (1995) Regulation of hepatic genes and liver transcription factors in

30: MATRIX

rat hepatocytes by extracellular matrix. Biochem Biophys Res Commun, 210, 38–43. 69. Nagaki, M., Sugiyama, A., Naiki, T., Ohsawa, Y. and Moriwaki, H. (2000) Control of cyclins, cyclindependent kinase inhibitors, p21 and p27, and cell cycle progression in rat hepatocytes by extracellular matrix. J Hepatol , 32, 488–96. 70. DiPersio, C.M., Jackson, D.A. and Zaret, K.S. (1991) The extracellular matrix coordinately modulates liver transcription factors and hepatocyte morphology. Mol Cell Biol , 11, 4405–14. 71. Brill, S., Zvibel, I., Halpern, Z. and Oren, R. (2002) The role of fetal and adult hepatocyte extracellular matrix in the regulation of tissue-specific gene expression in feta land adult hepatocytes. Eur J Cell Biol , 81, 43–50.

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72. Nagaki, M. and Moriwaki, H. (2008) Transcription factor HNF and hepatocyte differentiation. Hepatol Res, 38 (10), 961–9. 73. Giannelli, G., Bergamini, C., Marinosci, F., Fransvea, E., Quaranta, M., Lupo, L., Schiraldi, O. and Antonaci, S. (2002) Clinical role of MMP-2/TIMP-2 imbalance in hepatocellular carcinoma. Int J Cancer, 97, 425–31. 74. Farazi, P.A., Zeisberg, M., Glickman, J., Zhang, Y., Kalluri, R. and DePinho, R.A. (2006) Chronic bile duct injury associated with fibrotic matrix microenvironment provokes cholangiocarcinoma in p53-deficient mice. Cancer Res, 66, 6622–7. 75. Carr, B.I., Kar, S., Wang, M. and Wang, Z. (2007) Growth inhibitory actions of prothrombin on normal hepatocytes: influence of matrix. Cell Biol Int , 31, 929–38.

PART THREE : INTERRELATED CELL FUNCTIONS

31

Insulin Resistance Varman T. Samuel1,2, Kitt F. Petersen1 and Gerald I. Shulman1,3,4 1 Department

of Internal Medicine, Section of Endocrinology, Yale University School of Medicine, New Haven, CT, USA 2 Veterans Affairs Medical Center, West Haven, CT, USA 3 Department of Molecular and Cellular Physiology and Howard Hughes Medical Institute, 4 Yale University School of Medicine, New Haven, CT, USA

INTRODUCTION In the wake of the obesity epidemic, the world has witnessed a rising prevalence of associated conditions such as the metabolic syndrome, type 2 diabetes mellitus (T2DM), and non-alcoholic fatty liver disease (NAFLD). T2DM places a tremendous burden on patients and society. It is a leading cause of blindness in working adults, end-stage renal failure, non-traumatic limb loss, and a major risk factor for ischemic heart disease [1]. Similarly, our society is realizing the full significance of NAFLD. It is now considered the most common chronic liver disease and a major risk factor for the development of cirrhosis and liver-related death [2]. Insulin resistance is a common finding in patients with NAFLD and/or T2DM. Simply considered, resistance to insulin develops in the peripheral tissues (muscle and fat) and in the liver. Insulin resistance in the periphery impairs insulin-stimulated glucose uptake and muscle glycogen synthesis. In the liver, insulin resistance diminishes the ability of insulin to decrease gluconeogenesis and activate hepatic glycogen synthesis. This chapter will: (i) review the current model for the pathogenesis of insulin resistance in the muscle, (ii) discuss how insulin resistance in the muscle may promote the development of NAFLD, (iii) explore the mechanisms whereby NAFLD may lead to hepatic insulin resistance and (iv) increased gluconeogenesis, and finally (v) review the role of insulin-sensitizing therapies on correcting hepatic insulin resistance. The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

MECHANISM OF FAT-INDUCED SKELETAL MUSCLE INSULIN RESISTANCE A review of the mechanisms responsible for skeletal muscle insulin resistance serves as an important reference point from which to proceed to a discussion of hepatic insulin resistance. In skeletal muscle, insulin binds to its receptor, activating the receptor tyrosine kinase activity with subsequent phosphorylation and activation of insulin receptor substrate (e.g. IRS1). When phosphorylated, IRS1 activates phosphatidylinositol 3 kinase (PI3 kinase) and, through a cascade of signaling intermediaries, ultimately increases the translocation of glucose transporter (GLUT4)-containing vesicles to the plasma membrane. The current concept of muscle insulin resistance links intramyocellular fatty acid (FA) metabolite (i.e. diacylglycerol or DAG) accumulation with a blunting of insulin-mediated stimulation of glucose transport [3]. Specifically, intramyocellular fat accumulation prevents insulin-mediated activation of IRS1-associated PI3 kinase activity, ultimately limiting the ability of movement of GLUT4 from its intracellular compartment to the plasma membrane. This model has evolved over several decades. In the 1960s, Randle et al. first formed a hypothesis linking obesity with insulin resistance. He suggested that increased delivery of FAs promotes FA oxidation and inhibits

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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glucose oxidation. Fat oxidation increases intramitochondrial acetyl-CoA/CoA and NADH/NAD+, which in turn leads to the inactivation of pyruvate dehydrogenase (PDH), the enzyme responsible for the conversion of pyruvate into acetyl-CoA. Additionally, accumulation of intracellular citrate inhibits phosphofructokinase (PFK), a key glycolytic enzyme. The block in PFK would lead to accumulation of glucose 6-phosphate (G6P), which in turn inhibits hexokinase 2. The inhibition of hexokinase 2 activity results in an increase in intracellular glucose and subsequently a decrease in glucose uptake. In essence, Randle hypothesized that oxidation of fat prevents oxidation of glucose by inhibiting key glycolytic enzymes.

Intramyocellular lipid accumulation causes muscle insulin resistance The link between FAs and insulin resistance was examined in a cross-sectional study comparing lean normoglycemic offspring of patients with T2DM and age- and body mass index (BMI)-matched controls. An inverse correlation was seen between plasma FA and insulin sensitivity [4]. Using 1 H magnetic resonance spectroscopy (MRS) and the hyperinsulinemic-euglycemic clamp method, Krssak et al. measured intramyocellular lipid (IMCL) content and insulin sensitivity in normal individuals. They [5] and others [6] found IMCL concentration an even stronger predictor of insulin resistance than plasma FA. Rothman et al., using 13 C and 31 P MRS, studied the biochemical mechanisms responsible by measuring the changes in muscle glycogen and G6P both in patients with type 2 diabetes [7] and in non-diabetic first-degree relatives of patients with type 2 diabetes [8]. In contrast to Randle’s hypothesis, the accumulation of G6P was diminished in both the diabetic subjects and the first-degree relatives of diabetic patients. These data suggested the insulin resistance was due to impairments in either glucose transport or glucose phosphorylation and that these changes were present prior to the development of T2DM. Thus, the accumulation of fat within the muscle is associated with a decrease in insulin action, possibly due to a defect in either glucose transport or phosphorylation. To strengthen the causal link between lipids and insulin resistance, Roden et al. infused normal insulin-sensitive subjects with either glycerol or Intralipid/heparin. As the triglyceride in the Intralipid was hydrolyzed by lipoprotein lipase and released from the endothelium by heparin, plasma FA concentration increased. Using 13 C MRS to measure muscle glycogen content, insulin-stimulated muscle glycogen synthesis was then assessed during a euglycemic-hyperinsulinemic clamp [9]. After three hours of Intralipid/heparin infusion, the rate of insulin-mediated glycogen synthesis began to decrease. This decrease in glycogen synthesis was preceded by a drop in the concentration of G6P, as assessed by 31 P MRS. This finding

was in contrast to Randle’s model, which predicted an accumulation of G6P. Instead, these data suggested that the increase in plasma FAs resulted in muscle insulin resistance through inducing a defect in either glucose transport or phosphorylation activity. In order to distinguish between these two possibilities, Dresner et al. studied normal subjects with a similar glycerol vs. Intralipid/heparin infusion protocol in combination with 13 C MRS to look specifically at intracellular glucose concentrations. If the fat-induced muscle insulin resistance was due to a block in glucose phosphorylation, intracellular glucose would increase. In contrast, compared to the glycerol infusion, the Intralipid/heparin infusion was associated with lower concentrations of intracellular glucose and a failure to increase G6P concentrations [10]. This suggested that elevations in FAs led to insulin resistance through a defect in insulin-stimulated glucose transport activity. Furthermore, based on muscle biopsies obtained from these subjects, the apparent defect in insulin-stimulated glucose uptake was associated with impaired activation of IRS1-associated PI3 kinase activity. Thus, elevations in FAs could lead to peripheral insulin resistance through a defect in the insulin signaling pathway limiting the increase in muscle glucose transport activity.

How does lipid accumulate in muscle? Simply stated, IMCL accumulates when supply is in excess of utilization. On the supply side of the equation are the increased concentration of plasma lipids often seen in obesity and the metabolic syndrome. As stated above, increasing delivery of fat to the muscle via an infusion of Intralipid induces insulin resistance in normal, insulin-sensitive subjects. At the level of the muscle, flux into the myocyte may be increased by the action of various lipases. Pollare et al. measured liver-lipoprotein lipase (LPL) activity and insulin sensitivity, as assessed by the hyperinsulinemic-euglycemic clamp in insulin-sensitive controls and various insulin-resistant populations (obese/non-diabetic, obese/diabetic, etc.) [11]. They found that muscle LPL activity was inversely related to the glucose infusion rate during the clamp and positively correlated to fasting plasma insulin. That is, the higher the LPL activity, the more insulin-resistant the subject. Thus, increased delivery and uptake of FA promotes accumulation of IMCL and muscle insulin resistance. Balancing muscle lipid delivery is muscle lipid utilization, specifically fat oxidation by mitochondria. Muscle mitochondrial function has been shown to be impaired in two groups of patients who are at risk for developing type 2 diabetes: the offspring of patients with type 2 diabetes and the elderly [12–14]. Mitochondrial function can be assessed using a combination of 13 C MRS with 13 C-acetate

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infusion to quantify TCA cycle flux and 31 P MRS to quantify ATP synthesis. These techniques were used to study lean offspring of patients with type 2 diabetes who already had skeletal muscle lipid accumulation and insulin resistance, and were at high risk for developing type 2 diabetes later in life [12]. TCA cycle activity was decreased by 40% in these individuals compared to insulin-sensitive control subjects. Consistent with the decrease in mitochondrial oxidation, these patients were found to have ∼40% lower rats of basal mitochondrial ATP synthesis [14]. This decrease in muscle mitochondrial function was associated with a decrease in muscle mitochondrial content [15]. Thus, at least in this select cohort, a possible defect in mitochondrial biogenesis may lead to decreased mitochondrial oxidation, which favors the accumulation of lipids within the muscle, and the subsequent development of insulin resistance. The underpinnings for impaired mitochondrial biogenesis are still unclear. Though some microarray studies have pointed to decreased expression in peroxisome proliferator-activated receptor γ (PPARγ) coactivator 1α in the muscles of patients with T2DM [16, 17], this was not the case in lean, insulin-resistant first-degree relatives of patients with T2DM [15]. The development of insulin resistance is not limited to those with an inherent susceptibility toward insulin resistance. Indeed, as we age, most humans will manifest similar changes. Epidemiological studies such as the Framingham study, Baltimore Longitudinal Aging Study, and the Invecchiare in Chianti Study have clearly shown an association between aging and insulin resistance [18–20]. These epidemiological findings were experimentally confirmed with the hyperinsulinemic-euglycemic clamp study, where elderly subjects were reported as having both peripheral and hepatic insulin resistance [21, 22]. As in the younger insulin-resistant subjects, the presence of peripheral insulin resistance in the elderly is associated with accumulation of IMCLs [13, 23, 24]. Using 13 C and 31 P MRS techniques, Petersen et al. showed that age-related decreases in mitochondrial oxidation and phosphorylation were associated with increases in intramyocellular triglyceride content and decreases in insulin sensitivity [13]. These findings, along with the studies in the insulin-resistant offspring of patients with T2DM, suggest that impaired mitochondrial function, with an impaired capacity to oxidize fats, predisposes muscle fat accumulation and decreased insulin sensitivity.

How does muscle fat accumulation cause muscle insulin resistance? The prior discussion has presented the evidence demonstrating the association between IMCLs and muscle insulin resistance in humans. The mechanistic insights linking accumulation of lipid metabolites to impairments in insulin action were gleaned from rodent studies. Specifically, accumulation of a DAG within the muscle

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activates protein kinase C-θ (PKCθ) (Figure 31.1). The isoform belongs to the “novel” group of PKCs that are known to be activated by DAG. Once activated, PKCθ serine phosphorylates IRS1, which in turn prevents tyrosine phosphorylation of IRS1 by the insulin receptor. This proximal defect in insulin signaling impairs insulin-stimulated muscle glucose uptake. This model has been tested in several rodent models. Skeletal muscle DAG accumulation has been shown to temporally precede the activation of PKCθ and the development of muscle insulin resistance during an acute lipid infusion [25]. Conversely, muscle DAG accumulation can be prevented in fat-fed mice either by preventing entry or promoting oxidation. The former was accomplished by knocking out a key lipid transporter, fatty acid transport protein 1 (FATP1) [26], while the latter was accomplished by overexpression of uncoupling protein 3 [27] or knockout of acetyl-CoA carboxylase-2 [28]. In these very different models, the common findings were of lower skeletal muscle DAG and greater muscle insulin sensitivity. The necessity for PKCθ activation and IRS1 serine phosphorylation was also demonstrated in mice lacking PKCθ [29] and in mice carrying a Ser > Ala mutation in key residues of IRS1, thereby preventing serine phosphorylation by PKCθ [30]. Current data support a role for PKC activation in the pathogenesis of fat-induced insulin resistance in humans as well. Increased activity of PKCθ has also been seen in the muscles of patients with T2DM [31]. The development of insulin resistance following lipid infusion in normal, insulin-sensitive volunteers has been associated with increases in muscle PKC activity. However, in these studies, lipid infusion is associated with activation of both PKCβ2 and δ, but not θ [32].

PERIPHERAL INSULIN RESISTANCE CONTRIBUTES TO THE DEVELOPMENT OF HEPATIC STEATOSIS There is a close association between the development of NAFLD and insulin resistance. An attractive model is that the hyperinsulinemia in insulin-resistant states, combined with abundant substrate (i.e. plasma lipids), promotes the development of hepatic steatosis. This hypothesis has been examined by several investigators. Liver triglyceride can arise from either de novo lipogenesis or re-esterification of FAs released from adipose or absorbed after a meal. Donnelly et al. measured the relative contributions of these pathways in subjects with NAFLD [33]. While they demonstrated that in humans re-esterification of triglyceride released from adipose accounted for the largest portion of liver TG synthesis, in patients with NAFLD the most striking finding was the increase in de novo lipogenesis. Specifically, they found that de novo lipogenesis

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Muscle

Liver

IRS2

IRS1 IR

P P

PI3K

P

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P P

P P

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P

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Glucose

Pyruvate FOXO1 FOXA2 CREBBP Glucose GS

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PKC-q

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PKC-q Ser/Thr Kinases

P P P

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Figure 31.1 Pathogenesis of fat-induced insulin resistance. Normal insulin action in muscle (a) and liver (b). In the muscle, insulin acts to stimulate the translocation of GLUT4-containing vesicles to the plasma membrane, allowing for glucose entry into the myocyte. In the liver, insulin inhibits gluconeogenesis and activates glycogen synthesis, thereby decreasing hepatic glucose production. In the insulin-resistant state (c,d), accumulation of DAG in the muscle activates PKCθ, serine phosphorylation of IRS1, and impaired GLUT4 translocation to the membrane. Similarly, in the steatotic liver, DAG-activated PKCε leads to impaired insulin signaling and diminished regulation of hepatic glucose production.

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accounted for 26% of liver TAG. Moreover, they found that in these patients, de novo lipogenesis was consistently elevated without the normal rise and fall seen with under fed and fasted conditions in control subjects [34]. Diarison et al. utilized isotopic methods to assess the turnover of non-esterifided fatty acids and triglyceride as well as the rate of de novo lipogenesis [35]. In subjects with NAFLD, they found a approximately threefold increase in the rate of de novo lipogenesis, again suggesting that the increase in de novo lipogenesis distinguishes subjects with NAFLD from normal controls. However, is increased lipogenesis present before NAFLD has developed in subjects at risk, such as lean but insulin-resistant individuals? Petersen et al. measured lipogenesis in lean, healthy, young individuals that were either insulin-sensitive or insulin-resistant, as determined by oral glucose tolerance test. These subjects were matched for multiple factors, including age, BMI, % fat mass, blood pressure, and activity. Fasting plasma glucose and insulin, while still within normal range, were significantly higher in the insulin-resistant cohort. In addition, plasma TG was higher and plasma HDL lower. In contrast to other studies, in which visceral fat mass is found to be increased in insulin-resistant individuals, in these subjects abdominal fat mass was identical. At baseline, liver triglyceride content, as assessed by 1 H MRS, was normal ( type III [57]. The Kd for the type II InsP3 R is 27 nM, which is twice as great as the affinity of the type I isoform, and 10 times the affinity of the type III isoform [57]. InsP3 is absolutely required for Ca2+ release via the InsP3 R, but the concentration of Ca2+ in the cytosol modulates the open probability of the Ca2+ channel [39, 55, 56]. This dependence of the InsP3 R on the Cai 2+ concentration is important for orgasignals. nizing the spatial and temporal pattern of Ca2+ i For example, the open probability of the type I InsP3 R [39]. Thus, exhibits a bell-shaped dependence on Ca2+ i concentrations less than 1–2 µM exhibit a positive Ca2+ i feedback effect on the InsP3 R, while higher concentrations become progressively inhibitory. This relationship 2+ is responsible for regenerative Ca2+ i release and Cai oscillations [58], which can occur even in the presence of constant amounts of InsP3 [59]. In contrast to the type I InsP3 R, the open probability of the type III InsP3 R exhibits a sigmoidal dependency on Ca2+ i , which lacks an signaling via the inhibitory phase [55, 60]. Thus, Ca2+ i type III InsP3 R tends to be all-or-none, and activation of this isoform results in rapid and complete emptying of Ca2+ stores [55]. The type III InsP3 R is found in the apical region of epithelia, where Ca2+ i signals originate [61–64], and thus it may serve as the isoform that triggers Ca2+ release in these cells [55]. The role of the type II InsP3 R signaling is less clear. Single-channel recordings in Ca2+ i of this isoform suggest it displays a sigmoidal dependence [56], similar to what is observed with the type on Ca2+ i III InsP3 R. However, cells expressing only the type II isooscillations, similar to what form exhibit sustained Ca2+ i is observed in cells expressing only the type I InsP3 R [65]. concentraInhibition of the type I InsP3 R by high Ca2+ i tions results from interactions with calmodulin [66], so it is possible that calmodulin or other cofactors similarly interact with the type II InsP3 R but were omitted from single-channel studies. Further work will be needed to clarify the relationship between the single-channel properties of the type II InsP3 R and its behavior in intact cells. Calmodulin also may affect the binding of InsP3 to its receptor [67, 68]. This has been examined using peptides consisting of the N-terminal 581 amino acids of InsP3 R isoforms, which contain the full InsP3 binding domain (residues 226–578). In this system, calmodulin decreases the binding affinity of InsP3 to the type I and III InsP3 R, but only minimally affects binding to the type II InsP3 R.

These effects were observed in both the presence and absence of Ca2+ [68]. Together, these observations suggest that distinct InsP3 R isoforms may be involved in different intracellular functions. The activity of single channels or channels expressed in cell lines is highly dependent on postranslational modifications such as phosphorylation and binding of protein cofactors. The InsP3 R sequence has ∼2700 amino acids and includes sites for phosphorylation by kinases such as cyclic AMP-dependent protein kinase (PKA), cyclic GMP-dependent kinase (PKG), PKC, Ca2+ -calmodulin-dependent protein kinase II (CaMKII), and Akt kinase [69]. As expected, these sites are also targets for protein phosphatases and it has been show that protein phosphatase 1α (PP1α) is important for reversing the PKA-induced phosphorylation [70]. Calcineurin, a well-known Ca2+ -dependent phosphatase, is known to bind InsP3 Rs through its binding partner FKBP and to decrease either the PKC- or the CaMKII-mediated phosphorylation [71]. The effects of InsP3 R phosphorylation by PKA do not alter the sensitivity of the receptor to Ca2+ in liver [72], but PKA is inhibitory in other tissues i [46]. PKG and PKC both phosphorylate the InsP3 R and alter its function as well [46], but whether there are liver-specific effects of these kinases is not established. In cerebellar microsomes, which predominantly express the type I InsP3 R [73], phosphorylation by CaMKII increases the sensitivity to Ca2+ efflux. ATP increases the open probability of the types I and III InsP3 R up to a concentration of 2 mM, while ATP is inhibitory at concentrations above 4 mM [35]. More recently, yeast two-hybrid screenings have identified several novel InsP3 R-interacting proteins [52]. These interactions were shown to affect either the channel subcellular localization, the channel activity, or the channel’s association into signaling complexes. Protein 4.1N, initially described in neurons, has been show to direct basolateral sorting of InsP3 R I in MDCK cells [74], although it does not interact with any of the InsP3 R isoforms in a polarized liver cell line [75]. Among those binding partners leading to a reduction in InsP3 R-mediated Ca2+ release are DANGER [76] and IRBIT [77]. On the other hand, the proteins increasing the channel activity include chromogranins A and B [78, 79], NCS-1 [80], and Rack1 [81]. The assembly of signaling complexes that include InsP3 R has been show for the HOMER/Na+ K+ ATPase [82] and HOMER/TRPC3 [83]. The significance of these interactions for liver function awaits further investigation. Degradation of the InsP3 R can occur through the proteosome pathway [84], and provides yet another level of regulation of this Ca2+ release channel. Cells can express different InsP3 R isoforms, and many cell types express more then one isoform. For example, cerebellar purkinje neurons express almost exclusively type I InsP3 R, whereas the type II and III isoforms predominate in AR4-2j and RINm5F cells, respectively [73]. B-lymphocytes and pancreatic acinar cells express

32: Ca2+ SIGNALING IN THE LIVER

all three isoforms, while hepatocytes express only type I and type II InsP3 R [73]. The type III InsP3 R is not detected in hepatocytes, but is the predominant isoform in bile duct epithelia [85]. Moreover, isoforms may be expressed in distinct subcellular locations. Pancreatic acinar cells express all three isoforms in the apical region [63, 64], while the non-pigmented ciliary epithelium of the eye expresses the type I InsP3 basolaterally and the type III isoform apically, without expressing the type II isoform to a measurable extent [62]. The InsP3 R II is most concentrated in the apical region of hepatocytes, while InsP3 R I is dispersed throughout the cell (Plate 32.1a) [86, 87]. In contrast, in bile duct epithelia the InsP3 R III is most concentrated in the apical region, while InsP3 R I and II are dispersed throughout the cell (Plate 32.1b) [88]. The apical localization of the type II InsP3 R in hepatocytes depends upon lipid rafts in the canalicular membrane [89], but the mechanism by which InsP3Rs associate with lipid rafts is a topic of active investigation.

Ryanodine receptor The other major class of intracellular Ca2+ release channels is the ryanodine receptor (RyR). Like the InsP3 R, the RyR has three family members, and these channels contribute to Ca2+ signaling by releasing Ca2+ from the i lumen of the sarcoplasmic reticulum or ER into the cytosol [1]. RyR is a homotetramer consisting of four identical subunits, each of which contains ∼5000 amino acids [90]. The C-terminal regions of the RyR tetramer cooperate to form a Ca2+ channel, while the N-terminal regions project into the cytosol to form the so-called foot domain. RyR and InsP3 R share some sequence homology, especially in the C-terminal Ca2+ channel-forming region [91]. RyR isoforms have functional similarities as well as differences. All three isoforms autocatalytically release Ca2+ via a process known as Ca2+ -induced Ca2+ release (CICR). The type II and III RyR are sensitive to cyclic ADP-ribose (cADPR), while the type I RyR is not [92, 93]. The Ca2+ -mobilizing properties of cADPR were first demonstrated in sea urchin eggs [94]. It is now appreciated that cADPR mobilizes Ca2+ via the RyR in a wide range of mammalian cell types as well, including smooth [95] and cardiac [93] muscle, neuroendocrine cells [96], lymphocytes [97], and pancreatic acini [98]. In addition, the enzyme that catalyzes the formation of cADPR from NAD+ (ADP-ribosyl cyclase or CD38) is expressed in many tissues, including liver [99]. In rat hepatocytes this enzyme is mainly localized to the nuclear envelope, where it contributes to nuclear Ca2+ signaling [100]. RyR are of primary importance for Ca2+ signaling in i muscle, including vascular smooth muscle. Many other cell types express RyR as well, but there has been conflicting evidence regarding the presence and role of RyR in hepatocytes. Molecular studies suggest that none of the three RyR isoforms are expressed in liver [101]. However,

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pharmacological inhibition of the RyR reduces the speed of hormone-induced Ca2+ waves [86]. In addition, ryani odine binding sites have been identified in hepatic microsomes [102], and ryanodine releases Ca2+ from hepatic microsomal vesicles as well [103]. Finally, cADPR induces the release of Ca2+ from the nuclear envelope of isolated rat liver nuclei [33]. This controversy has been partially solved by a recent report demonstrating the presence of a truncated RyR I isoform in rat hepatocytes that by itself is not able to elicit Ca2+ release but can increase the frequency of InsP3 -induced Ca2+ oscillations [104]. These findings suggest there is a novel RyR-like protein which contributes to Ca2+ signaling in hepatocytes. Nicotinic acid adenine dinucleotide phosphate (NAADP) is another second messenger that mediates release of Ca2+ from intracellular stores. Like cADPR, this messenger molecule was discovered in sea urchin eggs [105] and has now been found to induce Ca2+ signaling i in mammalian cells as well [106, 107]. Both second messengers can be synthesized by the dual-function CD38 enzyme, but their formation is highly dependent on pH. In an acidic pH, similar to what is found in secretory granules, CD38 catalyzes the formation of NAADP. However, it promotes the formation of cADPR in neutral or slightly basic pH [108]. As suggested by studies in sea urchin eggs and pancreatic acinar cells, NAADP mobilizes Ca2+ from acidic compartments such as secretory granules through an unknown mechanism, although there have been reports showing NAADP-dependent Ca2+ release via RyR located at the ER [109]. The role of NAADP in hepatocyte Ca2+ signaling is not i clear, although there is in vitro evidence demonstrating NAADP-dependent Ca2+ release from reconstituted hepatocyte lysosomes and microsomes [110, 111].

Mitochondria Mitochondria are best known for the metabolic and respiratory role they play in cells. However, it is now clear that mitochondria strongly influence Ca2+ signaling as i well, by actively taking Ca2+ up from and releasing it back into the cytosol [112, 113]. Mitochondria have their own Ca2+ transport machinery, involving Ca2+ influx through a uniporter, and Ca2+ efflux via both an Na+ exchanger [114] and a H+ exchanger [115]. The uniporter is driven by the potential gradient across the mitochondrial membrane, while the Ca2+ efflux mechanisms are active transport systems [116]. Ca2+ efflux from mitochondria can also occur through the permeability transition pore (PTP). Formation of this pore results in a sudden, marked increase in the permeability of the mitochondrial inner membrane to ions and small molecules. Irreversible formation of the PTP can dissipate the potential gradient across the mitochondrial membrane, leading to mitochondrial swelling and irreversible cell injury [116], including necrosis [117]. However, the PTP also can form in a

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reversible fashion. This occurs in normal mitochondrial signaling [116, 118, function and plays a role in Ca2+ i 119]. Formation of the PTP is enhanced by factors including increases in mitochondrial free Ca2+ , membrane depolarization, and alkaline matrix pH [116, 120]. Activation of the PTP leads to rapid release of large amounts of mitochondrial Ca2+ into the cytosol [118, 121]. Formation of the PTP is inhibited by: antioxidants, reducing agents, PLA2 inhibitors, and the immunosuppressive peptide cyclosporin A [116, 122]. Endogenous inhibitors of the permeability transition include ADP, Mg2+ , and increased mitochondrial membrane potential [116]. Current evidence suggests that reversible formation of the PTP is typically involved in Ca2+ signaling through the foli lowing sequence of events [118]: (i) elevations in Ca2+ i are taken up by mitochondria through their uniporter, as a result of the mitochondrial membrane potential gradient; (ii) H+ efflux occurs to compensate for this influx of cations, resulting in increased mitochondrial pH; (iii) this increased pH activates formation of the PTP, which alters the membrane potential gradient; (iv) Ca2+ thus is released from the mitochondria, triggering mitochondrial Ca2+ -induced Ca2+ release (mCICR). Thus, mCICR depends on the transient depolarization of mitochondria resulting from the H+ conductive pathway provided by opening of the PTP. Ca2+ waves can propagate across the cytosol via mCICR in regions where mitochondria are densely distributed [118, 123]. Mitochondria, like the ER, are densely distributed in hepatocytes (Figure 32.2), and participate in hepatocyte Ca2+ signaling [123, 124]. i Recently, the discovery of interactions between apoptotic proteins, InsP3 R, and mitochondrial proteins helped

(a)

uncover an essential interplay between ER and mitochondrial Ca2+ signaling in the process of programmed cell death. The current view can be summarized as follows: anti-apoptotic proteins belonging to the Bcl-2 family, such as Bcl-XL and Bcl-2, induce ER Ca2+ leak through direct interaction with the InsP3 R I. This Ca2+ leak in turn reduces the ER Ca2+ available to be released into the cytosol and consequently decreases the amount of Ca2+ taken up by surrounding mitochondria. Since mitochondria are less likely to become overloaded with Ca2+ , the formation of the PTP is hampered and the overall result is a diminished cellular sensitivity to apoptotic stimuli [125]. Mcl-1 is another anti-apoptotic protein that may exert its effect in part by diminishing mitochondrial Ca2+ signals. Unlike other Bcl-2 family members, Mcl-1 is expressed in mitochondria and appears to inhibit mitochondrial Ca2+ signals directly [126]. Mcl-1 is the principal anti-apoptotic protein in cholangiocytes, and its overexpression is thought to promote the development of cholangiocarcinoma. In contrast, overexpression of the pro-apoptotic proteins Bax and Bak leads to ER Ca2+ overload and greater susceptibility to Ca2+ -mediated apoptosis [127]. Mitochondria are able to sequester significant amounts of Ca2+ from the cytosol. Free mitochondrial Ca2+ was first monitored in cells by targeting the Ca2+ -sensitive photoprotein aequorin to mitochondria [128, 129]. It was observed that mitochondrial Ca2+ can closely parallel the Ca2+ increase induced by receptor activation. It was also i observed that the increase in mitochondrial Ca2+ following release of Ca2+ from intracellular stores is faster and larger than the increase that follows influx of extracellular Ca2+ [129]. Based on this, it was hypothesized that mitochondria are close to intracellular Ca2+ release sites, and

(b)

Figure 32.2 Distribution of mitochondria and ER in hepatocytes. (a) A confocal image of isolated rat hepatocytes loaded with the mitochondrial dye rhodamine 123 demonstrates that mitochondria are densely distributed throughout each cell. Mitochondria influence Cai 2+ signaling in hepatocytes by taking up Ca2+ from nearby InsP3 receptors, then releasing it back into the cytosol. (b) Isolated rat hepatocyte couplet labeled with the ER membrane dye ER-Tracker visualized by two-photon microscopy shows that the ER is also densely distributed throughout both the apical and the basolateral regions. The ER is the principal Ca2+ store in hepatocytes, and InsP3Rs are localized to the ER membrane, where they act as InsP3-gated Ca2+ channels

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thus are exposed to local concentrations of Ca2+ much higher than those measured elsewhere in the cytosol. This hypothesis is consistent with electron microscopy data that show a close proximity between mitochondria and the ER [130], and a dynamic physical interaction between these two organelles [131]. Subsequent work has provided further functional evidence for microdomains where mitochondria and InsP3 Rs are in close apposition, so that mitochondria take up a significant fraction of the Ca2+ released by the InsP3 R [123, 124]. For example, in pancreatic acinar cells Ca2+ oscillations can be converted into global Ca2+ waves by means of pharmacological inhibition of the mitochondrial Ca2+ uniporter [132]. There is also evidence for microheterogeneity among mitochondria. It has been estimated that ∼30% of the total cellular mitochondrial pool is located in microdomains of high Ca2+ [133]. In addition, subcellular regions that i contain fewer mitochondria show greater sensitivity to InsP3 [123]. Since the uptake of Ca2+ by mitochondria decreases the positive feedback that Ca2+ may exert on the InsP3 R, mitochondria can locally modulate InsP3 R sensitivity and thus define the threshold for InsP3 to trigger Ca2+ i signals in different subcellular regions [123]. In fact, during T-cell receptor (TCR) activation, mitochondria migrate toward the site of Ca2+ entry, and once there create a buffer layer around the plasma membrane Ca2+ channels. This buffering allows for a lower Ca2+ concentration close to these channels, which slows down their Ca2+ -mediated inactivation. As a result, mitochondria migration prolongs Ca2+ entry, and doing so potentiates TCR signaling [134]. Although each InsP3R isoform may be able to transmit Ca2+ signals into mitochondria, the InsP3R III appears most tightly coupled to mitochondria and is most effective at transmitting apoptotic Ca2+ signals induced by either staurosporine or bile acids [135]. The uptake of Ca2+ by mitochondria affects multiple factors in cell metabolism, including the mitochondrial proton motive force, electron transport, and the activity of dehydrogenases associated with the TCA cycle, adenine nucleotide translocase, the F1 -ATPase, and PDH phosphorylase [136, 137]. Studies in isolated hepatocytes have revealed that slow or small Ca2+ elevations are not i transmitted effectively into mitochondria, and therefore are unable to activate mitochondrial metabolism [137]. In 2+ oscontrast, Ca2+ i oscillations trigger mitochondrial Ca cillations and sustained NAD(P)H formation [138]. Thus, the frequency rather than the amplitude of Ca2+ oscillai tions regulates mitochondrial metabolism [137, 138]. A recent study using the ATP-sensitive phosphoprotein luciferase to monitor the mitochondrial ATP concentration has shown that increases in mitochondrial Ca2+ furthermore trigger a long-term memory of the Ca2+ signal [139]. This allows ATP production to persist beyond the time period in which mitochondrial Ca2+ is elevated [86]. Thus, Ca2+ signaling in the cytosol and mitochondria are interdependent and together regulate cell metabolism in a complex fashion [119, 121, 136].

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ORGANIZATION OF Ca2+ SIGNALS Detection of Ca2+ signals in hepatocytes Our understanding of the way in which Ca2+ signals are i organized has evolved dramatically over the past 20 years, in large part as a result of two parallel technical advances. First, development of Ca2+ -sensitive fluorescent dyes has permitted Ca2+ i to be monitored continuously in live cells. Second, improvements in fluorescence imaging techniques have permitted Ca2+ i to be detected not only in populations of cells but in single cells, and in distinct subcellular regions of individual cells as well [3]. As observations move from cell populations to single cells to subcellular regions, the complexity of Ca2+ signaling patterns increases and i the time scale of signaling events decreases (Figure 32.3). It is now appreciated that Ca2+ signaling is regulated at i the subcellular level, and that this level of regulation is necessary for Ca2+ in turn to act as a second messenger i that regulates multiple cell functions simultaneously. The first Ca2+ -sensitive fluorescent dye to receive widespread use was Quin-2, which was developed more than 25 years ago [140]. An important innovation introduced at that time was the coupling of the dye to an acetoxymethyl ester, which rendered the dye cell-permeant and thus permitted it to be loaded easily into live cells [140, 141]. Other Ca2+ dyes were developed subsequently that provided additional advantages. For example, fura-2 and indo-1 could be used for ratiometric measurements of 2+ Ca2+ i , which permitted improved quantification of Ca 2+ concentrations in cytosol [142]. Other Ca dyes have been developed which allow preferential measurement of Ca2+ in organelles such as the mitochondria or ER [137, 143]. Fluorescence detection systems also have evolved considerably over the past 20 years. Initial studies used spectrofluorimetry to examine Ca2+ in cell populations [141]. i Improvements in low-light-level cameras, computers, and image processing then permitted fluorescence studies to be performed at the single-cell level [144, 145]. Further improvements in these technologies, along with the introduction of confocal, two-photon, and total internal reflection microscopy permitted subcellular Ca2+ signals i gradients to be appreciated [146–148]. The relaand Ca2+ i tive advantages and possible uses of each of these imaging modalities have been reviewed [3, 149, 150]. Although Ca2+ signaling is usually studied with Ca2+ sensitive fluorescent dyes, other approaches have been applied. These include radioactive 45 Ca2+ efflux studies, electrophysiological (patch clamp) studies which use Ca2+ dependent Cl− currents as a surrogate marker for Ca2+ i , and bioluminescence studies using the photoprotein aequorin. In fact, the first description of Ca2+ oscillations i

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Figure 32.3 Different views of Ca2+ i signaling in hepatocytes. In each case, isolated rat hepatocytes were stimulated with the α1B -adrenergic agonist phenylephrine. (a) In a population of hepatocytes, a single transient peak is observed, followed by a sustained elevation. (b) In a single hepatocyte, a series of repetitive peaks (oscillations) are observed. (c) In different regions of the same hepatocyte, the increase in Ca2+ occurs at different times. This represents a Ca2+ wave. Notice that these Ca2+ signaling events occur over progressively shorter time intervals as the level of focus moves from populations to single cells to subcellular regions. Reprinted from [3] with permission, Copyright  1994, with permission from Elsevier

in hepatocytes was based on luminescence measurements of microinjected aequorin [151]. Recombinant aequorin has been expressed in a targeted fashion to selectively detect Ca2+ signals in cellular compartments such as

the mitochondria [128], ER [152], nucleus [153], and beneath the plasma membrane [154]. More recently, newer genetically encoded fluorescent Ca2+ indicators have been developed based on modified versions of the green fluorescent protein (GFP) fused to calmodulin [155, 156], or on the principle of fluorescence resonance energy transfer (FRET) [157]. These indicators provide the advantage of better signal-to-noise ratio and photostability as compared to the aequorin reporters and yet retain the ability to be expressed in diverse tissues and subcellular locations. Transgenic mouse strains have been engineered to express some of these Ca2+ indicators, allowing in vivo neuronal Ca2+ measurements in brain slices and even through the skulls of anesthetized animals [158]. An additional Ca2+ measurement strategy has been developed that is specially useful for decoding fast and localized Ca2+ signals such as those arising from Ca2+ entry through plasma membrane voltage-gated channels or gap junctions [159]. Alternative methods of monitoring Ca2+ have been reviewed as well [3]. i

Ca2+ signaling patterns in hepatocytes Peptide hormones such as vasopressin or angiotensin generally induce biphasic increases in Ca2+ i in populations of isolated hepatocytes (Figure 32.3). These increases typically consist of two components. The first is the rapid peak then fall in Ca2+ i , which takes place over a period of seconds. This is due to release of Ca2+ from InsP3 -sensitive stores and occurs even in Ca2+ -free medium [145, 160]. The second component is the sustained plateau in Ca2+ i , which follows the rapid peak. This occurs only in the presence of extracellular Ca2+ and is due to influx of that Ca2+ to replenish depleted intracellular stores [161]. This biphasic Ca2+ signaling pattern is highly reproducible i among cell preparations. In contrast, Ca2+ signaling patterns vary markedly among single hepatocytes. Different stimuli evoke distinct responses [162], and additional variation can be seen among hepatocytes that are stimulated under identical conditions [145, 163]. The range of signaling patterns seen among single hepatocytes includes single transient or sustained Ca2+ increases and repetii spikes (i.e. oscillations). For example, lower tive Ca2+ i oscillations, concentrations of vasopressin induce Ca2+ i while higher concentrations induce sustained increases in Ca2+ [145]. In contrast, stimulation of hepatocytes i oscillations, with phenylephrine typically evokes Ca2+ i but the oscillation frequency is dose-dependent [145]. The duration of individual Ca2+ spikes also depends i spikes induced by upon the agonist. For example, Ca2+ i phenylephrine are short (∼7 seconds) compared to the duration of spikes induced by vasopressin (∼10 seconds) or angiotensin (∼15 seconds) [151, 162]. The frequency of vasopressin-induced Ca2+ oscillations tends to be greater i

32: Ca2+ SIGNALING IN THE LIVER

than that of phenylephrine-induced oscillations as well. oscillations have Differences in the frequency of Ca2+ i been shown to regulate gene transcription in some cell systems [8, 164, 165], although this has not yet been demonstrated in hepatocytes. oscillations is The mechanism responsible for Ca2+ i not completely understood. It was proposed initially that Ca2+ i oscillations result from oscillations in the InsP3 concentration. However, non-hydrolyzable InsP3 analogs can oscillations [59], and thapsigargin, which generate Ca2+ i releases Ca2+ from the ER through an InsP3 -independent oscillations [166]. Moremechanism, also triggers Ca2+ i over, measurements of InsP3 employing a fluorescent reporter show that Ca2+ oscillations are not driven by oscillations in InsP3 levels [167]. Thus, it is currently oscillations do not depend upon InsP3 thought that Ca2+ i oscillations. Instead, oscillations are thought to result from the bell-shaped dependence of the open probability of (see above). Experimental studies the InsP3 R on Ca2+ i which suggest this are further supported by mathematical models which predict that bimodal dependence of the InsP3 R would result in Ca2+ i oscillations [58]. Extracelluoscillations in hepatocytes, lar Ca2+ contributes to Ca2+ i oscillations gradually dissipate in Ca2+ -free since Ca2+ i 2+ medium [160]. Extracellular Ca thus serves to maintain internal Ca2+ stores, which are the primary source of Ca2+ for oscillations in hepatocytes [160]. In this regard, a new class of plasma membrane Ca2+ channels has gained importance, namely the transient receptor potential (TRP) channels. This family of non-selective cation channels is divided into six subfamilies: TRPC (canonical), TRPV (vanilloid), TRPM (melastatin), TRPP (polycystin), TRPML (mucolipin), and TRPA (ankyrin) groups [168]. These are implicated in diverse biological processes ranging from thermal perception to cell-volume regulation. Regarding Ca2+ signaling, there is now evidence that TRP isoforms function as store-operated Ca2+ channels (SOCs) located at the plasma membrane. Store-operated Ca2+ entry is a process characterized by the massive movement of Ca2+ from the extracellular space to the cytoplasm in response to the emptying of the intracellular stores. SOC activation is essential for the replenishment of the intracellular Ca2+ stores after an agonist challenge [169]. Recently, the ER protein STIM1 and the plasma membrane Ca2+ channel Orai1 have been identified in some cell systems as partners that come together to replenish ER Ca2+ stores when they are depleted [170]. Whether TRP channels or STIM1/Orai1 work as SOCs in the liver is not yet clear.

Subcellular Ca2+ signals and Ca2+ waves Ca2+ signals in single cells vary not only over time but i in over space as well. For example, increases in Ca2+ i

493

epithelia may occur as polarized waves [146, 171], or they may be restricted to specific subcellular regions [172, typically begin near the apical 173]. Increases in Ca2+ i membrane, where the InsP3 R is most concentrated [61, waves was obtained in 174]. The first evidence for Ca2+ i the medaka egg during fertilization [175]. Subsequently, Ca2+ i waves also were observed in non-polarized preparations of hepatocytes, and it was noted that the site of origin remained constant irrespective of the agonist [147]. Both waves that vasopressin and phenylephrine induced Ca2+ i began at a single locus, and these waves further spread in a non-diminishing fashion across the cell. Since these horthrough InsP3 mobilization, it has mones increase Ca2+ i been proposed that the InsP3 R would be concentrated in the region where Ca2+ i waves begin. Immunofluorescence studies in rat liver and in isolated rat hepatocyte couplets have demonstrated that the InsP3 R is concentrated in the pericanalicular region [86], and this localization might be dependent on intact lipid rafts (specialized membrane patches rich in cholesterol) [89] (Plate 32.1). Examination signaling in polarized preparations of isolated of Ca2+ i hepatocytes has suggested that the canalicular region is waves originate [86, 87]. Thus, the subcelwhere Ca2+ i lular distribution of InsP3 Rs in hepatocytes may serve to organize the spatial pattern of Ca2+ signals [176]. i Several studies have examined the mechanism by waves spread from their initiation site which Ca2+ i near the canalicular membrane to the rest of the cell. waves has been obApical-to-basal propagation of Ca2+ i served in other polarized epithelia, including lacrimal and pancreatic acinar cells [146, 171, 177]. As in hepatocytes, the InsP3 R is concentrated in the apical region of these cells as well. Unlike hepatocytes, though, these epithelia express RyR [98, 178, 179]. Several lines of evidence indicate that the RyR in these cells is distributed through most of the cell, but is excluded from the extreme apex where the InsP3 R is located [178]. Furthermore, inhibition of the RyR serves to inhibit the spread of Ca2+ i waves into the basolateral region [171]. Thus, the RyR may facilitate waves away from their initiation the spread of Ca2+ i point near the apical membrane [180]. The speed of Ca2+ i waves in hepatocytes is not altered in Ca2+ -free medium, waves decrease in such nor does the amplitude of Ca2+ i a medium [86]. These observations suggest that the spread of Ca2+ i waves across the hepatocyte depends only on release of Ca2+ from intracellular stores. Although the InsP3 R is concentrated apically in hepatocytes, lower-level expression of the receptor throughout the cell or even expression of a novel low-affinity InsP3 R waves isoform could account for the spread of Ca2+ i into the basolateral region. Alternatively, InsP3 could merely serve to initiate the Ca2+ release process at the canalicular region, followed by activation of neighboring Ca2+ stores through an as yet unidentified mechanism. In cholangiocytes the pattern of InsP3 R isoform expression is different from that observed in hepatocytes. Instead of InsP3 R I and InsP3 R II, cholangiocytes express

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all three InsP3 R isofoms. Moreover, in this polarized epithelium InsP3 R III rather than InsP3R II is concentrated near the apical surface, where Ca2+ waves originate [88].

Spread of Ca2+ signals from cell to cell Ca2+ signals occur asynchronously among isolated hepi atocytes. For example, the lag time between stimulation signaling varies with vasopressin and initiation of Ca2+ i among isolated hepatocytes by up to several seconds, while the frequency of Ca2+ oscillations can vary by up to 50% among isolated hepatocytes stimulated with phenylephrine [163]. However, hepatocytes that communicate via signals. gap junctions are able to coordinate their Ca2+ i For example, stimulation of isolated hepatocyte couplets with vasopressin induces a single Ca2+ i wave that crosses both of the cells, while stimulation with phenylephrine inoscillations that are synchronized in the two duces Ca2+ i cells [163]. In the isolated perfused rat liver (IPRL), Ca2+ i signaling displays an even higher level of organization, waves which cross the since vasopressin induces Ca2+ i waves cross entire lobule [181–183] (Plate 32.2). Ca2+ i individual hepatocytes at the same speed, regardless of whether the hepatocytes are isolated or within the liver waves cross the hep[181]. Vasopressin-induced Ca2+ i atic lobule in a pericentral-to-periportal direction [181, 183], presumably directed by the V1a vasopressin receptor gradient present from the pericentral to periportal region [181]. In contrast, ATP induces Ca2+ i signals in a random fashion across the hepatic lobule [183], consistent with evidence that there is no P2Y receptor gradient across the siglobule [184]. Thus, sophisticated patterns of Ca2+ i naling are induced in the intact liver, and these patterns appear to be agonist-specific. This may permit different Ca2+ agonists to have distinct effects in liver even though Ca2+ i signals induced by these agonists can appear similar in isolated hepatocytes. signals in liver has The basis for organization of Ca2+ i been studied in multicellular systems of hepatocytes. Ca2+ i waves can spread from hepatocyte to hepatocyte [163], and this type of communication depends critically on gap junctions [185]. Studies in isolated rat hepatocyte couplets demonstrate that hepatocytes communicate via gap junctions, and that both Ca2+ and InsP3 can cross these gap junctions [185]. In addition, hormone-induced Ca2+ i signaling is highly coordinated in such couplets, and this coordination depends upon gap junction conductance as well [163]. Hepatocytes express two gap junction isoforms, connexin 32 (Cx32) and connexin 26 (Cx26) [186]. Expression of both of these isoforms is dramatically reduced after bile duct ligation, and coordination of Ca2+ i signals is impaired under this condition as well [186]. Furthermore, cell-to-cell spread of InsP3 and Ca2+ i waves is markedly impaired in hepatocytes isolated from Cx32

KO mice [187, 188]. Moreover, the expression of Cx32 or connexin 43 (Cx43) in a liver cell line, where intercellular Ca2+ signals are not normally observed, causes propagation of Ca2+ from cell to cell [189]. Thus, gap junctions signals among play an essential role in organizing Ca2+ i adjacent hepatocytes. waves depends on Organization of cell-to-cell Ca2+ i other factors in addition to gap junctions. For example, increases in InsP3 are required in each cell across which a Ca2+ wave spreads [190]. Moreover, neither InsP3 nor Ca2+ alone is sufficient to support the spread of a Ca2+ i wave across a hepatocyte [191]. The presence of agonist binding to its specific receptor at the surface of the cell is also required to support the spread of Ca2+ i waves. This condition was demonstrated in experiments in which one or both cells of a hepatocyte couplet were microperfused with norepinephrine. Stimulation of individual cells oscillations only in the cell being perfused, evoked Ca2+ i and perfusion of the entire couplet was necessary to evoke oscillations in both cells [191]. Thus, the presence Ca2+ i of hormone at each cell ensures that sufficient levels of intracellular messengers are generated to reach the level of excitability necessary for supporting the propagation wave. This concept of cytosol as an excitable of a Ca2+ i medium was initially demonstrated in Xenopus oocytes signaling patterns ob[192], but likely accounts for Ca2+ i served in hepatocytes as well [182]. Other studies have focused on the mechanism by which intercellular Ca2+ i waves become oriented within the hepatic acinus. Vasopressin-induced waves begin in the region of the central venule, where the vasopressin V1a receptor is most heavily expressed, then spread to the portal region, where it is less heavily expressed [181, waves begin in a 183]. In contrast, ATP-induced Ca2+ i seemingly random pattern across the hepatic acinus [183]. Studies in isolated hepatocytes similarly show that pericentral hepatocytes are more sensitive to vasopressin but not to ATP [184]. Moreover, studies in isolated hepatocyte couplets and triplets stimulated with vasopressin or norepinephrine show that one cell generally has increased sensitivity to a particular hormone and acts as a pacemaker to drive Ca2+ i oscillations in neighboring cells. This increased sensitivity appears to be due to increased expression of hormone receptor rather than differences in downstream signaling components such as G-proteins or InsP3 R [193, 194]. Cells with increased expression of hormone receptor produce higher concentrations of InsP3 , so they respond sooner than other cells stimulated with the same concentration of hormone. As a result, cells with the greatest level of hormone receptor expression act as pacemakers for that hormone. Furthermore, different cells can act as the pacemaker for different hormones [194]. Thus, waves and oscillations in the intact the pattern of Ca2+ i liver depends upon multiple factors, which include (i) the establishment of pacemaker cells by virtue of increased expression of hormone receptors; (ii) simultaneous stimulation of both pacemaker and non-pacemaker cells; and

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(iii) communication of second messengers among these cells via gap junctions [195]. The liver also possesses paracrine mechanisms for generating and regulating Ca2+ i signals. Both hepatocytes and bile duct cells are able to secrete ATP [196–198], and both cell types express P2Y ATP receptors [199–201]. Since P2Y receptors are G-protein-coupled receptors that link to InsP3 -mediated Ca2+ signaling, secretion of ATP i signaling in neighborby hepatocytes stimulates Ca2+ i ing hepatocytes and bile duct cells [196]. This paracrine signaling mechanism thus permits increases in Ca2+ to i spread among neighboring cells independent of communication via gap junctions. Hepatocytes and bile duct cells express P2X receptors as well [202, 203], which are ATP-gated Ca2+ channels, but the physiological role of these receptors in liver is less clear. Ca2+ signaling i in liver can also be modified rather than initiated by paracrine pathways. For example, bradykinin does not mobilize Ca2+ in isolated hepatocytes, yet in the intact i waves induced liver it modifies the propagation of Ca2+ i by vasopressin [204]. Work in co-cultures of sinusoidal endothelial cells and hepatocytes, plus studies in the perfused liver, suggest this occurs through an NO-dependent mechanism. Specifically, bradykinin induces NO release from endothelial cells, which diffuses to hepatocytes, where it stimulates generation of cGMP. In hepatocytes, cGMP activates cGMP-dependent kinase, which in turn may phosphorylate and thus modulate the InsP3 R [204]. This last step is controversial, since some investigators have reported that NO donors and cGMP analogs potentiate InsP3 -induced Ca2+ mobilization in hepatocytes [205], while others have observed no such effect [206]. In any case, these studies demonstrate that there are various paracrine pathways for regulation of Ca2+ signaling i in liver.

Nuclear Ca2+ signaling The nucleus is separated from the cytosol by the nuclear envelope, which is a specialized region of the ER [207]. Like the ER, the nuclear envelope is able to store and release Ca2+ . The nuclear envelope accumulates Ca2+ via a Ca2+ -ATPase pump [208, 209], and releases Ca2+ via channels that are sensitive to InsP3 [33, 210], cADPR [33, 211], and NAADP [212]. These Ca2+ storage pumps and release channels have distinct distributions within the nuclear envelope. The Ca2+ -ATPase pump is located only in the outer leaflets of the envelope, while the InsP3 R is located only in the inner membrane [32]. Cyclic ADP ribose-sensitive channels appear to be present on both sides of the nuclear envelope [37]. In addition, both InsP3 R and RyRs are localized along invaginations of the nuclear envelope, denoted the nucleoplasmic reticulum, which work as a regulatory Ca2+ domain within the nucleus [36, 213]. The importance of nuclear Ca2+ is also highlighted by recent findings demonstrating that

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different transcription factors are directly or indirectly dependent on Ca2+ in the nucleus [214–216]. Several lines of evidence suggest that InsP3 releases Ca2+ directly from the nuclear envelope into the nucleus. InsP3 releases 45 Ca2+ into isolated hepatocyte nuclei [217], and InsP3 increases free nuclear Ca2+ [33] even if the nucleus is surrounded by a Ca2+ chelator [218]. Similarly, direct injection of InsP3 into the nucleus of Xenopus laevis oocytes results in increased nuclear Ca2+ , even if cytosolic InsP3 Rs are blocked [219]. Photorelease of caged InsP3 in the nuclei of starfish oocytes [218] increases nuclear Ca2+ as well. In addition, extracellular ATP was shown to preferentially activate nuclear Ca2+ release in HepG2 cells, via an InsP3 R-dependent mechanism [220]. The nuclear envelope possesses the machinery necessary to produce InsP3 , including PIP2 and PLC [38], and this machinery may be activated selectively through tyrosine kinase pathways [221]. In one study, IGF-1 and integrins caused PIP2 breakdown in the nucleus but not at the plasma membrane [221], while activation of G-protein-linked receptors caused breakdown of PIP2 in the cytosol, but not in the nucleus [222]. Similarly, activation of the HGF receptor c-Met in a liver cell line caused PIP2 breakdown in the nucleus, resulting in nuclear Ca2+ signals. Moreover, the activation of this highly localized cascade was dependent on the rapid translocation of the activated receptor to the nucleus [29]. It also has been hypothesized that relocation of MAP kinase to the nucleus activates nuclear PLC to generate InsP3 in the nucleus [37]. Like InsP3 , cADPR can elevate Ca2+ in isolated hepatocyte nuclei [33, 211]. Photorelease of caged cADPR furthermore induced nuclear Ca2+ oscillations in starfish oocyte nuclei [218]. These findings suggest that the machinery needed for production of cADPR exists in the nucleus. More recently, it has been demonstrated directly that ADP-ribosyl cyclase (CD38) is located on the inner membrane of the nuclear envelope, where it co-localizes with the RyR [211]. Thus, the nucleus has the independent capacity to generate both InsP3 - and cADPR-mediated Ca2+ signals. Nuclear Ca2+ signaling may also occur by transmission signals into the nucleus. The nuclear envelope of Ca2+ i contains pores that are permeable to molecules up to 60 kDa in size [223]. In the absence of a gating mechanism, a pore of this size would allow rapid equilibration of Ca2+ between the nucleus and cytosol. Under certain circumstances, free diffusion of Ca2+ through the nuclear grapore indeed occurs [153]. However, a nuclear-Ca2+ i dient has been demonstrated in a number of cell types [218, 224], suggesting that the permeability of nuclear pores can be regulated. Moreover, electrophysiological studies [207] suggest that Ca2+ permeability through the nuclear pores is severely restricted. Atomic force microscopy studies similarly suggest that nuclear-pore permeability is regulated, and that depletion of Ca2+ within the nuclear envelope closes the pores [225]. Other

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work using fluorescent dyes [226] or aequorin [227] also demonstrates that depletion of Ca2+ attenuates the permeability of the pores to intermediate-sized molecules that lack the nuclear localization sequence. Studies monitoring diffusion of photoactivatable GFP across the nuclear envelope suggest that permeability of the nuclear pore may be regulated by Ca2+ rather than by Ca2+ stores within i the nuclear envelope [228]. In either case, since EF-hand Ca2+ binding motifs are present in proteins of the nuclear pore, it is possible that these function as Ca2+ gating sensors for the pore [37]. Although nuclear Ca2+ signals can originate inside the nucleus, and the permeability of nuclear pores to Ca2+ is regulated, there is considerable evidence that nuclear Ca2+ can passively follow Ca2+ i [219, 229]. For example, stimulation of basophilic leukemia cells with antigen or photoreleased InsP3 results in a Ca2+ wave that appears to spread from the cytosol into the nucleus [230]. Similar observations have been made in hepatocytes stimulated with vasopressin [231]. Moreover, a mathematical analysis of Ca2+ waves in hepatocytes stimulated with vasopressin suggests that nuclear Ca2+ signals can be described simply by diffusion of Ca2+ inward from the nuclear envelope [232]. Nuclear Ca2+ can also contribute to Ca2+ signals in the cytosol. For example, localized increases of Ca2+ in the cytosol (Ca2+ puffs) can spread across the cell by diffusing across the nucleus. Ca2+ puffs are highly transient and localized Ca2+ i signals that result from the coordinated opening of small clusters of InsP3 Rs [233]. Puffs can be triggered by a subthreshold concentration of agonist and the resulting Ca2+ signal rapidly dissipates by diffusion i in the cytosol and sequestration of Ca2+ into intracellular stores. However, since the range of diffusion of Ca2+ in the nucleus can be much greater than in cytosol [232], Ca2+ puffs generated near the nuclear envelope can spread into and across the nucleus in order to spread to other, more distant regions of the cytosol [229]. Thus, the nucleus may function as a tunnel that helps distribute Ca2+ to the cytosol.

Ca2+ signaling in bile ducts cells Ca2+ signaling has been examined to a lesser extent in i bile duct epithelia than in hepatocytes. ATP and UTP both increase Ca2+ in the Mz-ChA-1 cholangiocarcinoma cell i line, a model for bile duct epithelium [201]. ATP and UTP also increase Ca2+ i in primary cultures of rat bile duct epin these ithelia, and acetylcholine (ACh) increases Ca2+ i cells as well [199]. As in hepatocytes and other epithelia, the range of patterns of agonist-induced Ca2+ signals i increases and Ca2+ includes sustained and transient Ca2+ i i 2+ oscillations. Cai spikes induced by ACh are longer in duration and lower in frequency than those induced by ATP [199]. Ca2+ signaling is mediated by InsP3 in bile duct i are blocked by InsP3 R ancells, since increases in Ca2+ i tagonists. Current evidence suggests that all three InsP3 R

isoforms are expressed in this cell type [85, 88]. Polarized Ca2+ waves in these cells begin in the apical region, where the type III InsP3R is concentrated, and then spread basolaterally via the types I and II InsP3Rs. Bile duct cells are coupled via Cx43 gap junctions, and expression of Cx43 synchronizes Ca2+ oscillations in these cells. Moreover, Cx43 permeability is under hormonal control, and activation of either protein kinase A or C decreases permeability and impairs intercellular communication [234].

FUNCTIONAL EFFECTS OF Ca2+ SIGNALS Ca2+ regulates a wide range of functions in liver and in other tissues. The following sections are meant to provide illustrative examples of the different ways in which Ca2+ regulates liver function, rather than to provide an exhaustive list of Ca2+ -mediated functions.

Glucose metabolism Storage and release of glucose was among the first functions of the liver shown to be regulated by Ca2+ i . Synthesis of glycogen is regulated by glycogen synthase, while phosphorylase is the rate-limiting enzyme for glycogenolysis. Both enzymes are regulated by phosphorylation and dephosphorylation [235], and an increase in Ca2+ is one of the most important signals i for regulating these events [236]. For example, hormones such as vasopressin and angiotensin increase InsP3 in hepatocytes, which mobilize Ca2+ , leading to phosphorylation and activation of glycogen phosphorylase, and then glycogenolysis. Similarly, in both rat and human hepatocytes, nucleotides activate glycogen phosphorylase and thus stimulate glycogenolysis by binding to P2Y nucleotide receptors [237]. Glucagon and beta-adrenergic agonists also stimulate glycogen phosphorylase activity in liver, but through an alternative, cAMP-dependent pathway. Ca2+ -mobilizing bile acids such as ursodeoxycholic acid (UDCA), taurolithocholic acid (TLCA), and lithocholic acid (LCA) activate phosphorylase to the same extent as hormones such as vasopressin [238]. These bile acids activate phosphorylase through a Ca2+ -dependent but InsP3-independent mechanism [238], consistent with the observation that they increase Ca2+ in an i InsP3 -independent fashion [239, 240]. Gluconeogenic enzymes are preferentially located in the periportal region [241], although other factors may also be involved in regional differences in glycogenolytic capacity. ATP mobilizes glucose mainly from the periportal zone, while norepinephrine and vasopressin preferentially release glucose from pericentral hepatocytes. This may in part reflect the fact that pericentral hepatocytes are more sensitive than periportal hepatocytes to vasopressin and norepinephrine [184]. When hepatocytes are not

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uniformly sensitive to a particular hormone, intercellular communication via gap junctions enhances glucose release. For example, glucose release is impaired in perfused livers from Cx32-deficient mice upon stimulation with either norepinephrine or glucagon [242]. Similarly, vasopressin- or glucagon-induced glucose release is impaired in perfused rat livers treated with the gap junction blocker 18α-glycyrrhetinic acid (αGA) [243]. However, glucose release is not altered in αGA-treated livers stimulated with dibutyryl cAMP or 2,5-di(tert-butyl)-1,4-benzohydroquinone (tBuBHQ), both of which stimulate glucose release in a receptorindependent fashion [243]. Hormone-induced glucose release is also impaired if gap junctions are blocked in isolated rat hepatocytes, or if hepatocytes are dispersed [244]. Therefore, hepatocytes may contribute differently to glucose metabolism across the hepatic lobule, although there is some integration of metabolic activity via gap junctions. This integration of metabolic function is particularly important in times of stress, because fasting induces hypoglycemia in cx32-deficient but not wild-type mice, and because endotoxin-induced hypoglycemia is exacerbated in KO mice as well [188].

Bile flow and paracellular permeability Ca2+ regulates fluid and electrolyte secretion in many i types of epithelia [3]. For example, in pancreatic acinar cells, apical-to-basal Ca2+ i waves direct vectorial transport of Cl− and Na+ [146, 245] . Although polarized Ca2+ i waves also occur in hepatocytes [86], the effects of these waves on bile secretion is not yet established. Nonetheless, it is established that Ca2+ i has multiple effects on bile flow. The net effect of Ca2+ i on bile acid-independent bile flow is inhibitory [246]. Studies in the IPRL have shown that a range of Ca2+ agonists, including vasopressin, the Ca2+ ionophore A23187, and the Ca2+ -ATPase inhibitor tBuBHQ, decrease bile flow. This inhibitory effect occurs independent of PKC activation or vasoconstriction [246]. Ca2+ i may inhibit bile flow in part by increasing tight-junction permeability, which would allow reflux of biliary constituents into the sinusoidal space, thereby dissipating the osmotic gradient that drives bile flow [247]. Evidence that Ca2+ increases paracellular permeability i comes from studies in both IPRLs and isolated rat hepatocyte couplets. In the IPRL, Ca2+ agonists such as vasopressin, angiotensin, and phenylephrine each increase paracellular permeability [248]. Subsequent studies in hepatocyte couplets confirmed that vasopressin increases paracellular permeability, but suggested that this effect actually is mediated by PKC [247]. NO also increases paracellular permeability in isolated rat hepatocyte couplets, but this effect is mediated by a direct effect of NO [206]. on PKC and occurs without any increase in Ca2+ i

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Thus, Ca2+ i signals may increase paracellular permeability in hepatocytes, but they appear to do so only by activating PKC.

Canalicular contraction Hepatocytes secrete bile across their apical membrane into a canalicular network that drains into the bile ducts. Time-lapse microscopy has demonstrated that bile canalicular segments within this network repeatedly expand and contract [249, 250]. Canalicular contraction is a Ca2+ -dependent process. Microinjection of Ca2+ into isolated rat hepatocyte couplets or stimulation of couplets with Ca2+ agonists such as vasopressin induces canalicular contraction [247, 251]. In permeabilized hepatocytes, Ca2+ plus ATP also induces canalicular contraction [252]. Canalicular contraction involves actin and calmodulin activation as well [252]. In hepatocytes, actin filaments are located beneath the cell membrane, and are most concentrated in the pericanalicular region (Plate 32.1). Increases in Ca2+ lead to phosphorylation of myosin light-chain i kinase [253], which in turn induces the pericanalicular actin network to contract. These pericanalicular filaments induce periodic contractions in individual pairs of hepatocytes [249], and these contractions are organized to form peristaltic waves within the hepatic lobule [250]. Canalicular peristalsis is necessary to maintain bile flow, and agents such as phalloidin are thought to induce cholestasis in part by preventing contractions of pericanalicular actin [254]. Canalicular peristalsis occurs at a frequency of ∼1.5–3.0 contractions per minute and in a pericentral-to-periportal direction [250], which matches the frequency and direction of Ca2+ i waves in the liver [86, 181, 182]. Based on these observations it has been suggested that canalicular peristalsis in vivo may be directed by propagating Ca2+ waves [6, 181]. i

Exocytosis Ca2+ regulates exocytosis directly in certain cell types. i The best-studied example is the release of neurotransmitters by neurons. In the presynaptic terminal, increases in Ca2+ are associated with release of neurotransmitters i in squid axons [255]. In the presynaptic membrane, the association between the v-SNAREs and t-SNAREs responsible for vesicle fusion with its target plasma membrane is triggered by high Ca2+ concentrations, although the precise molecular mechanism remains unsolved [256]. Also, localized apical increases in Ca2+ induce exocytic i release of zymogen granules in pancreatic acinar cells [245]. Interestingly, not every apical increase in Ca2+ ini signals that duces exocytosis in the acinar cell. Only Ca2+ i reach a concentration of 5–10 µM are sufficient to trigger exocytosis [245]. RyR-mediated Ca2+ release might be

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responsible for the premature intracellular enzyme activation during pancreatitis [257], but RyR-gated Ca2+ stores are in the basolateral rather than the apical region of the acinar cell [178, 180]. Current evidence suggests Ca2+ i plays an indirect role in regulating exocytosis in liver. For example, vasopressin stimulates biliary exocytosis in the IPRL [258]. However, this effect appears to be mediated by PKC and can be reproduced by PKC agonists that do not increase Ca2+ [247, 258]. Moreover, Ca2+ ionophores increase Ca2+ i yet decrease exocytosis in IPRL. Thus PKC activation rather than Ca2+ i may serve to stimulate exocytosis in liver, although hormone-induced Ca2+ influx can activate PKC in hepatocytes [247]. Tauroursodeoxycholic acid (TUDCA) also stimulates exocytosis in IPRL [259]. As with hormones such as vasopressin, TUDCA-induced exocytosis is thought to occur by inducing Ca2+ influx, which then leads to activation of the α isoform of PKC [259, 260]. TUDCA does not stimulate exocytosis after common bile duct ligation, even though the choleretic effect of TUDCA is maintained under these circumstances. However, Ca2+ influx is selectively impaired after bile duct ligation [259]. This provides additional evidence that Ca2+ influx plays a role in exocytosis in liver, and furthermore suggests that impairment of Ca2+ influx mechanisms may contribute to cholestasis.

Ca2+ signaling induced by bile acids Certain bile acids can induce Ca2+ signaling. Hydrophoi are LCA and TLCA. bic bile acids that increase Ca2+ i Relatively low concentrations of these bile acids cause a rapid, prolonged increase in Ca2+ in hepatocytes [239, i 240]. Bile acids such as taurodeoxycholic acid and taurochenodeoxycholic acid can also increase Ca2+ i , but much higher concentrations are required. In contrast, cholic acid and taurocholic acid do not affect Ca2+ i , even when these bile acids are present in millimolar amounts [240, 261]. LCA- and TLCA-induced Ca2+ i signaling occurs independently of extracellular Ca2+ . In addition, these bile acids mobilize Ca2+ from the same pool as InsP3 , but without InsP3 formation [239]. Therefore, it is thought that these bile acids increase Ca2+ i by permeabilizing the ER, which may in part account for their toxicity. Although LCA and TLCA do not increase Ca2+ in most cell types, this is i likely because most types of cell lack a mechanism to take up bile acids. For example, TLCA does not increase Ca2+ i in either platelets or neuroblastoma cells, but it potently increases Ca2+ in both of these cell types once they are i permeabilized [262]. Both LCA and TLCA are cholestatic, but it is unclear whether this is due to their effect on Ca2+ i [246, 263]. The therapeutic bile acids UDCA and TUDCA also increase Ca2+ i in hepatocytes, and promote rather than inhibit bile flow, suggesting that increases in Ca2+ i do not necessarily induce cholestasis. TUDCA induces a more prolonged increase in Ca2+ i than TLCA, and the sustained

signal depends upon phase of the TUDCA-induced Ca2+ i influx of extracellular Ca2+ [264]. TUDCA and TLCA both activate PKC in hepatocytes as well, but each of these bile acids preferentially activates distinct PKC isoforms [260, 265]. It remains to be established whether this differential activation of specific PKC isoforms can in part explain the different effects of these two bile acids on hepatocyte function.

Regulation of cell volume Most cells have the ability to regulate their volume. Since hepatocytes and biliary epithelia are frequently exposed to osmotic stress generated by portal shifts in glucose, amino acids, and bile acids, the ability to regulate cell volume is particularly important in liver. In general, increases in cell volume promote K+ and Cl− efflux, whereas Na+ influx occurs in response to cell shrinkage [266]. Several agonists alter rat liver volume. For example, adrenaline perfusion caused a 12% volume increase and ATP stimulation led to a 9% volume decrease [267]. Ca2+ may be i involved in volume regulation in two ways. First, regulatory volume changes in liver are mediated by extracellular ATP [268]. Specifically, cell swelling induces release of ATP, which then activates hepatocyte P2Y receptors. This leads to activation of Cl− channels, which are responsible for regulatory volume decreases [268]. Stimulation of P2Y receptors likely activates these Cl− channels via increases in Ca2+ [201, 269], although this has not yet i been demonstrated directly. This autocrine pathway for cell-volume regulation has been demonstrated in primary hepatocytes [270] as well as in liver cell lines [268]. In addition, both swelling-induced and constitutive ATP release occur in the Mz-ChA-1 bile duct cell line [271]. ATP release in this cell line similarly is responsible for P2Y receptor-mediated activation of Cl− channels and volume regulation [269, 271, 272]. A related observation is that inhibition of P2Y receptors under isotonic conditions leads to cell swelling in these cells [271]. Stimulation of bile duct cells with ATP or other Ca2+ agonists under isotonic conditions does not induce a change in cell volume, however [199]. Thus, Ca2+ may be important for autocrine i regulation as well as maintenance of cell volume in hepatocytes and bile duct epithelia. Hypo-osmotic cell swelling also activates a number of MAP kinase signaling pathways in the liver [273]. Cell swelling has been associated with activation of Erk-1 and Erk-2, p38MAPK , and the c-Jun-N-terminal kinases (JNKs). Ca2+ plays an intermediate step in this MAPK i activation in certain cell types. For example, Erk activation in astrocytes requires Ca2+ influx [274]. It is less clear whether Ca2+ is similarly involved in hepatocytes [273]. i Ca2+ may also influence cell volume decrease in liver, since PKCα, a Ca2+ -dependent kinase, is known to mediate cell shrinkage via activation of K+ and Cl− channels and inactivation of non-selective Na+ channels [275].

32: Ca2+ SIGNALING IN THE LIVER

Cell proliferation It has been long recognized that Ca2+ signals are associated with progression through the cell cycle [276]. A well-characterized model system in which to study cell cycle-related Ca2+ signals is the sea urchin embryo. In this cell, two prominent Ca2+ transients have been detected. The first Ca2+ transient occurs just prior to entry into mitosis, and the second occurs during the metaphase–anaphase transition [277]. Intracellular injection of Ca2+ chelators such as BAPTA or an InsP3 R antagonist such as heparin abolishes these Ca2+ signals and prevents entry into mitosis. Moreover, introduction of InsP3 or Ca2+ in the cytosol has the opposite effect, to accelerate entry into mitosis. Ca2+ transients are also observed during cell-cycle progression in somatic cells [278], although the relationship between these Ca2+ signals and progression through the cell cycle is less established. In Swiss 3T3 cells, serum withdrawal suppresses these Ca2+ transients but does not affect progression though mitosis. However, mitosis is blocked by specific inhibition of Ca2+ transients through injection of BAPTA plus incubation in calcium-free media. Moreover, photorelease of caged Ca2+ is able to induce premature entry into mitosis. Downstream targets of Ca2+ have also been implicated in cell-cycle progression. For example, calcineurin has been show to be essential for Xenopus laevis embryonic development [279]. In addition, pharmological inhibition of the CaMKII arrests cells at the G2/M transition [280]. Moreover, calmodulin overexpression accelerates the cell cycle in mouse C127 cells and its downregulation leads to an extended cell cycle [281]. Heterologous expression of the Ca2+ binding protein parvalbumin has also been used to study the role of Ca2+ signaling in the regulation of the cell cycle. This protein is normally expressed in skeletal muscle and neurons [282]; in myocytes it modulates relaxation of fast twitch muscle fibers due to its Ca2+ buffering capacity. The first report using this protein as a molecular tool showed that buffering Ca2+ slowed progression through the cell cycle in mouse C127 cells [283]. More recently, parvalbumin variants targeted to the nucleus or the cytoplasm [214] were used to investigate the relative importance of Ca2+ signals in each of these cellular compartments for regulation of the cell cycle in a liver cell line. It was found that nucleoplasmic rather than cytosolic Ca2+ is essential for cellular proliferation, and is necessary in particular for progression through early prophase [284]. Very recent findings suggest that HGF and insulin, two potent growth factors in liver, selectively form InsP3 in the nucleus to initiate nuclear Ca2+ signals [29, 30]. These findings suggest that certain growth factors may stimulate proliferation of hepatocytes by selectively inducing Ca2+ signals in the nucleus. Perhaps the most common model of hepatocyte proliferation in vivo is hepatic regeneration after partial hepatectomy [285]. Following 70% hepatectomy, the liver

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undergoes a coordinated regenerative response involving all hepatic cell types, but this is punctuated by marked proliferation of hepatocytes. In rats, 24 hours after partial hepatectomy there is a decrease in InsP3 R II expression accompanied by a decrease in the frequency of Ca2+ oscillations. This decrease in receptor expression normalizes within four days and is then associated with an increase in the frequency of Ca2+ oscillations. This remodeling of the Ca2+ signaling machinery is thought to be essential for the initial regenerative response [286]. The paracrine and hormonal signals involved in this response are still under study but at least two growth factors, HGF and epidermal growth factor (EGF), are known to mobilize Ca2+ i in hepatocytes and are required for liver regeneration [23, 287]. The role played by these growth factors in cell proliferation is further highlighted by their involvement in the development of hepatocellular carcinoma [288].

Ductular secretion Ca2+ i directly regulates fluid and electrolyte secretion in a number of epithelia, including bile duct epithelia. Cholangiocytes express apical, Ca2+ -activated Cl− channels [269, 272], and it is thought that this is the primary mechanism for Ca2+ -stimulated secretion in these cells. The other principal mechanism for regulating cholangiocyte secretion is via the apical, cAMP-activated cystic fibrosis transmembrane conductance regulator (CFTR) Cl− channel [289]. Together, these two mechanisms create the Cl− − gradient responsible for driving HCO− 3 /Cl exchange that mediates bicarbonate secretion into the bile [290]. Ca2+ potentiates adenylyl cyclase activity and cAMP formation in cholangiocytes via a calcineurin-dependent pathway [291], so this cross-talk between the Ca2+ and cAMP signaling pathways has been considered a secondary, indirect mechanism for Ca2+ -stimulated secretion. However, current evidence suggests that cAMP- and CFTR-mediated secretion may rely on Ca2+ in a more direct way as well [197]. Activation of CFTR in cholangiocytes promotes ATP release into the bile, which stimulates apical P2Y receptors [197, 198]. These G-protein-coupled receptors link to InsP3 formation and then Ca2+ release from apical, type III InsP3 Rs, which in turn drives Ca2+ -dependent Cl− channels and then HCO− 3 secretion [197, 203]. Therefore, neurotransmitters such as acetylcholine stimulate basolateral M3 muscarinic receptors on cholangiocytes, leading to Ca2+ release from InsP3 R I and InsP3 R II. This Ca2+ in turn activates apical Ca2+ -dependent Cl− channels, which ultimately stimulate biliary HCO− 3 secretion. In contrast, peptide hormones such as secretin stimulate basolateral secretin receptors, which induce the formation of cAMP, activating CFTR and leading to paracrine, ATP-mediated HCO− 3 secretion. This putative universal role of InsP3R-mediated Ca2+ signaling in ductular secretion is in agreement with the observation that loss of InsP3 Rs is a common molecular event in cholestatic

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disorders including primary biliary cirrhosis, primary sclerosing cholangitis, extrahepatic biliary obstruction, and biliary atresia [292]. Cholangiocytes express primary cilia on their apical membranes, and this sensory organelle also links to Ca2+ signaling and secretion. For example, primary cilia can sense and transduce mechanical stimuli from within the bile duct lumen, so that increases in bile flow activate both cAMP production and Ca2+ release [293]. Cholangiocyte cilia also sense increases in osmolarity within the bile duct lumen, and this links to activation of TRPV4 channels, which allow entry of extracellular Ca2+ into the cytoplasm [294]. The role of cilia and ciliary Ca2+ signaling in cholestatic disorders remains an area of investigation.

CONCLUSION The importance of Ca2+ as a second messenger in liver and in other tissues is well known. Most of the cellular components that are involved in generating Ca2+ signals i have been identified and were reviewed here. It is now becoming apparent, however, that Ca2+ signaling also i depends on interactions between these components. Thus, Ca2+ signaling in one region of the cytosol is dependent i signaling in other subcellular regions, as well upon Ca2+ i as upon interactions between the cytosol and organelles, and among neighboring cells. Future advances in this field likely will result from an increased understanding of how these various aspects of Ca2+ signaling are integrated i to regulate signaling in the intact liver. This in turn may increase our understanding of the complex way in which liver function is regulated by Ca2+ in health and disease.

ACKNOWLEDGMENTS This work was supported by NIH grants DK57751, DK34989, DK61747, and DK45710, and by grants from CNPq, FAPEMIG, and HHMI.

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Role of Intracellular Iron Movement and Oxidant Stress in Hepatocellular Injury John J. Lemasters1 , Akira Uchiyama2, Jae-Sung Kim3, Kazuyoshi Kon2 and Hartmut Jaeschke4 1 Center

for Cell Death, Injury and Regeneration, Departments of Pharmaceutical & Biomedical Sciences and Biochemistry & Molecular Biology, Medical University of South Carolina, Charleston, SC, USA 2 Department of Gastroenterology, Juntendo University School of Medicine, Tokyo, Japan 3 Department of Surgery, University of Florida, Gainesville, FL, USA 4 Department of Pharmacology, Toxicology & Therapeutics, University of Kansas Medical Center, Kansas City, KS, USA

CHELATABLE IRON, OXIDATIVE STRESS, AND CELL DEATH The liver is often a target of injury by ischemia reperfusion, oxidative stress, and toxic chemicals [1–6]. Reactive oxygen species (ROS) and reactive nitrogen species (RNS) frequently underlie hepatic injury. Xanthine oxidase utilizing xanthine and hypoxanthine generated after ATP degradation, NADPH oxidase in Kupffer cells and inflammatory cells attracted to the sites of tissue injury, and dysregulated respiration in • mitochondria generate superoxide (O2 − ) and hydrogen

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

peroxide (H2 O2 ). In the presence of transition metal ions, • such as iron and copper, O2 − and H2 O2 react to form • the highly reactive and toxic hydroxyl radical (OH ) through the Fenton reaction (Figure 33.1). In addition, transition metals catalyze a lipid peroxidation chain reaction sustained by lipid alkyl and peroxyl radicals. • Nitric oxide (NO ) is an important signaling molecular • produced by nitric oxide synthase (NOS). NO reacts at •− diffusion-limited rates with O2 to form peroxynitrite (OONO− ) (Figure 33.1). Peroxynitrite causes nitrosation of tyrosyl residues in proteins and also decomposes to a hydroxyl radical-like species. Increasingly, peroxynitrite and other RNS are recognized as important toxic intermediates in toxicity to liver and other tissues [7].

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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Figure 33.1 Scheme of iron-catalyzed free-radical generation. Oxidative stress causes oxidation of NAD(P)H and GSH, • important cofactors for antioxidant defenses, which promotes increased net formation of superoxide (O2 − ) and hydrogen •− •− to H2 O2 , which peroxidases and catalase degrade to water for peroxide (O2 ). Superoxide dismutase converts O2 • detoxification. In the presence of chelatable iron, O2 − reduces ferric iron (Fe3+ ) to ferrous iron (Fe2+ ), which reacts with • H2 O2 to form the highly reactive hydroxyl radical (OH ). Cellular ferrireductases may also reduce Fe3+ to ferrous iron Fe2+ . • • OH reacts with lipids to form alkyl radicals (L ) that initiate an oxygen-dependent chain reaction, generating lipid peroxides • • • (LOOH) and peroxyl radicals (LOO ). Iron also catalyzes a chain reaction, producing alkoxyl radicals (LO ) and more LOO . • • •− Nitric oxide synthase catalyzes nitric oxide (NO ) formation from arginine. NO reacts with O2 to form peroxynitrite anion • • (ONOO− ), which decomposes to nitrogen dioxide (NO2 ) and OH . These reactive oxygen, nitrogen, and lipid species also attack nucleic acid and proteins

In cells and tissues, iron exists in two pools. Ferritin, hemosiderin, and iron-containing prosthetic groups in proteins (e.g. heme, iron–sulfur complexes) sequester “non-chelatable” iron that conventional iron chelators like desferal cannot bind. “Chelatable” iron represents free unbound iron plus loosely bound iron. Free iron actually constitutes only a small fraction of all chelatable iron in cells because of binding of ferrous and ferric iron to polyorganic anions such as citrate. Overall, chelatable iron is estimated to be 5 µM in hepatocytes [8]. Although an essential nutrient, excess iron is toxic and causes acute hepatocellular necrosis after accidental overdose and chronic hepatic injury in hereditary hemochromatosis [9]. Iron in excess also appears to aggravate diabetes, cardiovascular disease, cancer, and alcoholic and non-alcoholic steatohepatitis [10–15]. Promotion of cell death by increased intracellular chelatable iron and cytoprotection by iron chelators like desferal in various models of oxidative stress and hypoxia/ischemia infer a role for iron in the pathogenesis of injury, most likely by catalyzing ROS formation [16–23]. The catalase mimetic, TAA-1/Fe, blocks iron-dependent killing of hepatocytes, as expected if Fenton chemistry is underlying in iron-dependent toxicity [24]. However, the permeable superoxide dismutase (SOD) mimetic, MnTBAP, which dis• mutates O2 − to oxygen and H2 O2 , does not protect [25].

This finding suggests that conversion of Fe3+ to Fe2+ by a ferrireductase occurs inside cells, which bypasses the • need for iron reduction by O2 − (Figure 33.1). In other systems, H2 O2 potentiates iron toxicity, consistent with • a pathophysiological role for OH formation from H2 O2 2+ and Fe [26].

IRON UPTAKE BY RECEPTOR-MEDIATED ENDOCYTOSIS AND RELEASE INTO THE CYTOSOL In plasma, transferrin binds almost all non-heme iron as Fe3+ . At a plasma transferrin concentration of 5–10 µM, 30% iron occupancy, and two iron binding sites per transferrin molecule, plasma iron is 3–6 µM. Extracellular transferrin continuously delivers iron to the endosomal/lysosomal compartment by receptor-mediated endocytosis [27], but how iron is released from this compartment into the cytosol for cellular needs, such as synthesis of iron-containing proteins, is incompletely understood [28]. A membranous iron transporter, divalent metal transporter-1 (DMT1), mediates H+ /Fe2+ symport by enterocytes across the plasma membrane and early

33: ROLE OF INTRACELLULAR IRON MOVEMENT AND OXIDANT STRESS IN HEPATOCELLULAR INJURY

endosomes into the cytosol, and may mediate iron release from lysosomes and late endosomes [28, 29]. However, recent data indicates that type IV mucolipidosis-associated protein (TRPML1) is the cation channel responsible for release of Fe2+ and other trace metals like Mn2+ and Zn2+ from late endosomes and lysosomes [30]. Both DMT1 and TRPML1 are specific for Fe2+ and not for the Fe3+ that is delivered to endosomes by transferrin. Thus, endosomal iron release requires reduction of Fe3+ to Fe2+ by an endosomal ferrireductase, which is identified as the product of the six transmembrane epithelial antigen of the prostate 3 (Steap3) gene [30]. Proteolysis in lysosomes and proteosomes also recycles iron for biosynthetic needs. Similarly, heme oxygenase releases free iron as heme is degraded, and heme oxygenase-2 is at least in part localized to endosomes [31]. Ferritin and hemosiderin store iron in a highly chelated and unreactive form. Ferritin and hemosiderin reversibly accumulate in iron-overload states [32].

MITOCHONDRIAL PERMEABILITY TRANSITION In the mitochondrial permeability transition (MPT), permeability transition (PT) pores open to make the mitochondrial inner membrane permeable to all solutes up to a molecular weight up of about 1500 Da [33–35]. Ca2+ , ROS, peroxynitrite, and numerous reactive chemicals promote pore opening, whereas cyclosporin A, high Mg2+ , and pH less than 7 inhibit PT pores. When PT pores open, mitochondria depolarize, and matrix swelling occurs driven by colloid osmotic forces. Additionally, mitochondria release matrix solutes, including calcium and pyridine nucleotides, and oxidative phosphorylation becomes uncoupled. Swelling of the inner-membrane-matrix compartment leads to rupture of the outer membrane and release of cytochrome c and other proapoptotic factors from the intermembrane space. PT pores have a very large single-channel conductance, and patch clamping studies suggest that opening of a single PT pore may be sufficient to cause mitochondrial depolarization and swelling [36]. Onset of the MPT is a key pathophysiological event inducing necrotic and apoptotic cell death after ischemia/reperfusion, oxidative stress, and many instances of hepatotoxicity [6, 37]. The composition of PT pores remains uncertain. In one model, PT pores are composed of a supramolecular complex of the voltage-dependent anion channel (VDAC) from the outer membrane, the adenine nucleotide translocator (ANT) from the inner membrane, cyclophilin D (CypD) from the matrix, and possibly other proteins (reviewed in [38, 39]). Although once widely accepted, genetic knockout studies question the validity of the model by showing that an MPT can still occur in mitochondria that are deficient in ANT and VDAC [40–42]. Moreover, a cyclosporin A-insensitive MPT occurs in mitochondria

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from CypD-deficient mice [34, 43]. In an alternative model, PT pores arise when damaged, misfolded membrane proteins altered by oxidative and other stresses aggregate to form aqueous channels regulated by CypD and other molecular chaperones [44]. ANT is one but not the only membrane protein that can undergo such misfolding and aggregation.

RELEASE OF CHELATABLE IRON FROM THE LYSOSOMAL/ENDOSOMAL COMPARTMENT Fluorescent probes like phen green SK and calcein visualize changes of chelatable iron [45, 46]. Ferrous (Fe2+ ) but not ferric (Fe3+ ) iron quenches these fluorophores (Table 33.1), as do divalent transition metal ions of Cu, Ni, and Co [47]. In calcein-loaded hepatocytes, calcein fluorescence decreases preceding cell death in models of oxidative stress, hypoxia, ischemia reperfusion, and drug toxicity [48–50], as shown in Figure 33.2 for a mouse hepatocyte exposed to acetaminophen [51]. Calcein quenching is not caused by passive leak of calcein across the plasma membrane, since calcein fluorescence decreases to less than that of calcein free acid placed in the extracellular medium. The iron chelator dipyridyl, which competes with calcein for binding of Fe2+ , restores calcein fluorescence (Figure 33.2), and desferal, another iron chelator, is protective in these injuries [19, 52–54]. During hypoxia/ischemia and oxidative stress, lysosomes rupture [48, 55–57]. Iron release appears to occur with lysosomal rupture to promote pro-oxidant cell injury [58–60]. Lysosomal breakdown also occurs in models of hepatocellular apoptosis [61, 62]. The intriguing possibility from these studies is that chelatable iron increases in these various injuries and contributes to iron-dependent ROS formation, onset of the MPT, and cell death. In calcein-loaded hepatocytes, inhibition of the vacuolar (endosomal and lysosomal) proton-pumping ATPase with bafilomycin (50 nM) causes calcein fluorescence to decrease (Figure 33.3) [63]. Bafilomycin-induced calcein Table 33.1 Ferrous iron but not ferric iron quenches calcein fluorescence Addition None DPD Fe(NH4 )2 (SO4 )2 Fe(NH4 )2 (SO4 )2 + DPD FeCl3 FeCl3 + DPD

Fluorescence 28 32 1.2 13 26.5 30.5

Fluorescence in arbitrary units of caIcein (2 µM) was measured in culture medium before and after addition of 10 µM FeCl3 (ferric iron), 10 µM Fe(NH4 )2 (SO4 )2 (ferrous iron), and the iron chelator dipyridyl (DPD, 20 mM), as indicated.

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Calcein

3h

6h

DPD

Figure 33.2 Acetaminophen-induced quenching of calcein. Mouse hepatocytes were ester-loaded with calcein acetoxymethylester (1 µM) for 30 minutes before exposure to acetaminophen (10 mM) in the presence of fructose (20 mM) and glycine (5 mM) to prevent onset of necrotic cell death but not toxic acetaminophen metabolism. Note strong quenching of calcein fluorescence after between 3 and 6 hours exposure to the hepatotoxicant. Loss of calcein fluorescence was not due to passive leak since calcein (300 µM) was also present in the extracellular medium. The iron chelator dipyridyl (20 mM, DPD) reversed calcein quenching [51]

Figure 33.3 Bafilomycin-induced intracellular calcein quenching: inhibition by desferal and starch-desferal. Mouse hepatocytes were loaded with calcein-AM (1 µM) and incubated in culture medium containing calcein free acid (300 µM) with no further addition (control), 50 nM bafilomycin (Baf), bafilomycin plus 1 mM desferal (Baf + Dsf), and bafilomycin plus 1 mM desferal equivalency of starch-desferal (sBaf + Dsf). Note marked decrease of intracellular calcein fluorescence after bafilomycin, which did not occur during the control incubation and which was suppressed by desferal and starch-desferal [63]

33: ROLE OF INTRACELLULAR IRON MOVEMENT AND OXIDANT STRESS IN HEPATOCELLULAR INJURY

quenching is evident within 60 minutes and even more marked after 2 hours (Figure 33.3). Desferal blocks this bafilomycin-induced calcein quenching nearly completely. Al3+ is the only other biologically relevant metal ion that is chelated by desferal [64], but Al3+ does not quench calcein fluorescence [47]. Thus, calcein quenching specifically indicates an increase of Fe2+ . The magnitude of increase of Fe2+ is substantial, since the decrease of intracellular calcein fluorescence is approximately equal to the fluorescence of calcein free acid (300 µM) placed in the extracellular medium. Assuming a one to one stoichiometry of chelatable Fe2+ to quenched calcein, bafilomycin causes chelatable Fe2+ to increase by about 300 µM. High-molecular-weight starch-desferal also prevents bafilomycin-dependent calcein quenching (Figure 33.3) [63]. Starch-desferal is membrane-impermeable. Starch and related glucans like dextran can only enter hepatocytes via endocytosis to accumulate in the lysosomal/endosomal compartment [48]. Thus, the source of the increase of chelatable iron in the cytosol after bafilomycin is most likely iron released from the lysosomal/endosomal compartment secondary to lysosomal/endosomal alkalinization.

CHELATABLE IRON AND CYTOTOXICITY AFTER OXIDATIVE STRESS Glutathione peroxidase in mitochondria reduces tert-butylhydroperoxide (TBH) to t-butanol, which promotes mitochondrial oxidative stress by depleting mitochondrial glutathione and NADPH [30]. When hepatocytes are exposed to a low dose of TBH (25 µM), little cell killing occurs (Figure 33.4) [63]. Similarly, bafilomycin alone does not cause cell killing. However, the combination of TBH plus bafilomycin causes substantial cytotoxicity (Figure 33.4). Cell killing is accompanied by increased ROS production (Figure 33.4) and mitochondrial depolarization, signifying onset of the MPT. Desferal and starch-desferal both prevent almost completely enhanced ROS production, mitochondrial depolarization, MPT onset, and cell death after low-dose TBH plus bafilomycin (Figure 33.4 and data not shown) [63].

UPTAKE OF CHELATABLE IRON BY MITOCHONDRIA Using a cold ester loading/warm incubation protocol, mitochondria and lysosomes of hepatocytes can be loaded selectively with calcein [49, 65]. After bafilomycin, mitochondrial calcein fluorescence decreases (Figure 33.5), whereas calcein fluorescence in rhodamine-dextran-colabeled lysosomes increases

515

Figure 33.4 Synergistic cell killing and free-radical generation after bafilomycin plus t -butylhydroperoxide: protection by desferal and starch-desferal. Loss of viability of mouse hepatocytes was determined by PI fluorometry, and ROS formation was assessed with the fluorogenic substrate, chloromethyldihydrodichlorofluorescein diacetate (cmH2 DCF-DA, 10 µM). As indicated, hepatocytes were exposed to 50 nM bafilomycin (Baf) and TBH (25 µM) with and without desferal (Des, 1 mM) and starch-desferal (sDes, 1 mM desferal equivalency) [63]

(Figure 33.5, arrows) [63]. These changes are consistent with the conclusion that mitochondrial chelatable iron increases after bafilomycin, while lysosomal chelatable iron decreases. Desferal and starch-desferal both suppress bafilomycin-induced quenching of mitochondrial calcein fluorescence without preventing the increase of lysosomal fluorescence (Figure 33.5). Overall, these results suggest that mitochondria take up chelatable iron released by lysosomes. In confirmation, the highly specific inhibitor of the mitochondrial calcium uniporter, Ru360 [66], blocks mitochondrial calcein quenching after bafilomycin (Figure 33.5).

TWO-HIT HYPOTHESIS OF IRON-CATALYZED HYDROXYL RADICAL FORMATION DURING OXIDATIVE STRESS •

•

Fe2+ catalyzes OH formation from H2 O2 (and O2 − dismutating to H2 O2 catalyzed by SOD), leading to lipid peroxidation and other deleterious effects [67]. After bafilomycin, an increase of chelatable Fe2+ is not suffi• cient by itself to enhance OH production, the MPT, and cell killing. Similarly, mild oxidative stress is insufficient to cause injury. Rather, two “hits” of oxidant stress and increased chelatable Fe2+ act synergistically to promote cellular damage (Figure 33.6). Lack of cytotoxicity after bafilomycin alone is not due to iron chelation by calcein, since calcein loading by itself is not cytoprotective [49]. Indeed, most Fe2+ chelates remain redox-active, unlike desferal which stabilizes chelatable iron as less-reactive

516

THE LIVER: CONCLUSION

Figure 33.5 Calcein quenching in mitochondria after bafilomycin: prevention by desferal and Ru360. Mitochondria and lysosomes of rat hepatocytes were loaded with calcein by cold ester loading/warm incubation, and 70 kDa rhodamine-dextran (Rhod-Dex) was used to identify lysosomes. As indicated, hepatocytes were exposed to bafilomycin (Baf, 50 nM), bafilomycin in the presence of desferal (Dsf, 1 mM), and bafilomycin in the presence of 20 µM Ru360, an inhibitor of the mitochondrial calcium uniporter. Green fluorescence of calcein and red fluorescence of rhodamine-dextran were imaged by laser scanning confocal microscopy. Note that mitochondrial calcein was quenched markedly after bafilomycin, whereas lysosomal calcein fluorescence co-localizing with rhodamine-dextran increased. In the presence of desferal, mitochondrial calcein quenching was suppressed, and the increase of lysosomal calcein fluorescence became more marked. Ru360 also suppressed mitochondrial calcein quenching but did not increase lysosomal calcein fluorescence. Arrows identify representative lysosomes [63]

ferric iron (Fe3+ ) [68]. Chelatable iron as a first hit in combination with a second hit of oxidative stress may thus explain aggravation of injury to liver and other organs by high tissue iron. Isolated mitochondria accumulate Fe2+ electrogenically via the mitochondrial Ca2+ uniporter [69]. Fe3+ is not taken up. Similarly, in bafilomycin-treated hepatocytes, mitochondria accumulate Fe2+ by the Ru360-sensitive calcium uniporter (Figure 33.5). Thus, mitochondria take up at least some of the chelatable Fe2+ released from lysosomes by bafilomycin. Moreover, in isolated mitochondria, Fe2+ induces the MPT at concentrations comparable to the ∼300 µM increase of chelatable iron observed after bafilomycin [70]. During oxidative stress and ischemia reperfusion, ROS formation occurs primarily within mitochondria and precedes onset of the MPT and cell death [19, 52]. Desferal blocks this ROS formation and the ensuing cytotoxicity. Thus, the two hits of ROS generation and increased chelatable iron may be taking place within mitochondria to promote onset of the MPT and cell death (Figure 33.6).

induction, and activation of death pathways [61, 62, 72]. In Wilson’s disease, similar mechanisms may play a role, with copper replacing iron as the transition metal promoting oxidative stress [73]. Chelatable iron is an emerging dynamic regulator of cellular function and mediator of cytotoxicity. Like free Ca2+ , chelatable iron may represent both a signal-regulating normal cellular response and an intracellular mediator of toxicity when iron homeostasis is dysregulated. In Kupffer cells, chelatable iron increases transiently after stimulation by lipopolysaccharide, and iron chelators block activation of nuclear factor κB (NFκB) and cytokine formation, suggesting a signaling role for chelatable iron [74–76]. Iron chelation also induces hypoxia inducible factor-1α transactivation [77, 78]. Although the mechanisms of iron release from the lysosomal/endosomal compartment are still being worked out, an Fe2+ /H+ exchange mechanism may be important for lysosomal iron release at high lysosomal pH and for iron retention at low lysosomal pH, since the primary effect of bafilomycin is to block acidification of lysosomes and endosomes.

RELEVANCE TO LIVER PHYSIOLOGY AND DISEASE

CONCLUSION

Mitochondrial iron overloading may contribute to a variety of hepatic diseases [10–12, 14, 71], and direct addition of membrane-permeable iron complexes induces the MPT and killing of hepatocytes [22]. Additionally, in disease settings associated with lysosomal fragility and breakdown, such as lipotoxicity and high cytokine exposure, release of Fe2+ from lysosomes and uptake into mitochondria may also contribute to oxidative stress, MPT

Storage of chelatable iron in the lysosomal/endosomal compartment and the mobilization of this iron into mitochondria by various stressors may be important in exacerbating liver disease. Specifically, two “hits” to mitochon• dria of O2 –/H2 O2 formation and increased chelatable iron • likely promote formation of toxic OH . Understanding of these mechanisms may lead to better therapies for liver diseases.

33: ROLE OF INTRACELLULAR IRON MOVEMENT AND OXIDANT STRESS IN HEPATOCELLULAR INJURY

517

ACKNOWLEDGMENT This work was supported, in part, by Grants DK037034, DK070195, DK070844, DK073336, and AA016011 from the National Institutes of Health.

REFERENCES

Figure 33.6 Scheme of two-hit hypothesis of mitochondrial hydroxyl radical formation. Chelatable iron is delivered to the lysosomal/endosomal compartment primarily as ferric iron (Fe3+ ), some of which is reduced to ferrous iron (Fe2+ ) by ferrireductase. Inhibition of the vacuolar proton-pumping ATPase by bafilomycin, as well as ATP deficiency, causes luminal alkalinization, leading to Fe2+ release by an apparent Fe2+ /H+ exchanger. Fe2+ entering the cytosol is taken up into mitochondria via the mitochondrial calcium uniporter as a first hit, promoting injury. Mitochondrial oxidative stress, such as that caused by TBH, • provides a second hit of superoxide (O2 − ) and hydrogen peroxide (H2 O2 ) generation, with some H2 O2 formed from • O2 − by SOD. Fe2+ and H2 O2 react to form hydroxyl rad• ical (OH ) by the Fenton reaction. The ensuing damage to proteins and lipids leads to onset of the cyclosporin A (CsA)-sensitive MPT, which causes mitochondrial swelling and uncoupling of oxidative phosphorylation. Swelling induces release of cytochrome c and activation of caspases 3 and apoptosis. With severe mitochondrial dysfunction, cellular ATP depletion develops due to disruption of oxidative phosphorylation and activation of the mitochondrial uncoupler-stimulated ATPase. With ATP depletion, plasma membrane failure and necrosis then occur

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34

Regulatory Pathways of Liver Gene Expression: The Central Role of Cyclic AMP Giuseppe Servillo1, Maria Agnese Della Fazia1 and Paolo Sassone-Corsi2 1 Department

of Clinical and Experimental Medicine, School of Medicine, University of Perugia, Perugia, Italy 2 Department of Pharmacology, School of Medicine, University of California, Irvine, CA, USA

INTRODUCTION All physiological and metabolic functions in mammals are orchestrated by the need of homeostasis. A remarkable variety of these functions occur in the liver, which therefore is one of the central homeostatic tissues in mammals. The adaptive plasticity of liver physiology is possible through an exquisite control of gene expression that is influenced by a variety of signaling factors. A number of agents, including hormones, growth factors, and cytokines, influence liver gene transcription, through the triggering of intracellular signaling pathways and the consequent activation of transcription factors [1, 2]. A central signaling molecule in liver physiology is cyclic adenine monophosphate (cAMP) [3]. For example, hormones critical in liver physiology (e.g. glucagon, adrenaline, noradrenaline) bind to specific receptors and via G-proteins regulate the intracellular cAMP concentration [4]. The deciphering of the signaling pathways and relative control

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

of gene expression in the liver represents a critical step toward the understanding of the role that cAMP perturbation may have in liver physiopathology.

THE cAMP TRANSDUCTION PATHWAY Of all the several second messengers that transduce signals from the external environment into the cell, cAMP appears to occupy a privileged position in the liver. In 1957, Earl W. Sutherland isolated a previously unknown compound, called cyclic adenine monophosphate, and proved that it had an intermediary role in many hormonal functions. cAMP derives from ATP by the action of the adenylate cyclase (AC). AC is an enzyme located in the cell membrane and composed of two clusters of six transmembrane segments spacing two intracellular catalytic domains [5, 6]. Nine related isoforms of

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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THE LIVER: COUPLING cAMP TO LIVER GENE TRANSCRIPTION

AC with an extensive homology in their structure have been described in mammals. The different AC isoforms (AC1–AC9) display a tissue-specific distribution. In the liver, the isoforms AC4–AC7 and AC9 are present, AC6 being the most abundant [7]. Different external stimuli, which act on specific receptors coupled to trimeric G-proteins, change AC activity [8, 9]. G-proteins are constituted by three chains: α, β, and γ. G-protein subunits exhibit a number of isoforms, some displaying differential tissue distribution, particularly in the liver [10]. Regulation of the intracellular concentration of cAMP depends on what type of receptor is activated; indeed, depending on the coupling of receptors to G-proteins, these will elicit either activation or inhibition of AC function [11, 12]. AC activation causes increased cAMP levels in the cell, which in turn trigger the cAMP-dependent protein kinase A (PKA). PKA is a tetrameric holoenzyme constituted of two regulatory (R) and two catalytic (C) subunits [13, 14]. Multiple isoforms of both R and C subunits have been identified, namely RIα, RIβ, RIIα, RIIβ, Cα, Cβ, and Cγ, generating a variety of combinatorial associations with not yet fully characterized functional differences [15–17]. Once cAMP binds to the two regulatory subunits (RI and RII), the PKA protein complex dissociates. The activated C-subunits are released from cytoplasm and Golgi anchoring sites and phosphorylate specific substrates in the nucleus and cytoplasm. They do so on specific serine residues, preferentially on the canonical peptide sequence X-Arg-Arg-X-Ser-X (where X is any amino acid) [18, 19]. Importantly, RI subunits contain binding sites for MgATP. Binding of MgATP stabilizes the holoenzyme by raising the threshold of cAMP concentration required to cause activation and enhancing holoenzyme reassociation. RII subunits do not bind MgATP but are targets of autophosphorylation, which destabilizes and activates the holoenzyme. Each subunit isoform displays a characteristic pattern of tissue-specific expression. In the liver, the Cα subunit is most abundant [16].

COUPLING cAMP TO LIVER GENE TRANSCRIPTION The complex processes of cell growth and differentiation rely on highly regulated programs of gene expression. The transcription of a gene is controlled by regulatory sequences that, in most cases, are present 5′ from the start site of transcription. The dissection of the promoter regions of many cAMP-responsive genes allowed the identification of a distinctive element that mediates transcriptional activation in response to increased levels of intracellular cAMP; the cAMP responsive element (CRE) [19, 20]. The CRE is constituted by a consensus palindromic sequence of eight base pairs (TGACGTCA; see Table 34.1) with higher conservation in the 5′ half of the palindrome with respect to the 3′ sequence [19]. CRE sites have been identified in numerous promoters [21], among which many display a neuroendocrinespecific expression. Mutations of these elements cause a significant decline in both cAMP inducibility, and in the basal transcriptional activity [22]. CRE consensus sequences have also been found in promoters, (where they appear to confer transcriptional properties of tissue-specificity [23] and of non-endocrine expression [as for the fibronectin, the MHC class II genes [24, 25]]), and in viral transcription units (as for the enhancer of the HTLV-I LTR [26] and some adenoviral promoters) [27]. In the liver, the cAMP signaling pathway controls many physiological functions, in part by controlling the expression of many genes whose promoters contain CRE consensus sequences. The CRE is often located proximal to the transcriptional start site. The use of genome-wide analysis of CRE-binding protein (CREB) target genes has shown that positional conservation of CREB binding sites is essential for the identification of target genes, and their proximity to the TATA box seems to facilitate CREB-mediated activity.

Table 34.1 Expression profiles of some liver genes and location of CREs in their promoters Gene

CRE Location

Fetal Life

Normal Liver

Induction Upon Liver Regeneration (h after PH)

TAT PEPCK SDH G-6-Pase

−3651/−3644 −90/−83 −1129/−1122 −136/−129 −161/−152 −105/−98 −116/109 −136/−129 −148/140 −66/−59 −80/−73

− − − +

+ + + +

2–10 1–4 2–5 30 min to 24

?

+

2–5

? ?

− −

30 min to 2 h 38–72

CREM (ICER)

c-fos Cyclin A

TAT, tyrosine aminotransferase; PEPCK, phosphoenolpyruvate carboxykinase; SDH, serine dehydratase; G-6-Pase, glucose-6phosphatase; CREM, cyclic AMP-responsive element modulator; ICER, inducible cyclic AMP early repressor; PH, partial hepatectomy.

34: REGULATORY PATHWAYS OF LIVER GENE EXPRESSION: THE CENTRAL ROLE OF CYCLIC AMP

523

nevertheless CREB, ATF-1 (activating transcription factor 1), and CREM (cAMP-responsive element modulator) display similar domains in their structural organization [29, 30] (Figure 34.1). They are able to heterodimerize with other family members in determinate combinations, seemingly by using a specific “dimerization code.” All the CREB family members share the common feature of generating various isoforms by alternative splicing. CREM presents as more than 10 isoforms, which display different functions. The CREM gene also presents alternative polyadenylation, alternative use of two promoters, and alternative translation, determining functionally different transcripts [21, 30]. Indeed, depending on which domain is present in the encoded protein, the function could be as different as activation and repression. For example, the transcription factors CREMτ, CREB, and ATF-1 act as

THE CRE-BINDING PROTEIN (CREB) FAMILY The characterization of transcription factors binding specifically to the CRE site revealed a group of proteins that display common structural and functional characteristics [21, 28]. The CREB belongs to the bZip (basic domain-leucine zipper) class. The bZip has a basic domain with about 50% of lysine and arginine residues, necessary for direct interaction with DNA, located near a leucine-zipper domain. The leucine-zipper domain is an α-helical coiled-coil structure, with the leucines, variable in number from four to six, set within the zipper in heptad repeats [19, 21]. The CREB family members share little similarity in primary amino acid sequence outside of the bZip region;

Ligands

AC G R

cAMP

ATP

PKA C

R R

C + CBP −

+

Phosphatase

+ CREB CREM −

+

CRE cAMP-inducible genes

−

ICER P1

CAREs

CREM

P2

Figure 34.1 Schematic representation of the cAMP signal transduction pathway and its nuclear effectors controlling gene expression. Ligands binding to specific membrane receptors activate coupled G-proteins (G), which in turn stimulate the activity of the membrane-associated adenylyl cyclase (AC), which converts ATP to cyclic AMP. Increased intracellular levels of cAMP cause the dissociation of the inactive tetrameric protein kinase A (PKA) complex into the active catalytic subunits (C) and the regulatory subunits (R). Catalytic subunits migrate into the nucleus, where they phosphorylate (P) and induce transcriptional activators such as CREB and CREM. Phosphorylation allows the recruitment of CBP (CREB binding protein), a large co-activator. DNA binding to cAMP-responsive elements (CREs) found in promoters of cAMP-responsive genes is a key step in transcriptional activation. Products of cAMP-inducible genes are involved in the hormonal response, differentiation, and proliferation. An intronic, cAMP-inducible promoter (P2) of the CREM gene directs the synthesis of ICER (inducible cAMP early repressor), which downregulates the expression of CRE-containing promoters, including P2, generating a negative auto-regulatory transcriptional loop

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THE LIVER: MECHANISMS OF TRANSCRIPTIONAL ACTIVATION

activators [30–32], while CREMα, β, γ, and CREB-2 have a repressor function [30, 33, 34]. A remarkable repressor of CREB is ICER (inducible cAMP early repressor), the only inducible repressor of the class, which is generated by an alternative promoter within the CREM gene ([21, 35]; see below).

MECHANISMS OF TRANSCRIPTIONAL ACTIVATION Transcriptional activation by CREB family members is mediated by two protein domains, independent from each other but conserved among species. The first domain, named the kinase inducible domain (KID) or phosphorylation box (P-box), contains a serine residue within a consensus phosphorylation site for PKA at position 133 in CREB and 117 in CREM [19, 31, 36, 37]. The second region is constituted by two glutamine-rich domains, termed Q1 and Q2, which flank the P-box on both sides [19, 21].

Regulation by phosphorylation The phosphorylation event is a prerequisite for turning CREB family members into powerful activators. The serine placed in the center of the P-box is the direct link between the cAMP signal transduction pathway and related gene expression. Indeed, several studies have demonstrated that the serine in the P-box is phosphorylated not only by PKA, but also by other kinases, Activation domain

representing a convergence site of various signaling pathways [21, 38–40] (Figure 34.2). For example, in Hep-G2 and 3T3-L1 cells, insulin-induced signaling causes CREB phosphorylation and stimulates transcriptional activity [41]. Several examples describe how CREB phosphorylation may be independent of the cAMP pathway. The question is particularly relevant in the liver, where various growth factors and cytokines may induce CREB phosphorylation and thereby CRE-dependent transcription, possibly mimicking the physiological effect of cAMP on gene expression. A revealing illustration of signaling cross-talk leading to CREB phosphorylation relates to the nerve growth factor (NGF) and its receptor Trk, whose stimulation results in the activation of several kinases. Trk is a receptor tyrosine kinase which, once activated, stimulates the activity of the small GTP-binding protein Ras [42]. Activation of Ras triggers the MAPK pathway, which includes the MAP kinase kinase (MEK) and the ribosomal S6 kinase pp90rsk [43]. Interestingly, constitutively activated expression of MAPK and MEK is sufficient to induce neurite outgrowth in PC12 cells [44], indicating a direct role of this pathway in eliciting the changes in gene expression required for the neuronal differentiation program. Although MAPK and MEK have not been shown to directly phosphorylate CREB, the use of cells expressing a dominant interfering Ras mutant has revealed the involvement of this pathway in CREB phosphorylation upon NGF-induction [39]. Indeed, the involvement of a CREB kinase with characteristics similar to pp90rsk has been proposed. pp90rsk is likely to be responsible for CREB phosphorylation in human melanocytes [45], while a DNA Binding domain

133 Q1

P-Box

Q2

Basic Leucine 341aa Domain Zipper

CREB

341aa

CREM

271aa

ATF1

117

63

(a)

AESEDSQESVDSVTDSQKRREILSRRPSYRKILNDLSSDAPGVPRIEEEKSEEETSA

CREB

AETDDSADS..EVIDSHKRREILSRRPSYRKILNELSSDVPGIPKIEEEKSEEEGTP

CREM

SESEESQDSSDSIGSSQKAHGILARRPSYRKILKDLSSEDTRGRKGDGENSGVSAAV

ATF1

(b)

Figure 34.2 Structure of CREB, CREM, and ATF-1. (a) The glutamine-rich domains (Q1 and Q2), the DNA-binding region (basic domain and leucine zipper), and the P-box are shown. The position of the serine residue phosphorylated by the cAMP-dependent protein kinase (PKA) and other kinases (see Figure 34.3) is also indicated. (b) Sequence alignment of the P-box domain from these proteins. The serine corresponding to the PKA-phosphoacceptor site is shown

34: REGULATORY PATHWAYS OF LIVER GENE EXPRESSION: THE CENTRAL ROLE OF CYCLIC AMP

distinct member of the RSK family, p70s6k , also possesses CREB phosphorylation activity [46]. Cells originate from patients with the Coffin–Lowry syndrome, who carry mutations in the gene encoding the RSK-2 protein, one of the four isoforms of pp90rsk [47]. It has been demonstrated that RSK-2 is responsible for CREB phosphorylation in response to epidermal growth factor (EGF) and for the consequent transcriptional induction of c-fos [40]. Finally, CREB has been shown to be phosphorylated upon activation of the stress pathway involving the p38/MAPKAP-2 kinases [48]. Thus, various signaling pathways may converge to modulate gene expression via the same transcriptional regulator, CREB. The complexity of the signaling pathways controlling transcription factors is a demonstration of the pleiotropic functions played by these molecules in the regulation of the physiology and metabolism of the cell.

Modularity of the activation domain The two glutamine-rich domains, Q1 and Q2, are crucial for the activation function of CREB family members. Q-rich domains are also present in other transcription factors, such as AP-2 and SP-1 [49, 50]. Glutamine-rich domains are supposed to mediate the interaction between the activation domain and other components of the transcriptional machinery, in order to allow an efficient initiation. The contribution of the Q2-rich domain to the transcription seems more significant than that of the Q1-rich domain [21]. The role of the Q2-rich domain is demonstrated by splicing isoforms present in CREM and ATF-1 [51, 52], both of which lack Q1 but are still able to activate transcription [32]. This shows that the P-box region and the Q2-rich domain are sufficient for the induction. The identification and characterization of the CREB-binding protein (CBP), a protein of 265 kDa that interacts selectively with the phosphorylated form of CREB, has increased understanding of the mechanism by which the cAMP pathway controls transcription [53]. CBP presents two zinc finger domains, a glutamine-rich domain, and PKA consensus sites. Following phosphorylation, CREB binds to CBP, promoting the binding of CBP to TFIIB, a factor directly linked to RNA polymerase II [54]. p300 [55] shares similar regulatory functions with CBP in differentiation [56], cell growth [57], apoptosis, and DNA repair [58]. Both p300 and CBP interact with the basal transcription factors, including TFIIB, TBP, and an RNA helicase. Several other factors involved in transcription (e.g. Jun, Fos, MyoD, p53, NF-κB) and nuclear receptors [59] interact with CBP/p300. CBP has an intrinsic or associated histone acetyltransferase activity (HAT), establishing a direct link with chromatin remodeling [54, 60–62]. The HAT function of CBP establishes a link between the signal transduction pathway and chromatin modifications. While epigenetic control of gene expression, specifically

525

directed by chromatin remodeling, is an attractive area of research, its implication in liver physiology is as yet ill-defined.

Additional routes of transcriptional activation In addition to the “canonical” phosphorylation-dependent mechanism of activation, other pathways have been described that lead to activation of CRE-mediated transcription in a phosphorylation-independent manner. Transcription factor CREM was shown to be unphosphorylated in male germ cells, a finding that led to the identification of activator of CREM in testis (ACT). ACT is a testis-specific protein that displays in its structure four complete LIM domains and one amino-terminal half-LIM motif. ACT is a co-activator that shares a high degree of homology with a family of proteins expressed in heart and skeletal muscle. ACT exerts its function independently of CREM phosphorylation at Ser117—and thus of the interaction with CBP. This finding indicated a new way to elicit activation by the transcription factors of the CREB class, possibly bypassing classical signaling pathways [63]. As ACT defines a novel class of tissue-specific co-activators, it would be of great interest to define whether proteins with similar features may be present in the liver, activating CREB-related members bypassing phosphorylation. Recently, the mechanism by which the co-activators known as TORCs (transducers of regulated CREB activity) stimulate CREB activity in a phosphorylationindependent way has been extensively studied. TORCs interact with the bZip domain of CREB, enhancing the recruitment of TAFII 130 to the promoter [64]. Moreover, the phosphorylation state of TORCs determines their binding to CREB and their activation. The link between CREB and TORCs suggests a role played by TORCs in modulating the active and inactive cellular pool of CREB. In the liver, an important role of this mechanism has been described in the control of glucose homeostasis [65].

MECHANISMS OF TRANSCRIPTIONAL REPRESSION Dephosphorylation One of the most important mechanisms that lead to down-regulation of CREB family member activity is the dephosphorylation of the serine placed in P-box. While the mechanisms and pathways by which CREB family members are phosphorylated have been well studied and defined, little is known about their

526

THE LIVER: THE cAMP PATHWAY IN THE LIVER

dephosphorylation [21, 28]. Protein phosphatases such as PP-1 and PP-2a seem to be involved in the process of dephosphorylation, as demonstrated by in vitro studies [66, 67]. However, the pathways and the signals that are activated to trigger the phosphatases and their regulation are as yet poorly defined. Thus, the role played by phosphatases in the in vivo regulation of CREB function still needs further investigation.

CRE-binding repressors Several studies have revealed that alternative splicing isoforms of CREM play a key role in down-regulating the cAMP-induced transcriptional activation. Contrary to CREB and all the activators of the CREB family, whose expressions appear to be mostly constant and ubiquitous [21, 68], differential transcript processing leads to unique profiles of CREM expression [38, 43]. The isoforms CREMα, β, γ generated from the GC-rich housekeeping promoter (P1) lack the activation domain and therefore act as antagonists of cAMP-responsive transcription by either competing for binding to CREs or blocking CREB by heterodimerizating with it [30]. Another important repressor is the CREM isoform ICER. The synthesis of ICER is cAMP-inducible, peaking at about 1 hour after the stimulus. This allows rapid downregulation of the expression of early-response genes

Hormones

ADENYL CYCLASE

following activation. ICER is a small protein of 120 aa consisting uniquely of the bZip domain and is synthesized by an alternative P2 promoter lying within an intron located upstream of the 3′ end of the CREM gene (Figure 34.3). ICER is a very small transcription factor and functions as a powerful repressor of cAMP-induced transcription [30, 35]. An essential feature of ICER is its inducibility. The P2 promoter is strongly inducible by cAMP as it contains two pairs of closely-spaced CREs (CAREs) [35]. Following cAMP treatment there is a rapid and transient increase of ICER expression characteristic of the early-response gene class [35]. ICER then turns off its own expression by repressing the activity of the P2 promoter, establishing a negative autoregulatory loop. Of course, ICER expression represses other cAMP-inducible genes whose promoters contain a CRE. ICER represents a dynamic and versatile protein that has a tissue- and developmental-specific expression pattern, suggesting different functions in physiological and pathological contexts. Its inducible expression in the liver appears to be a critical event for liver physiology (see below).

THE cAMP PATHWAY IN THE LIVER A survey of the genes under control of the cAMP signaling pathway expressed in the liver reveals that

Synaptic activity

Ca2+

Growth factors

Inflammatory cytokines

Stress

PLCg RAS Pl-3K

cAMP

Calmodulin

PKA

CaMKIV

p38

ERK

p70S6K

RSK2

MSK

MAPKAP-K2

CREB CREM

Figure 34.3 Signal transduction pathways that lead to phosphorylation of the activators CREM and CREB. Various signaling cascades are shown according to the external stimulus by which they are activated. Cross-talk between pathways is indicated by arrows. Dashed lines indicate the presence of intermediate kinases not shown. CaMKIV, Ca2+ -calmodulin-dependent kinase IV; ERK, extracellular regulated kinase; MAPKAP-K2, MAP-kinase-activated protein kinase 2; MSK, mitogen- and stress-activated kinase; p70S6K, p70 S6 kinase; PI-3K, phosphoinositide-3 kinase; PKA, cyclic AMP-dependent protein kinase; PLCγ, phospholipase C; RSK2, ribosomal S6 kinase 2

34: REGULATORY PATHWAYS OF LIVER GENE EXPRESSION: THE CENTRAL ROLE OF CYCLIC AMP

Table 34.2 A partial list of liver cAMP-responsive genes and their biological function. All these genes have been described as containing one or more CRE sequence in their regulatory promoter sequences ATF-3 c-fos ICER Krox-20 Pit-1 C/EBP-β EGR-1 JunD Nurr-1 STAT-3 NF-1 iNOS PGS-2 Ornitine decarboxylase SOD2 Tyrosine amino transferase (TAT) Cox-2 Glutamine synthetase PEPCK β1-adrenergic receptor β2-adrenergic receptor Somatostatin receptor E-cadherin CFTR Chromogranin A TGFβ2 Cyclin A Cyclin D1 mPer-1 mPer-2 GLUT-2 Bcl-2 Cytochrome c Fibronectin IGF-1

Transcription factors

Metabolic enzymes

Transmembrane receptors Transmembrane proteins Secreted proteins Cell-cycle regulators Circadian rhythm regulators Transmembrane carrier Apoptosis Cytochrome Extracellular matrix glycoprotein Growth factor

they all encode proteins that play critical roles in liver’s physiology (Table 34.2). For example, genes that play an important role in gluconeogenesis, such as tyrosine-aminotransferase (TAT), phosphoenolpyruvate carboxykinase (PEPCK), serine dehydratase (SDH), and glucose-6-phosphatase (G-6-P), contain CREs in their promoters and have cAMP-inducible expression [69–72]. In particular, TAT and PEPCK display common characteristics and represent two useful models for the study of cAMP-responsive transcription in the liver. These proteins are gluconeogenetic enzymes localized in periportal hepatocytes; their expression increases upon fasting and decreases during diets rich in glucose. Glucagon stimulates the expression of both genes by acting through the cAMP pathway [73–75]. TAT is the rate-limiting enzyme of tyrosine catabolism; it is synthesized exclusively in hepatocytes and is involved

527

in gluconeogenesis [76]. The enzyme synthesis is absent during the fetal life and rises within the first hours after birth. Administration of glucagon starting from the seventeenth day of fetal life results in the transient induction of TAT synthesis [76]. In addition, TAT is highly expressed during liver regeneration following partial hepatectomy (PH) [77]. The TAT gene promoter contains a functional CRE 3.6 kb upstream of the transcription start site [69]. Moreover, TAT is a liver-specific enzyme and presents a circadian rhythm with an increased expression during the night. Intriguingly, TAT circadian expression is modified during aging [78]. PEPCK is the enzyme that catalyzes the synthesis of phosphoenol-pyruvate from oxaloacetate. Unlike TAT, PEPCK is not a tissue-specific gene; its expression is high in the liver, kidney, and adipose tissues and low in skeletal muscle, heart, mammary gland, ovary, and lung [79]. An interesting feature of the PEPCK promoter is that distinct segments confer differential activity in different tissues. For example, sequences between positions –2088 and –888 are essential for expression in fat, heart, ovary, and muscle tissues, while for expression in kidney, sequences located between –460 and –355 appear crucial. In the liver, PEPCK expression is controlled by sequences located between –460 and +73 [80]. Interestingly, this region contains two CREs, named CRE-1 and CRE-2, CRE-1 being more active than CRE-2 [80]. However, it has been shown that CRE-1 exerts its full activity only if the integrity of other flanking elements is preserved [81]. Indeed, in addition to the CREs, the promoter contains multiple critical binding sites for liver-enriched nuclear proteins, which direct liver-specific expression. In support of this observation, the cooperative action of various transcription factors on this essential region of the PEPCK promoter has been shown. Other transcription factors, such as C/EBPα, C/EBPβ, and DBP bind to the promoter of PEPCK and seem to cooperate with CREB [82]. The observations reported above underscores one central issue in liver gene expression: the interplay between cAMP-dependent transcriptional regulators and other transcription factors controlling liver-specific promoters. This aspect has not been extensively explored to date, but future studies are likely to reveal as yet unappreciated regulatory pathways in the control of gene expression. In this respect, the link between CREB and PGC-1 (peroxisome proliferator-activated receptor (PPAR) γ coactivator) in the liver represents a revealing example. In fasted animals, following activation of the gluconeogenic pathway, CREB regulates PEPCK expression mediated by catecholamines and glucagon. In protracted condition of neoglucogenesis, CREB activates neoglucogenic genes by overexpressing PGC-1 [83]. Furthermore, in the liver of fasted animals CREB seems to control lipid mobilization. In the liver, CREB controls the metabolism of lipids by repressing the expression of PPARγ. Thus, CREB seems to operate as a sort of molecular balance to equilibrate liver

528

THE LIVER: THE ROLE OF cAMP IN PROLIFERATION AND LIVER REGENERATION

physiology during fasting gluconeogenesis and fatty-acid oxidation [84]. Importantly, CREB activation results in the induction of ICER, another transcriptional player that appears to have a central role in liver physiology. Rats treated with intraperitoneal injection of cAMP show a remarkable induction of CREB phosphorylation with a subsequent ICER expression at two, four, and eight hours following treatment [85]. Similarly, in vitro in a hepatoma cell line induced to proliferate or treated with forskolin, the results show a significant increase in CREB phosphorylation and ICER expression [85]. The powerful repressive function of ICER on cAMP-inducible transcription begs the question of the physiologically relevant targets of this repressor. Interestingly, all putative target genes encode proteins involved in metabolism, cell cycle, and signaling (Table 34.2). Finally, the function of CREB in the liver is not restricted to the hepatocytes as it appears to play an interesting role in stellate cells and cholangiocytes. Hepatic stellate cells are strictly involved in the pathogenesis of hepatic fibrosis. When activated, they proliferate, producing an excess of collagen type I. Studies performed on the proliferation of stellate cells indicate that activation of CREB by phosphorylation results in cell-cycle progression and inhibition of proliferation [86]. CREB appears to also operate in bile duct cells. These cells are unique in expressing cystic fibrosis transmembrane conductance regulator (CFTR). The CTFR is a Cl− channel mediating the transport in bile duct epithelium. Genetic mutations in CFTR cause the cystic fibrosis disease, which develops hepatobiliary disease, one of the most important causes of death in cystic fibrosis patients. CFTR has a CRE in its promoter, controlling its expression. These studies underscore the role played by CREB in the control of CFTR in cholangiocytes [87].

THE ROLE OF cAMP IN PROLIFERATION AND LIVER REGENERATION The role of cAMP signaling in proliferation has been debated for a long time. Depending on the cell type, cAMP appears to act as a promoting factor or to suppress proliferation. For example, it has been observed that glucagon increases cAMP levels in primary rat hepatocytes, resulting in increased DNA synthesis in cooperation with EGF and insulin [88]. In the same cells, prolonged exposure to cAMP synergizes with glucocorticoids to inhibit DNA replication [89, 90]. The differential effect of cAMP seems to be not only time-dependent, but also concentration-dependent. In vitro, low concentrations of glucagon treatment induce DNA synthesis, while high concentrations of the hormone show opposite effects. Indeed, it has been demonstrated that elevated levels of cAMP control hepatocyte proliferation acting at

two cell-cycle checkpoints. cAMP facilitates the transition from G0/early G1 to the pre-replicative period and it has an inhibitory action before G1/S. While the molecular pathways implicated in these effects still need in-depth analysis, some recent results may provide promising clues. The protein HOPS was characterized as translocating from the nucleus to the cytoplasm in hepatocytes, depending on the timing of the cell cycle. HOPS is mostly cytoplasmic in proliferating hepatoma cells, while it is prominently in the nucleus when cells are arrested. Similarly, HOPS is present mostly in the nucleus of the hepatocytes and is rapidly shuttled from nucleus to cytoplasm during liver regeneration, acting as a sort of switch during proliferation. Treatment of hepatoma cells with cAMP, or its intraperitoneal injection, rapidly induces translocation of HOPS from nucleus to cytoplasm (Figure 34.4). These notions suggest that the increase in cAMP levels that accompanies the first hours of liver regeneration could be responsible for HOPS shuttling from nucleus to cytoplasm, thereby allowing the residual hepatocytes to proliferate [91, 92]. Varied hormones and growth factors act on hepatocyte proliferation [93–96], suggesting a convergence and/or a cross-talk between cAMP signaling and other transduction pathways. In some cases these effects result in the increasing or delaying of the DNA synthesis. β-adrenergic agonists potentiate the effect of cAMP following PH and stimulate DNA synthesis, although it seems that the balance between the stimulatory and inhibitory effects of cAMP is strictly related to the presence of other factors (e.g. glucocorticoid, prostaglandins, vasopressin, insulin, etc.) [93–96]. Indeed, cAMP levels peak in hepatocytes at two distinct times that involve proliferation: at the birth during liver-cell proliferation and in the first hours of liver regeneration [3, 96]. The notions outlined above underscore the central role played by cAMP in the control of proliferation of residual hepatocytes following PH [3, 96] (Figure 34.5). During liver regeneration, two distinct intracellular peaks of cAMP take place in residual hepatocytes. The first peak occurs two to six hours following PH, while the second precedes the first round of mitosis and has been associated with hepatocyte proliferation [3, 96]. Interestingly, during liver regeneration the expression of PKA subunits changes in residual hepatocytes. The protein levels of RIα and RIIα subunits increase during the first hours following PH, concomitant to an increase in cAMP intracellular concentration, while the expression of the catalytic subunits remains constant [97]. Similar results have been obtained in hepatocytes stimulated in vitro with cAMP [90]. In residual hepatocytes, paralleling the peaks in cAMP levels, there is a rapid and transient induction of ICER expression in the first eight hours following PH (Figure 34.6), suggesting a regulatory function of this transcription repressor in the early phase of proliferation [85]. The critical role of CREM in liver regeneration has been confirmed by using mutant mice in which the CREM gene was deleted by homologous recombination. A significant delay in the

34: REGULATORY PATHWAYS OF LIVER GENE EXPRESSION: THE CENTRAL ROLE OF CYCLIC AMP

DAPI

Merge

(a)

(b)

(c)

(d)

(e)

(f)

(g)

(h)

(i)

HOPS

529

control

cAMP (60 min)

cAMP (120 min)

Figure 34.4 HOPS localization in hepatocytes after cAMP treatment. Intraperitoneal injection with cAMP rapidly induces the translocation of HOPS from nucleus to cytoplasm. (a–c): in normal liver (control) HOPS appears in nucleus of hepatocytes. (d–f): HOPS shuttles from nucleus to cytoplasm 60 minutes after cAMP treatment. (g–i): HOPS is relocalized in nucleus 120 minutes after cAMP treatment

first round of mitosis was observed in the regenerating liver of CREM-deficient mice [98]. The expression of cyclins such as cyclin A, B, D1, E, and cdc2 is shifted in accord with the observed delay of mitosis. Moreover, the expression of proteins involved in neoglucogenesis that are liver-specific markers, such as TAT or PEPCK, is modified. Similarly, there is an altered expression profile of c-fos, an immediate early gene activated during liver regeneration. The protein c-Fos dimerizes with c-Jun to constitute the transcription factor AP-1, a key regulator of the proliferative process. These data suggest that lack of CREM could cause AP-1 deregulation, leading to a delay in hepatocyte proliferation. Finally, it is worth noting that all the genes whose expression is aberrant in the CREM-deficient mice, including cyclin A and D1, TAT, PEPCK, and c-fos, have CRE sequences in their promoter regions [99, 100]. CREB was shown to mediate the response to prostaglandins in liver regeneration [101], and to be involved in the control of tumor progression in hepatocellular carcinoma. Indeed, CREB appears to control angiogenesis and induces cell survival. Importantly, in the liver, CREB has been identified as a privileged target of the pX protein of hepatitis B virus in hepatitis disease. The pX protein is able to bind the unphosphorylated form of CREB and activate transcription, bypassing

the canonical requirement of Ser-133 phosphorylation. The result of this aberrant regulation is that CREB transcription and its downstream targets are continuously activated. Constitutive upregulation of cAMP-responsive genes has been suggested to be responsible for the development of liver cancer after viral infection [102, 103]. Finally, it has been shown that functionally impaired mitochondria activate a signaling pathway that has as its terminal target CREB phosphorylation [104]. Thus, CREB involvement in proliferation may be linked to aberrant energy metabolism and mitochondrial dysfunction.

CONCLUSION Accumulating evidence has demonstrated that the cAMP transduction pathway plays an essential role in the control of liver function. Through the regulation of gene expression cascades, members of the CREB family of transcriptional regulators govern a remarkable array of physiological and pathophysiological functions in the liver. In addition, these regulators participate in a functional interplay with transcription factors of different classes to finely control the expression of liver genes. The proliferation of residual hepatocytes during liver regeneration

530

THE LIVER: CONCLUSION

125 TAT PEPCK SDH

100

G-6-Pase CREM c-fos

A.U.

75

50

25

0 0

5

10

15

20

25

30

Time after hepatectomy (hours)

Figure 34.5 Expression profiles of genes bearing at least one CRE site in their promoter regulatory region at different times after partial hepatectomy. This representation reveals the various kinetics of gene expression in response to the proliferative stimulus. The relative level of expression of the various genes is not respected and data are expressed in arbitrary units (A.U.). TAT, tyrosine aminotransferase; PEPCK, phosphoenolpyruvate carboxykinase; SDH, serine dehydratase; G-6-Pase, glucose-6-phosphatase; CREM, cAMP-responsive element modulator dBt-cAMP 0

2

4

8

Partial Hepatectomy 0 2 5 8 12 18 24 48 72

hours

CREM

Figure 34.6 CREM gene induction in the liver. Expression of ICER in rat liver was analyzed after dBtcAMP i.p. injection (lane 2–4) (lane 1 saline injection) and in the regenerating liver at different times after partial hepatectomy (lane 5 non-operated control). The arrowhead corresponds to the ICER-specific transcript generated from the CREM P2 promoter

relies on an orchestration of various signaling pathways and the consequent specialized and timed gene expression. At this level, the timing of gene expression appears to be crucial to directing the synchronized mitotic waves of hepatocytes proliferation. Although the role of cAMP in a number of physiological and pathological functions is now established, many open questions remain: how does cAMP-responsive transcription intersect with circadian

regulation? How is chromatin remodeling achieved synchronously during liver regeneration? What is the role played by cross-talks among signaling pathways? In what way does cAMP influence the cell cycle of proliferating hepatocytes? Various experimental approaches will be needed to further elucidate the molecular mechanisms governing normal and pathological liver physiology. The available

34: REGULATORY PATHWAYS OF LIVER GENE EXPRESSION: THE CENTRAL ROLE OF CYCLIC AMP

technology of targeted disruption of specific genes in the mouse by homologous recombination, especially if coupled to tissue-specific and conditional mutagenesis, will very likely provide a great deal of novel, essential information about the specialized role played by cAMP and CREB family members in the liver.

ACKNOWLEDGMENTS We wish to thank all the members of Sassone-Corsi and Servillo’s laboratories for help and discussion. Work in G. Servillo’s laboratory is supported by grants from Associazione Italiana Ricerca sul Cancro (AIRC), Fondazione Guido Berlucchi, and Only the Brave Foundation. Work in the laboratory of P. Sassone-Corsi is supported by the Cancer Research Coordinating Committee of the University of California, the Institut National de la Recherche Scientifique et Medicale (France), and the National Institute of Health.

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35

AMPK: Central Regulator of Glucose and Lipid Metabolism Maria M. Mihaylova and Reuben J. Shaw Molecular and Cell Biology Laboratory, The Salk Institute for Biological Studies, La Jolla, CA, USA

AMPK STRUCTURE AND MECHANISM OF ACTIVATION A fundamental requirement of all cells is that they couple the availability of nutrients to signals emanating from growth factors to drive proliferation only when nutrients are in sufficient abundance to guarantee successful cell division. Even in non-dividing cells, nutrients in the environment supply the necessary building blocks for cellular metabolism and survival, and fuel the bioenergetic needs of the cell by providing substrates to produce intracellular 5′ -adenosine triphosphate (ATP) to be used for all cellular processes. When nutrient levels fall, ATP levels fall and, unless ATP-consuming biosynthetic processes are curtailed, a critical shortage of ATP will cause catastrophic cellular demise. Eukaryotic cells all share a highly conserved metabolic checkpoint that acts as a sensor of ATP levels in the cell – the 5′ -adenosine monophosphate (AMP)-activated protein kinase (AMPK). As intracellular

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

ATP levels fall due to pathological stresses such as glucose or oxygen shortages, osmotic stress, or disruptions of glycolysis or mitochondrial oxidative phosphorylation, these will all result in decreased ATP. Upon activation under low-ATP conditions, AMPK acts a metabolic checkpoint in the cell, suppressing ATP-consuming biosynthetic processes while stimulating ATP-generating processes to repair the initiating loss of ATP [1]. Upon activation, AMPK initiates acute effects on metabolic enzymes and also prolonged adaptions in glucose and lipid metabolism through modulation of transcriptional programs of metabolic enzymes. In addition to ubiquitous roles as an energy checkpoint, AMPK also plays additional key roles in glucose and lipid metabolism in specialized metabolic tissues in mammals and higher eukaryotes such as liver, muscle, and adipose [2]. Thus AMPK governs not only cellular energetics, but indeed overall organismal bioenergetics by coordinating the response between tissues to nutritional input. AMPK was first discovered as a mammalian protein kinase that is activated by changes in intracellular adenosine

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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THE LIVER: AMPK STRUCTURE AND MECHANISM OF ACTIVATION

Length T172

α1

βγ Binding

Kinase domain

Major Site Chromosome of Expression Location

α-CTD

550

ubiquitous

5p11-14

α-CTD

552

liver, muscle widespread

1p31

T172

Kinase domain

α2

glycogen binding β1

GBD

270

ubiquitous

12q24.1-.3

β2

GBD

272

ubiquitous?

1q21.1

Bateman domain 1 AMP γ1 γ2 long

γ3 short

Bateman domain 2 AMP AMP

CBS1

CBS2

CBS3

CBS4

331

ubiquitous

12q12-14

CBS1

CBS2

CBS3

CBS4

569

ubiquitous

7q36

γ2 short

CBS1

CBS2

CBS3

CBS4

328

ubiquitous

7q36

γ3-NTD

CBS1

CBS2

CBS3

CBS4

489

skeletal muscle

2q35

CBS1

CBS2

CBS3

CBS4

464

skeletal muscle

2q35

γ2-NTD

γ3 long

αγ Binding

γ3-NTD

Figure 35.1 Human AMPK subunit isoforms. Domain structure, expression pattern, and alternative splice isoforms of the two catalytic kinase (α) isoforms, the two beta regulatory subunits which contain a glycogen-binding domain (GBD), and the three genes encoding the gamma subunits, each of which contains four CBS domains which directly bind to AMP as drawn

nucleotide levels [3]. However, it was not until later that the yeast ortholog SNF-1 (sucrose non-fermenting complex) was annotated from a Saccharomyces cerevisiae mutant screen for cells that failed to grow on non-fermentable carbon sources or sucrose [4, 5]. The AMPK is an obligate heterotrimeric kinase complex composed of a catalytic (α) subunit and two regulatory (β and γ) subunits. AMPK is activated under conditions of energy stress, when intracellular ATP levels decline and intracellular AMP increases, as occurs during nutrient deprivation or hypoxia [2]. Upon energy stress, AMP binds directly to tandem repeats of crystathionine-β-synthase (CBS) domains in the AMPK γ subunit. Binding of AMP is thought to prevent dephosphorylation of the critical activation loop threonine in the α subunit [6]. In mammals, there are seven mammalian genes encoding each of the α, β, and γ subunits, allowing for 12 distinct heterotrimeric variations (Figure 35.1). There are two genes encoding catalytic subunits, α1 and α2, two regulatory β, and three γ subunits that participate in the heterotrimer. Of these, γ3 appears mostly skeletal muscle specific and α2 appears to be most highly expressed in key metabolic tissues including muscle and

liver. The catalytic α1 and α2 subunits contain a kinase domain within their N-terminus, and also a critical region for binding β and γ subunits within their C-terminus. All kinases possess an activation loop which is often a target for upstream kinases, creating a conformation change that allows substrate access to the catalytic pocket. The activation loop of the AMPKα subunits contains a single threonine (Thr172 in mammalian AMPKα) that is the key regulatory site whose phosphorylation is absolutely required across all species for AMPK activation. Its been shown that α1 catalytic isoform is found mainly in the cytoplasm, whereas the α2 isoform appears to be nuclear in some cell types. In liver, phosphorylation of both the α1 and α2 subunits accounts for half of the total AMPK activity and there appears to be no preferential binding of the α1 and α2 subunits with the different β or γ subunits [7]. Although the crystal structures of Saccharomyces cerevisiae Snf1 and the human α2 kinase domains have been revealed [8, 9], efforts for crystallizing the remaining components of the enzyme still remain. The regulatory β subunits contain conserved glycogen-binding domains (GBDs) and a region required for binding the α and γ subunits in the C-terminus. The γ subunits contain four

35: AMPK: CENTRAL REGULATOR OF GLUCOSE AND LIPID METABOLISM

repetitive CBS domains. Each CBS domain binds one molecule of AMP [10]. When AMP levels rise within the cell, AMP binds allosterically to the CBS domains of the γ subunit, which is hypothesized to change the conformation of the heterotrimer protecting the phosphorylation on Thr172 by dephosphorylation which it actively undergoes at all other times.

STRESS, HORMONES, AND THERAPEUTICS ACTIVATE AMPK Animals, in their multi-cell complexity and multi-organ utilization, have evolved intricate mechanisms to sense and initiate immediate responses to energy requirement or nutrient deprivation. Multiple studies have now shown that cellular stress caused by starvation or exercise can activate AMPK. In single-cell eukaryotes and also mammalian cell culture conditions, AMPK is activated in response to nutrient depletion of glucose or oxygen (hypoxia). It is now evident that as complex organisms developed, various circulating hormones gained function to act as whole-organism sensors and are capable of turning on AMPK in response to metabolic stresses such as starvation. One well-known adipokine, adiponectin, has been shown to activate AMPK in liver, leading to fatty acid oxidation and a decrease in blood glucose levels, consistent with previous findings that adiponectin is able to suppress hepatic glucose production [11]. In addition to hormonal input from adiponectin, AMPK activity has been shown to be modulated by leptin, resistin, ghrelin, adrenaline, and cannabinoids [12]. Exercise is another metabolic stress that has been shown to activate AMPK in response to muscle contraction. Such activation may be due to increased AMP to ATP ratios caused by movement and muscle contraction, correlating with studies in mice where electrical muscle stimulation increased AMP levels and turned on AMPK [13]. Two different classes of drugs for treatment of diabetes mellitus type 2, biguanides such as metformin [14] and thiazolidinediones (TZDs) such as rosiglitazone [15] and pioglitazone [16] have been shown to activate hepatic AMPK, most likely through perturbation of mitochondrial ATP output via mild inhibition of complex I of the mitochondrial respiratory chain. Metformin is a biguanide that has been structurally modified from guanide, a naturally occurring compound found in the French lilac (Galega officinalis). Although it was not until 1918 that biguanides were discovered to have a blood glucose-lowering effect, people have been using the French lilac to ameliorate diabetes-like symptoms since the Middle Ages [17]. Bitter melon or Momordica charantia, like the French lilac, has been used for hundreds of years in traditional Chinese medicine to treat many different ailments, and it

537

was not until recently that scientists isolated triterpenoid compounds from Momordica charantia that are able to activate AMPK and facilitate fatty acid oxidation and glucose utilization when administrated in mice [18]. In a recent study, AMPK has also been shown to become active in response to resveratrol, a polyphenol that is found in the skin of red grapes, certain nuts, and berries and has been linked to longevity in model organisms such as yeast, nematodes, fruit flies, fish, and mice. Further, it was shown that resveratrol could mimic the benefits of dietary restriction in mice by ameliorating the fatty liver phenotype and increasing insulin sensitivity in animals fed a high-calorie diet [19]. Although direct mutations in AMPK have not been found in diabetic patients, studies have shown that AMPK is implicated in various pathways deregulated in such metabolic disorders and makes it an attractive therapeutic target in the treatment of such diseases.

UPSTREAM REGULATORS OF AMPK: LKB1 AND CAMKK There was evidence for an upstream kinase-activating AMPK as early as 1978 [20] and in the years that followed scientists labored intensively to identify the kinase responsible for phosphorylation and activation of catalytic subunits α1 or α2. However, it was not until 1996 that an “AMP-activated protein kinase kinase” (AMPKK) was partially purified from rat liver and shown to phosphorylate AMPK on Thr172 [21]. Quickly after the budding yeast genome was completed, three upstream kinases, Sak1 (Pak1), Elm1, and Tos3, were identified by whole genome screening methods and were shown to act upstream of the yeast AMPK ortholog, the SNF1 complex. When these kinases were genetically knocked out in yeast, the results yielded the same phenotype as an snf1 mutant, again placing them upstream of the SNF1 complex [22, 23]. In the human genome, the closest related kinases to the yeast ones were found to be calmodulin-dependent protein kinases, CaMKKα and CaMKKβ, and the protein kinase LKB1 (liver kinase B1). Several groups simultaneously showed that LKB1 [1, 24, 25] and CAMKKβ [26–28] were indeed the upstream kinases acting on AMPK in mammals and were capable of phosphorylating both AMPKα1 and AMPKα2 subunits on Thr172. Interestingly, LKB1 was first identified in humans as a serine/threonine tumor suppressor kinase that is defective in the cancer-predisposing Peutz–Jeghers syndrome [29]. In mammalian cells, LKB1 exists in a complex with two other proteins, STRAD (sterile-20-related adaptor) and MO25 (mouse protein-25) [24], and when bound to these accessory proteins it is stabilized and constitutively activated. Recent data suggest that when AMP nucleotides bind allosterically to the Bateman domains in the γ subunit of AMPK complex, conformational changes occur that protect the LKB1-mediated phosphorylation Thr172

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THE LIVER: REGULATION OF ACUTE METABOLIC RESPONSE–ENZYMES IN LIPOGENESIS

in the α subunit [6], although effects on localization and complex formation between LKB1 and AMPK following energy stress remain to be fully explored. In 2005, scientists showed that LKB1 is indeed the major upstream AMPK kinase in liver [30] and that genetic deletion of hepatic LKB1 almost completely reduces hepatic AMPK activity. Lack of LKB1 in mouse liver rapidly leads to hyperglycemia and increased levels of gluconeogenic and lipogenic gene expression. It was also shown in these animals that activation of AMPK by the anti-diabetic therapeutic metformin is dependent on LKB1 and, in the absence of hepatic LKB1, metformin is unable to lower blood glucose levels [30]. However, it is important to note that AMPK is not the only substrate of LKB1. LKB1 similarly phosphorylates the activation loop of a family of 14 kinases all related to AMPK, also resulting in their activation [31]. Importantly, of these 14 LKB1-dependent kinases, only AMPKα1 and AMPKα2 are activated under low-ATP conditions, probably because only they interact with AMPKγ which contains the AMP-binding domains [32]. Little is known about what stimuli direct LKB1 towards any of these AMPK-related kinases and current evidence suggests that LKB1 is constitutively active and these other kinases may be regulated through phosphorylation at other sites outside their activation loops. Collectively, these findings map AMPK on the axis of a major tumor suppressor pathway and provide an interesting link between cancer and metabolism. In addition to LKB1, the CaMKKs, CaMKKα and CaMKKβ, also regulate AMPK activity, although only in response to calcium flux and not in response to changes in AMP, which appears to work completely through LKB1 based on genetic knockout and RNA interference (RNAi) studies. Genetic deletion of LKB1 dramatically reduces AMPK activation in liver, suggesting little role for CAMKK in this tissue, although in hypothalamic neurons controlling food intake CAMKKβ (CAMKK2) appears to be the dominant upstream kinase for AMPK [33]. This is consistent with the fact that CaMKKα and CaMKKβ are most highly expressed in neurons, whereas LKB1 is more ubiquitously expressed.

DOWNSTREAM TARGETS I: REGULATION OF ACUTE METABOLIC RESPONSE– ENZYMES IN LIPOGENESIS In the liver, AMPK phosphorylates and regulates multiple downstream targets involved in lipogenesis and lipid homeostasis (Figure 35.2). One of the first identified downstream targets of AMPK was acetyl-coenzyme A carboxylase (ACC) [34]. ACC is an enzyme involved in the generation of fatty acid precursor malonyl-CoA, a key metabolite in the regulation of energy homeostasis. Two genes encoding two different ACC isoforms are found in

mammals – ACC1 and ACC2 – and they appear to have distinct tissue specificity. It has been shown that ACC1 and ACC2 control the synthesis of two different pools of malonyl-CoA production. ACC1 is thought to suppress the production of malonyl-CoA used in fatty acid synthesis whereas ACC2 stimulates fatty acid oxidation (reviewed in [35]). AMPK inhibits both ACC1 and ACC2 through direct phosphorylation of their homologous residues Ser79 in ACC1 and Ser218 in ACC2. Down-regulation of ACC1 activity leads to reduced malonyl-CoA levels and a decrease in lipogenesis. AMPK phosphorylation of ACC2 inhibits its enzymatic activity and decreases cellular levels of malonyl-CoA levels, which leads to direct inhibition of mitochondrial fatty acid uptake and increased fatty acid oxidation and ATP production through carnitine palmitoyltransferase 1 (CPT-1) (reviewed in [12]). Interestingly, it has also been shown that calorie restriction can increase AMPK activity, causing a decrease in fatty acid synthesis or up-regulation of fatty acid oxidation through inhibition of ACC. In a recent study, various AMPK-activating polyphenol compounds led to lowering of lipid accumulation in HepG2 cells grown in high glucose and inhibited atherosclerosis in diabetic LDLR−/− (low-density lipoprotein receptor) mice treated with the various polyphenols [36]. In addition to reporting ACC as a downstream target of AMPK, HMG-CoA reductase was also found to be one of the first key downstream substrates [34, 37]. It had already been known for over 10 years at that point that HMG-CoA reductase kinase [3-hydroxy-3-methylglutaryl-CoA reductase (HMGR)] activity was regulated by an upstream kinase [20], however that kinase had not yet been discovered. Today, it is known that HMG-CoA reductase is a rate-limiting enzyme involved in the production of cholesterol and other isoprenoids, and more specifically functions in converting HMG-CoA to mevalonic acid. By phosphorylating HMGR, AMPK blocks anabolic or ATP-consuming processes such as cholesterol synthesis in order to preserve intracellular ATP levels. Strikingly, the AMPK phosphorylation site in HMGR is conserved throughout eukaryotes, including plants. Glycerol-3 phosphate acetyltransferase (GPAT) is the first designated enzyme in the glycerolipid synthesis pathway and its regulation by AMPK can ultimately lead to inhibition and a decrease in overall triacylglycerol synthesis. It has been shown that 5aminoimidazole 4-carboxamide-1β-4-ribofuranoside (AI CAR), a cell-permeable AMP mimetic that binds directly and activates AMPK, can inhibit the activity of mitochondrial GPAT in cultured hepatocytes by increasing fatty acid oxidation. In the same study, it was determined that when recombinant AMPK is incubated with hepatically derived mitochondria, there is a decrease in GPAT activity, demonstrating that the effect of AICAR on GPAT activity originally observed is most likely due to AMPK activation [38]. In a later study, another

35: AMPK: CENTRAL REGULATOR OF GLUCOSE AND LIPID METABOLISM

Low Nutrients (glucose, O2 )

Exercise adiponectin

LKB1

Mitochondrial inhibitors: TZDs, biguanides

STRAD MO25

Ghrelin, cannibinoids Resistin

AICAR (amp mimetic) AMP

Resveratrol, polyphenols

539

CAMKKβ Ca2+

Glycolysis inhibitors: 2-DG γ

Abbott A769662

β

P AMPKα

Glycolysis/ Glucose Uptake iPFK2 Ser461 TBC1D1 Ser237

Fatty Acid Oxidation ACC2 Ser221

Transcriptional Control of Glucose Metabolism

Lipid Synthesis HMG CoR Ser872 ACC1 Ser79

Insulin Sensitivity Protein Synthesis TSC2 Ser1387 Raptor Ser792 IRS1 Ser794

CRTC2 FOXO3 p300 AREBP HNF4a Chrebp

Ser171 Ser413 Ser89 Ser470 Ser313 Ser568

Figure 35.2 The AMPK signaling pathway. AMPK is phosphorylated on its kinase activation loop threonine and activated by two distinct upstream kinases in response to different stimuli. LKB1 activates AMPK in response to all stimuli that lower intracellular ATP and increase AMP. CAMKKb activates AMPK in response to calcium flux in an AMP-independent manner. AMPK is activated physiologically by exercise, low nutrients such as lowered glucose or lowered oxygen, and hormones including ghrelin, leptin, adiponectin, and cannabinoids. Leptin is reported to activate AMPK in peripheral tissues but to inhibit AMPK in the central nervous system through poorly understood mechanisms. In addition, AMPK is activated by agents that disrupt ATP production by inhibiting or poisoning the mitochondria, including uncouplers, or agents that inhibit glycolysis such as the glucose analog 2-deoxyglucose that is a competitive inhibitor for hexokinase. Additional pharmacological agents that activate AMPK include resveratrol and related polyphenols and also the cell-permeable AMP-mimetic AICAR and the first small-molecule direct activator Abbott A-769662. Upon activation, AMPK serves to inhibit anabolic, ATP-consuming biosynthetic processes such as protein, lipid, and glucose synthesis, while up-regulating catabolic processes to generate ATP production, including increased glycolysis, glucose uptake, and fatty acid oxidation. AMPK modulates cell growth and insulin sensitivity through the mTOR signaling pathway. All current best established in vivo direct AMPK substrates and their AMPK phosphorylation sites are listed. All the sites listed conform to the identified optimal AMPK substrate motif

group demonstrated that a decrease in GPAT activity occurs in adipose and liver tissues, but not muscle, in rats following 30 minutes of exercise on a treadmill, concurrent with changes in AMPK, ACC, and MDC (malonyl-CoA decarboxylase) activity [39]. Recently, it has been shown that the increase in ACC, decrease in MDC, and increase in GPAT activity correlate with the decrease in AMPK activity in rats starved for 48 hours and re-fed a carbohydrate chow diet [40]. The activity of another enzyme involved in the glycerolipid synthesis pathway, diacylglycerol O-acyltransferase (DGAT), was also shown to increase in rat livers upon re-feeding. Importantly, however, a direct AMPK phosphorylation site in GPAT, DGAT, or MDC has not yet been identified, so the observed effects may be mediated indirectly. Taken together, these studies suggest that AMPK could play an important role in changes in hepatic lipid metabolism and, more specifically, promote a shift from catabolic to anabolic processes in response to cues from a starved to fed transition state in animals. This further reinforces

the idea that AMPK acts as a “fuel gauge” and, when it senses that the cell is running low on ATP, it protects the cell by turning on processes that generate ATP and turning off processes that consume ATP.

DOWNSTREAM TARGETS II: REGULATION OF METABOLIC ADAPTATION: CONTROL OF TRANSCRIPTION In response to changes in AMP:ATP ratios, AMPK can rapidly regulate downstream targets via phosphorylation. However, in addition to these fast post-translational modifications, AMPK can also promote long-term transcriptional changes and re-program transcription of certain genes in response to the cellular state. These effects are thought to be mediated by the direct phosphorylation of

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THE LIVER: REGULATION OF METABOLIC ADAPTATION: CONTROL OF TRANSCRIPTION

14-3-3 P

LKB1

CRTC2 cytoplasm

P P

SIK P

AMPK

p300

PGC1α

P P ChREBP

nucleus

AMPK

14-3-3

P

CRTC2 FASN

CREB

PGC-1α

Inhibition of Lipogenesis

P FoxO3

HNF4a

P AREBP

PEPCK G6Pase

Inhibition of Gluconeogenesis

Figure 35.3 LKB1- and AMPK-mediated control of metabolic transcriptional programs. AMPK and its related family members SIK1 and SIK2 (not shown) phosphorylate a common set of substrates including the CREB coactivator CRTC2 (CREB-regulated transcriptional coactivator 2), previously known as TORC2 (transducer of regulated CREB 2), and the histone acetyltransferase p300. Phosphorylation of CRTC2 creates a 14-3-3 docking site which then results in 14-3-3-mediated nuclear export of CRTC2. This cytoplasmic sequestration of CRTC2 by AMPK or SIK kinases causes an inhibition of CREB-dependent transcriptional targets including key mediators of gluconeogenesis such as the PGC-1α coactivator. In addition to suppressing PGC-1a mRNA expression, AMPK also directly phosphorylates a handful of transcription factors (FOXO3, HNF4a, and AREBP) that directly bind to the promoters of the two key gluconeogenic enzymes PEPCK and G6Pase. In addition to these gluconeogenesis regulators, AMPK is also reported to phosphorylate Chrebp, a key lipogenic transcription factor that controls levels of fatty acid synthase (FASN), acetyl-CoA carboxylase 1 (ACC1), and l-pyruvate kinase mRNA

sequence-specific transcription factors and transcriptional co-activators by AMPK. The two best-studied transcriptional programs controlled by AMPK in liver are gluconeogenesis and lipogenesis. A number of direct substrates for AMPK in gluconeogenesis have been identified. Interestingly, two different co-activators which modulate CREB (cyclic AMP-responsive element binding protein)-dependent transcription in response to glucagon induced by fasting are the histone acetyltransferase p300 and the CREB-regulated transcription coactivator 2 (CRTC2) (also known as TORC2) [41–43]. CRTC2 is a critical rate-limiting transcriptional regulator of gluconeogenesis in mice via effects on hepatic CREB targets, including the PGC-1α promoter. When AMPK is active, it can phosphorylate CRTC2 on residue Ser171, which in turn allows CRCT2 to associate to 14-3-3 proteins and be sequestered from the nucleus to the cytoplasm, an event that shuts off transcription of gluconeogenic enzymes [44, 45]. If AMPK activity is decreased, hypophosphorylated CRTC2 can conversely move into the nucleus where it binds to transcriptional co-activator CREB and promotes

transcription of PGC1α and downstream gluconeogenic targets phosphoenolpyruvate carboxykinase (PEPCK) and glucose 6-phosphatase (G6Pase) (Figure 35.3). Strikingly, both p300 and CRTC2 have been shown to be phosphorylated at the same regulatory sites by either AMPK or its related family member SIK1 (salt-inducible kinase 1), both of which are LKB1 dependent. In mice lacking hepatic LKB1, CRTC2 is hypophosphorylated and predominantly nuclear compared with wild-type mice [30]. Furthermore, LKB1–/– mice in liver showed dramatic increases in fasting blood glucose levels, which were greatly attenuated upon introduction of a short hairpin RNA (shRNA), reducing CRTC2 levels in the liver, reinforcing the idea that gluconeogenesis in liver is controlled by LKB1-dependent kinases and that CRTC2 is a key downstream target of these kinases. Consistent with these findings in the LKB1 liver-specific knockout mice, animals bearing a liver-specific knockout of the AMPKα2 isoform also exhibited elevated hepatic glucose output, glucose intolerance, and impaired leptin- and adiponectin-regulated hepatic glucose production [46]

35: AMPK: CENTRAL REGULATOR OF GLUCOSE AND LIPID METABOLISM

Table 35.1 Mouse models of AMPK/LKB1 function in liver Mouse model

Metabolic phenotype

Reference

AMPKα1 knockout AMPKα2 knockout

None Glucose uptake in muscle Hyperglycemia, low insulin Hyperglycemia

[48] [49]

AMPKα2 liver knockout LKB1 liver knockout

Glucose intolerance Hyperlipidemia Hyperglycemia

[48] [46]

[30]

Glucose intolerance Hyperlipidemia

(Table 35.1). Similarly, short-term adenoviral expression of constitutively active AMPKa2 in murine liver resulted in hypoglycemia [47]. Future studies will be needed to dissect the temporal and spatial regulation the contexts in which AMPKα2 or SIK1 controls these key modulators of CREB and gluconeogenesis. Interestingly, SIK1 itself is a CREB target, providing a time-delayed mechanism to attenuate chronic CREB-dependent transcription. Phosphorylation of p300 and CRTC2 by AMPK and SIK kinases may be key effectors of metformin in the control of type 2 diabetes. How AMPK phosphorylation of p300 may regulate its many other downstream interacting transcription factors key to hepatic metabolism remains to be examined, although mice lacking LKB1 or AMPK in liver provide excellent tools for such future studies. In addition to direct phosphorylation of co-activators, AMPK also phosphorylates the hepatocyte nuclear factor 4 alpha (HNF4α), which is a key transcription factor of the nuclear receptor superfamily that binds to the promoters of the two key gluconeogenic enzymes, PEPCK and G6Pase. It is hypothesized that AMPK phosphorylation of HNF4α on residue Ser304 decreases the protein’s stability by interfering with its ability to dimerize and bind to DNA and may further promote its degradation [50, 51]. In addition to PEPCK and G6Pase, HNF4α controls gene expression of glucose transporter GLUT2 and glycolytic enzymes such as aldolase B and liver-type pyruvate kinase (L-PK), which are diminished when AMPK is activated by AICAR in hepatocytes [50]. It has been also shown that a subset of patients with MODY (maturity-onset diabetes of the young) form of diabetes harbor mutations in HNF4α [52]. Interestingly, HNF4α liver-specific knockout mice do not develop hyperglycemia, unlike MODY patients, but do develop lipid accumulation, consistent with altered triglyceride levels in MODY patients [53]. These findings further reinforce the possible role of AMPK in lipid and glucose homeostasis through modulation of downstream transcriptional regulators such as HNF4α. In addition to HNF4α, in a recent report AMPK was shown to phosphorylate directly another transcription

541

factor, named the AICAR response-element binding protein (AREBP), that directly binds the PEPCK promoter [54]. AMPK phosphorylation of AREBP on Ser470 reduces its ability to bind DNA and in turn lowers expression of PEPCK. Thus, like HNF4α, AMPK phosphorylation of AREBP controls its ability to bind to DNA and promote transcription of downstream target genes. In addition to effects on gluconeogenesis, AMPK activation is also known to inhibit hepatic lipogenesis. While some of that is due to acute effects on lipogenic enzymes, as previously discussed, it is also known that AMPK activation leads to decreased transcription of key hepatic lipogenic enzymes, including fatty acid synthase (FASN) and ACC1. Two sequence–specific transcription factors are known to co-regulate many of these lipogenic enzymes: the sterol-responsive binding protein (SREBP-1c) and the carbohydrate-response binding protein (ChREBP). ChREBP is highly expressed in liver and has essential roles in glucose-induced transcription of L-PK, in addition to its effects on lipogenic enzyme promoters. Phosphorylation by AMPK on Ser568 of ChREBP [55] promotes decreased DNA binding, which causes a decrease in the transcription of glycolytic and lipogenic genes. It is known now that glucose metabolism can be repressed by fatty acids, which act as another source of energy when glucose needs to be preserved. This effect has been named the fatty acid “sparing effect” on glucose. Like ChREBP, SREBP-1 is also regulated by AMPK, although whether this is direct or not is unclear at present [14, 56]. Due to its rate-limiting effects on lipogenic enzyme expression, SREBP-1 has been linked to insulin resistance, dyslipidemia, and type 2 diabetes [57, 58]. In one study, treatment of rat hepatocytes with AMPK activators such as metformin or AICAR led to suppressed SREBP-1 mRNA expression, and also lowered hepatic mRNA levels for SREPB-1-controlled genes FAS and S14 [14]. This suggests that AMPK activation through metformin can inhibit the expression of lipogenic genes. Indeed, metformin treatment or overexpression of an activated allele of AMPK was found to be sufficient to reduce triglyceride content in insulin-resistant HepG2 cells [59]. Mice lacking hepatic AMPK function due to liver-specific LKB1 deletion show elevated SREBP1 and SREBP1 target genes, resulting in lipid accumulation and hepatic steatosis [30]. As seen from the above-mentioned studies, AMPK plays a major role in the gluconeogenic transcriptional program through p300, CRTC2, HNF4α, and AREBP, and also the lipogenic transcriptional program through ChREBP and SREBP-1. AMPK has a unique interaction with peroxisome proliferator-activated receptor α and δ. Peroxisome proliferator-activated receptor α (PPARα) was the first PPAR out of the three isotypes (PPARα, -γ, and -δ) to be isolated and belongs to the nuclear receptor superfamily of ligand-inducible transcription factors. PPARs act as lipid sensors and are activated by a number of ligands derived from dietary intake or cellular biosynthesis

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pathways. When ligands or agonists such as eicosanoids and fatty acids bind to PPARα–retinoid X receptor (RXR) complex, conformational change occurs and PPARα–RXR receptor heterodimer dissociates to from the co-repressor complex, allowing it to bind to the promoter region of transcription targets [60]. AMPK binding to the PPARα appears to enhance downstream transcription of PPARα targets, which in turn promotes fatty acid oxidation [61]. Similarly, AMPK associates with PPARδ- and AMPK activation promotes PPARδ-dependent transcription in skeletal muscle [62]. The regulation of PPARδ-dependent transcription by AMPK has been shown to be involved in the reprogramming of muscle fiber type during training, and small molecular activators of PPARδ and AMPK were shown to promote endurance running in mice [62]. A recent study suggested that AMPK-dependent regulation of PPARα in liver controls the ability of glucagon to suppress hepatic lipogenesis and to promote fatty acid oxidation [63]. Thus AMPK activation of PPARα-dependent transcription likely plays a key role in the metabolic reprogramming towards fat utilization in the liver during fasting, although the direct substrate that AMPK phosphorylates to control PPAR activation remains unknown at this time. Finally, studies have hinted that AMPK may play a more global role in transcription through direct phosphorylation and regulation of transcriptional repressors histone deacetylases class IIa (HDACs IIa) enzymes [64] and transcriptional activator histone acetyltransferase p300 [41]. It has been shown that AMPK can phosphorylate class IIa HDAC5 on residues Ser259 and Ser498 in human primary myotubes, promoting 14-3-3 binding and dissociation from DNA binding transcription partner myocyte enhancer factor-2 (MEF2), which in turn allows expression of downstream target genes [64]. Although direct regulation of class IIa HDACs by AMPK has not yet been implicated in liver metabolism, this provides an attractive postulate where AMPK can simultaneously control multiple downstream transcriptional events in response to upstream metabolic stresses. Such analysis will not happen without challenges, since it has been shown that multiple upstream kinases are capable of phosphorylating the same critical residues of HDAC 5 [65].

DOWNSTREAM TARGETS III: REGULATION OF CELL GROWTH AND INSULIN SIGNALING VIA mTOR Environmental factors cue cells to cease growing and dividing when conditions are unfavorable. These mechanisms are conserved from the smallest and simplest of eukaryotes to the most complex ones. When nutrients are scarce, cellular energy sensor AMPK becomes activated and inhibits energy-demanding processes such as protein

synthesis and cell growth. One of the ways in which AMPK accomplishes that task is by negatively regulating the mammalian target of rapamycin (mTOR) pathway. mTOR is a serine/threonine kinase highly conserved in all eukaryotes, and is a central regulator of cell growth. Whereas AMPK is active under nutrient-poor conditions and inactive under nutrient-rich conditions, mTOR is activated in the inverse pattern. In higher eukaryotes, mTOR activation requires positive signals from both nutrients (glucose, amino acids) and growth factors. mTOR, like its budding yeast orthologs, is found in two biochemically and functionally discrete signaling complexes [66]. In mammals, the mTORC1 complex is composed of four known subunits: raptor (regulatory-associated protein of mTOR), PRAS40, mLST8, and mTOR. The mTORC2 complex contains rictor (rapamycin-insensitive companion of mTOR), mSIN1, PRR5/Protor, mLST8, and mTOR [67]. Signaling from mTOR complex 1 (mTORC1) is nutrient sensitive, acutely inhibited by the bacterial macrolide rapamycin, and controls cell growth, angiogenesis, and metabolism. In contrast, mTORC2 is not sensitive to nutrients, nor acutely inhibited by rapamycin, and its known substrates include the hydrophobic motif phosphorylation sites in AGC kinases including Akt and PKC family members. Downstream of the AMPK- and rapamycin-sensitive raptor–mTOR (mTORC1) complex are its two wellcharacterized substrates, 4EBP1 (eIF4E-binding protein 1) and the p70 ribosomal S6 kinase. Phosphorylation of 4EBP-1 by mTORC1 suppresses its ability to bind and inhibit the translation initiation factor eIF4E. mTORC1 mediates phosphorylation of S6K at a Thr residue in a hydrophobic motif at the C-terminus of the kinase domain. A specific motif (TOS motif) found in both 4EBP1 and S6K was shown to mediate direct binding of these proteins to raptor, allowing them to be phosphorylated in the mTORC1 complex. Mechanistic details of how mTORC1 regulates the assembly of translational initiation complexes via a number of ordered phosphorylation events were recently discovered [68]. mTORC1-dependent translation is known to control a number of specific cell growth regulators, including cyclin D1, the HIF-1α transcriptional factor, and c-myc, which in turn promote processes including cell cycle progression, cell growth, glycolysis, and angiogenesis, all contributing to enhanced tumorigenesis [67]. Upstream components of the mTORC1 complex were initially discovered through classical cancer genetics. The TSC2 tumor suppressor tuberin and its obligate binding partner hamartin (TSC1) are mutated in a familial tumor syndrome called tuberous sclerosis complex (TSC). TSC patients are predisposed to widespread benign tumors termed hamartomas in kidney, lung, brain, and skin. Genetic studies in Drosophila and mammalian cells identified the TSC tumor suppressors as critical upstream inhibitors of the mTORC1 complex. TSC2 (also known as

35: AMPK: CENTRAL REGULATOR OF GLUCOSE AND LIPID METABOLISM

tuberin) contains a GTPase activating protein (GAP) domain at its C-terminus that inactivates the small Ras-like GTPase Rheb, which has been shown to associate with and directly activate the mTORC1 complex in vitro [69]. Loss of TSC1 or TSC2 therefore leads to hyperactivation of mTORC1. Phosphorylation of TSC1 and TSC2 serves as an integration point for a wide variety of environmental signals that regulate mTORC1 [70]. One of the key activators of the mTORC1 pathway is PI3-kinase, which plays a key role in promoting cell growth and insulin-mediated effects on metabolism. PI3-kinase activates the serine/threonine kinase Akt, which directly phosphorylates and inactivates both TSC2 and an inhibitor of the mTORC1 complex named PRAS40 [69, 71]. In addition to these growth factor cues that activate mTORC1, the complex is rapidly inactivated by a wide variety of cell stresses, thereby ensuring that cells do not continue to grow under unfavorable conditions. One of the unique aspects of the mTORC1 complex is that unlike many of the aforementioned growth factor activated kinases, it is dependent on nutrient availability for its kinase activity. Withdrawal of glucose, amino acids, or oxygen leads to rapid suppression of mTORC1 activity even in the presence of full growth factors [70]. Upon LKB1and AMP-dependent activation of AMPK by nutrient loss, AMPK directly phosphorylates the TSC2 tumor suppressor on conserved serine sites distinct from those targeted by other kinases, which constitutes one mechanism by which glucose and oxygen control mTORC1 activation [72–75]. Interestingly, TSC2 orthologs are absent from

543

lower eukaryotes such as S. cerevisiae and C. elegans and mammalian cells lacking TSC2 still remain partially sensitive to AMPK activation, indicating that there may be an alternative and more ancient back-up mechanism which allows AMPK to inhibit cell growth and proliferation through the mTORC1 pathway. Collectively, these observations prompted the discovery of yet another novel mechanism of inhibition. The mTOR kinase exists in a complex consisting of mLST8/Gbl, PRAS40, and the scaffold protein raptor. In a recent study, it was shown that AMPK is able to phosphorylate raptor directly on two conserved residues, Ser722 and Ser792, which in turn induces binding to 14-3-3 proteins and inactivation the mTORC1 complex [76]. Taken together with previous studies, these findings indicate that AMPK directly phosphorylates both TSC2 and raptor to inhibit mTORC1 activity by a dual-pronged mechanism (Figure 35.4). Importantly, metformin treatment of mice leads to robust phosphorylation of raptor Ser792 in murine liver, an effect which is ablated in LKB1 liver-specific knockout mice [76]. LKB1 liver-specific knockout mice which lack AMPK activity in liver exhibit hyperactivation of mTORC1 signaling in the liver, including increased phosphorylation of S6K1 and 4EBP1 under ad libidum fed conditions [30]. In addition, hormones that activate AMPK in liver, including glucagon [77] and adiponectin [78], have been reported to suppress mTORC1 signaling. Beyond effects on cell growth, mTOR also has effects on lipid metabolism that may be particularly important in liver. One key regulator of lipogenesis is the

Insulin R IRS1/2 PI3K mTOR rictor

LKB1

PIP3

AMP

FOXO GSK3

Akt TSC2 TSC1

AMPK

-3 14

Srebp1

-3 -3

Rheb raptor

rapamycin

14

-3

metformin

PTEN

PRAS40 mTOR

4EBP1

S6K

EIF4E

S6

Figure 35.4 AMPK regulates the mammalian target of rapamycin TOR pathway to control cell growth and insulin sensitivity. AMPK directly phosphorylates the TSC2 (tuberous sclerosis 2) tumor suppressor and the key mTOR scaffold subunit raptor to inhibit the activity of the mTOR–raptor kinase complex (mTORC1) towards its downstream substrates 4EBP1 and S6K1. PI3K activity activates mTORC1 by Akt-dependent phosphorylation of TSC2 and the mTORC1 inhibitor PRAS40. Under conditions of nutrient excess, overstimulated mTORC1 and its substrate S6K both directly phosphorylate the insulin receptor substrate 1 and 2 proteins, resulting in their degradation. Thus too much mTORC1 activity attenuates insulin signaling, leading to cellular insulin resistance. AMPK reverses this resistance by inactivating mTORC1 and can also directly phosphorylate IRS1 itself. mTORC1 has also recently been shown to play a role in the control of lipogenesis through regulation of SREBP1

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aforementioned SREBP-1 transcription factor. Recently, mTORC1 signaling was shown to be required for nuclear accumulation of SREBP1 and the induction of SREBP1 target genes [79]. Consistent with previous results with metformin, treatment with the AMPK activators AICAR and 2DG or the mTORC1 inhibitor rapamycin resulted in suppression of nuclear SREBP1 accumulation [79]. In future studies, it will be important to define how much of the lipid-reducing effects of AMPK are due to direct phosphorylation of lipogenic enzymes such as ACC, and how much are due to effects on SREBP-1 or ChREBP-dependent transcription through effects of AMPK on mTORC1. One final aspect of liver physiology that may be under control of the AMPK–mTOR axis is insulin signaling. Insulin sensitivity is compromised in a number of metabolic disorders such as type 2 diabetes and obesity and is linked to hyperglycemia, artherosclerosis, and hypertension. Patients with insulin resistance present with a decreased ability to uptake and utilize glucose in peripheral tissues such as skeletal muscle, leading to an increase in total blood glucose levels and over-production of pancreatic insulin to compensate this imbalance. One mechanism through which metformin or other AMPK activators can lead to increased insulin sensitivity in the whole animal is by reducing hepatic gluconeogenesis and increasing glucose uptake in the muscle, thereby reducing circulating blood glucose levels and lowering the demand on the pancreas to produce more insulin. This reduction in circulating insulin levels is often accompanied by a decrease in insulin resistance [80]. In addition to this organismal hormone response, AMPK activation has also been shown to increase insulin sensitivity in cultured cells through a recently discovered molecular mechanism whose full details are yet to be defined. A major route by which excess nutrients down-regulate insulin signaling leading to cellular insulin resistance is through hyperactivation of the mTORC1 complex. Excess glucose leads to hyperactivation of mTOR via suppression of AMPK, and excess fats and excess amino acids also act to hyperactivate mTORC1 [81]. The mTOR–raptor complex, along with its key downstream substrate S6K, have been shown to phosphorylate directly insulin receptor substrate-1 and -2 (IRS1 and IRS2), leading to their proteosome degradation. The same is observed under conditions of hyperinsulinemia, as insulin signaling itself also leads to increases in mTORC1, activity as described above. The net effect is a negative feedback loop whereby too much mTOR–raptor activity leads to hyperphosphorylation of IRS1/IRS2 and suppression of PI3-kinase and Akt signaling downstream of the insulin receptor [81]. This nutrient-induced hyperactivation of mTORC1 and consequent down-regulation of Akt signaling is observed in a cultured cell systems and also in peripheral tissues of mice on a high-fat diet. Illustrating its importance downstream of mTORC1 in the IRS1/IRS2 inhibition, this effect is lost in peripheral tissues from an S6K1−/− mouse [82].

As one of the key direct substrates of AMPK is raptor and TSC2, AMPK activation leads to inhibition of mTORC1 and its phosphorylation of IRS1 Ser636/639 and negative feedback loop on PI3-kinase/Akt signaling. Exogenous LKB1 expression in human embryonic kidney 293 (HEK293) cells and metformin treatment of human hepatocellular carcinoma HepG2 cells can suppress phosphorylation of IRS-1 on Ser636/639 and induce Akt phosphorylation [83]. In addition to suppressing phosphorylation of IRS-1 by mTORC1, AMPK has also been shown to phosphorylate IRS1 itself directly, although the functional consequence on that phosphorylation event on IRS function remains uncertain [83, 84]. Taken altogether, these studies demonstrate a mechanism by which AMPK activation can promote PI3-kinase/Akt activity while simultaneously reducing mTORC1 activity. This provides cells with a negative feedback switch that integrates upstream signals from both nutrients and growth factors and allows the cells maintain energy homeostasis and insulin sensitivity.

THERAPEUTICS AND FUTURE PERSPECTIVES It is remarkable that therapeutics aimed at AMPK activation can modulate so many distinct and key targets involved in the metabolic syndrome and type 2 diabetes. Results from the past 5 years have begun to decode some of the critical upstream and downstream components of the AMPK pathway, and many of the therapeutic effects of the most widely used type 2 diabetes therapeutic in the world—metformin—can be traced to effects downstream of AMPK in the liver. The genetic proof that AMPK is a key target for metformin came from studies of the LKB1 liver-specific knockout mice. As noted previously, these mice lack nearly all AMPK activity in liver, although they retain normal AMPK activation in other organs, and metformin does not stimulate their hepatic AMPK activity. These mice or their wild-type littermates were placed a high-fat diet to induce a diabetic-like state of hyperglycemia and, upon treatment with a metformin regiment, the wild-type mice displayed reduced fasting blood glucose levels – an effect completely lost in the LKB1 liver-specific knockout mice [30]. Given that AMPK and LKB1 are expressed throughout the body, why might hepatic AMPK be so central to metformin action? A potential answer was derived from studies of type 2 diabetics who were non-responders to metformin therapy. Single nucleotide polymorphism (SNP) analysis led to a genetic locus encoding a solute carrier named Oct1 (organic cation transporter 1), which the authors went on to show is critical for metformin uptake into hepatocytes and happens to be almost exclusively expressed in hepatocytes [85]. Collectively, these studies suggest that the ability of metformin to regulate blood glucose may be largely due to effects on hepatic gluconeogenesis, and this requires the

35: AMPK: CENTRAL REGULATOR OF GLUCOSE AND LIPID METABOLISM

LKB1 (and presumably AMPK) pathway. It remains to be determined how much of a role the LKB1-dependent AMPK-related kinases also play in suppressing hepatic gluconeogenesis or lipogenesis. Future studies will also be needed to detail which kinases downstream of LKB1 are most critical for hepatic lipogenesis and what the precise substrates downstream of these kinases are that control lipogenesis. By activating AMPK, metformin suppresses many key targets in hepatic lipogenesis and hepatic gluconeogenesis, reducing the need for the pancreas to over-produce insulin, in addition to restoring cell-autonomous insulin sensitivity through direct effects on IRS1 phosphorylation and PI3K signaling downstream of insulin. Looking into the future, a more directly targeted AMPK activator may prove the most valuable therapeutically. Recently, the first small-molecule activator of AMPK was reported, originating from a screen of over 700 000 compounds using purified AMPK heterotrimeric complexes [86]. The compound Abbott A-769662 induced AMPK activation in cultured cells and in mice and a 5 day administration in diabetic ob/ob mice decreased plasma glucose and plasma and hepatic triglyceride levels. These effects were mirrored by a reduction in hepatic gluconeogenic and lipogenic enzyme mRNA expression. Thus, as one single therapeutic biochemical target, AMPK-activating compounds such as metformin, AICAR, and Abbott A-769662 can re-program glucose and lipid metabolism with 20 downstream effectors the same as would require 20 separate compounds targeted to each of these effectors. These effects of these therapeutics, taken together with the phenotypes of genetic loss of AMPK or LKB1 function in mice, suggest that AMPK holds great promise as a target for future diabetes drug development.

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Liver Regeneration Nelson Fausto and Jean S. Campbell Department of Pathology, School of Medicine, University of Washington, Seattle, WA, USA

HISTORICAL NOTES AND DEFINITIONS Liver regeneration is popularly associated with the legend of Prometheus in ancient Greece. The legend, as it refers to the liver, is best summarized by this sentence from Hesiod’s Theogony: “. . . [Zeus] sent a long-winged eagle to feed on [Prometheus’] liver, which was immortal; but whatever this long-winged bird ate during the day, grew during the night again to perfection.” Prometheus’ ordeal lasted for 30 years according to some, and 100 years according to others, and ended only when Herakles (Hercules) had slain the eagle with an arrow. Daily regeneration for 30 years or a full century is too good to be true, and most scholars doubt that the Greeks had any real knowledge of liver regeneration [1], although there are dissenting opinions [2]. Was the myth of Prometheus then an extremely successful instance of pre-scientific serendipity? Apparently not, as the myth was just one of many that dealt with immortal gods, and the replacement and multiplication of their immortal organs [3]. The best known of these myths include the regenerative capacity of the heads of the nine-headed Hydra (an amazing example of compensatory growth), the tale of the three hags who swapped a single detachable eye and a tooth between them (the earliest example of organ transplantation), and the regeneration of a whole new Dionysus from his heart (after his successful cloning, Dionysus became known as “the twice-born”). Poor Tityus, the giant son of Zeus, had it worse than Prometheus, as his liver was eaten by two vultures, but somehow his story is rarely used as an example of liver regeneration. Note that the period when these legends were written, somewhere between 800 and 400 BCE (Before Christian Era), was the golden age of the liver; it

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

had a privileged position as the site of the soul, which it eventually relinquished to the heart, and as an instrument of divination in the practice of hepatoscopy. Nevertheless, this exalted image of the liver was not associated with knowledge of its anatomy and function. Studies about the anatomy, blood circulation, and function of the human liver started much later, with the performance of autopsies, and, according to the records, human vivisection, by Herophilus in Alexandria, around 300 BCE, about five centuries after Hesiod’s writing. These historical comments point out that, contrary to the popular view, real knowledge about liver regeneration is not ancient. It actually dates from animal experiments, mostly with rabbits, which were initiated during the last 20 years of the nineteenth century. Regeneration may mean different things to different people, but we should consider two types of regeneration: epimorphic regeneration and tissue regeneration [3, 4]. Epimorphic regeneration consists of the regrowth of amputated structures, and generally involves the regrowth of appendages, or the growth of fragments of a dissociated organism into a complete organism. The hallmark of appendage regrowth is the formation of a blastema at the amputation site. The blastema can be thought of as a regeneration bud, composed of stem and dedifferentiated cells, which give rise to all of the tissues of the lost appendage [5]. In contrast, tissue regeneration is a process of growth that occurs in wounded tissues without the formation of a blastema. Although most mammalian tissues have the capacity to undergo some level of compensatory growth after damage, only the liver, and to some extent the pancreas, can restore its mass after major tissue loss. This is best illustrated by the growth of the liver after resection of two-thirds of its mass, the standard partial hepatectomy (PH) procedure commonly used to study liver regeneration.

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Noting that a salamander can regenerate almost any structure that has been cut off, such as limbs, tail, lower jaw, lens, retina, heart ventricules, small intestine, and other tissues [4, 6], Tanaka places the issue into perspective by asking a basic question about regeneration: “if they can do it, why can’t we?” Based on the brief description of the two types of regeneration, some hypotheses can be proposed, but no clear answers are available. It is thought that if a blastema could develop in the amputation stump of the leg or arms of a mammal, regrowth of the appendage would take place. As to the reasons why blastemas do not form in the amputation surface of mammals, several possibilities exist. For instance, cells in the amputation stump may have lost the capacity to dedifferentiate; dedifferentiation, if it takes place, may fail to reactivate a developmental pattern of gene expression; or the stroma at the amputation site may prevent cell proliferation and the regrowth of tissues. Another issue is the failure of tissue regeneration to restore the original organ architecture after injury, as most healing processes in mammalian tissues result in fibrosis and not in the restoration of the original tissue. The problem most likely lies in the degree of damage to the tissue stroma. In the liver, acute injury that does not damage the stroma heals perfectly without fibrosis, even if the injury causes widespread necrosis of hepatocytes. In contrast, fibrosis develops after repeated injuries that produce only a small degree of necrosis but damage the hepatic stroma. From these observations, it would appear that lack of robust regeneration in other organs besides the liver may in part be a consequence of the rapid activation of stromal cells and the deposition of collagen after injury, which may inhibit the replication of parenchymal and progenitor cells in these tissues. Several general reviews on liver regeneration are available [7–10]. In this chapter, we discuss some selected aspects of the cellular and molecular biology of liver regeneration after PH.

REGENERATIVE CAPACITY AND THE LIVER AND BODY WEIGHT EQUILIBRIUM Unless specified otherwise, PH consists of the removal of two-thirds of the liver of mice or rats. Liver regeneration is a process of compensatory growth and not of true regeneration, as the liver regains its mass after PH by the increase in the mass of the lobes remaining after the operation, without regrowth of the resected lobes. The increase in liver mass is usually a consequence of hepatocyte hyperplasia, but other mechanisms may occur (described below). A remarkable feature of liver regeneration is that it involves synchronized waves of DNA replication in hepatocytes and non-parenchymal cells, and that it

terminates when the liver weight regains its original weight (±10%). The relationship between liver weight and body weight has been convincingly illustrated by experiments in dogs that received liver transplants whose sizes were either large or small relative to the body weight of the recipient animal [11]. The transplanted small livers grew until an optimal liver weight/body weight ratio was achieved, whereas the large livers did not show a growth response, and even decreased in size after transplantation. A similar decrease in the mass of human livers has been described after the transplantation of large livers that do not fit well into the abdominal cavity. These livers generally decrease in size and permit the perfect closure of the abdominal cavity in about 1 week. In human liver transplantation, small livers grow faster than large livers, but there is a lower limit for the volume of the transplanted liver, and also for the volume of the residual liver after tumor resection, below which metabolic complications and liver failure ensue, producing the “small for size syndrome” [12–14]. In both rats and mice, regeneration is mostly completed by 10–14 days after PH. Human livers have a great proliferative capacity, as indicated by the gain in volume of the liver of the donor and of the transplanted liver in living donor transplantation. After right lobe resection, the donor liver representing 45–50% of the original organ grows to 70–79% at 1 month and 80–90% 6 months after transplantation [15, 16]. For unknown reasons, the growth of the transplanted liver is more rapid than that of the liver of the donor, and the latter generally does not grow beyond 85–90% of the original volume. If the liver can grow so well after PH, would it maintain this capacity after repeated hepatectomies? This type of experiment presents difficult technical challenges. Ingle performed eight PHs with 21 day intervals between operations, but liver failure and infection were a major problem [17]. However, Simpson and Finckh performed a successful experiment in which they carried out five partial hepatectomies in rats with intervals of 5–7 weeks between operations [18]. The amount of the original liver remaining after each operation corresponded to 34% after the first PH, and then to 20, 14, 8, and 4% after subsequent PHs (Figure 36.1). Despite the anatomical changes created by the repeated surgeries, the liver regained its original weight after each operation (for technical reasons, the liver growth after the fifth operation reached 72% of the original weight). This is one of the best illustrations of the enormous regenerative capacity of the liver, which was maintained regardless of its size and anatomical form. These older observations are in a sense a prelude to the hepatocyte transplantation experiments of Grompe and colleagues [19], who showed that adult hepatocytes could repopulate mouse livers during six serial transplantations, going through a minimum of 69 doublings (corresponding to a 7.3 × 1020 -fold expansion) at the end of the serial transplants.

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and number of lobules, in about equal proportions. However, after subsequent PHs, growth was due to an increase in the number of lobules, which were estimated to have multiplied about 18-fold after five operations.

DNA REPLICATION IN HEPATIC CELLS AND HEPATOCYTE PLOIDY

Figure 36.1 Liver growth after repeated hepatectomies. Simpson and Finckh [18] performed five partial hepatectomies in rats with intervals of 5–7 weeks between operation. After each operation the liver regained its original weight (weight regain was about 75% at the fifth operation). The figure shows a drawing of the original liver (red) and the portion of the original liver remaining after the third operation (yellow). The remaining portion corresponds to about 14% of the non-operated liver

LIVER GROWTH: BIGGER LOBULES OR MORE LOBULES? From the early days of studies on liver regeneration, there has been a debate on whether regeneration after PH consists of an enlargement of liver lobules or an increase in the number of lobules. In rodents, the first wave of hepatocyte proliferation after PH produces clusters of 10–14 cells, without the formation of sinusoids and additional extracellular matrix. Penetration of these clusters by stellate and sinusoidal cells occurs 2–4 days later, and is associated with laminin deposition [20], but it is not clear whether there is formation of new lobules. Simpson and Finckh addressed this issue in the repeated hepatectomy experiments already described [18]. After each operation, the histology of the liver was similar, but analysis of the vascular elements of the liver revealed that they increased progressively after each operation. A detailed analysis of the growth of liver vascularization after repeated hepatectomies showed that after a single two-thirds PH the growth of the liver involved increases in both the size

Liver regeneration after PH is dependent on the proliferation of hepatocytes, and does not involve the activation of hepatic progenitor cells (oval cells). After PH, DNA replication first occurs in hepatocytes, with peaks at 34–36 hours after the operation in mice, and 20–24 hours in rats. The timing of these peaks varies in different strains of rats and mice, and also with the age of the animals. In 3–4-week-old rats, a second peak of DNA replication in hepatocytes is detectable approximately 13 hours after the first replication peak [21]. Hepatocyte mitosis follows hepatocyte DNA replication, reaching a maximum approximately 6–10 hours after the peak of DNA replication. In rats, DNA replication of “littoral cells” (a population composed of Kupffer and endothelial cells) occurs later, starting at least 1 day after PH, and reaches a maximum at approximately 16–28 hours after the peak of DNA replication in hepatocytes [22]. Studies using peroxidase markers revealed that Kupffer cells show a sharp peak of mitosis at 48 hours after PH, while the mitotic peak for endothelial cells occurs at 4 days after PH [23]. No clear explanation has been found for the delay in DNA replication of Kupffer and sinusoidal cells after PH. It is possible that these cells respond to mitogens released by hepatocytes, but it has not been determined whether Kupffer and endothelial cell replication after PH is dependent on hepatocyte DNA replication. The liver is a major reservoir of cells of the innate immune system, namely natural killer(NK) cells, natural killer T cells (NKT) and γδT cells. In mouse liver, NKT cells constitute 20–30% of lymphocytes, and this proportion increases to 50–60% after PH [24]. Both NK and NKT are believed to inhibit liver regeneration, but it has also been shown that T cells are the source of lymphotoxins (LTs) that may initiate liver regeneration (see below). Changes in hepatocyte ploidy occur after PH. In young rats, the overall hepatocyte ploidy is distributed as 19.5% diploids, 75% tetraploids, and 5.3% octoploid cells. Approximately 20% of hepatocytes are binucleated (about 18% with two diploid nuclei and 2% with two tetraploid nuclei). The ploidy of mononucleated hepatocytes is distributed as 20% diploid, 56% tetraploid, and 3.5% octoploid. After PH, the proportion of binucleated cells decreases to less than 5%, while there is an overall increase in ploidy characterized by a decrease in 2n hepatocytes and an increase in 4n and 8n mononuclear hepatocytes

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[25–28]. Recent data demonstrate that mononucleated tetraploid hepatocytes in fumarylacetoacetate hydrolase (FAH)-deficient mice may undergo reduction to 2n ploidy, and it has been proposed that this ploidy reduction may increase hepatocyte diversity [29]. The opposite appears to occur in the regenerating liver, in which there is a decrease in 2n hepatocytes and an overall ploidy increase, suggesting that after PH there is a strong pressure for DNA replication, but not for cytokinesis. Hepatic weight gain and restoration of the liver mass after PH can, however, occur without hepatocyte hyperplasia. Two interesting examples are the recovery of liver mass after PH in mice deficient in Skp2, which results in the accumulation of the cell cycle inhibitor p27, and in mice lacking separase, a protein that triggers sister chromatid disjunction at anaphase. In both models, there is little hepatocyte proliferation after PH, but the liver regains its mass by an enlargement and polyploidization of hepatocytes, with cells with DNA content of 18n, 32n, or higher constituting almost 20% of the total [30, 31]. Similarly, hepatocytes with telomere dysfunction caused by a deficiency in the shelterin component TRF2 restore liver mass after PH through endoreduplication without cell proliferation. This defect causes an increase in the proportion of cells in prophase compared to wild type mice, and an increase in ploidy such that hepatocytes with 8n or higher DNA content constitute more than 40% of the total [32]. Hepatic function appears to be similar regardless of the mechanism by which the liver regains its weight after PH, that is, by cell proliferation or endoreduplication. This is not entirely surprising, because global patterns of gene expression in 2n, 4n, and 8n hepatocytes do not differ among ploidy classes, and quantitative changes in gene expression between hepatocytes of different DNA content vary by less than twofold [33]. The functional equivalency between tetraploid and octoploid hepatocytes has also been demonstrated by their similar capacities to repopulate injured livers [34]. In the regenerating liver after PH, not only is liver function in a hypertrophic, highly polyploid liver not significantly different from that in a hyperplastic liver, but also the mechanisms that terminate regeneration after mass restoration appear to function equally well in the hypertrophic liver.

STAGES OF LIVER REGENERATION: PRIMING AND PROGRESSION A very large number of genes are activated after PH, which could imply that most of what is needed for hepatocyte DNA replication might be simultaneously activated shortly after the operation. However, many genes that are reported to be activated within minutes after PH do not show a sustained response at these early times, but their sustained expression occurs many hours later. Our

own studies and those from other laboratories have indicated that liver regeneration is a multi-stage process that unfolds gradually, and that different mediators act at different stages of regeneration. It proved to be particularly useful for the analysis of liver regeneration to divide the process into a priming phase, corresponding to the passage of hepatocytes from G0 to G1, and a progression phase, that corresponds to the progression of hepatocytes through the G1 phase of the cell cycle. Throughout these phases, and continuing during the first 2 days after PH, there is wide expression of metabolic genes that are required to maintain liver functions. Some of these functions, and in particular ribosome synthesis and protein translation, are also required for hepatocyte growth and replication. We consider priming as a reversible phase, mostly regulated by cytokines, which is necessary but not sufficient for DNA replication after PH. Progression leading to DNA replication is regulated by growth factors, and is irreversible. However, its various stages have complex regulatory mechanisms needed to overcome restriction points in the cell cycle. Hepatocyte entry into the S phase in the regenerating liver is a cell autonomous process, that is, it is regulated by a program intrinsic to the hepatocyte. Weglarz and Sandgren studied DNA replication in partially hepatectomized mice with chimeric livers composed of mouse hepatocytes and transplanted rat hepatocytes, and relied on the difference in the timing of the peak of DNA replication after PH in rats, which is about 12–16 hours earlier than in mice [35]. In the chimeric mouse livers, the transplanted rat hepatocytes replicated about 12 hours before the mouse hepatocytes, despite being in the same environment as the mouse cells. The implication of these data is that entry into S phase from late G1 is regulated by an intrinsic program of the hepatocyte that does not depend of extrinsic factors.

Priming Bucher and colleagues [36] reported many years ago that rats that had been pretreated with growth hormone, or been sham-operated for as long as 3 days before PH, showed an acceleration of liver regeneration, with an earlier onset of DNA replication. They concluded that hepatocytes of these rats were prepared (“primed”) for replication, and were capable of responding faster to PH. Webber et al. observed a similar acceleration of liver regeneration in rats treated with tumor necrosis factor (TNF) for as long as 24 hours before PH [37]. These experiments showed that “primed” hepatocytes had initiated the regenerative process without advancing to DNA replication, and suggested that these cells were no longer in G0 quiescence but were ready to enter G1. Hepatocytes in the normal liver show very weak responses to growth factors such as hepatocyte growth factor(HGF), transforming growth factor alpha (TGFα),

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activation, the initiation of a cascade of events leading to activation of the transcription factors NFκB and STAT3, and the immediate early gene response that has the notable participation of the protooncogenes fos, c-jun, and myc. These changes prepare hepatocytes for replication, but they are reversible, and are not a determinant of replication. Moreover, in contrast to DNA replication, which is proportional to the extent of the tissue deficit after PH, most early changes of gene expression also occur after a one-third PH, a procedure that does not lead to appreciable hepatocyte replication [40, 41]. Priming is not necessary for hepatocyte DNA replication mediated by some chemical agents, as demonstrated by Columbano and colleagues [42]. These agents probably act on G1 cell cycle components, or directly on DNA, and can bypass cytokine and protooncogene activation. It is of great practical interest that whereas the extent of liver regeneration after PH decreases in older animals, old rats injected with the hepatic mitogen TCPOBOB do not show a decrease in hepatocyte replicative capacity [42].

and EGF. Although it is possible to induce hepatocyte DNA replication in normal mouse liver by HGF infusion, high doses are needed. For instance, Patijn et al. obtained BrdU labeling in 20% of normal mouse liver hepatocytes by continuous infusion of HGF into the portal vein for 5 days at a dose of 5 mg kg−1 per day [38]. However, some cytokines, and in particular TNF, can greatly enhance the sensitivity of hepatocytes in the normal liver to the replicative effect of growth factors. Rats given a single injection of TNF and infused with 40 µg of either HGF or TGFα during a 24 hour period show a 4–5-fold increase in BrdU labeling of hepatocyte DNA over controls, and rats that receive both growth factors and TNF can reach labeling indices of more than 50% [37]. Thus, in vivo, hepatocytes do not seem to be able to jump directly into the cell cycle, but need a transition period that we have called priming, that corresponds to the G0 to G1 transition and makes them “competent” to enter into G1 and progress to DNA replication. The term “competence,” as it applies to the replication of quiescent cells, was coined by Pardee from his extensive studies of fibroblast proliferation in culture [39]. He also showed that the G0 to G1 transition and the progression through G1 required external factors, and that DNA replication would only occur when growth factors were activated sequentially, and in a particular order. The notion that liver regeneration involves priming and progression phases is thus consistent with carefully controlled cell cycle analysis experiments. Among many other changes in gene expression (see below), priming after PH is associated with cytokine

CYTOKINES AND THE INITIATION OF LIVER REGENERATION Using mouse knockout models, it has been shown that the tumor necrosis factor type 1 receptor (TNFR1), lymphotoxin β receptor (LTβR), and the C3a and C5a components of the complement system (Figure 36.2) are important

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Figure 36.2 Diagram indicating the involvement of components of the innate immune system in the initiation of liver regeneration. Shown in the diagram are TNF, lymphotoxin (LT) and complement pathways and their receptors, and also their distribution in Kupffer cells, lymphocytes, and hepatocytes. Also indicated is that activation of these pathways converges into NFκB and STAT3 activation, which is inhibited by SOCS3

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for the triggering of the priming phase of liver regeneration and the increase in IL-6 after PH [43–45]. All of these systems initiate a cascade of events that activate NFκB, increase the production of IL-6, promote STAT-3 phosphorylation, and activate suppressor of cytokine signaling (SOCS)3 during the first 6–8 hours after PH. The same sequence of events is also activated after PH by the Toll-like receptor (TLR) intermediate Myd88 (myeloid differentiation factor 88), as Myd88-deficient mice show practically no IL-6 elevation after PH [46, 47]. It is not clear whether these multiple systems act independently or through interactive modes to stimulate IL-6 production. However, given the overlap between the components of these pathways, it is most likely that there is active cross-talk between them during the early steps of liver regeneration.

TNF, lymphotoxins, and the complement system TNF and LTs are members of the TNF superfamily that comprises more than 20 ligand–receptor systems [48]. There are two distinct forms of LTs, LTα, a component of the soluble, homotrimeric form LTα3, and LTβ, a component of the membrane-anchored heterotrimeric form LTα1β2 (for simplicity the two forms will be referred to as LTα and LTβ). Both TNF and LTα can bind TNFR1 (Figure 36.3), which may explain why mice lacking TNF generally show no defects in liver regeneration, whereas defective regeneration occurs in TNFR1 knockout mice, and also in TNF−/− , LTα−/− double knockout mice [49]. LTβR is activated by LTβ, which interacts with TNF to stimulate IL-6 production and STAT-3 phosphorylation. The sharing of various ligands by TNFR1 and LTβR suggests that there is a LTa1b 2

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Figure 36.3 Lymphotoxin and TNF ligands and receptors. Both TNF and LTα, which is a soluble trimer, bind to TNFR1 and TNFR2. LTβ (LTα1β2) is membrane-anchored, and binds only to the LTβ receptor. Redrawn from [18]

close connection between the activities of these receptors. One interesting difference in the activation of NFκB from these receptors is that in lymphocytes, TNFR1 activation involves the formation of the p50–p65 NFκB1 complex, while signaling through LTβR results in the formation of p52–p65 NFκB2. It is not known whether these different modes of NFκB activation occur in the liver, and if they may have biological significance. Regarding components of the complement system, earlier work reported a significant defect in DNA replication and high mortality of C3- and C5-deficient mice after PH [43]. More recent results showed that C3 knockout mice have no abnormal mortality after PH, and have a defect in DNA replication that is less severe than originally described [50]. The proteolytic activation of C3 after PH does not involve the “traditional” upstream pathways of C3 convertase activation. Instead, it has been proposed that C3 activation after PH may be mediated through plasmin or other components of the coagulation system [50].

The toll-like receptor system The TLR system plays an important role in host defense against injury, and is a strong activator of the innate immune system (Figure 36.4). So far, 13 TLR receptors have been identified; with the exception of TLR3, TLRs 1–9 signal through Myd88 after ligand binding. As already stated, Myd88 knockout mice fail to activate the IL-6 cascade after PH [46, 51], and it is then logical to hypothesize that defects in one or more TLRs would cause a similar defect. In particular, it is of special interest to determine whether the lipopolysaccharide (LPS)-binding receptor TLR4 is involved in the triggering of liver regeneration after PH, as LPS has been considered an important agent in this process [52]. A delay in liver regeneration occurs in mice (C3H/HeJ strain) that have a natural mutation in TLR4 [53], but mice made deficient in TLR4 or CD14 (a TLR4 co-receptor) by genetic techniques did not show defects in IL-6 production. Production of IL-6 after PH was also unaffected in TLR2, TLR9, and TLR2/4 double knockouts [46, 47]. These results led to a paradoxical situation in which deficiency in Myd88, the intermediate signaling molecule for most TLRs, causes an almost total inhibition of IL-6 production after PH, but deficiency of TLRs, the receptors that provide the signals for Myd88 activation, does not. However, recent data indicates that signaling through TLR4 may be responsible for about 50% of the IL-6 increase after PH. In any case, despite the lack of knowledge about the mechanisms of Myd88 activation after PH, it is clear that it has a key role in the control of cytokine production at the start of liver regeneration. Also, despite the uncertainties described, it is possible that LPS participates in the triggering of cytokine production and complement activation after PH.

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Figure 36.4 TNF and LPS signaling. The figure shows the steps involved in LPS binding to TLR4, with CD14 as a co-receptor. Ligand binding activates Myd88, eventually leading to activation of NEMO in the IKK complex. Signaling through TNFR1 also activates the IKK complex through a different route that does not involve Myd88. In addition, TNFR1 activation can lead to caspase activation and apoptosis through the TRAD/FADD pathway

The NFκB system The role of NFκB during liver development and regeneration has been extensively investigated using genetically modified mice. NFκB forms an inactive complex with IκB, and its activation requires the phosphorylation of IκB, which is accomplished by the IκB kinase complex known as IKK (Figure 36.4). The IKK complex contains IKK1, IKK2, and NEMO. NFκB activation in the liver has multiple effects, including antiapoptotic, proliferative, and pro- or anti-inflammatory outcomes. Mice deficient in the p65 component of NFκB (known as RelA) die at embryonic day 14–15 with massive apoptosis. However, apoptosis is prevented in animals that are deficient for both RelA and TNF [54] or RelA and TNFR1 [55], demonstrating that NFκB protects the embryonic liver against TNF-mediated apoptosis. In adult livers, inhibition of NFκB activation in hepatocytes by a dominant-inhibitor IκB mutant did not interfere with liver regeneration after PH [56] but led to apoptosis after TNF injection. In cultured oval cells, the IκB mutant blocked the TNF-mediated production of IL-6, STAT3 activation, and DNA replication [57]. Mice in which partial NFκB inhibition was produced by hepatocyte-specific deletion of IKK2 showed an acceleration of liver regeneration after PH, probably caused by the early activation

of NFκB in non-parenchymal cells, and a strong innate immune system response [58]. Complete NFκB inactivation in hepatocytes by direct genetic methods, or by deletion of NEMO, greatly increased the sensitivity of hepatocytes to TNF-mediated apoptosis. These data show that in the regenerating liver after PH, NFκB is activated in both hepatocytes and non-parenchymal cells, and that it transmits proliferative, antiapoptotic, and innate immunity signals. The parallel activation of NFκB in hepatocytes and non-parenchymal cells in the first few hours after PH is an important feature of the cross-talk between different cell types at the early stages of liver regeneration [58, 59].

The IL-6 system The role of IL-6 in the regenerating liver is a subject of debate. It has been reported that IL-6 knockout mice have severe defects in DNA replication after PH [10]. However, other experiments showed a decrease in apoptosis in IL-6 knockouts at 60–96 hours after PH with only a small defect in DNA replication [60], or increased mortality at 2–3 days after PH without replicative defects [61]. Injections of IL-6 can restore DNA replication in partially hepatectomized TNFR1 knockout mice [45], but IL-6 hyperstimulation inhibits cell cycle progression after

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PH [62]. On the other hand, “trans signaling” by soluble IL-6R (a “super” IL-6 agonist) has a strong stimulatory effect on hepatocytes [63], and systemic exposure to IL-6 can cause massive liver growth [64]. Although IL-6 increases in patients with chronic liver diseases, it is thought that it has a protective role in experimental liver injury [65]. One way to make sense of these often contradictory data is to conclude that IL-6 has both protective and proliferative effects in the liver, but that these effects are strictly dependent on dosage and mode of administration. As an example of the influence of some of these variables on IL-6 effects, subcutaneous injections (long acting) can protect IL-6 knockout mice from mortality after PH, but intravenous injections (short acting) did not [61]. Adding to the complexity of the mechanisms that regulate IL-6 activity in the regenerating liver is the possibility that stem cell factors acting through its c-kit receptor may account for some of the effects of IL-6 [66, 67].

The IL-6 family and the JAK/STAT pathway IL-6 signals by forming a complex with the receptor subunit gp80 (Il-6R), which then binds to the gp130 receptor, promoting the dimerization of the intracellular domains of two gp130 molecules (Figure 36.5). Receptor-associated Janus kinases (JAK1, JAK2, and TYK) bind to and phosphorylate tyrosine residues of the activated gp130 receptor. The phosphotyrosine residues of the receptors provide

docking sites for signaling molecules such as STAT1, STAT3, and ras. Although all of these and also other molecules may be activated by the phosphorylated gp130 complex, STAT3 appears to be one of the main products of this pathway in the regenerating liver after PH. STAT3 is phosphorylated at specific tyrosine residues, and undergoes homodimerization. The dimer migrates to the nucleus, binds to specific DNA sequences, and stimulates the transcription of a large number of genes that encode protooncogenes and cell cycle components (c-jun, c-myc, cyclin D1, among others), acute-phase proteins (such as serum amyloid A, the C3 component of complement, and fibrinogen), antiapoptotic proteins (such as Bcl-2 and BCl-xL). STAT3 also reduces gluconeogenesis, and decreases insulin levels [68, 69].

The suppressor of cytokine signaling (SOCS) system Given the widespread effects of the activation of the IL-6/STAT3 pathway (also referred to as the JAK/STAT pathway) during the first 6–8 hours after PH, it is logical to ask how the pathway is regulated. In the regenerating liver after PH, Campbell et al. [70] and Riehle et al. [71] showed that the IL-6/STAT3 pathway is regulated primarily by SOCS3, which binds to phosphotyrosine 759 of the gp130 receptor [72]. SOCS3 messenger RNA (mRNA) is transiently induced by about 40-fold

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Figure 36.5 IL-6 pathways and SOCS3 inhibition. Binding of IL-6 to the gp130 receptor activates both the JAK/STAT system and MAP kinases. SOCS3 represses gp130 signaling by functioning as a feedback inhibitor. Reproduced from Heinrich et al., Biochem J , 2003, 374, 1

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Figure 36.6 SOCS3 deficiency affects multiple pathways. The figure shows pathways that are activated or inhibited after PH in SOCS3 knockout mice. Data were generated by KEGG analysis of cDNA microarrays [71]

starting at 2 hours after PH, and SOCS3 protein follows the same timing of activation from 2 to 8 hours after PH. This time course correlates with a decrease in STAT3 DNA binding and loss of STAT3 tyrosine 705 phosphorylation. SOCS3 production is markedly reduced in IL-6 knockout mice, and can be restored by IL-6 but not TNF injections, demonstrating the IL-6 requirement for SOCS3 induction. Liver regeneration is accelerated in hepatocyte-specific SOCS3 knockout mice (Soc3 h-KO), through prolonged and enhanced STAT3 phosphorylation, and ERK 1 and 2 stimulation. Socs3 h-KO mice showed enhanced phosphorylation of phospho-S6 protein, a downstream target of the mTOR pathway, but no alterations in Akt phosphorylation, which, however, was highly induced by IL-6 in cultured Socs3 h-KO hepatocytes. Absence of SOCS3 resulted in profound changes in gene expression in the regenerating liver measured by oligonucleotide microarray analyses, particularly in TLR, cytokine, and JAK/STAT pathways, but also in the Wnt and MAPK pathways (Figure 36.6). Promoter analysis revealed an enrichment for transcriptional regulatory elements (TREs) in the SOCS3 h-KO gene set, particularly for NFκB, ELK 1 and 2, c-jun, CREB, USF, and Pax-6 binding sites. SOCS3 expression during the first 8–12 hours after PH appears to be an essential element for the termination the cytokine response and acute-phase protein expression during the priming phase of liver regeneration. The termination of the priming phase by SOCS3 is coordinated with the effect of growth factors that regulate cell-cycle progression. Both HGF and EGFR ligands appear to act on SOCS3 to induce late STAT3 phosphorylation, which is independent of the gp130/IL-6 pathway [51]. Lack of SOCS3 enhances the hepatocyte proliferative capacity, and increases the risk of hepatocellular carcinoma [71, 73]. Oncostatin M, a member of the IL-6 family of cytokines, may act as a key mediator of IL-6 functions. It can protect mice after carbon tetrachloride injury, prevent tissue digestion by metalloproteases, and enhance cell proliferation, as revealed by studies of mice deficient of their receptor [74]. In humans and mice, oncostatin M(OSM)

signals through its own OSM receptor, and in humans also through the leukemia inhibitory factor (LIF) receptor. Receptor binding stimulates liver development through a paracrine effect, and produces effects on inflammation and hematopoiesis. In the human cirrhotic liver, OSM is uniformly expressed in Kupffer cells, the OSM receptor does not appear to be expressed, but the LIF receptor is expressed in reactive ductules and in perisinusoidal areas, colocalizing with Cytokeratin 7(CK7) but not Alpha smooth muscle actin(αSMA) staining [75]. It remains to be determined whether OSM functions as an IL-6 substitute or in conjunction with IL-6 in reactive ductular cells and progenitor cells.

PROGRESSION PHASE: GROWTH FACTORS AND CELL CYCLE GENES Although there is no sharp demarcation between the priming and progression phases, it is clear that SOCS3 expression terminates priming and the main phase of cytokine activation after PH. At the same time, it opens the way to the entry of hepatocytes into the G1 phase of the cell cycle, and the transition through G1 checkpoints, indicated by the expression of D cyclins. In the regenerating liver, as in cells in culture, the cell cycle is controlled by the sequential activation of complexes between cyclins and cyclin-dependent kinases (cdks) [76]. Growth factors control the expression of D cyclins and formation of complexes between these cyclins and cdk4 and cdk6, which act on targets such as Rb and p107 to liberate E2F transcription factors required for DNA replication. The progression through G1 and the S and G2 phases are associated with the sequential activation of cyclins E, A, and B, and appear to depend on a cell autonomous process, as already discussed [35]. In addition to its essential role in the hepatocyte cell cycle, cyclin D also participates in the transcriptional regulation of genes involved in lipid, amino acid, and carbohydrate metabolism [77].

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We will consider here only the most important growth factors that are involved in cell cycle progression, and DNA replication in the regenerating liver.

Hepatocyte growth factor HGF is a member of the plasminogen-related growth factor family. It has approximately 40% homology with plasminogen, but no proteolytic activity. HGF has a multiplicity of effects, through the binding of its receptor tyrosine kinase c-Met, which is expressed in epithelial cells, hepatocytes, neurons, and endothelial cells. HGF is stored in the extracellular matrix of various organs, particularly the liver and the lungs, but also in the spleen and various other organs, and is produced by mesenchymal cell types, and in the liver, mostly in stellate cells. HGF (sometimes referred to as scatter factor, or more often as HGF/SC) is synthesized as an inactive precursor consisting of a single chain. Active HGF is formed by proteolytic cleavage of the precursor, resulting into a heterodimeric molecule linked by a disulfide bond. Several proteases have been implicated in the cleavage process, including urokinase-type plasminogen activator, a soluble protease known as HGF activator (HGFA), the blood coagulation factors XIIa and XIa, thrombin, kallikrein, and matriptase/MT-SP1, an integral-membrane serine proteinase localized on the baso-lateral membrane of epithelial cells that can cleave inactive HGF and also pro-urokinase-type plasminogen activator. Of all of the activators, HGFA has the highest HGF processing activity, which is about 1000 times more potent than that of urokinase-type plasminogen activator [78, 79]. In any case, urokinase-type plasminogen activator is thought to be involved in the cleavage of inactive HGF after PH [9]. After PH, HGF may first be released from the ECM into the plasma through the effect of metalloproteases, whereas its synthesis in the liver occurs later [80–82]. Data are needed to indicate clearly what proportion of total HGF is converted into the heterodimeric active HGF molecule (or into the splice variant HGF/NK1), during liver regeneration after PH, given the discrepancy between some of the published data [78, 83–85]. The development of mice deficient in c-met signaling provided important insights into the role of c-met in liver homeostasis, injury and regeneration. As null mice for HGF or c-met die during gestation, mostly because of placental defects, Huh et al. produced liver-specific c-met knockout mice using Cre/lox gene targeting [86]. These animals grew normally, suggesting that c-met pathways are not essential for normal liver physiology. However, c-met knockout mice were highly sensitive to Fas-induced apoptosis, and showed delayed healing after CCl4 injection, which was associated with hepatic inflammation and calcification. Hepatocyte DNA replication after CCl4 injection did not differ between knockout and control mice,

but the knockout mice showed a deficiency in DNA replication after combined treatment with CCl4 and phenobarbital. Delayed healing occurred in the knockout mice regardless of the presence of a defect in hepatocyte DNA replication, demonstrating an important role for c-met in hepatocyte survival and tissue remodeling. Borowiak et al. developed conditional c-met knockout mice which carried an Mx -cre-inducible c-met deletion [87]. Induction of the deletion (recombination) was obtained by injections of pIpC to activate the Mx promoter. The mode of performance of PH had a major influence in the result of the experiments, in that most knockout mice did not survive when a transversal rather than a vertical incision was used. Under good survival conditions, the weight gain of the knockout mice was slower than that of controls for at least 7 days, and there was a 60% decrease in BrdU incorporation into hepatocytes at 48 hours after PH. In the knockout mice, both HGF and IL-6 were continuously high from 6 to 48 hours after PH, the cell cycle inhibitor p21 was strongly induced, cyclin D1 was delayed, and ERK 1 and 2 phosphorylation was practically absent. Data from these animals suggest that ERK phosphorylation during liver regeneration may depend exclusively on c-met, and that c-met deficiency impaired the exit of hepatocytes from quiescence and decreased their entry into the S phase. The exclusive regulation of ERK pathways implied by these experiments is difficult to reconcile with a large amount of data showing that EGF-family factors may stimulate the same pathways, unless one assumes that the EGF family factors may also act through c-met (see below). Recent data (S. Thorgeirsson, unpublished work) show that c-met deficiency decreases the magnitude of the peak of DNA replication after PH, without altering the time of replication. Moreover, the deficiency strongly inhibited mitosis, blocking hepatocytes in the G2/M phases, and abolishing the second peak of ERK1/2 phosphorylation occurring between 36 and 48 hours after PH. Paranjpe, et al. used small hairpin RNA (shRNA) to block HGF and c-met in the regenerating liver after PH [88]. Blockage of HGF produced only a modest decrease in cell proliferation; blockage of c-met caused inhibition of hepatocyte DNA replication and mitosis at 24 hours after PH, which was restored to normal values at later times. Another potentially important effect of HGF is the phosphorylation of β-catenin, promoting its nuclear translocation [89]. Translocation of β-catenin has been reported to occur rapidly after PH in rats [90] and to remain elevated for 2 days in rats and mice [5]. Several conclusions can be derived from all of these different types of observations that c-met (i) may exclusively regulate some pathways in the regenerating liver, but that it has not been shown that HGF is solely responsible for the activation of these pathways; (ii) may maintain hepatocyte viability by preventing apoptosis and oxidative stress [91]; (iii) has an effect on the extent of hepatocyte replication, and inhibits mitosis (S. Thorgeirsson, unpublished observations).

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EGFR ligands: amphiregulin, heparin-binding EGF, and transforming growth factor alpha The EGF family includes, among others, amphiregulin (AR), heparin binding EGF-like growth factor (HB-EGF), TGFα, betacellulin, epiregulin, and epigen. We will discuss AR, HB-EGF, and EGF/TGFα in relation to liver regeneration after PH. AR, HB-EGF, and TGFα are produced as pro-molecules anchored in the cell membrane, which are released from the cell surface by proteolysis [92]. The main enzymes involved in this process are the ADAMs (a disintegrin and metalloproteinase), particularly ADAM 17, also known as TACE or TNFα converting enzyme. It is not known at this time whether TACE may regulate the interactions between EGFR and its various ligands during liver regeneration. 1. Amphiregulin: AR, like EGF, TGFα, and HGF, stimulates replication of hepatocytes maintained in serum-free medium. It can be induced in cultured hepatocytes by IL-1β and prostaglandin E2, but not by HGF, TNF, or IL-6. AR expression increases within 1 hour after PH, reaching a maximum at about 6 hours [93]. DNA replication is severely impaired in partially hepatectomized AR-deficient mice, suggesting that AR has unique mechanisms of action that are not shared by other EGF ligands. By comparison, hepatocytes of TGFα knockout mice show no deficiency in DNA replication, because of the overlapping activity of other members of the EGF family. 2. Heparin-binding EGF-like factor: HB-EGF appears to act in the transition between the priming and progression phase. This conclusion derives in part from studies of mice that had only one-third of the liver resected by PH [94]. These animals go through most of the priming phase steps, but have negligible DNA replication. A single injection of HB-EGF but not of other growth factors stimulates hepatocytes of one-third PH mice to enter the G1 phase and undergo DNA replication. HB-EGF is produced as a pro-HB-EGF precursor molecule anchored into the cell membrane, which releases mature HB-EGF by proteolytic cleavage, probably through the action of ADAM 17. HB-EGF participation in liver regeneration after two-thirds PH has been described in several publications [95, 96]. 3. TGFα and EGF: In contrast to HGF, TGFα acts only as a proliferative agent. It is produced by hepatocytes, and functions as an autocrine factor through its binding to hepatocyte EGFR1. EGF is produced in the duodenal and salivary glands but not in the liver. TGFα acts on the progression phase of the cell cycle and stimulates hepatocyte DNA replication through its effect on MAPK pathways, and in part on PI3K and Akt signaling. In hepatocytes in culture, addition of

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either EGF or TGFα stimulates cell cycle progression and the transition through a restriction point in late G1 leading to DNA replication [97]. TGFα interacts with HGF in that HGF can stimulate TGFα production, while blockage of TGFα suppresses HGF-mediated DNA replication [98]. TGFα and EGF also interact with TNF in enhancing hepatocyte proliferation and producing multiple division cycles in cultured hepatocytes [99]. Studies of the regulation of MAPK/ERK pathways by EGF ligands demonstrated that ERK2 has a key role in regulating the entry of hepatocytes into the S phase, whereas ERK1 acts on survival [100].

Transforming growth factor beta TGF-β inhibits hepatocyte replication after PH, and it was expected that because of this feature it would be involved in the termination of liver regeneration. However, this does not seem to be the case. The expression of TGFβ and Smads is increased early after PH, and the termination of regeneration is not affected by lack of TGFβ signaling [101–103]. TGFβ may be antagonized by SnoN/Ski signals during liver regeneration [104], but the biological relevance of this antagonism regarding TGFβ effects is unknown. Activin A, a member of the TGFβ family, also inhibits liver regeneration, but there is disagreement on whether activin expression increases after PH [105]. On the other hand, follistatin, a glycoprotein that binds and inactivates activin A, produces hepatic enlargement, but more detailed studies are needed to show that the functional capacity of follistatin-enlarged livers is intact [106]. Hence at present there is a lack of data to demonstrate that inhibitors of hepatocyte replication participate in the termination of liver regeneration.

Other agents A number of agents, such as growth and thyroid hormones, serotonin [107] and norepinephrine [108], can influence liver regeneration. These agents are generally considered to be adjuvants of the regenerative process rather than primary drivers of regeneration. Although caveolin increases after PH, there are contradictory reports about the requirement for caveolin during liver regeneration [109, 110]. Huang et al. reported that mice lacking the nuclear receptor FXR (farnesoid X receptor) had deficient liver regeneration [111]. This receptor, which belongs to a family of nuclear receptors that include the constitutive androstane receptor (CAR) and the vitamin D receptor, functions as a transcription factor that regulates the transcription of genes involved in bile acid homeostasis, lipid metabolism, and glucose balance [112]. Its main ligands are bile acids, chenodeoxycholic acid being the most effective. Although the FXR-dependent

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pathways in the regenerating liver need further study, the identification of FXR as a participant of the regenerative process is of importance because it provides a connection between proliferative and metabolic events in liver regeneration. This is further highlighted by the finding that Foxm1b, a transcription factor associated with DNA replication, may be one of FXR targets [112].

GLOBAL PATTERNS OF GENE EXPRESSION AND TRANSCRIPTION FACTORS Microarray analyses of the global patterns of gene expression after PH show that there are widespread increases and decreases in gene expression, after the operation [41, 113, 114]. Although there is some discrepancy between published data, the pattern of gene expression during liver regeneration does not seem to recapitulate the transcriptional patterns of developing livers [115, 116]. Many of the gene expression changes that occur after PH are controlled by the transcription factors NFκB, STAT3, jun/AP1, and C/EBPβ. More recent work has also identified the transcription factors PAX-6, GATA-1, and HNF-1 (TCF-1) as participants in the control of gene expression during the first 6 hours after PH [116, 117]. In all of these analyses, patterns of gene expression in the regenerating liver have been compared with that of sham-operated or non-operated mice. This type of comparison makes it difficult to identify gene expression changes in the regenerating liver that are related to DNA replication and cell proliferation. Li et al. performed an extensive analysis of gene expression after two-thirds PH in comparison with that of one-third PH, a procedure that causes negligible DNA replication [41]. One of the main findings of this study was that one-third PH causes widespread changes in gene expression, despite the lack of DNA replication. Moreover, there was great overlap between genes expressed after one-third and two-thirds PH. Surprisingly, the decrease in the expression of genes of lipid metabolism that occurs after two-thirds PH [113] is also present after one-third PH. A set of genes was, however, preferentially expressed after two-thirds PH, that is, changes in the expression of these genes after two-thirds PH were of higher magnitude than those found after one-third PH. In terms of functional categories, at 12 hours after two-thirds PH there was preferential expression of genes associated with cell adhesion and angiogenesis. At 20 hours after two-thirds PH, preferential gene expression was associated with genes related to protein synthesis, and at 30 hours, with genes involved in cell growth and DNA replication. Genes expressed during the first 4 hours after PH did not fit into any functional category, but their expression was associated with the transcription factors FOXD3, FOXI1 (HNF-3), CCAAT displacement protein (Cutl1 or Cux1 cut-like homeobox 1), ER, and E2F-1. A

major switch in gene expression occurred at 12 hours after two-thirds PH with the activation of genes that have TREs for c-jun, CCAAT box, Myb, Ets-1, Elk-1, and USF. These analyses agree with and extend published data suggesting that changes in gene expression during the first 4–6 hours after PH are not related to the amount of liver tissue resected, and do not reflect an irreversible commitment to cell proliferation [40]. The decision to replicate after PH is apparently made hours later, when hepatocytes are progressing through G1. The forkhead transcription factors FOXD3 and FOX1I, and the homeodomain protein Cux1, which are associated with gene transcription during the priming phase of liver regeneration, have been shown to act as chromatin remodeling agents. The chromatin remodeling effect of these transcription factors at the initiation steps of liver regeneration might involve the creation of new binding sites that are occupied at later times by transcription factors required for DNA replication.

PERSPECTIVES Until a few years ago, it was possible to create diagrams that summarized almost all of the events in liver regeneration. In retrospect, constructing these diagrams was made possible because of the paucity of information, and the lack of knowledge of some basic aspects of the process. During the last few years, an enormous amount of new data has been generated, bringing new insights into various aspects of liver regeneration. Thus, diagrams can now only cover steps of the process. However, the newly published work exposed deficiencies in experimental design and techniques, particularly in the performance of PH that might raise questions about the reliability of some of the data. To avoid these pitfalls, it is essential to control variables such as feeding schedules, light exposure, Helicobacter contamination, types of anesthesia, and operative technique, as each of these variables can have a significant effect on the timing and magnitude of events occurring during liver regeneration. Another important point is that the mortality of wild-type mice and rats after PH should be less than 5%. As most knockout and transgenic mice are sensitive to anesthesia and operative stress, their mortality after PH may not, in some cases, be directly related to the genetic defect carried by these animals. Finally, instead of a summary, we highlight some key aspects of liver regeneration after PH: 1. Liver regeneration after PH occurs in stages, starting with a priming phase that prepares hepatocytes to enter the cell cycle. This phase is mostly regulated by cytokines and components of the innate immune system, depends on the interaction between non-parenchymal cells and hepatocytes, and is reversible. 2. During the priming phase, there is activation of a set of transcription factors that respond to cytokines, and also of forkhead and homeobox transcription factors that

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can remodel chromatin. A change in the transcriptional program of the regenerating liver occurs 12 hours or later after PH, which is associated with the activation of transcription factors required for DNA replication. 3. Cell cycle progression is controlled by growth factors, mostly HGF, and the EGFR ligands HB-EGF and TGFα. Lack of c-met causes major changes in hepatocyte homeostasis, and inhibits regeneration by decreasing the extent of DNA replication and blocking mitosis. 4. Hepatocytes progress through the cell cycle and transit through restriction points, as indicated by the expression of cyclin D1. Entry into S does not depend on external signals, and relies on a cell autonomous program.

ACKNOWLEDGMENTS The authors’ work cited in this chapter was supported by grants CA23226 and CA74131. We thank past and present members of the laboratory for their contributions.

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42. Ledda-Columbano, G.M., Pibiri, M., Cossu, C., Molotzu, F., Locker, J. and Columbano, A. (2004) Aging does not reduce the hepatocyte proliferative response of mice to the primary mitogen TCPOBOP. Hepatology, 40 (4), 981–88. 43. Strey, C.W., Markiewski, M., Mastellos, D., Tudoran, R., Spruce, L.A., Greenbaum, L.E. et al. (2003) The proinflammatory mediators C3a and C5a are essential for liver regeneration. J Exp Med , 198 (6), 913–23. 44. Tumanov, A.V., Koroleva, E.P., Christiansen, P.A., Khan, M.A., Ruddy, M.J., Burnette, B. et al. (2009) T cell-derived lymphotoxin regulates liver regeneration. Gastroenterology, 136 (2), 694–704, e4. 45. Yamada, Y., Kirillova, I., Peschon, J.J. and Fausto, N. (1997) Initiation of liver growth by tumor necrosis factor: deficient liver regeneration in mice lacking type I tumor necrosis factor receptor. Proc Natl Acad Sci U S A, 94 (4), 1441–46. 46. Campbell, J.S., Riehle, K.J., Brooling, J.T., Bauer, R.L., Mitchell, C. and Fausto, N. (2006) Proinflammatory cytokine production in liver regeneration is Myd88-dependent, but independent of Cd14, Tlr2, and Tlr4. J Immunol , 176 (4), 2522–28. 47. Seki, E., Tsutsui, H., Iimuro, Y., Naka, T., Son, G., Akira, S. et al. (2005) Contribution of Toll-like receptor/myeloid differentiation factor 88 signaling to murine liver regeneration. Hepatology, 41 (3), 443–50. 48. Ware, C.F. (2005) Network communications: lymphotoxins, LIGHT, and TNF. Annu Rev Immunol , 23, 787–819. 49. Knight, B. and Yeoh, G.C. (2005) TNF/LTalpha double knockout mice display abnormal inflammatory and regenerative responses to acute and chronic liver injury. Cell Tissue Res, 319 (1), 61–70. 50. Clark, A., Weymann, A., Hartman, E., Turmelle, Y., Carroll, M., Thurman, J.M. et al. (2008) Evidence for non-traditional activation of complement factor C3 during murine liver regeneration. Mol Immunol , 45 (11), 3125–32. 51. Seki, E., Kondo, Y., Iimuro, Y., Naka, T., Son, G., Kishimoto, T. et al. (2008) Demonstration of cooperative contribution of MET- and EGFR-mediated STAT3 phosphorylation to liver regeneration by exogenous suppressor of cytokine signalings. J Hepatol , 48 (2), 237–45. 52. Cornell, R.P. (1985) Restriction of gut-derived endotoxin impairs DNA synthesis for liver regeneration. Am J Physiol , 249 (5Pt 2), R563–69. 53. Cornell, R.P., Liljequist, B.L. and Bartizal, K.F. (1990) Depressed liver regeneration after partial hepatectomy of germ-free, athymic and lipopolysaccharide-resistant mice. Hepatology, 11 (6), 916–22. 54. Doi, T.S., Marino, M.W., Takahashi, T., Yoshida, T., Sakakura, T., Old, L.J. (1999) Absence of tumor necrosis factor rescues RelA-deficient mice from embryonic lethality. Proc Natl Acad Sci U S A, 96 (6), 2994–99.

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55. Rosenfeld, M.E., Prichard, L., Shiojiri, N. and Fausto, N. (2000) Prevention of hepatic apoptosis and embryonic lethality in RelA/TNFR-1 double knockout mice. Am J Pathol , 156 (3), 997–1007. 56. Chaisson, M.L., Brooling, J.T., Ladiges, W., Tsai, S. and Fausto, N. (2002) Hepatocyte-specific inhibition of NF-kappaB leads to apoptosis after TNF treatment, but not after partial hepatectomy. J Clin Invest , 110 (2), 193–202. 57. Kirillova, I., Chaisson, M. and Fausto, N. (1999) Tumor necrosis factor induces DNA replication in hepatic cells through nuclear factor kappaB activation. Cell Growth Differ, 10 (12), 819–28. 58. Malato, Y., Sander, L.E., Liedtke, C., Al-Masaoudi, M., Tacke, F., Trautwein, C. et al. (2008) Hepatocytespecific inhibitor-of-kappaB-kinase deletion triggers the innate immune response and promotes earlier cell proliferation during liver regeneration. Hepatology, 47 (6), 2036–50. 59. Tacke, F., Luedde, T. and Trautwein, C. (2009) Inflammatory pathways in liver homeostasis and liver injury. Clin Rev Allergy Immunol, 36 (1), 4–12. 60. Sakamoto, T., Liu, Z., Murase, N., Ezure, T., Yokomuro, S., Poli, V. et al. (1999) Mitosis and apoptosis in the liver of interleukin-6-deficient mice after partial hepatectomy. Hepatology, 29 (2), 403–11. 61. Blindenbacher, A., Wang, X., Langer, I., Savino, R., Terracciano, L. and Heim, M.H. (2003) Interleukin 6 is important for survival after partial hepatectomy in mice. Hepatology, 38 (3), 674–82. 62. Wustefeld, T., Rakemann, T., Kubicka, S., Manns, M.P. and Trautwein, C. (2000) Hyperstimulation with interleukin 6 inhibits cell cycle progression after hepatectomy in mice. Hepatology, 32 (3), 514–22. 63. Scheller, J. and Rose-John, S. (2006) Interleukin-6 and its receptor: from bench to bedside. Med Microbiol Immunol , 195 (4), 173–83. 64. Zimmers, T.A., Pierce, R.H., McKillop, I.H. and Koniaris, L.G. (2003) Resolving the role of IL-6 in liver regeneration. Hepatology, 38 (6), 1590–91; author reply 1. 65. Bansal, M.B., Kovalovich, K., Gupta, R., Li, W., Agarwal, A., Radbill, B. et al. (2005) Interleukin-6 protects hepatocytes from CCl4 -mediated necrosis and apoptosis in mice by reducing MMP-2 expression. J Hepatol , 42 (4), 548–56. 66. Ren, X., Hogaboam, C., Carpenter, A. and Colletti, L. (2003) Stem cell factor restores hepatocyte proliferation in IL-6 knockout mice following 70% hepatectomy. J Clin Invest , 112 (9), 1407–18. 67. Ren, X., Hu, B. and Colletti, L. (2008) Stem cell factor and its receptor, c-kit, are important for hepatocyte proliferation in wild-type and tumor necrosis factor receptor-1 knockout mice after 70% hepatectomy. Surgery, 143 (6), 790–802. 68. Inoue, H., Ogawa, W., Ozaki, M., Haga, S., Matsumoto, M., Furukawa, K. et al. (2004) Role of STAT-3 in regulation of hepatic gluconeogenic genes and carbohydrate metabolism in vivo. Nat Med , 10 (2), 168–74.

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69. Gao, B. (2005) Cytokines, STATs and liver disease. Cell Mol Immunol , 2 (2), 92–100. 70. Campbell, J.S., Prichard, L., Schaper, F., Schmitz, J., Stephenson-Famy, A., Rosenfeld, M.E. et al. (2001) Expression of suppressors of cytokine signaling during liver regeneration. J Clin Invest , 107 (10), 1285–92. 71. Riehle, K.J., Campbell, J.S., McMahan, R.S., Johnson, M.M., Beyer, R.P., Bammler, T.K. et al. (2008) Regulation of liver regeneration and hepatocarcinogenesis by suppressor of cytokine signaling 3. J Exp Med , 205 (1), 91–103. 72. Lehman, J.A., Calvo, V. and Gomez-Cambronero, J. (2003) Mechanism of ribosomal p70S6 kinase activation by granulocyte macrophage colony-stimulating factor in neutrophils: cooperation of a MEK-related, THR421/SER424 kinase and a rapamycin-sensitive, m-TOR-related THR389 kinase. J Biol Chem, 278 (30), 28130–38. 73. Ogata, H., Kobayashi, T., Chinen, T., Takaki, H., Sanada, T., Minoda, Y. et al. (2006) Deletion of the SOCS3 gene in liver parenchymal cells promotes hepatitis-induced hepatocarcinogenesis. Gastroenterology, 131 (1), 179–93. 74. Nakamura, K., Nonaka, H., Saito, H., Tanaka, M. and Miyajima, A. (2004) Hepatocyte proliferation and tissue remodeling is impaired after liver injury in oncostatin M receptor knockout mice. Hepatology, 39 (3), 635–44. 75. Znoyko, I., Sohara, N., Spicer, S.S., Trojanowska, M. and Reuben, A. (2005) Expression of oncostatin M and its receptors in normal and cirrhotic human liver. J Hepatol , 43 (5), 893–900. 76. Nelsen, C.J., Hansen, L.K., Rickheim, D.G., Chen, C., Stanley, M.W., Krek, W. et al. (2001) Induction of hepatocyte proliferation and liver hyperplasia by the targeted expression of cyclin E and skp2. Oncogene, 20 (15), 1825–31. 77. Mullany, L.K., White, P., Hanse, E.A., Nelsen, C.J., Goggin, M.M., Mullany, J.E. et al. (2008) Distinct proliferative and transcriptional effects of the D-type cyclins in vivo. Cell Cycle, 7 (14), 2215–24. 78. Kaibori, M., Inoue, T., Oda, M., Naka, D., Kawaguchi, T., Kitamura, N. et al. (2002) Exogenously administered HGF activator augments liver regeneration through the production of biologically active HGF. Biochem Biophys Res Commun, 290 (1), 475–81. 79. Miyazawa, K., Shimomura, T. and Kitamura, N. (1996) Activation of hepatocyte growth factor in the injured tissues is mediated by hepatocyte growth factor activator. J Biol Chem, 271 (7), 3615–18. 80. Mars, W.M., Liu, M.L., Kitson, R.P., Goldfarb, R.H., Gabauer, M.K. and Michalopoulos, G.K. (1995) Immediate early detection of urokinase receptor after partial hepatectomy and its implications for initiation of liver regeneration. Hepatology, 21 (6), 1695–701. 81. Mars, W.M., Zarnegar, R. and Michalopoulos, G.K. (1993) Activation of hepatocyte growth factor by the plasminogen activators uPA and tPA. Am J Pathol , 143 (3), 949–58.

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95. Kiso, S., Kawata, S., Tamura, S., Higashiyama, S., Ito, N., Tsushima, H. et al. (1995) Role of heparin-binding epidermal growth factor-like growth factor as a hepatotrophic factor in rat liver regeneration after partial hepatectomy. Hepatology, 22 (5), 1584–90. 96. Kiso, S., Kawata, S., Tamura, S., Inui, Y., Yoshida, Y., Sawai, Y. et al. (2003) Liver regeneration in heparin-binding EGF-like growth factor transgenic mice after partial hepatectomy. Gastroenterology, 124 (3), 701–7. 97. Loyer, P., Cariou, S., Glaise, D., Bilodeau, M., Baffet, G. and Guguen-Guillouzo, C. (1996) Growth factor dependence of progression through G1 and S phases of adult rat hepatocytes in vitro. Evidence of a mitogen restriction point in mid-late G1. J Biol Chem, 271 (19), 11484–92. 98. Scheving, L.A., Stevenson, M.C., Taylormoore, J.M., Traxler, P. and Russell, W.E. (2002) Integral role of the EGF receptor in HGF-mediated hepatocyte proliferation. Biochem Biophys Res Commun, 290 (1), 197–203. 99. Serandour, A.L., Loyer, P., Garnier, D., Courselaud, B., Theret, N., Glaise, D. et al. (2005) TNFalpha-mediated extracellular matrix remodeling is required for multiple division cycles in rat hepatocytes. Hepatology, 41 (3), 478–86. 100. Fremin, C., Ezan, F., Boisselier, P., Bessard, A., Pages, G., Pouyssegur, J. et al. (2007) ERK2 but not ERK1 plays a key role in hepatocyte replication: an RNAi-mediated ERK2 knockdown approach in wild-type and ERK1 null hepatocytes. Hepatology, 45 (4), 1035–45. 101. Braun, L., Mead, J.E., Panzica, M., Mikumo, R., Bell, G.I. and Fausto, N. (1988) Transforming growth factor beta mRNA increases during liver regeneration: a possible paracrine mechanism of growth regulation. Proc Natl Acad Sci U S A, 85 (5), 1539–43. 102. Jakowlew, S.B., Mead, J.E., Danielpour, D., Wu, J., Roberts, A.B. and Fausto, N. (1991) Transforming growth factor-beta (TGF-beta) isoforms in rat liver regeneration: messenger RNA expression and activation of latent TGF-beta. Cell Regul, 2 (7), 535–48. 103. Oe, S., Lemmer, E.R., Conner, E.A., Factor, V.M., Leveen, P., Larsson, J. et al. (2004) Intact signaling by transforming growth factor beta is not required for termination of liver regeneration in mice. Hepatology, 40 (5), 1098–105. 104. Macias-Silva, M., Li, W., Leu, J.I., Crissey, M.A. and Taub, R. (2002) Up-regulated transcriptional repressors SnoN and Ski bind Smad proteins to antagonize transforming growth factor-beta signals during liver regeneration. J Biol Chem, 277 (32), 28483–90. 105. Rodgarkia-Dara, C., Vejda, S., Erlach, N., Losert, A., Bursch, W., Berger, W. (2006) The activin axis in liver biology and disease. Mutat Res, 613 (2–3), 123–37. 106. Endo, D., Maku-Uchi, M. and Kojima, I. (2006) Activin or follistatin: which is more beneficial to support liver regeneration after massive hepatectomy? Endocr J , 53 (1), 73–78.

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107. Lesurtel, M., Graf, R., Aleil, B., Walther, D.J., Tian, Y., Jochum, W. et al. (2006) Platelet-derived serotonin mediates liver regeneration. Science, 312 (5770), 104–7. 108. Cruise, J.L., Knechtle, S.J., Bollinger, R.R., Kuhn, C. and Michalopoulos, G. (1987) Alpha 1-adrenergic effects and liver regeneration. Hepatology, 7 (6), 1189–94. 109. Mayoral, R., Fernandez-Martinez, A., Roy, R., Bosca, L. and Martin-Sanz, P. (2007) Dispensability and dynamics of caveolin-1 during liver regeneration and in isolated hepatic cells. Hepatology, 46 (3), 813–22. 110. Fernandez, M.A., Albor, C., Ingelmo-Torres, M., Nixon, S.J., Ferguson, C., Kurzchalia, T. et al. (2006) Caveolin-1 is essential for liver regeneration. Science, 313 (5793), 1628–32. 111. Huang, W., Ma, K., Zhang, J., Qatanani, M., Cuvillier, J., Liu, J. et al. (2006) Nuclear receptor-dependent bile acid signaling is required for normal liver regeneration. Science, 312 (5771), 233–36. 112. Wang, Y.D., Chen, W.D., Moore, D.D. and Huang, W. (2008) FXR: a metabolic regulator and cell protector. Cell Res, 18 (11), 1087–95.

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Ribosome Biogenesis and its Role in Cell Growth and Proliferation in the Liver Stefano Fumagalli and George Thomas Department of Cancer and Cell Biology, Genome Research Institute, University of Cincinnati, Cincinnati, OH, USA

INTRODUCTION Ribosome biogenesis is the process that leads to the assembly of ribosomal ribonucleic acid (rRNA) and ribosomal proteins into mature 40S and 60S ribosomal subunits. These ribosomal subunits are employed in the cytoplasm in the process of protein synthesis. The biosynthesis of ribosomes takes place in discrete subregions inside the nucleus, the nucleoli, which form around sites of transcription of rRNA [1]. Because ribosome biogenesis is a process that consumes a considerable amount of energy, it is important for the cell to regulate ribosome production tightly according to the availability of nutrients and energy sources in the extracellular milieu [2]. This is very well illustrated in the liver, where fasting that results in a decrease in the mass of the hepatocytes leads not only to a reduction in ribosome biogenesis, but also to the degradation of ribosomes through autophagy, in order to provide the cells with amino acids and nucleotides in periods of nutrient shortage. Conversely, refeeding results in the stimulation of ribosome biogenesis and inhibition of

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

ribosome degradation that support an increase in the rate of protein synthesis and reestablishes an adequate content of ribosomes for the cell to perform its metabolic functions once it has grown back to its original size [3, 4]. The rate of ribosome biogenesis also rises during cell proliferation. For example, during the proliferative response that is triggered by partial hepatectomy, the rates of ribosome synthesis in the cells of the liver are increased dramatically [5–7]. It is generally believed that cells must increase in size to proliferate. Because cell mass also depends on the quantity of intracellular proteins, upregulation of ribosome biogenesis during liver regeneration may contribute to the increase in protein synthesis which is required for cell growth and eventually cell cycle progression. Most importantly, enhanced ribosome biogenesis is believed to ensure that, upon cell division, daughter cells are provided with the appropriate amount of ribosomes. In recent years, mainly through the use of in vitro systems, considerable progress has been made in the understanding of how ribosome biogenesis is regulated by both nutrients and mitogenic signals. Furthermore, evidence has emerged

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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for a link between ribosome biogenesis and components of the cell cycle machinery. Although important findings have been obtained on this subject in the liver, it can be said that, in general, the in vivo study of these processes has been lagging behind in vitro studies because of the difficulties and time constrains inherent to animal systems. It is the aim of this review to summarize general concept of ribosome biogenesis and its regulation, with an emphasis on findings that are relevant to liver biology, and to discuss important data that have linked ribosome biogenesis to cell cycle progression in the liver.

THE NUCLEOLUS AND RIBOSOME BIOGENESIS The steps of ribosome biogenesis largely take place in specialized nuclear domains, the nucleoli [1]. Nucleoli form around so-called nucleolar organizer regions (NORs), which are heterochromatic regions found on a subset of chromosomes (five in humans). Each NOR contains 20–40 copies of the rRNA gene arranged in tandem repeats in a head-to-tail configuration. Only about 50% of the rRNA genes contained in an NOR are found in a euchromatic configuration and can, therefore, be transcribed. The remaining rRNA genes are instead found in heterochromatic regions and are, thus, transcriptionally silent. This type of configuration seems to be important for the maintenance of nucleolar integrity and to limit the chance of recombination

between transcriptionally active rRNA genes (discussed in [8]). The presence of the nucleolus is dependent on active transcription of rRNA genes by a specialized RNA polymerase complex, RNA polymerase I (Pol I) and ongoing ribosome biogenesis. For this reason nucleoli are clearly visible during interphase, when ribosomes are being synthesized, disappear when rRNA transcription ceases during mitosis, and become visible again when rRNA transcription resumes at the onset of G1 [1]. Pol I transcribes the precursor rRNA transcript, 47S rRNA, which undergoes a series of processing events that include its cleavage into mature 18S, 28S, and 5.8S rRNA [9] (see Figure 37.1 for the description of the two main cleavage pathways of rRNA in mouse cells); nucleotide modifications, namely pseudouridynilation and methylation; and assembly with ribosomal proteins to form small (40S) ribosomal subunits that contain 33 ribosomal proteins and 18S rRNA, and large (60S) ribosomal subunits that contain 47 ribosomal proteins and, in addition to 28S and 5.8S rRNA, 5S rRNA which is transcribed by RNA polymerase III at sites distinct from the nucleolus [10]. The complex series of events that results in the synthesis of mature ribosomal subunits involves the activity of diverse enzymes, including helicases, acetic anhydride (AAA) adenosine triphosphates (ATPases), guanosine-triphosphates (GTPases), endonucleases, exonucleases, chaperones, and RNA-modifying activities. Although most of the studies of the trans-acting factors involved in ribosome biogenesis have been performed in the yeast Saccharomyces

5’-ETS

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Figure 37.1 Representation of the two main pre-rRNA processing pathways in mouse cells. The arrowheads represent cleavage sites. ETS, external transcribed sequence; ITS, internal transcribed sequence

37: RIBOSOME BIOGENESIS AND ITS ROLE IN CELL GROWTH AND PROLIFERATION IN THE LIVER FC DFC

GC

Figure 37.2 The morphology of the nucleolus in mammalian cells. On the left is represented the nucleolus of a cell actively engaged in ribosome biogenesis. On the right is the nucleolus of a cell undergoing inhibition of rRNA transcription. Indicated are the three nucleolar compartments: the fibrillar center (FC), the dense fibrillar component (DFC), and the granular component (GC). See text for details

cerevisiae [11], recent proteomic analysis shows that homologs of many of them are found in the nucleolus of higher eukaryotes, where they likely perform a function similar to the one described for their yeast counterparts [12]. The different steps of ribosome biogenesis take place in different regions of the nucleolus, which have been classified through ultrastructural analysis as three different nucleolar compartments: the fibrillar centers (FCs), the dense fibrillar components (DFCs) that surround FCs, and the granular component (GC) in which FCs and the adjacent DFCs are embedded (Figure 37.2) [13]. FCs are enriched with Pol I; however, transcription of rRNA seems to take place mainly at the interface between the FCs and the DFCs. In the DFCs most rRNA cleavages take place. The DFCs are also the sites of nucleotide modification of rRNA by snoRNPs, small ribonucleoproteic particles composed of RNAs, known as small nucleolar ribonucleic acids (snoRNAs), and RNA-modifying enzymes. The RNA component of each snoRNP hybridizes to a different region of the rRNA molecules, thereby directing the associated RNA-modifying activity toward specific nucleotide residues [14]. The GC is enriched in ribosomal proteins and assembly factors and is the sites where ribosome assembly is completed. The spatial distribution of the different nucleolar compartments changes dramatically when ribosome biogenesis ceases, for example following inhibition of rRNA transcription with low doses of actinomycin D. In this case, the nucleolus undergoes a process of segregation in which the three compartments remain in contact but are no longer intermingled (Figure 37.2) [13].

REGULATION OF RIBOSOME BIOGENESIS The rates of ribosome biogenesis are greatly increased by growth and proliferative stimuli, whereas they are severely diminished by nutrient deprivation. The effects of these factors on ribosome synthesis are exerted mainly through

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the production of the building blocks of ribosomes, namely rRNA and ribosomal proteins. For example, it is known that following partial hepatectomy in the rat, liver cells upregulate ribosome biogenesis by increasing the translation and transcription of ribosomal protein genes as well as the transcription of rRNA genes [6, 15, 16]. Similarly, it has been shown that in the liver of rats fed after a period of fasting, rRNA transcription increases [3] and ribosomal protein mRNAs are recruited onto actively translating polysomes [17]. The knowledge of how nutritional and mitogenic signals affect intracellular signaling pathways to stimulate the synthesis of ribosomal components is derived mainly from studies of tissue culture systems, whereas in this respect little is known for the liver.

Ribosomal protein translation and mTOR An important regulator of ribosome biogenesis in response to nutrient signaling, in particular amino acids, is the mammalian target of rapamycin (mTOR). mTOR is a serine threonine kinase that is found in two different complexes known as mammalian target of rapamycin complex 1 (mTORC1) and mammalian target of rapamycin complex 2 (mTORC2). Whereas both complexes respond to serum stimulation and to growth factors, stimulation by nutrients, in particular amino acids, seems to be specific to mTORC1. Furthermore, of the two complexes, only mTORC1 is a target of the immunosuppressant rapamycin that, in a gain-of-function complex with the peptidyl-prolyl isomerase FKBP12, binds and inhibits the kinase activity of mTOR [18]. The main targets of mTORC1 are eIF4E-binding protein (4EBPs) (eukaryotic initiation factor 4E (e-IF4E) binding proteins) and S6K (S6 kinase). 4EBPs, by binding to the cap-binding protein e-IF4E, inhibit its association with the scaffold e-IF4G and, therefore, inhibit formation of an active eIF4F complex, which is required for recruitment of mRNAs to 40S subunits during initiation of protein synthesis. Phosphorylation of 4EBPs by mTORC1 relieves the binding of 4EBPs to e-IF4E and allows the assembly of the e-IF4F complex [19]. mTORC1 also phosphorylates and activates the Ser/Thr kinase S6K1, which is known to phosphorylate a number of substrates that are involved in translation [20]. A link between mTOR and ribosome biogenesis came from studies showing that mTOR stimulates the translation of mRNAs encoding ribosomal proteins in resting cells that were stimulated with mitogens to reenter the cell cycle. In vertebrates, all the mRNAs encoding ribosomal proteins belong to the 5′ -TOP (5′ -terminal oligopyrimidine tract) family of mRNAs. 5′ -TOP mRNAs are characterized by an uninterrupted stretch of pyrimidines at their 5′ end, the 5′ -TOP, which has an average length of 12.2 nucleotides in humans and 8.2 nucleotides in mice, and which starts invariably with a C [21]. 5′ -TOP mRNAs are poorly

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translated in resting cells; however, they are abruptly recruited on to actively translating polysomes when cells are stimulated in the presence of nutrients with mitogens and growth factors [22]. Mutations of the 5′ -TOP abolish regulation and render the translation of the mutated transcript constitutive [23]. It was shown that rapamycin treatment prevents translational upregulation of 5′ -TOP mRNAs in resting cells stimulated with serum [24, 25]. The inhibitory effects of rapamycin were determined to be mediated through the 5′ -TOP because the translation of transcripts bearing a mutated 5′ -TOP was insensitive to rapamycin treatment [26]. Interestingly, upregulation of the translation of 5′ -TOP mRNAs also takes place during liver regeneration in the rat; however, it is not known if mTOR is involved in that case [15]. With regard to the response to feeding, experimental evidence supports a role for mTOR in regulating the translation of 5′ -TOPs mRNA. In fact, it has been shown that, concomitant with upregulation of 5′ -TOP mRNA translation, the mTOR pathway becomes activated in the liver of rats in response to nutrient intake. Furthermore, in the liver of rats fed with leucine, the mTOR pathway and translation of 5′ -TOP mRNAs are activated, and rapamycin prevents both responses [17]. Likewise, rapamycin impairs liver growth during refeeding in rats, concomitant with inhibition of both activation of the mTOR pathway and upregulation of ribosomal protein-mRNAs translation [27].

Regulation of the transcription of rRNA genes Like the translation of ribosomal proteins, transcription of rRNA has also been shown to respond to nutrient availability. Feeding increases the rates of rRNA transcription about twofold in the liver [3]. Furthermore, there is ample evidence of rRNA transcription being upregulated during liver regeneration (see, for example, [6]). Transcription of rRNA genes by Pol I requires the formation of a preinitiation complex, which forms through the binding of the promoter selectivity factor 1/transcription initiation factor-IB (SL1/TIF-IB), to the core promoter (CP) of rRNA genes. SL1/TIF-IB is a multiprotein complex containing the TATA-binding protein (TBP) and TATA-binding protein-associated factors (TAFs) [28–30]. Binding of SL1/TIF-IB to the CP is facilitated by its interaction with upstream binding factor (UBF). UBF contains high mobility group (HMG) boxes, through which it binds as a dimer to a distal region in the rRNA promoter known as the upstream control element (UCE). The interaction of UBF with SL1/TIF-IB is mediated by the carboxy-terminal activation domain of UBF and requires the phosphorylation of Ser residues in this domain by casein kinase 2 (CK2) [31]. After binding to the CP, SL1/TIF-IB recruits Pol I, to form a productive initiation complex (Figure 37.3). This process is mediated through the interaction of SL1/TIF-IB with

UBF UCE

SL1/TIF-IB

TIF-IA A43

PoI I

CP

Figure 37.3 Schematic representation of the transcription initiation complex at the promoter of the rRNA gene. See text for details

transcription initiation factor-IA (TIF-IA), that is associated with the A43 subunit of Pol I (reviewed in [28]). The link between transcription of rRNA and nutrients suggests that mTOR, in addition to regulating translation of ribosomal proteins, also regulates the activity of Pol I. In fact, it was shown first that treatment of lymphosarcoma cells with rapamycin inhibits rRNA transcription [32]. On the basis of other experiments with rapamycin, it was later proposed that the effects of mTOR on rRNA transcription are mediated through phosphorylation of TIF-IA [33]. In particular, mTOR, by promoting phosphorylation of S44, possibly through inhibition of protein phosphatase 2A (PP2A) and dephosphorylation of S199 of TIF-IA, promotes the association of TIF-IA with Pol I and components of SL1/TIF1-B complex and the formation of the initiation complex at rRNA promoters [33]. In addition, rapamycin treatment results in the translocation of TIF-IA to the cytoplasm, which suggests that mTOR may also regulate Pol I transcription by promoting accumulation of TIF-IA in the nucleus [33]. Unfortunately, these studies did not determine if phosphorylation of S44 and S199 was influenced by changes in nutrient concentration in the extracellular environment or the proliferative status of the cells. In tissue culture systems, transcription of rRNA has been shown to be enhanced by mitogenic stimulation with either serum or purified growth factors like epidermal growth factor (EGF). Thus, it has been shown that in resting cells stimulated with serum, rRNA transcription is mediated by the mTOR pathway through S6K1 that phosphorylates Ser residues in the UBF carboxy-terminal activation domain, promoting binding of UBF to SL1/TIF-IB [34]. The microtubule associated protein (MAP) kinase (MAPK) pathway is also involved in the stimulation of rRNA transcription by mitogens. In response to serum, these effects are mediated through phosphorylation of S633 and S649 of TIF-IA suggesting that, like mTOR, the MAPK pathway increases the rate of rRNA transcription at the level of the initiation step [35]. Others have shown that, in contrast, in response to EGF stimulation, increases in the rates of elongation rather than initiation are responsible for increased transcription of rRNA by Pol I. Also, the effects of EGF stimulation seem to be mediated by the MAPK pathway through phosphorylation of UBF [36]. UBF has been proposed to pack the DNA in a nucleosome-like structure, called an enhancesome (reviewed in [29]). As already mentioned, UBF facilitates the binding of SL1/TIF-IB to the CP and formation of

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the preinitiation complex. On the other hand, UBF is found associated not only to the promoter but also to the transcribed region of rRNA genes, where it promotes the formation of enhancesomes, that slow the progression of Pol I. In response to EGF stimulation, phosphorylation by extracellular-regulated kinase (ERK) 1/2 of T117 and T201 in HMG box 1 and 2, respectively, of UBF, has been proposed to lead to disruption of the enhancesome, resulting in an increase in the rate of elongation of Pol I transcription [37].

The role of MYC in ribosome biogenesis The proto-oncogene MYC has emerged as a regulator of ribosome biogenesis. c-MYC is a transcription factors found deregulated in a variety of cancers [38]. It has been shown to control the transcription of genes involved in cell proliferation and cell growth [39]. Interestingly, the analysis of MYC-dependent changes in the pattern of transcription, performed by several laboratories, showed that c-MYC and its related homolog N-MYC drive transcription of ribosomal proteins genes [40–42]. In line with those observations, it has been shown that overexpression of c-MYC in mouse liver leads to hypertrophy of hepatocytes associated with enlargement of the nucleoli, an indication of elevated rates of ribosome biogenesis. Concomitant with these changes, the levels of mRNAs encoding ribosomal proteins were found to be upregulated, suggesting that c-MYC overexpression increases the translational capacity of the hepatocytes by enhancing ribosome biogenesis, which leads to cell hypertrophy [43]. In this respect, it has to be noted that c-MYC also drives the transcription of genes encoding translation initiation factors [39]. Interestingly, overexpression of c-MYC in the liver did not result in increased proliferation. As mentioned above, during liver regeneration the transcription of genes encoding ribosomal proteins increases by an unknown mechanism [15]. Because expression of c-MYC is also induced by partial hepatectomy [44], it is tempting to speculate that c-MYC could contribute to the increase in mRNAs encoding ribosomal proteins in proliferating hepatocytes. In addition to regulating the synthesis of ribosomal proteins, c-MYC has also been shown to drive Pol I transcription by associating with components of the SL1/TIF-IB complex and promoting formation of the preinitiation complex at the promoter of rRNA genes [45, 46]. Interestingly, c-MYC, by interacting with the Pol III-specific factor TFIIIB, can drive transcription of, among others, the gene encoding the 5S rRNA, a component of 60S ribosomal subunits [47]. In summary, by regulating transcription of all cellular RNA polymerase, c-MYC ensures the coordinated accumulation of ribosomal components to make ribosome biogenesis a highly efficient process [48]. In this respect, c-MYC is reminiscent of mTOR, which regulates both translation of ribosomal

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protein mRNAs and rRNA transcription. In contrast to c-MYC, mTOR has not been shown to drive Pol III transcription in mammals; however, data in budding yeast suggest that it may do so in the yeast system [49].

Inhibition of rRNA transcription by members of the pocket protein family In addition to being activated by growth and mitogenic stimuli, ribosome biogenesis can be actively inhibited by the products of tumor suppressor genes. A well-characterized example is the activity of the RB gene product, pRB. In addition to functioning to modulate the transcriptional activity of E2F transcription factors, pRB has been shown to be localized in the nucleolus where it inhibits transcription of rRNA in differentiating cells [50] and resting cells [51]. The inhibitory effect of pRB on Pol I transcription has been proposed to be exerted through binding to UBF and the consequent impairment of SL1/TIF-IB recruitment to the CP [52]. The interaction with UBF seems to be mediated through the pocket region of pRB, a domain which is required for its tumor suppressive activity [50]. Interestingly, pRB has also been shown to inhibit Pol III transcription [53]. Similar functions in both Pol I and Pol III transcription have been ascribed to the pRB-related pocket proteins p130 and p107 [54, 55].

THE ROLE OF RIBOSOME BIOGENESIS IN CELL GROWTH AND PROLIFERATION IN THE LIVER In the previous sections, we described how ribosome biogenesis is controlled by signaling pathways that mediate the response to growth and proliferative stimuli. In this section, we will discuss how alterations in ribosome biogenesis affect those processes. Important insights into these issues have come from studies in mice where liver-specific deletion of the gene encoding the small subunit ribosomal protein S6 (rpS6), engineered to contain flox sequences, could be triggered in adults through interferon-dependent induction of CRE [56]. rpS6 is incorporated into the 40S subunit at an early stage of ribosome biogenesis [57], and it was assumed that its depletion would affect the assembly of 40S ribosomes. Indeed, it was shown that deletion of the rpS6 gene in the liver specifically inhibited 40S ribosome biogenesis, but did not affect the production of 60S ribosomes. The inhibition of 40S ribosomes synthesis was concomitant with a failure to produce 18S rRNA and the accumulation of the 34S precursor of the 18S rRNA (Figure 37.1 and Figure 37.4a), suggesting that rpS6 is required for the processing of rRNA to its mature form. The requirement

THE LIVER: THE ROLE OF RIBOSOME BIOGENESIS IN CELL GROWTH AND PROLIFERATION IN THE LIVER

S6 −/−

S6 + /+

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45S 34S 28S 18S (a)

Fasting

S6 +/+

Refeeding

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Partial hepatectomy

S6 −/−

S6+/+

S6−/−

Ribosome biogenesis

Ribosome biogenesis

Ribosome biogenesis

Checkpoint Liver regrowth

Liver regrowth

(b)

Liver regeneration

Liver regeneration

(c)

Figure 37.4 (a) 32 P-labeled RNA from the liver of wild-type mice (S6+/+) and mice where deletion of the rpS6 gene was induced (S6−/−). Note that deletion of the rpS6 gene leads to impairment of 18S rRNA synthesis and the accumulation of the 34S precursor of the 18S rRNA. (b) Deletion of the rpS6 gene and the concomitant inhibition of 40S ribosome biogenesis do not affect liver regrowth induced by refeeding following a period of fasting. (c) Deletion of the rpS6 gene and the concomitant inhibition of 40S ribosome biogenesis activate a checkpoint that halts cell cycle progression and impairs liver regeneration following partial hepatectomy

for ribosome biogenesis in cell growth in the liver was tested in an experiment involving refeeding after fasting (see the Introduction). Surprisingly, deletion of rpS6, and consequent inhibition of 40S ribosome biogenesis, did not impair liver growth in that setting (Figure 37.4b). Due to the high stability of ribosomes [58], deletion of the rpS6 gene affected only partially the total content of 40S ribosomes in the liver, yet, judging from the profile of the polysome distribution at 24 hours after refeeding, translation rates were severely affected [56]. This suggests that the rate of protein synthesis sustained by the population of pre-existing ribosomes was sufficient to ensure liver growth. Moreover, the fact that cell growth took place in the absence of ribosome biogenesis ruled out the possibility of the existence a system that coupled cell growth to the proper execution of ribosome biogenesis in the liver. The requirement for ribosome biogenesis in the proliferative response was tested in an experiment involving liver

regeneration (see the Introduction). Interestingly, deletion of rpS6 impaired liver regeneration after partial hepatectomy because hepatocytes failed to enter S phase [56]. This could not be attributed simply to loss of translational capacity, because, as mentioned above, the remaining ribosomes were sufficient to allow the liver cells of fasted animals to regrow in response to feeding. The analysis of the expression and activity of cell cycle regulators revealed that accumulation of cyclin D1 (CYCD1) and formation of active CYCD1/CDK4 kinase complexes were not affected by rpS6 deletion [56]. The CYCD1/CDK4 complex is involved in progression through the G1 phase of the cell cycle. Its main function is to phosphorylate the pocket proteins pRB and p130 to promote the transcription of genes that encode positive effectors of cell cycle progression [59]. This transcription is dependent on members of the E2F family of transcription factors [60]. Interestingly, rpS6-deficient liver cells failed to induce

37: RIBOSOME BIOGENESIS AND ITS ROLE IN CELL GROWTH AND PROLIFERATION IN THE LIVER

cyclin E (CYCE) mRNA and protein and, therefore, failed to form CYCE/CDK2 complexes [56], whose activity is required for the cells to initiate DNA synthesis [61]. Because CYCE transcription is driven by E2F transcription factors, it was speculated that inhibition of ribosome biogenesis by depletion of rpS6 would suppress the activation of E2F-dependent transcription. Indeed, a DNA hybridization microarray analysis revealed that, following partial hepatectomy, hepatocytes bearing the deletion of the rpS6 gene fail to activate the expression of several other genes that are targets of E2F transcription factors (unpublished results). In agreement with these findings, it was found that, in contrast to what happens in wild-type liver cells, the pocket protein p130 does not undergo phosphorylation following partial hepatectomy in rpS6-depleted cells (unpublished results). These findings clearly argue that CYCD1/CDK4 activity is not sufficient to relieve pocket protein-mediated inhibition of E2Fs when 40S ribosome biogenesis is inhibited. This raises the possibility that phosphorylation and inhibition of pocket proteins by CYCD1/CDK4 require prior phosphorylation of pocket proteins by an as yet unidentified kinase that fails to be activated when ribosome biogenesis is inhibited. Indeed, it has been shown in an in vitro system that pRB can be phosphorylated by a cyclin C/CDK3 complex as soon as cells arrested in G0 are stimulated to re-enter the cell cycle, and that this phosphorylation is necessary for the G0/G1 transition [62]. If a similar mechanism is disrupted in cells depleted of rpS6, this would suggest that inhibition of ribosome biogenesis impairs proliferation by targeting cell cycle regulators that act very early during cell-cycle re-entry. In conclusion, the observations of Volarevic et al. [56] imply that, during the proliferative response of hepatocytes, a checkpoint operates that target the expression of genes whose products promote cell proliferation in order to halts cell-cycle progression when ribosome biogenesis is impaired (Figure 37.4c).

CONCLUSION The paper by Volarevic et al. [56] emphasizes the importance of the proper execution of ribosome biogenesis for cell-cycle progression to take place. A number of questions remain to be answered concerning the components of the surveillance mechanism that promotes cell-cycle arrest when ribosome biogenesis is inhibited. First, it is not known how a defect in ribosome biogenesis is detected by the cell. One possibility is that an rRNA precursor that accumulates at abnormally high levels, the 34S rRNA in the case of S6 depletion, is recognized by sensing machinery that, in turn, transduces a signal to an effector system to inhibit cell cycle progression. Findings in other systems have clearly shown that the tumor suppressor p53 is a key player in promoting cell cycle arrest when ribosome biogenesis is inhibited [63–65]. It will be important, therefore, to determine whether inhibition of ribosome biogenesis in

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the liver results in induction of p53 activity. Also, knowledge is lacking concerning the pathways that regulate ribosome biogenesis in response to cytokines and growth factors during the different phases of liver regeneration. The observation that protein deprivation impairs liver regeneration [66, 67] underscores the importance of the role played by nutrients during liver regeneration, possibly by controlling the rates ribosome biogenesis through the mTOR pathway. In the future, the utilization of genetic models and the verification in the liver of findings gathered from other systems promises to increase our knowledge of the regulation of ribosome biogenesis and of its crosstalk with the cell cycle machinery during liver regeneration.

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Liver Repopulation by Cell Transplantation and the Role of Stem Cells David A. Shafritz1, Michael Oertel1 , Mariana D. Dabeva1 and Markus Grompe2 1 Marion

Bessin Liver Research Center, Albert Einstein College of Medicine, Bronx, NY, USA 2 Oregon Health Sciences University, Portland, OR, USA

INTRODUCTION The major impetus for trying to reconstitute the liver by cell transplantation derives from the very high regenerative capacity of this organ, which has been of interest since biblical times (ancient Egypt, Babylonia, and Greece). Studies of liver tissue transplantation in the modem era began in the early 1900s, when liver fragments were transplanted into the anterior chamber of the eye, but the transplanted liver tissue degenerated rapidly and disappeared within a few days [1]. The first successful report of liver cell transplantation was in the 1970s, when isolated hepatocytes were transplanted to the liver, leading to transient reduction in serum bilirubin in the Gunn rat model for Crigler–Najjar syndrome, Type 1 [2]. Since these initial studies, much progress has been made defining the cell types that can repopulate the liver and restore function, and the host liver conditions (physiological or pathophysiological) under which effective repopulation can be achieved. Both adult hepatocytes and hepatic stem-like progenitor cells (stem/progenitor cells), and The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

also stem and progenitor cells of non-hepatic origin, have been used for transplantation into the liver. These issues will be covered in the text, including final sections dealing with studies conducted to date in humans and future horizons.

HEPATOCYTE TRANSPLANTATION: RATIONALE AND EARLY STUDIES Currently, orthotopic liver transplantation (OLT) is the only method available to treat acquired and genetic hepatic disorders effectively, especially when these diseases reach their end stages [3–5]. However, the number of patients who can benefit from whole liver transplantation is limited by the availability of donor organs. In addition, liver transplantation is expensive, carries significant morbidity and mortality, and requires long-term immunosuppression. Since many disorders treated by liver

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transplantation are caused by simple dysfunction of hepatocytes, in principle it should not be necessary to replace the entire organ. This is particularly true for genetic deficiencies of proteins produced specifically by hepatocytes in which selective replacement by normal hepatocytes would clearly be therapeutic. Examples include hemophilia, hypercholesterolemia, and phenylketonuria (diseases in which there is no liver injury) and Wilson’s disease, α1 -antitrypsin deficiency, and hemochromatosis (diseases in which there is extensive liver injury). In inherited disorders, cell transplantation could, in principle, also be performed in an autologous setting in which the patient’s own cells are genetically manipulated ex vivo and then transplanted back into the liver without the need for immunosuppression (and indeed this has been done) [6]. However, only a small percentage of the total hepatocellular mass (∼1–2% maximum) can be replaced by hepatocyte transplantation without causing portal hypertension and hepatic infarction. Therefore, in most instances, it will be necessary to expand the therapeutic cells after they have been transplanted. Attempts to increase the proportion of transplanted hepatocytes in the liver simply by stimulating liver regeneration (for example through the use of partial hepatectomy (PH) or carbon tetrachloride (CCl4 )-induced hepatic necrosis) have generally failed to show significant benefit. This is not surprising if one considers that, on average, hepatocytes need to undergo only one or two rounds of cell division to replace the mass of liver removed by two-thirds PH [7, 8], and both endogenous and transplanted hepatocytes contribute equally to this very limited proliferative response. Performing repeated PH or CCl4 administration is also not effective [9], since both transplanted and host hepatocytes can again respond similarly to this regenerative stimulus. Repeated cell transplantations have also been performed [10, 11], but this has not significantly increased the efficacy of liver replacement by transplanted cells.

BASIC REQUIREMENTS FOR LIVER REGENERATION In the normal adult liver, hepatocytes are in a quiescent state and turn over very slowly (only two to three times per year). However, following surgical reduction of liver mass or extensive acute toxic liver injury, hepatocytes rapidly enter the cell cycle and proliferate to restore liver mass. Many years ago, it was shown that during liver regeneration, 70–90% of the residual mature hepatocytes engage in DNA synthesis and undergo cell division [7]. Liver regeneration is a highly organized, complex, multistep process that involves growth factors and cytokines, gene transcription factors, cell signaling pathways and

expression of cell cycle regulatory genes [12, 13; see Chapter 36]. From these studies, it has been concluded that the proliferative activity of adult hepatocytes is sufficient to regenerate the liver following two-thirds PH and that participation of stem cells is not required [14].

MOLECULAR REGULATION OF LIVER MASS For many years, it has been known that the liver size (mass) is proportional to total body weight, ranging from 3 to 5% in different mammalian species. After two-thirds PH, cellular proliferation begins within 12–18 hours and the liver size returns to normal within 1–2 weeks in rodents and after 4–12 months in humans. When an undersized liver is transplanted, it grows to the expected full size for the host, and when an oversized liver is transplanted, its size is reduced to the expected mass compared with total body weight. However, until very recently, nothing was known concerning how this process is controlled. Pan and colleagues [15] have recently shown that mammalian genes comparable to those in the Drosophila Hippo kinase signaling cascade, that regulates wing mass during development, can control hepatocyte proliferation. When YAP, the mammalian counterpart to Yorki, the last gene in the Drosophila Hippo kinase cascade, is overexpressed in a transgenic mouse model, hepatocyte proliferation becomes unchecked and there is massive liver hyperplasia and hepatic carcinogenesis. When YAP hyperexpression is turned off or blocked, liver size returns to normal. YAP is a transcriptional activator, whose function is regulated by phosphorylation of a specific serine residue (S127). Several important questions regarding this pathway remain to be explored. Most important is the question of whether YAP is a physiological regulator of liver size or whether the reported phenotype is an aberrant result of YAP overexpression. Certainly, other factors such as HGF, can also produce extensive reversible liver hypertrophy when over-expressed. Loss of function studies will clarify this problem. In addition, it is unknown (i) whether YAP expression or Yap S127 phosphorylation increases and decreases during normal liver regeneration, consistent with it being a regulator of hepatocyte proliferation, (ii) what the downstream targets of YAP that regulate hepatocyte proliferation are, and (iii) whether YAP-regulated transgenic hepatocytes will be able to repopulate a normal liver after their transplantation. Answers to these questions could have major implications in terms of finding ways to stimulate host liver cell proliferation. This could be important in restoring liver regeneration in various pathological states, and also in regulating proliferation of transplanted cells during liver repopulation.

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uPA transgenic mice, and they observed extensive liver repopulation (Figure 38.1b), estimating that each transplanted hepatocyte that had engrafted into the uPA host liver, underwent 12–14 cell divisions [17]. Another mouse model for liver repopulation was generated by targeted disruption of the last gene in tyrosine catabolism, fumarylacetoacetate hydrolase (Fah) [18]. Deletion of Fah leads to accumulation of upstream intermediates in tyrosine catabolism, some of which (namely, fumaryl acetoacetate) are toxic and cause extensive and continuous liver injury. The Fah null mouse represents an animal model for the human metabolic disorder hereditary tyrosinemia, Type 1 (HT1), which causes extensive liver injury, hepatocellular carcinoma, and death at an early age in affected individuals. Administration of 2-(2-nitro-4-trifluoromethylbenzoyl)cyclohexane-1,3dione (NTBC), a pharmacological inhibitor of tyrosine catabolism upstream of homogentisic acid, prevents accumulation of maleyl acetoacetate and fumaryl acetoacetate and is partially successful in treating patients with HT1 [19], although it does not prevent hepatocarcinogenesis [20]. NTBC also allows Fah null mice to survive and liver failure occurs in these mice only when NTBC is discontinued. After transplanting syngeneic wild-type (wt) hepatocytes into Fah null mice maintained on NTBC, only scattered small clusters of transplanted hepatocytes are detected (Figure 38.2). However, if NTBC treatment is

ANIMAL MODELS TO AUGMENT LIVER REPOPULATION BY TRANSPLANTED HEPATOCYTES For many years, it was thought that mature hepatocytes could undergo only two or three divisions, after which they become terminally differentiated and are incapable of further proliferation. However, during the last decade, it has been shown in several rodent model systems that under specialized circumstances hepatocytes retain their high proliferative capability and can extensively repopulate the liver. Initially, Sangren et al. [16] developed a transgenic mouse model in which a protease, urokinase plasminogen activator (uPA), is expressed exclusively in hepatocytes under control of the albumin (Alb) promoter. In this model, tissue protease activity caused continuous and extensive liver injury and sub-fulminant liver failure, leading to death of the mice at 4–6 weeks of age. However, some mice survived and, in these mice, there were scattered nodules of normal liver tissue distributed throughout the hepatic parenchyma (Figure 38.1a). This occurred by deletion of the uPA transgene from individual hepatocytes, which then clonally expanded into large clusters that replaced damaged tissue. These findings prompted these investigators to transplant normal hepatocytes (containing only a β-galactosidase marker gene) into

(a)

R W

(b)

Figure 38.1 Liver regeneration and hepatocyte transplantation in the uPA transgenic mouse. (a) Wt mouse liver (left), regenerating uPA transgenic mouse liver by deletion of uPA transgene (middle), uPA transgenic liver (right). (b) LacZ transgenic mouse liver (left), uPA transgenic mouse liver (middle), uPA transgenic mouse liver repopulated with LacZ hepatocytes (right)

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THE LIVER: ANIMAL MODELS TO AUGMENT LIVER REPOPULATION BY TRANSPLANTED HEPATOCYTES

(a)

(b)

(c)

Figure 38.2 Repopulation of the Fah null mouse liver by wt hepatocytes: (a) 2 days, (b) 3 weeks, and (c) 6 weeks after cell transplantation. The same results as in (a) were obtained at 6 weeks after hepatocyte transplantation in the absence of NTBC withdrawal

discontinued shortly after cell transplantation, liver injury resumes and transplanted cells proliferate extensively, forming large clusters within 3 weeks and replacing most of the liver mass within 6 weeks. Fah null mice with repopulated livers remain healthy, have normal liver function tests and show a relatively normal liver structure for many months after wt hepatocyte transplantation [18]. These studies provide proof of principle that liver repopulation can effectively cure a metabolic disease, namely, the mouse equivalent to HT1. Transplanted repopulating hepatocytes integrate into the host hepatic architecture and express all functions necessary for normal health of the animals. In the Fah null mouse, not only do transplanted wt hepatocytes replace Fah null hepatocytes, but the transplanted cells can also be serially transplanted through seven consecutive Fah null mice, while retaining full ability to proliferate and replace host hepatocytes [21]. In these studies, it was calculated that each serially transplanted hepatocyte underwent an average of at least 69 cell divisions. Thus, murine hepatocytes exhibit essentially infinite capacity to proliferate and restore liver function under circumstances in which there is both massive and continuous liver injury and the transplanted hepatocytes have a significant selective advantage for survival compared with host hepatocytes [18, 21]. Therefore, under the specialized circumstances existing in the liver of Fah null mice, hepatocytes exhibit many properties of stem cells, except for the ability to differentiate into more than one lineage, in this case into both hepatocytes and bile duct epithelial (BDE) cells. However, the ability of hepatocytes to differentiate into BDE cells has been demonstrated in another rodent animal model system in which extensive bile duct injury is produced by administration of methylenedianiline (4,4′ -diaminodiphenylmethane, DAPM) [22]. This change in cellular phenotype from an hepatocyte to a BDE cell is referred to as “transdifferentiation” or cellular plasticity and is thought to occur through changes in the microenvironment in which a particular target cell (in this case, the hepatocyte) is located. Thus, mature adult hepatocytes through an epigenetic phenomenon, induced by a specific set of pathophysiological circumstances, exhibit

properties heretofore not previously imagined for “terminally differentiated” or near “terminally differentiated” hepatocytes. In both the uPA transgenic and Fah null mouse models, successful liver repopulation by adult hepatocytes required two experimental conditions: (1) the liver was under massive and continuous liver injury and (2) the transplanted hepatocytes had a strong selective advantage for survival in the host liver. In the Fah model, it has been shown that this selective advantage is based on a p21-dependent cell cycle arrest in the host hepatocytes, resulting from p21 induction caused by DNA damage [23]. Thus, an alternative strategy to obtain a high level of liver repopulation by transplanted hepatocytes is to block proliferation of endogenous hepatocytes using exogenous agents and then transplant normal hepatocytes in conjunction with a liver proliferative stimulus. The first method described to achieve this effect was to treat rats with retrorsine, a plant alkaloid that is taken up and metabolized selectively by hepatocytes to produce a DNA alkylating agent that cross-links cellular DNA and disrupts hepatocyte division [24]. When retrorsine or a related compound, monocrotaline, is administered to rats or mice, there is a long-lived inhibition of hepatocyte proliferation. However, the basic metabolic functions of DNA damaged hepatocytes are maintained and the animals survive. After the effects of acute chemical injury have subsided (2–4 weeks), the animals are subjected to two-thirds PH or CCl4 administration in conjunction with transplantation of hepatocytes from normal animals. This leads to a brisk regenerative response by transplanted hepatocytes and there is near total repopulation of the liver in 3–6 months [24]. Another method to achieve effective liver repopulation by transplanted hepatocytes is to induce DNA damage with selective liver irradiation in conjunction with hepatocyte transplantation and either two-thirds PH, CCl4 administration, or ischemic liver injury [25, 26]. Most recently, in X-irradiated mice, administration of HGF through introduction of a recombinant adenovirus vector expressing this growth factor has been used to replace PH as a liver regenerative stimulus [27]. With retrorsine or monocrotaline treatment of the host liver, transplanted hepatocytes have a proliferative advantage over host hepatocytes. With

38: LIVER REPOPULATION BY CELL TRANSPLANTATION AND THE ROLE OF STEM CELLS

retrorsine-induced DNA damage, host hepatocytes exhibit an increased level of apoptosis [28], which could also contribute to liver repopulation by transplanted cells. Although the above procedures cause massive liver damage and are unlikely to be used in patients, the hope of cell transplantation investigators is to develop less toxic methods that will be usable clinically.

“OVAL CELLS” AS HEPATOCYTE PROGENITORS Although the term “oval cells” is widely used in the field, it is important to emphasize that this term is used for a highly heterogeneous population of cells. No uniformly accepted definition exists and therefore it is not surprising that reports on the properties of “oval cells” are sometimes conflicting. Not only do multiple different cell types emerge in livers undergoing “oval cell” activation, but it is also not clear whether “oval cells” from different animal species or from different hepatic injuries are comparable. The reagents to understand heterogeneity better within these injury-activated populations are only now becoming available. In the future, it is likely that the term “oval cell” will be replaced by more specific designations, based on expression of clearly defined markers. The term “oval cells” was first coined by Farber [29] to describe non-parenchymal cells in the periportal region that were present after treatment of rats with carcinogens: ethionine, α-acetaminofluorene (2-AAF), and 3-methyl-4-diethylaminobenzene. Other methods to induce proliferation of “oval cells” are to treat rats with d-galactosamine [30, 31], a choline-deficient (CD)/ ethionine-substituted diet [32, 33], or allyl alcohol [34], or to treat mice with dipin [35] or 3,5-diethoxycarbonyl-1,4-dihydrocollidine (DDC) [36]. In each of these models, cells are induced that have a small, ovalshaped, pale-stained blue nucleus and very scant, lightly basophilic cytoplasm. Farber did not believe that “oval cells” were hepatocyte progenitors, but Thorgeirsson and co-workers [37] demonstrated that “oval cells” induced to proliferate in the periportal region after treatment of rats with 2-AAF followed by two-thirds PH subsequently differentiated into distinct clusters of basophilic hepatocytes. This was demonstrated by pulse labeling of the liver with [3 H]thymidine and following the progression of label from periportal “oval cells” to hepatocytic clusters in the mid-parenchyma in conjunction with the kinetic pattern of expression of bile ductular (CK-7 and CK-19) and hepatocytic [α-fetoprotein (AFP) and Alb] markers over time [37, 38]. Other indirect evidence suggesting that “oval cells” are hepatic progenitors is their expression of c-kit [39], CD34 [40], flt3 receptor [41], and LIF [42], all known to be expressed in hematopoietic stem cells or their immediate derivatives. Sca-1, another cell surface marker gene expressed by hematopoietic stem cells of the mouse, is also expressed in fetal liver epithelial

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cells [43] and “oval cells” in the adult mouse liver [44, 45]. In most recent lineage tracing studies using a double transgenic mouse expressing a β-galactosidase reporter gene under control of the Foxl1 promoter, Greenbaum and co-workers [46] have shown the production of both hepatocytes and BDE cells from progenitor cells induced to proliferate by bile duct ligation or feeding mice a DDC-containing diet, thus extending and strengthening conclusions from earlier studies in 2-AAF/PH treated rats [37, 38]. A role for “oval cells” in normal liver physiology and cell turnover has not been established. However, as indicated previously, “oval cells” are induced to proliferate when there is liver injury superimposed on circumstances in which hepatocyte proliferation is impaired. These cells exhibit many features of progenitor cells, dividing rapidly and appearing to differentiate into both hepatocytes and BDE cells. Thorgeirsson and colleagues [47] performed extensive immunohistochemical and ultrastructural studies in which they demonstrated that “oval cells,” induced to proliferate by 2-AAF/PH, are derived from undifferentiated cells in the Canals of Hering, after which they pass through discontinuities in the laminar basement membrane of the ductal limiting plate and join together with stellate cells as they enter the hepatic parenchyma, proliferate, and differentiate into hepatocytes. Attempts to establish specific markers for “oval cells” to distinguish them from mature hepatocytes and BDE cells, and to determine their lineage origin (mesoderm or endoderm), have led to conflicting findings. All investigators agree that “oval cells” express common liver epithelial progenitor cell markers, such as AFP and Alb for hepatocyte progenitors and CK-19 (and OV6 in the rat) for bile duct progenitor cells. They were initially thought to express hematopoietic stem cell markers, c-kit, CD34, and Thy 1 [39, 40, 45, 48, 49], but several recent studies have reported that both fetal liver progenitor cells and “oval cells” are negative for these markers [43, 50–53].

TRANSPLANTATION OF “OVAL CELLS” If “oval cells” are indeed stem cells or hepatic progenitor cells, they should be able to restore liver mass after their transplantation. About 20 years ago, Faris and Hixson [54] reported that “oval cells” isolated from the liver of rats fed a CD diet, treated with 2-AAF, and transplanted into the liver of secondary hosts produced “colonies” or clusters of cells with an hepatocytic phenotype in recipients that had also been subjected to the CD diet but not in recipients that had received a normal diet. However, the level of liver repopulation by transplanted CD/2-AAF “oval cells” was not determined. “Oval cells” isolated from the liver of rats treated with d-galactose (d-Gal) also proliferate and differentiate into hepatocytes after their transplantation into rats undergoing two-thirds PH [55]. However, in

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THE LIVER: STEM CELLS IN THE ADULT LIVER AND THE HEPATIC STEM CELL NICHE

these studies, liver repopulation by transplanted d-Gal induced “oval cells” was also fairly low and not quantified. Duct-like epithelial cells isolated from the atrophic pancreas of rats undergoing treatment with a copper chelating agent (trien) also proliferate modestly and differentiate into hepatocytes after transplantation into normal rat liver [55]. Isolated pancreatic cells from normal mice also repopulate the liver of Fah null mice [56]. “Oval cells” isolated from the liver of DDC-fed mice also repopulate the liver of Fah null mice, albeit with less efficiency than with mature hepatocytes [57]. Similarly, “oval cells” from GFP transgenic mice maintained on a DDC diet also repopulate the liver of wt mice treated with monocrotaline in conjunction with PH [58]. Other recent studies have also shown effective repopulation of the liver by purified “oval cells” in both rats treated with retrosine [53] and in Fah null mice [59], but not in animals with a normal liver. Most recently, “oval cells” have been isolated from normal mouse and dog liver [44, 60]. These cells exhibit hepatic progenitor cell characteristics in culture, but in vivo repopulation data are very limited. Numerous studies have reported the isolation of “oval cell” lines from mice and rats, and also from humans, that are clonal, bipotent, and exhibit other stem and progenitor cell characteristics in vitro and in vivo (for a review, see [61]). This has provided valuable information concerning the basic biological properties of these cell lines, but, in general, in vivo repopulation by “oval cell” lines has been very low.

STEM CELLS: THEIR ORIGIN AND PROPERTIES Considerable attention has been focused on the use of stem cells to reconstitute liver mass. The earliest stem cells originate from the inner cell mass of the blastocyst during development and are pluripotent (capable of differentiating into all the cell types in mammalian species); they are generally referred to as embryonic stem (ES) cells [62]. These cells give rise to somatic stem cells that subsequently differentiate into multipotent tissue-specific stem cells [62–64]. The latter give rise to lineage-committed progenitor cells that proliferate and differentiate into mature phenotypes that ultimately become the somatic, tissue-specific, working cells in different organs (Figure 38.3). Through studies of cell transplantation to reconstitute the hematopoietic system, and also studies of cell turnover in other tissues undergoing rapid and continuous regeneration, such as skin and intestinal epithelium, stem cells have been shown to exhibit four essential properties: (1) they have the capacity to maintain themselves (self-renew), while at the same time they generate progeny that differentiate into mature cellular phenotypes; this is referred to as asymmetric cell division [65, 66]; (2) they are multipotent, that is, capable of producing differentiated cells in at least two lineages; (3) their progeny are stable, reconstitute organ mass and

Inner cell mass Blastocyst

Embryonic stem (ES) cell

Tissue-specific multipotent stem cell

Lineage committed progenitor cells

Differentiated mature cells

Bone

Liver

Heart

Lung

Pancreas

Brain

Figure 38.3 Stem cells and tissue differentiation

remain functional in the tissue for a long time; and (4) by virtue of their capability of self-renewal, stem cells can be transplanted serially through successive hosts.

STEM CELLS IN THE ADULT LIVER AND THE HEPATIC STEM CELL NICHE Although recent studies claim to have identified, isolated, and purified hepatic epithelial stem cells from the adult liver, most of the evidence is through in vitro cell culture studies. None have achieved long-term, functional reconstitution of liver tissue or serial transplantability of these cells. Since unique markers for hepatic specified endodermal stem cells have not been identified, whether such cells exist in the adult liver still remains to be established. New monoclonal antibodies to mouse progenitor cells may prove useful in this regard [67]. If such liver stem cells do exist, the question remains as to where they reside (i.e. where is their “niche”?) (Figure 38.4). The original idea of a stem cell “niche” evolved from the concept that stem cells reside in tissues within an “inductive microenvironment” that directs their differentiation [68, 69]. More recently, the stem cell niche has been described further as “a specific location in a tissue where stem cells can reside for an indefinite period and produce progeny cells, while self-renewing” [70]. Local

38: LIVER REPOPULATION BY CELL TRANSPLANTATION AND THE ROLE OF STEM CELLS

“Oval cells”

Canal of Hering Hepatocytes Bile ducts

Figure 38.4 Canal of Hering as the proposed liver stem cell “niche”

stromal cells and other extracellular environmental factors attract stem cells to these niches and affect their behavior (i.e. gene expression program, proliferation, and/or differentiation), and such niches have been identified in the bone marrow, brain, skin, and intestinal mucosa [71–75]. At present, the most likely candidate for a liver stem cell niche is the Canals of Hering. The Canals of Hering were originally identified more than 100 years ago as luminal channels linking the hepatocyte canalicular system to the biliary tree [76]. These channels contain small undifferentiated epithelial cells [77, 78] that are in direct physical continuity with hepatocytes at one membrane boundary and bile duct cells at another boundary to form duct-like structures, enclosing a lumen (i.e. the canal; Figure 38.5). In most models of liver structure, the Canals of Hering are depicted as very limited structures confined to the portal space. However, recent studies in humans have demonstrated that the Canals of Hering are much more elaborate than previously thought and extend far into the hepatic parenchyma [79, 80]. These structures contain epithelial cells with dual expression of bile ductular and fetal hepatocytic markers (AFP, HepPar1, CK-19)

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and were thus considered to represent “facultative hepatic stem cells” [79]. Recent studies in the rat 2-AAF/PH model have demonstrated that proliferating “oval cells” are indeed located in the Canals of Hering [47]. These proliferating cells are thought to pass into the parenchyma through discontinuities in the ductal basement membrane, continue to proliferate in the parenchyma in conjunction with stellate (Ito) cells and then differentiate into hepatocytes [47] or undergo apoptosis [81]. Because the very first cell to undergo proliferation and subsequent differentiation into hepatocytes is found in the Canals of Hering, this structure is thought to contain both stem cells and their initial progeny, “oval cells”. A separate population of less well-differentiated progenitor cells has also been identified in the periductular space and these cells proliferate following allyl alcohol-induced liver injury [34]. It has been suggested that these cells comprise a second progenitor or stem cell compartment and that they may originate from hematopoietic stem cells [82]. Hence the liver may have several distinct hepatic stem and progenitor cell niches. A specific method developed recently to identify stem cells in vivo in rapidly turning over tissues (such as skin epithelium) is to pulse-label them with BrdU, which is taken up by the cells and becomes stably incorporated into DNA. Over time, most of the cells that are pulse labeled with BrdU turn over and BrdU is lost from the tissue. However, some cells may retain the label. These are thought to be stem cells that have become quiescent, while still maintaining their undifferentiated state. These cells, referred to as “label-retaining cells”, have been identified by H2B-GFP labeling in hair follicles of the skin [83], that is, the so-called “bulge cells,” that have been shown to regenerate complete new hair follicles [84]. However, the validity of the label-retaining method to identify stem cells has recently been challenged [85]. Nonetheless, recent preliminary studies have reported the presence of four distinct populations of “label-retaining cells” in the liver (i.e. cells in the Canals of Hering, intralobular bile duct cells, periductular “null” or undifferentiated mononuclear cells, and peribiliary hepatocytes) [86]. These cells were identified by BrdU pulse labeling during acute liver injury with acetaminophen and 2 weeks later “washing out” the label by a second liver regenerative stimulus with acetaminophen. The animals were then examined for cells retaining BrdU label at 4 and 8 weeks following the second acetaminophen administration. However, to determine which of these cells represent hepatic stem cells will require further study.

HEPATIC STEM CELLS IN THE FETAL LIVER Figure 38.5 Electron micrograph of the Canal of Hering. The Canal of Hering (*) is bounded by two mature hepatocytes (Hcs) and one undifferentiated epithelial cell (Ec)

Classical embryological studies have traced the proliferation and differentiation of tissue-specified stem cells

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THE LIVER: LIVER REPOPULATION BY FETAL LIVER STEM/PROGENITOR CELLS

ED8.0

ED8.5–9.5

ED11–15

~ED16

Birth A1b+

AFP+

AFP+/AIb+/CK-19+

H

OS

, GF

M

Hepatocytes (unipotent)

No

sig (FGFs & BMP-4)

(bipotent)

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na

tch

CK-19+

lin

g

Bile duct cells (unipotent)

Figure 38.6 Schematic diagram of liver development

into the hepatocytic and biliary epithelial (cholangiocytic) lineages during normal development in the fetal liver (Figure 38.6). In the mouse, endodermal stem cells begin to proliferate when the ventral wall of the endoderm becomes positioned next to the developing heart on embryonic day (ED) 8.0 [87, 88]. Specification toward the hepatic epithelial lineages occurs at ED8.5 and requires fibroblast growth factor (FGF) signaling from the cardiogenic mesoderm [89] and bone morphogenic protein (BMP) signaling from the septum transversum mesenchyme [90]. By ED 9.0–9.5, these cells begin to express GATA4 and liver-enriched, nuclear transcription factor HNF4α, and also liver-specific genes, AFP followed by Alb [88, 91]. The hepatic-specified cells are now referred to as hepatoblasts and proliferate massively. Cords of hepatoblasts invade the septum transversum mesenchyme that contains stellate cells and sinusoidal endothelial cells. These cells secrete a variety of cytokines and growth factors, such as EGF, FGF, HGF, TGFβ, TNFα, and IL-6, that are known to be involved in liver development, and also in the hepatocyte proliferative response during liver regeneration [12, 88–90]. At ED11, hematopoietic stem cells invade the liver bud to form a visible liver structure that is primarily a hematopoietic organ. Hepatoblasts continue to expand rapidly and begin to express numerous liver-specific genes [92]. Just prior to ED16, hepatoblasts diverge along two lineages, hepatocytes and cholangiocytes [91, 93]. Differentiation along the cholangiocytic lineage is promoted by Notch signaling and is antagonized by HGF, which in conjunction with oncostatin M promotes hepatocytic differentiation [94]. After ED16, there is a massive change in the gene expression profile of rat fetal liver epithelial cells to a more differentiated phenotype [95] and the percentage of bipotent cells, that is, those expressing genes in both the hepatocytic and cholangiocytic lineages (e.g. AFP, Alb, and CK-19), is markedly reduced [93, 96, 97]. At

this point, most of the cells are unipotent and irreversibly committed to either the hepatocytic or the cholangiocytic lineage [96, 97]. As organogenesis proceeds, intrahepatic bile ducts are formed in the vicinity of large portal vein branches, beginning on ∼ED17 [98]. Thus, the fetal liver contains cells that are in different stages of hepatic epithelial lineage progression. These cells have been isolated, cultured, and transplanted into various animal model systems and fetal liver epithelial cells exhibit superior properties to mature hepatocytes in terms of durability, function, and repopulation of the normal adult liver.

LIVER REPOPULATION BY FETAL LIVER STEM/PROGENITOR CELLS The ultimate test for a putative stem cell is to demonstrate its ability to self-renew in vivo and to repopulate functionally a tissue or organ, long-term. Sandhu et al. [97] reported 5–10% repopulation of DPPIV− mutant F344 rat liver by transplanting wt ED14 fetal liver epithelial cells in conjunction with two-thirds PH. Liver repopulation by transplanted cells increased progressively over 6 months, and the bulk of repopulating clusters contained both hepatocytes and mature bile ducts. The transplanted cells were integrated into the host parenchyma and formed hybrid bile canaliculi with host hepatocytes. Thus, transplanted rat ED14 fetal liver epithelial cells exhibited three major properties of liver stem cells: (1) extensive proliferation, (2) bipotency, and (3) long-term repopulation in vivo [97]. Liver repopulation by transplanted rat fetal liver cells was achieved in a non-selective host liver environment but required PH to initiate the process. This is consistent with studies in hematopoietic stem cells, in which hematopoietic reconstitution does not occur unless there is near total bone marrow ablation in the host.

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with fetal liver epithelial cells, they are referred to as fetal liver stem/progenitor cells (FLSPCs), rather than stem cells. The mechanism of liver repopulation by rat FLSPCs has been shown to be cell competition between the transplanted cells and host hepatocytes [99], a process that was originally described in Drosophila during wing development [101, 102]. These cells have been cryopreserved with full ability to repopulate the normal adult liver after thawing [103] and, most recently, rat FLSPCs have been enriched to 95% purity by selection with immunomagnetic beads [100].

In ED14 rat fetal liver, there are three distinct populations of epithelial cells, those positive for AFP and Alb but negative for CK-19, those positive for AFP, Alb, and CK-19 and those positive for CK-19 but negative for AFP and Alb [96]. The number of AFP+ /Alb+ /CK-19+ cells decreased dramatically at ED16, after which liver repopulation potential of rat fetal liver cells also decreased dramatically [97]. The level of liver repopulation by ED14 fetal liver cells under non-selective conditions (i.e. in a normal liver) can also be increased to 20–25% by increasing the number of ED14 fetal liver cells transplanted (Figure 38.7) [99]. Repopulation continues to increase for up to 1 year, reaching an average of ∼30% for the total liver, and remains stable for the life of the animal (Oertel M. et al., unpublished data). This represents a several thousand-fold amplification of transplanted fetal liver epithelial cells in the host organ. Both hepatic parenchymal cords and mature bile ducts are formed by transplanted fetal liver cells, and the progeny of the transplanted cells express normal levels of hepatocytic and cholangiocytic genes in the respective cell types [99, 100]. Since serial transplantation has not yet been demonstrated

LIVER REPOPULATION BY EXTRAHEPATIC AND EMBRYONIC STEM CELLS Various studies have reported that cells are released from the bone marrow (BM) into the circulation, migrate to the liver and differentiate into hepatocytes. However, the

(a)

(b)

Figure 38.7 Repopulation of the normal adult rat liver by fetal liver stem/progenitor cells. (a) Two examples of whole rat liver sections repopulated by ED14 fetal liver cells. (b) Selected areas of repopulated liver at higher magnification showing both hepatocytes and mature bile duct structures

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THE LIVER: OTHER ROLES FOR BM CELLS IN LIVER REGENERATION

extent to which this occurs and the mechanism(s) involved remain highly controversial (for reviews, see [104–107]). Estimates of liver repopulation by hematopoietic cells vary widely, ranging from < 0.01 to 40%. Originally, Petersen et al. reported that BM stem cells from DPPIV+ F344 rats transplanted into sublethally irradiated DPPIV− F344 rats repopulate the BM and then migrate to the liver and “transdifferentiate” into hepatocytes through the liver “oval cell” progenitor pathway [108]. This mechanism was generally accepted until studies by Wang et al. [57] using lacZ marking showed that BM cells did not enter the “oval cell” pool in wt mice treated with DDC or contribute to liver repopulation by “oval cells” in secondary Fah −/− mouse recipients. Menthena et al. [109] also showed in rats that DPPIV+ BM cells transplanted into DPPIV− rats contributed < 1% to “oval cells” expanded by three different methods: (1) 2-AAF/PH, (2) retrorsine/PH, or (3) d-Gal induced liver injury. In Fah −/− mice and other model systems, it has been shown that cell fusion and reprogramming, rather than transdifferentiation, is the mechanism by which hematopoietic cells acquire an hepatocytic phenotype. Initial studies in cell culture showed that BM and neuronal cells can fuse with ES cells [110, 111]. Wang et al. [112] and Vassilopoulos et al. [113] subsequently showed that hematopoietic stem cells fuse with hepatocytes in Fah null mice to produce cells expressing the deficient enzyme, which then expand massively to restore liver mass and function [112, 113]. Fusion also occurs between hematopoietic cells and neurons or muscle cells [114, 115] and it has been shown that myelomonocytic cells can fuse with hepatocytes [116, 117] or muscle cells [118] to produce somatic hybrids expressing genes from both parental cell types. Other studies have reported that fusion does not appear to be required for BM-derived cells to differentiate into hepatocytes [119–121]. Unfractionated or CD34+ enriched cells from human cord blood [122–125], multipotent adult progenitor cells (MAPCs) [126, 127], or mesenchymal stem cells [128–133] have been transplanted into the liver of immunodeficient mice. These transplanted cells express a differentiated hepatocytic phenotype [122–133], but liver repopulation and physiological restoration were once again very low. Several studies have reported that mesenchymal stem cells, isolated from adipose tissue and differentiated in culture along the hepatocytic lineage, can also engraft in the liver parenchyma and contribute to liver regeneration [134, 135]. One study [135] reported large repopulation clusters with hepatocyte-differentiated mesenchymal stem cells, but this required retrorsine treatment. These studies are promising, but the ability of mesenchymal stem cells to repopulate the adult liver under more normal, clinically viable circumstances will need to be established. ES cells in culture can be induced along the endodermal and hepatocytic lineages by addition of specific

cytokines and growth factors [136–143]. The first step typically involves the generation of embryoid bodies, followed by the induction of definitive endoderm using activin A. The endodermal population can then be further specified toward the hepatic lineage using BMP-4 and basic FGF. Cells produced in this fashion express typical markers such as Alb, but usually also express AFP. They can then be transplanted into the liver with differentiation into both mature hepatocytes [142, 143] and BDE cells [143]. The level of liver repopulation obtained with hepatocyte-differentiated ES cells is very low but is somewhat higher when the cells are transplanted into MUP-uPA/SCID mice [143]. To date, all ES differentiation protocols generate “hepatocyte-like” cells, but not fully functional, mature, and transplantable equivalents of hepatocytes isolated from adult liver. It appears possible to select for more mature “hepatocyte-like” cells using surface markers, such as the asialoglycoprotein receptor for purification [144]. In the future, it is hoped that conditions will be developed in which lineage-specified ES cells will be therapeutically effective.

INDUCED PLURIPOTENT STEM CELLS (iPS) Because of their extensive proliferative capacity, pluripotent stem cells are an attractive potential source of transplantable hepatocytes. Not only can these cells divide extensively, but they also retain the ability to differentiate into multiple different mature cell types [62]. Until recently, ES cells were the sole source of pluripotent human cells and are ethically controversial. Now, however, pluripotent cells can be derived from direct genetic reprogramming of somatic cell types, such as dermal fibroblasts [145–147]. Studies of liver repopulation by these induced pluripotent stem cells (iPS) have not yet been reported, but this would have a major impact in advancing cellular therapy.

OTHER ROLES FOR BM CELLS IN LIVER REGENERATION Several studies have reported that injections of BM-derived stem cells can stimulate liver regeneration and restore liver function during chronic liver injury by enhancing the degradation of liver fibrosis in mice [148, 149]. Such events are associated with induction of metalloproteinases, especially MMPs 2, 9, and 13 [150]. Most recently, it has been reported that BM-derived endothelial progenitor cells (EPCs), injected into the spleen during liver injury, engraft in the liver, form new blood vessels, and secrete growth factors, such as HGF, TGFα, EGF, and VEGF, that stimulate liver regeneration and improve survival of animals with massive liver

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injury [151]. Thus, the role of BM stem cells in liver regeneration may be supportive in generating new parenchymal mass and, under some circumstances, in ameliorating hepatic fibrosis.

HUMAN “OVAL CELLS” AND STEM CELLS A human counterpart to “oval cell” activation has been described in liver tissue obtained from patients with extensive chronic liver injury or submassive hepatic necrosis, that is, the so-called “ductular reaction” (for a detailed description, see [152]). “Ductular reactions” are comprised of collections of cells in ductular arrays with the morphological appearance and immunohistochemical markers comparable to those found in rodent “oval cells.” These cells are present primarily in the portal tracts with extension into the parenchyma and express both hepatocytic and bile ductular markers, and also certain neuroendocrine genes [152–155]. Using double and triple label immunohistochemistry, Zhou et al. [156] have shown that “ductular reactions” are bipolar structures with cells at one pole exhibiting hepatocytic morphology and gene expression (HepPar1 or HepPar1/NCAM) and cells at the other pole exhibiting biliary morphology and gene expression (CK-19 or CK-19/NCAM), with undifferentiated epithelial cells in the center expressing only NCAM. Cells with similar morphological and immunohistochemical properties have also been identified in the human fetal liver beginning at 4 weeks gestation [157]. A number of investigators have isolated, cultured, and/or passaged human fetal liver epithelial cells with bipotent properties, and several of these studies have demonstrated their differentiation into hepatocytes after transplantation into SCID or nude mice [158–160]. Schmelzer et al. [161] have identified two populations of hepatic progenitor cells from human fetal, neonatal, and pediatric liver that exhibit stem cell properties. One population is thought to represent a hepatic stem cell (AFP− /Alb+ ) and the other a slightly more differentiated hepatoblast (AFP+ /Alb+ ). More recently, they reported data suggesting that these cells may reside in the Canals of Hering [162]. In culture, they can differentiate into hepatoblasts, possibly requiring cues from other cells that copurify with them during immunoselection. One curious question, which remains unanswered, is that the presumed hepatic-specified stem cells in human neonatal, pediatric, and adult liver are Alb positive but AFP negative, which is opposite to what one might expect based on expression of AFP before Alb during liver development [91]. Nonetheless, these studies suggest that a human liver somatic stem cell might exist and hopefully future studies will demonstrate in vivo self-renewal and long-term repopulation of the liver by these cells, proving that they are indeed stem cells.

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CLINICAL TRIALS OF HEPATOCYTE TRANSPLANTATION Allogeneic hepatocyte transplantation has had limited clinical success. In 1998, a child with Crigler–Najjar syndrome Type I, suffering from severe hyperbilirubinemia, was infused with 7.5×109 allogeneic donor hepatocytes via a portal vein catheter [163]. This resulted in a reduction, but not normalization, of serum bilirubin. Although the procedure was only partially effective, the patient still showed glucuronosyltransferase activity in the liver 3 years after cell transplantation (J. Roy Chowdhury, personal communication). Studies have also been conducted in patients with chronic liver disease and cirrhosis, again with only marginal, if any, success [164, 165]. Hepatocyte transplantation has also been used in conjunction with ex vivo retroviral gene therapy in five patients with a defect in the low-density lipoprotein (LDL) receptor [6]. In this study, the proportion of liver cell mass replaced was estimated to be ∼1%. The procedure resulted in a very modest decrease in plasma cholesterol in several of the patients. These results are encouraging, but also highlight another important problem of tissue replacement with standard hepatocyte transplantation protocols, namely that only limited (or no) division of transplanted cells takes place in the recipient liver under most circumstances. Therefore, augmented expression of the transferred gene through proliferation of transplanted cells, will be necessary for effective ex vivo gene therapy in most instances. The worldwide experience with human hepatocyte transplantation has been reviewed [166]. Data, compiled up to December 2005, represented experience in 78 patients who had received human hepatocyte infusions. Usually, suspensions of adult hepatocytes (autographs or allographs) were utilized in a single injection, although some patients received multiple infusions. Twenty-one patients had inherited metabolic disorders, including familial hypercholesterolemia (five patients), Crigler–Najjar syndrome Type 1 (four patients), factor VII deficiency (two patients), Refsum’s disease (one patient), progressive familial intrahepatic cholestasis (two patients), α1 -antitrypsin deficiency (one patient), and urea cycle defects (four patients). Nearly all patients showed initial improvement of metabolic function. Some patients subsequently received a liver transplant, so that the long-term effects of hepatocyte transplantation could not be assessed. However, in the remaining patients, the function of transplanted hepatocytes was not sustained. Whether this resulted from rejection of transplanted cells, their lack of proliferation, or eventual apoptosis has not been determined. In no patient was expansion of transplanted hepatocytes or progressive improvement of metabolic function documented. Twenty patients with chronic liver disease of varying etiologies have also

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been transplanted with adult hepatocytes, but results in these patients have not been encouraging. However, in short-term studies (33 days), some patients with alcoholic cirrhosis showed significant reduction in blood ammonia and encephalopathy [166]. Many additional trials are in progress, but the results of these studies are not yet available.

TREATMENT OF FULMINANT HEPATIC FAILURE Hepatocyte transplantation therapy for fulminant hepatic failure as a bridge to OLT has met with more encouraging results [164, 167]. Although the numbers of patients treated by this modality are low, the diseases are of varying etiologies and it is not possible to analyze the results by statistical methods. Data from 37 such cell transplants have been reported, many with reductions in both blood ammonia and encephalopathy and successful OLT. In several cases, the patients recovered without the need for a transplant, but others died. Two patients with fulminant hepatic failure received fetal liver cells and interestingly both survived.

XENOREPOPULATION MODELS The further development of hepatic cell therapy requires robust animal models to test potential sources of transplantable cells for their functional efficiency. Since human cells are rejected by other animals, this requires the use of immune-deficient animals or pharmacological immune suppression. Small rodents capable of harboring human hepatocytes are not only of interest as a test bed for cell therapy, but also for pharmaceutical research, including

(a)

testing of drug metabolism and virology (modeling of viral hepatitis). Two models capable of supporting extensive repopulation of the mouse liver with human hepatocytes have been reported. In 2001, Dandri et al. first showed that immune-deficient uPA transgenic mice could be engrafted with human hepatocytes and used as a model for hepatitis B [168]. Subsequently, this model was further developed to permit extensive repopulation, reaching levels as high as 90% human cells [169, 170]. Fah knockout mice can also be repopulated extensively with human cells from a variety of sources when they are crossed on to severely immune deficient backgrounds (Figure 38.8) [171]. This model has the advantage that the hepatic injury is not constitutive but can be titrated by NTBC. Both systems are likely to be important in testing the ability of stem cell derived “hepatocytes” for use in cell therapy.

FUTURE HORIZONS Many questions regarding the basic biology of stem cells in the liver remain unresolved, including the all-important question of whether hepatic stem cells exist at all in the adult liver. Most data suggest that it is possible to generate new hepatocytes from cells other than hepatocytes themselves and that there is an intrahepatic source of these precursors. Their precise nature and location, however, remains unclear, as do the molecular mechanisms that lead to their activation. Much of this uncertainly is related to the lack of markers that would permit the dissection of the complex cellular heterogeneity of the liver. However, cell surface markers, and also genetic lineage tracing tools, are rapidly becoming available. Thus, biological studies of antigenically defined and genetically marked cells will be feasible and are likely to solve some of these dilemmas. In addition, pluripotent stem cells now afford the opportunity for “developmental biology in a dish,”

(b)

Figure 38.8 Repopulation of the Fah−/− /Rag2−/− /IL2rg−/− mouse liver by primary human hepatocytes at 6 weeks after cell transplantation, shown at (a) low and (b) high magnification

38: LIVER REPOPULATION BY CELL TRANSPLANTATION AND THE ROLE OF STEM CELLS

making attractive models to study molecular mechanisms of hepatic lineage development. Although substantial progress has been made during the past 10–15 years concerning the possibility of liver repopulation by transplanted cells, much still needs to be learned. Factors governing engraftment of transplanted cells into the liver and their homing to the correct niche, factors regulating proliferation and differentiation of transplanted cells into specific phenotypes required for organ function, and specific host conditions under which effective liver replacement can be achieved, all need to be determined. The best starting point for therapeutic liver repopulation will probably be a genetic disorder with ongoing liver injury, which will hopefully induce or augment proliferation of transplanted cells in the host liver. A good example of such a condition is Wilson’s disease, in which transplanted cells might also have a modest selective advantage [172], since they will not store high levels of copper, which is toxic to the hepatocyte and causes liver injury. Another example is α1 -antitrypsin deficiency, in which a mutated form of α1 -antitrypsin is not secreted from the cell and, like urokinase in uPA transgenic mice, causes liver injury, but not nearly as severe as in uPA transgenic mice. To date, very few patients have been transplanted with fetal liver cells. Studies in rats have shown that these cells have the capacity to proliferate in the host, replace hepatic mass with functional hepatocytes, and maintain differentiated hepatic function, long-term [91]. This requires a liver proliferative stimulus or liver injury at the time the cells are transplanted, but no selective advantage other than their ability to replace host hepatocytes by cell competition. Use of stem cells from pediatric or adult cadaveric liver or from other sources, such as bone marrow, cord blood, or ES cells, is a possibility, and also cultured human fetal cells or adult hepatocytes modified to favor engraftment and proliferation in the host, but this will require substantial additional research. Cell lines have been established, including ES cells, fetal liver cells, and “oval (progenitor) cells” that also exhibit stem cell properties and differentiate into hepatocytes and/or bile ducts in vitro and in vivo. However, these cell lines have shown only limited repopulation of the normal liver at the current state-of-the-art. In order to advance the field of liver cell therapy further, it will be necessary to find conditions under which cells and cell lines derived from ES, fetal liver, or adult liver can be expanded in culture and successfully repopulate the liver under conditions that will be clinically acceptable. Although we are not yet there, restoration of liver function by therapeutic cell transplantation holds great promise for the future.

ACKNOWLEDGMENTS The authors would like to thank Anna Caponigro and Emily Bobe for assistance in preparing the manuscript

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and Dr Irmin Sternlieb, a former colleague at Albert Einstein College of Medicine, who provided the electron micrograph illustrating a Canal of Hering to D.A.S.

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161. Schmelzer, E., Wauthier, E. and Reid, L. (2006) The phenotypes of pluripotent human hepatic progenitors. Stem Cells, 24, 1852–58. 162. Schmelzer, E., Zhang, L., Bruce, A., Wauthier, E., Ludlow, J., Yao, H.L., Moss, N., Melhem, A., McClelland, R., Turner, W., Kulik, M., Sherwood, S., Tallheden, T., Cheng, N., Furth, M.E. and Reid, L.M. (2007) Human hepatic stem cells from fetal and postnatal donors. J Exp Med , 204, 1973–87. 163. Fox, I.J., Chowdhury, J.R., Kaufman, S.S., Goertzen, T.C., Chowdhury, N.R., Warkentin, P.I., Dorko, K., Sauter, B.V. and Strom, S.C. (1998) Treatment of the Crigler-Najjar syndrome type I with hepatocyte transplantation. N Engl J Med , 338, 1422–26. 164. Strom, S.C., Chowdhury, J.R. and Fox, I.J. (1999) Hepatocyte transplantation for the treatment of human disease. Semin Liver Dis, 19, 39–48. 165. Mito, M. and Kusano, M. (1993) Hepatocyte transplantation in man. Cell Transplant , 2, 65–74. 166. Fisher, R.A. and Strom, S.C. (2006) Human hepatocyte transplantation: worldwide results. Transplantation, 82, 441–49. 167. Bilir, B.M., Guinette, D., Karrer, F., Kumpe, D.A., Krysl, J., Stephens, J., McGavran, L., Ostrowska, A. and Durham, J. (2000) Hepatocyte transplantation in acute liver failure. Liver Transpl , 6, 32–40. 168. Dandri, M., Burda, M.R., T¨or¨ok, E., Pollok, J.M., Iwanska, A., Sommer, G., Rogiers, X., Rogler, C.E., Gupta, S., Will, H., Greten, H. and Petersen, J. (2001) Repopulation of mouse liver with human hepatocytes and in vivo infection with hepatitis B virus. Hepatology, 33, 981–88. 169. Tateno, C., Yoshizane, Y., Saito, N., Kataoka, M., Utoh, R., Yamasaki, C., Tachibana, A., Soeno, Y., Asahina, K., Hino, H., Asahara, T., Yokoi, T., Furukawa, T. and Yoshizato, K. (2004) Near completely humanized liver in mice shows human-type metabolic responses to drugs. Am J Pathol , 165, 901–12. 170. Meuleman, P., Libbrecht, L., De Vos, R., de Hemptinne, B., Gevaert, K., Vandekerckhove, J., Roskams, T. and Leroux-Roels, G. (2005) Morphological and biochemical characterization of a human liver in a uPA-SCID mouse chimera. Hepatology, 41, 847–56. 171. Azuma, H., Paulk, N., Ranade, A., Dorrell, C., Al-Dhalimy, M., Ellis, E., Strom, S., Kay, M.A., Finegold, M. and Grompe, M. (2007) Robust expansion of human hepatocytes in Fah−/− /Rag2−/− /Il2rg−/− mice. Nat Biotechnol , 25, 903–10. 172. Yoshida, Y., Tokusashi, Y., Lee, G.H. and Ogawa, K. (1996) Intrahepatic transplantation of normal hepatocytes prevents Wilson’s disease in Long–Evans cinnamon rats. Gastroenterology, 111, 1654–60.

PART FOUR : RELATION TO OTHER ORGANS

39

Hepatic Encephalopathy Roger F. Butterworth and Javier Vaquero Neuroscience Research Unit, Hˆopital Saint-Luc, University of Montreal, Montreal, Canada

HEPATIC ENCEPHALOPATHY Hepatic encephalopathy (HE) refers to a wide spectrum of psychiatric and neurological alterations that occur in patients with liver disease and/or portal–systemic shunting. Even in its mildest form, HE is a manifestation of severe liver failure and has somber prognostic implications both in acute and in chronic liver disease (CLD). In CLD, HE seriously interferes with quality of life and remains a major clinical problem. In acute liver failure (ALF), progression of HE is associated with increased risk of developing brain edema, high intracranial pressure (ICP), and brain herniation, a major cause of death. Exposure of the brain to blood-borne toxins that are not adequately cleared by the liver is the common pathogenetic basis for all forms of HE.

Type A HE or HE associated with acute liver failure

A CONSENSUS CLASSIFICATION OF HE The syndrome of HE comprises a wide variety of symptoms, signs, and underlying liver abnormalities. Such complexity together with the lack of a consensus classification has traditionally hampered both clinical and experimental research on HE. In order to unify the nomenclature and facilitate the performance of clinical studies and therapeutic trials, a new classification of HE was proposed in 2002 by a Working Party commissioned by the XI World Congresses of Gastroenterology (see Table 39.1) [1]. Notwithstanding the advance, the need The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

for further refinement of this classification system has been acknowledged [2]. HE is currently defined as the spectrum of neuropsychiatric abnormalities seen in patients with liver dysfunction after exclusion of other known brain diseases. A multiaxial classification of HE was adopted to reflect both the type of underlying liver disease and the characteristics of the neurological manifestations, while still recognizing a common pathogenetic basis. Three major types of HE were distinguished: type A or HE associated with ALF, type B or HE associated with portal–systemic bypass without intrinsic hepatocellular disease, and type C or HE associated with cirrhosis and portal hypertension/or portal–systemic shunts. Minimal HE was recognized as part of the spectrum of HE, being classified as a subtype of type C HE.

In a patient with an acute deterioration of liver function and no previous liver disease, appearance of the mildest sign of HE defines the syndrome of ALF [3]. HE in these patients may rapidly progress from mild cognitive alterations to a state of coma, with worsening of HE being associated with worse prognosis and increased risk of developing brain edema and intracranial hypertension. The risk of these complications is higher when there is a short interval (less than 7 days) between the onset of liver dysfunction (ictericia, prolongation of coagulation times) and the appearance of the first signs of HE, probably reflecting

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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Table 39.1 Proposed classification and nomenclature of HE Underlying disease Category and Subcategory, based on duration and characteristics

Type A

Type B

Type C

Acute liver failure N/Aa

Portal–systemic bypass without intrinsic hepatocellular disease Minimal Episodic Spontaneous Precipitated Recurrent (more than two episodes/year) Persistent Mild Severe Treatment dependent

Cirrhosis and portal hypertension or portal–systemic shunts Minimal Episodic Spontaneous Precipitated Recurrent (more than two episodes/year) Persistent Mild Severe Treatment dependent

a N/A, not applicable.

Adapted from [1], with permission.

an insufficient time for developing brain compensatory mechanisms. In ALF, brain dysfunction is mainly due to the impairment of liver function, with portal hypertension and extrahepatic portal–systemic shunting being absent or minor factors. Common alterations of ALF, such as the presence of an inflammatory response, infection, or the development of hydro-electrolytic alterations in plasma, may largely influence the appearance and severity of HE [4]. Agitation is particularly common as a manifestation of HE in ALF, frequently requiring the sedation and mechanical ventilation of the patient.

Type B HE or HE associated with portal–systemic bypass without intrinsic hepatocellular disease The presence of extensive portal–systemic shunting in the absence of intrinsic hepatocellular disease is uncommon, but it can occur in conditions such as non-cirrhotic portal vein thrombosis or in congenital vascular malformations. These patients may present manifestations of HE indistinguishable of those observed in patients with CLD [5, 6]. The extent of portal–systemic shunting and remaining liver perfusion is a major factor determining the development of HE. Due to the absence of liver disease, these patients may be erroneously diagnosed of other neuropsychiatric entities such as dementia or depression. Determination of ammonia in arterial blood may orientate the diagnosis. Imaging techniques such as echography, abdominal computed tomography (CT) scans, magnetic resonance imaging (MRI), and vascular radiology are important to detect the presence of portal–systemic shunts or hepatic vascular abnormalities. In this type of HE, the clinical manifestations, the neurophysiological examinations, and the findings from other techniques [hyperintensity of globus pallidus in T1 MRI, increased glutamine +

glutamate/creatine (Glx/Cr) ratio in nuclear magnetic resonance (NMR) spectroscopy] are similar to those observed in the HE of CLD, which emphasizes the pathogenetic relevance of portal–systemic shunting in other forms of HE. Portal–systemic shunts may be amenable to correction by surgery or interventional radiology.

Type C HE or HE associated with cirrhosis and portal hypertension/or portal–systemic shunts Underlying CLD is the most common setting in patients with HE. This type of HE is further subdivided into minimal, episodic, and persistent HE, a subdivision that may also apply to type B HE [2].

Minimal HE Minimal HE (previously called subclinical HE) refers to individuals with an otherwise normal neurological examination who present subtle abnormalities that are detected only by using specific neuropsychometric and/or neurophysiological tools [1]. Neuropsychometric tests are considered the gold standard for the diagnosis of this entity, which may represent an initial form of HE. There is no consensus, however, on which specific tests should be used to make the diagnosis. Minimal HE in patients with compensated cirrhosis is associated with an increased risk of developing overt HE and with a shorter survival, particularly in those with an altered response to an oral glutamine challenge [7]. Minimal HE is also associated with worse indices of quality of life [8] and may impair fitness to drive a car [9].

Episodic HE Episodic HE is the most classical presentation of HE in patients with CLD, and refers to intermittent periods

39: HEPATIC ENCEPHALOPATHY

of overt alterations of brain function (from impaired attention, memory or temporo-spatial orientation to stupor and coma) that usually resolve within hours or days. These episodes are frequently triggered by one or more precipitating factors (precipitated HE), but in some cases no precipitating factor can be identified (spontaneous HE). Recurrent HE is diagnosed when two or more episodes of HE occur within one year.

Persistent HE Persistent HE distinguishes a subset of patients who present overt neuropsychiatric alterations that may fluctuate without ever manifesting an asymptomatic stage. Persistent HE can be further subdivided into mild and severe grades.

NEUROPATHOLOGY OF HEPATIC ENCEPHALOPATHY The astrocyte manifests the most conspicuous neuropathological alterations in liver failure, but neuropathological changes may also involve other neural cells (neurons, microglia, endothelial cells). The extent and nature of these changes is a function of the type of liver injury (acute versus chronic), the extent of portal–systemic shunting and the number of episodes of HE that the patient has manifested. Examination of material from patients with end-stage ALF reveals signs of severe cytotoxic brain edema and the cell primarily involved is the astrocyte [10]. Figure 39.1a depicts an electron micrograph of material from a patient with ALF due to acetaminophen overdose showing massive swelling of the astrocyte end foot, a

(a)

601

feature also demonstrated in animal models of ALF [11]. Consequent brain edema may ultimately result in the development of high ICP and brain herniation. In contrast to the situation in ALF, the cardinal neuropathological feature of end-stage CLD is an astrocytic alteration known as “Alzheimer Type II astrocytosis.” The Alzheimer Type II phenotype is characterized by swelling of both the cell and its nucleus, a prominent nucleolus and margination of the chromatin pattern (Figure 39.1b). Although present in all brain regions, the severity of Alzheimer Type II changes shows regional selectivity, particularly affecting cerebral cortex, basal ganglia, and cerebellum [12]. Although mild compared with ALF, astrocyte swelling has been demonstrated by electron microscopy in the portacaval-shunted rat, an animal model of the HE of CLD [13]. Along the same lines, a number of studies using novel MRI techniques have reported a decrease in magnetization transfer ratios and an increase in water apparent diffusion coefficients in basal ganglia and in several white matter regions (particularly the corticospinal tract) of the brain of patients with HE and CLD [14–16], suggesting the presence of low-grade brain edema in this condition. Such abnormalities were correlated with alterations in neuropsychometric [14, 16, 17] and neurophysiological [17] performance, and were reversible after liver transplantation. Mild astrocytic swelling may be sufficient to influence osmosensitive signaling pathways and to interfere with the expression of cell-surface receptor and transporter proteins leading to disturbances of cell–cell signaling [18]. Together, these findings suggest a causal relationship between low-grade brain (astrocytic) edema and the development of HE in CLD. Recent studies suggest that abnormalities affecting brain cells other than the astrocyte may also be present in some types of HE. Although it was previously

(b)

Figure 39.1 Characteristic ultrastructural changes in the brain in acute and in chronic liver failure. (a) Electron micrograph from a patient with acute liver failure due to acetaminophen overdose showing cytotoxic brain edema with marked swelling of perivascular astrocyte (A), and dilatation of endoplasmic reticulum (arrows) and mitochondria (M). (b) Electron micrograph from a 51-year-old cirrhotic patient who died in hepatic coma presenting Alzheimer type II astrocytes with a large, pale nucleus and margination of chromatin (Alz) compared with normal astrocytes with normal chromatin pattern (N). Reproduced from [10] with permission

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THE LIVER: PATHOGENESIS OF HEPATIC ENCEPHALOPATHY

suggested that neuronal cell loss was not a feature of HE, a recent review of the literature reveals at least three distinct neuropathological entities that may occur in end-stage CLD including non-Wilsonian hepatocerebral degeneration, cerebellar degeneration, and the so-called post-shunt myelopathy affecting the spinal cord [19]. Although uncommon, these neurodegenerative conditions are most often encountered in patients with long-standing HE or following surgical portacaval shunting. CLD may also contribute to the diencephalic lesions characteristic of Wernicke’s encephalopathy in alcoholic patients [20]. Similarly, activation of microglial cells has been shown in experimental animal models of ALF [21], but to date this feature has not been studied in patient material. Finally, discrete neuropathological changes to endothelial cells of brain capillaries consisting of swelling and increased vacuolization have also been noted in patients with ALF, suggesting alterations in permeability of the blood–brain barrier (BBB) [10]. Recent reports in animal models of ALF also describe altered expression of some key BBB proteins suggesting functional changes to the cellular components barrier [22, 23], in addition to altered permeability to diverse small molecules [24]. Circulating factors derived from the necrotic liver, such as pro-inflammatory cytokines or matrix metalloproteinases, could be involved in these alterations [24, 25]. In apparent contradiction with the notion of edema being of a cytotoxic origin, a recent MRI study suggested the presence of reversible interstitial brain edema in patients with CLD and minimal or overt HE [26]. Even though these studies support the presence of altered BBB permeability, there remains little convincing evidence for a physical breakdown of the BBB in HE associated with either acute or chronic liver failure [10].

PATHOGENESIS OF HEPATIC ENCEPHALOPATHY Consequences of liver failure include hypoglycemia, hypothermia, hyponatremia, hyperammonemia, increased blood manganese concentrations, and inflammation, all of which have the capacity to compromise central nervous system function. Alterations of blood glucose, electrolytes, and infection are generally considered as “precipitating factors,” and once recognized they should be effectively corrected. Circulating toxins that are more difficult to remove rapidly include ammonia, manganese, and proinflammatory cytokines resulting from inflammatory processes caused by dying liver cells.

Ammonia Hyperammonemia without concomitant liver failure, such as that observed in patients with congenital urea cycle

disorders or in experimental models of pure hyperammonemia, resembles most of the cellular and biochemical alterations present in the brain of patients with HE. Arterial ammonia concentrations are increased two- to threefold in cirrhotic patients with mild to moderate HE [27] and are increased up to 10-fold in patients with ALF [28]. In the latter, increased arterial ammonia concentrations present an independent risk factor for encephalopathy and intracranial hypertension [29]. Although proposed as the major toxin in the pathogenesis of HE for over a century, it is only recently that the precise mechanisms responsible for ammonia’s neurotoxic actions and, by extension, its role in the pathogenesis of HE have been elucidated. Hyperammonemia in liver failure is the consequence of alterations of inter-organ trafficking of ammonia and of its metabolite, glutamine (see Figure 39.2). Under normal conditions, glutamine metabolism via glutaminase in intestinal epithelial cells accounts for 50% of the ammonia produced by the portal-drained viscera, the remaining 50% being produced by the colon, most of which is derived from the breakdown of urea [30]. Hepatic ammonia removal is compartmentalized, involving two distinct hepatocyte cell types. Urea is synthesized from ammonia via the urea cycle in periportal hepatocytes whereas perivenous hepatocytes convert ammonia into glutamine via glutamine synthetase (Figure 39.2a). Loss of these classes of hepatocytes in liver failure together with portal–systemic shunting of venous blood results in decreased ammonia removal by the liver and increased ammonia delivery to the systemic circulation (Figure 39.2b) [31]. In addition, patients with CLD may also present an increased intestinal production of ammonia due to enhanced glutaminase activity in intestinal epithelium [32]. In contrast to the liver, both muscle and brain are devoid of a urea cycle and rely almost entirely upon glutamine synthesis for effective ammonia removal. In liver failure, muscle becomes the principal organ with the requisite metabolic machinery to remove blood-borne ammonia and, to this end, adapts due to a post-translational increase in glutamine synthetase [33]. Brain ammonia accumulates in liver failure but, unlike muscle, brain does not adapt by glutamine synthetase induction [33, 34]. This lack of induction in brain may be the consequence of nitrosative stress and subsequent nitration of tyrosine residues of the protein [35]. NMR spectroscopic studies confirm a lack of increase of glutamine synthesis at late stages of HE [36]. Despite the lack of induction of glutamine synthetase, brain glutamine has consistently been reported to be increased in liver failure [37, 38]. Increased brain glutamine in liver failure may partly result from decreased brain output or from ammonia-induced inhibition of glutaminase. In patients with CLD and minimal HE, positron emission tomography (PET) reveals increased brain ammonia concentrations and increased brain ammonia utilization [27]. Increased brain ammonia in CLD has been ascribed to multiple mechanisms including increased arterial ammonia concentrations, increased cerebral blood

39: HEPATIC ENCEPHALOPATHY

603

BRAIN

BRAIN

LIVER

urea

Periportal hepatocytes

Periportal hepatocytes

Perivenous hepatocytes

Perivenous hepatocytes NH3

NH3

GUT

GUT

MUSCLE

MUSCLE

GLUTAMINE

LIVER

urea

KIDNEY

Normal

GLUTAMINE

KIDNEY

Liver Failure

Figure 39.2 Schematic representation of inter-organ trafficking of ammonia and glutamine under normal physiological conditions and in liver failure. Gut-derived ammonia is normally removed as urea (periportal hepatocytes) or glutamine (perivenous hepatocytes). In liver failure, ammonia removal by both types of hepatocytes is decreased. Brain ammonia uptake increases but there is limited capacity for further increases in brain glutamine synthesis. In contrast, skeletal muscle becomes the principal route for ammonia removal as a consequence of a post-translational increase in the enzyme glutamine synthetase

flow (CBF), and increased permeability of the BBB to ammonia [39, 40]. At the physiological pH of plasma, most ammonia (∼99%) circulates as the NH4 + ion and only ∼1% as its gaseous form (NH3 ). Even though only the latter form can readily cross hydrophobic cellular membranes, the contribution of channels at the BBB specifically transporting the ionic form of ammonia has recently been postulated [41]. Differences in blood and brain ammonia concentrations resulting from pH gradients, from regional variations of ammonia production/removal and from alterations in the BBB permeability to ammonia partly explain the absence of a clear correlation between circulating (particularly venous) ammonia concentrations and the severity of HE in liver failure. Ammonia has multiple effects on brain function that include direct effects of the ammonium ion (NH4 + ) on excitatory and inhibitory neurotransmission, effects on brain energy metabolism and mitochondrial function, selective effects on neurotransmitter systems, and effects mediated via proinflammatory mechanisms.

Brain organic osmolytes and cell volume regulation The brain has a remarkable ability to avidly metabolize circulating ammonia during hyperammonemic conditions, as the concentration of ammonia is a limiting factor in the astrocytic glutamine synthetase reaction under physiological conditions. The concentration of brain glutamine readily increases after the creation of a portacaval anastomosis in the rat [42], and three- to fourfold elevations of brain glutamine levels have been repeatedly demonstrated in acute and in chronic liver failure both in humans and in experimental models [12, 36, 38]. Because glutamine is an organic osmolyte, its rapid increase restricted to a

single cerebral compartment (the astrocyte) has been suggested to generate an osmotic gradient responsible for the development of astrocyte swelling and brain edema. Impairment of cell volume control in astrocytes is thought to underlie the pathogenesis of all types of HE, but the characteristics of this alteration depend on the extent and acuteness of the insult. In patients with CLD, studies using in vivo NMR spectroscopy have consistently reported a prominent disturbance of brain organic osmolytes consisting of an increase in the glutamine + glutamate signal and a decrease in the myo-inositol peak [18, 43]. In addition to myo-inositol, experimental models of hyperammonemia and/or liver failure demonstrate a decrease in other organic osmolytes such as taurine that may also be released by astrocytes [44], which is interpreted as a compensatory response to the increase in glutamine directed to preserve normal cell volume. These abnormalities are present even in cirrhotic patients who present no alterations in neuropsychometric tests, suggesting an underlying impairment in the capacity of the brain to face osmotic challenges in patients with cirrhosis [38]. In accordance with this notion, administration of an ammonia-generating solution induced greater deteriorations of neuropsychometric performance in patients with cirrhosis who had lower levels of brain myo-inositol [43]. Importantly, inhibiting the synthesis of glutamine prevents brain swelling and most of the cerebral metabolic alterations induced by hyperammonemia in rats [45, 46]. Altogether, these studies indicate the presence of a severe disturbance of organic osmolytes and astrocytic cell volume regulation in the brain of patients with CLD, which is relevant for explaining the increase in brain water demonstrated in patients with minimal and overt HE by recent MRI studies. Overt brain edema and intracranial hypertension, uncommon in CLD, are frequent in patients with ALF.

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THE LIVER: PATHOGENESIS OF HEPATIC ENCEPHALOPATHY

Extrusion of potassium is a first-line mechanism of regulatory volume decrease by which astrocytes respond to swelling, and inhibition of glutamine synthesis in rats infused with ammonia has been shown to attenuate the increase in potassium in brain extracellular space [47]. Reduction of organic osmolytes is a medium/long term mechanism of regulatory volume decrease that requires enough time to be complete [44]. In ALF, a previously healthy brain faces a rapid increase in brain glutamine levels due to a sudden exposure to high circulating levels of ammonia. Compensation of such increases of brain glutamine would require decreases in myo-inositol or other organic osmolytes to an extent that is not observed in experimental models of ALF or acute hyperammonemia [44, 48, 49]. Release of glutamine would be a potential alternative and, indeed, patients with ALF have been reported to present an elevation of extracellular brain glutamine and an increased cerebral glutamine efflux, which were higher in those developing high ICP and brain herniation [50, 51]. The levels of brain glutamine, however, do not correlate with the development of coma and overt brain edema in experimental ALF, as they are similarly elevated at earlier time points when these complications are not present [49]. Furthermore, maneuvers that prevent the development of brain edema and high ICP, such as hypothermia or indomethacin, are not accompanied by attenuation of the increase in brain glutamine [52, 53]. In both acute and chronic liver failure, therefore, the elevation of brain glutamine appears to be a necessary step for the complete development of the metabolic and cell volume alterations characteristic of HE, but its osmotic effects are not a sufficient explanation. Additional osmotic effects derived from the elevation of other organic compounds such as alanine and lactate, which present a better correlation with the neurological deterioration and the development of brain edema and high ICP in human and experimental ALF, may be required [36, 53, 54]. Alternatively, consequences of glutamine accumulation independent of its osmotic effects or the requirement for additional factors not related to cell osmolarity are other (non-mutually exclusive) explanations [55].

Brain glucose and energy metabolism Results of biochemical, physiological, and spectroscopic investigations demonstrate that there is no convincing evidence to support the notion that HE is the consequence of primary energy failure, as early reductions in brain levels of high-energy phosphates are absent [56, 57]. Important alterations of brain glucose uptake and of pathways involved in brain glucose and energy metabolism, however, have been consistently reported in HE both in acute and in chronic liver failure. For example, PET studies

using [18 F]fluorodeoxyglucose in patients with mild HE reveal decreased glucose uptake by anterior cingulate cortex [58], a brain structure implicated in the control of the anterior attention system responsible for the monitoring of tasks such as responses to visual stimuli. Decreased glucose utilization was correlated with impaired psychometric performance in these patients. In another study, decreased glucose and oxygen uptake by anterior cingulate cortex was accompanied by increased glucose uptake by thalamic structures, indicating a redistribution of brain glucose utilization in HE [59]. These differences in regional brain glucose and oxygen uptake at mild to moderate stages of HE are generally considered to be a consequence (rather than the cause) of HE, indicating altered brain fuel (glucose) requirements. Not in contradiction with the previous concept, a substantial body of evidence indicates that some alterations of brain glucose and energy metabolism are pathogenetically relevant in HE. Studies performed in animal models and in vitro systems suggest that brain glucose metabolism switches from the predominantly oxidative pathway to anaerobic glycolysis. Ammonia, in pathophysiologically relevant concentrations, is a potent inhibitor of the tricarboxylic acid (TCA) cycle enzyme α-ketoglutarate dehydrogenase (Figure 39.3) [60]. The consequent decrease in pyruvate oxidation together with ammonia-induced stimulation of enzymes of the glycolytic pathway [61] leads to shuttling of pyruvate carbon into alanine and lactate [62]. In addition, the inhibition of α-ketoglutarate dehydrogenase by ammonia may deplete the TCA cycle intermediates resulting in an impairment of the malate–aspartate shuttle, a major system for transferring reducing equivalents from cytosol to mitochondria [63]. Malfunction of this shuttle would also enhance the glycolytic flux to lactate. In accordance with these situations, precipitation of severe encephalopathy and deep coma following the administration of ammonium salts in portacaval-shunted rats leads to brain lactate accumulation and, at terminal stages, to loss of ATP [56]. The finding that cerebrospinal fluid lactate concentrations correlate well with the severity of neurological impairment in cirrhotic patients suggests that increased glycolytic flux is also mechanistically relevant in humans with mild- to moderate-type C HE [64]. The disturbance of brain glucose metabolism is particularly prominent in ALF, where it presents a close relation with clinical manifestations. The global consumption of glucose and oxygen is decreased in the brain of these patients as a consequence of the reduced electrical activity of the comatose state [65, 66]. In contrast, the concentration of lactate in brain extracellular space is generally increased, a finding that was initially taken as an indication of cerebral hypoxia [66]. Subsequent studies ruled out this possibility, showing that CBF and oxygen delivery generally exceeded the brain demands and that the increased lactate coexisted with unchanged lactate-to-pyruvate ratios in many patients [54, 65]. Increased cerebrospinal

39: HEPATIC ENCEPHALOPATHY

605

Glucose Glycolytic Pathway Alanine

LDH Pyruvate Lactate

NH3

Acetyl CoA

Aspartate

Citrate

Oxaloacetate TCA Cycle

NH3

α-Ketoglutarate

Succinate

NH3 Glutamate

GlutamIne

αKGDH

Figure 39.3 Schematic representation of the potential effects of ammonia on glucose metabolism. Ammonia stimulates the amidation of glutamate by glutamine synthetase and possibly of α-ketoglutarate (solid arrows) leading to increased synthesis of glutamine. Ammonia has also been shown to inhibit the enzyme α-ketoglutarate dehydrogenase (dashed line) and to enhance the activity of glycolytic enzymes (solid arrow), both of which can result in increased glycolytic flux. In the presence of an inhibition of the TCA cycle, pyruvate is redirected towards the formation of alanine and lactate, both of which are known to be increased in experimental acute liver failure and hyperammonemia

fluid and brain lactate have consistently been reported also in experimental ALF [36, 67]. The direct effects of ammonia on glucose-metabolizing enzymes and the malfunction of the malate–aspartate shuttle are potential explanations, although neither of them has been conclusively demonstrated. In patients with ALF, increased concentrations of lactate in brain microdialysate were associated with an increased risk of developing surges of ICP, which were frequently preceded by lactate peaks [54]. Similarly, spectroscopic studies in experimental ALF have demonstrated a progressive increase of de novo synthesis of lactate correlated in time and magnitude with the severity of encephalopathy and the presence of brain edema [36, 68]. Independently of its cause, therefore, both clinical and experimental studies suggest a potential role of the lactate elevation in the development of brain edema in ALF.

Oxidative/Nitrosative stress and mitochondrial dysfunction Recent studies have provided indirect evidence of the presence of oxidative and nitrosative stress in the brain in diverse animal models of both acute and chronic liver failure. Increased brain expression of heme oxygenase 1, a gene highly induced in the presence of oxidative stress, and of the neuronal isoform of nitric oxide synthase (NOS-1), and also decreased expression of the antioxidant enzyme Cu,Zn-superoxide dismutase, have been reported in portacaval-shunted rats [69, 70]. Three interdependent

mechanisms were proposed to explain the increase in NOS-1, namely (i) ammonia-induced increase in uptake of l-arginine, the obligate substrate for NOS, (ii) stimulation of NOS-1 gene expression, and (iii) stimulation of the NMDA (N -methyl-d-aspartate) receptor–NO–cGMP signal transduction pathway (see [71] for a review). In the presence of superoxide, the production of NO can result in the formation of peroxynitrite, a highly reactive compound associated with the nitration of proteins. The presence of such a situation is supported by the observation of an increased cerebral efflux of NO and the formation of nitrotyrosine in the brain of portacaval-shunted rats receiving an acute infusion of ammonia [72, 73]. Nitrotyrosine predominantly localized to the astrocytic end-feet and endothelial cells of brain capillaries, suggesting a relation with the increase in CBF that generally precedes the appearance of intracranial hypertension in this model. However, administration of either a specific inhibitor of NOS-1 or an unspecific inhibitor of all NOS isoforms did not prevent the development of cerebral hyperemia [74]. Studies performed in cultured cells provide further insights into the mechanisms and consequences of oxidative and nitrosative stress in HE. Exposure of astrocytes to ammonia leads to increased production of oxygen free radicals, and lipid peroxidation, and reproduces the induction of heme oxygenase 1 and NOS, the formation of nitrotyrosine, and the astrocyte swelling observed in animal models [75]. Importantly, antioxidants and NOS inhibitors are effective in the prevention of ammonia-induced swelling in cultured astrocytes [75]. Antioxidants also prevent the induction of the mitochondrial permeability

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transition (MPT) by ammonia, a Ca2+ -dependent process that leads to the opening of the permeability transition pore in the inner mitochondrial membrane. Opening of this pore allows the gradient-driven movement of protons, ions, and other solutes, ultimately resulting in the collapse of the inner membrane potential, mitochondrial dysfunction, and further free radical generation. Impairment of mitochondrial energy metabolism and activation of mitogen-activated protein kinases and other signaling pathways are mechanisms by which oxidative stress can induce astrocyte swelling. Cyclosporine A, an inhibitor of the MPT, attenuates many of the alterations induced by ammonia in astrocytes, including the generation of free radicals and astrocyte swelling. The mechanism by which ammonia induces the MPT is still unclear, but one potential cause is the increase in intracellular calcium that occurs when astrocytes are exposed to ammonia [76]. These findings strongly suggest that mitochondrial dysfunction and oxidative/nitrosative stress play major roles in the pathogenesis of HE.

Neurotransmitter systems Liver failure leads to the alteration of several neurotransmitter systems in the brain. These alterations may contribute to the pathogenesis of astrocyte swelling and CBF disturbances that are characteristic of HE, and ultimately determine the neuropsychiatric symptoms of the syndrome.

Glutamate Glutamate is the major excitatory neurotransmitter of the central nervous system in mammals, but its brain extracellular concentration must be maintained low in order to ensure efficient synaptic neurotransmission and to avoid excitotoxicity. Astrocytes avidly uptake the glutamate released in the synaptic cleft during neurotransmission and shuttle it back as glutamine to neurons (Figure 39.4). Because astrocytes are altered in HE and because the metabolisms of glutamate and ammonia are linked, it is not surprising to find alterations of brain glutamate in HE. Alterations of glutamate reported in HE may differ in acute and in chronic liver failure, but they affect virtually all elements of the glutamatergic system, including its concentration, its metabolism, and the expression and function of its transporters and receptors (see [77] for a review). Most alterations are also observed in experimental models of pure hyperammonemia. Whereas the total concentration of brain glutamate appears to be decreased, its concentration in brain extracellular space has generally been found to be increased in animal models of ALF and hyperammonemia and also in humans with ALF and severe HE [54]. In these situations, the increase in extracellular glutamate correlates with the

ASTROCYTE PRESYNAPTIC NERVE TERMINAL Glutamine Glutaminase

Glutamine NH3

Glutamate

NH3

GS

Glutamate Glutamate POSTSYNAPTIC CELL

EAAT-2

NMDAR Ca2+ NOS

NO

Figure 39.4 Simplified schematic representation of neuron–astrocyte trafficking of ammonia, glutamine, and glutamate; the so-called “glutamate–glutamine cycle.” Glutamate released from the presynaptic nerve terminal stimulates the glutamate receptor on the postsynaptic cell membrane [only the NMDA subclass of glutamate receptor (NMDAR) is depicted here]. Excess glutamate is removed by transport into the adjoining astrocyte via one of the high-affinity glutamate transporters (EEAT-2 shown here). This step represents the major glutamate-deactivating mechanism. Glutamate transported into the astrocyte is then converted to glutamine by the enzyme glutamine synthetase (GS) with removal of NH3 . Glutamine is then available for release from the astrocyte, and uptake into the presynaptic nerve terminal as the immediate precursor of the neurotransmitter pool of the glutamate. Conversion of glutamine to glutamate occurs by the action of glutaminase and the cycle is complete. One turn of the cycle leads to the transfer of ammonia from the astrocyte to the neuronal compartment. Ammonia may act on several components of the glutamate–glutamine cycle. Ammonia inhibits glutaminase (product inhibition), down-regulates the glutamate transporter EAAT-2 and activates the glutamate (NMDA) receptor. This latter action leads to entry of Ca2+ into the postsynaptic cell, activation of nitric oxide synthase (NOS), and production of nitric oxide (NO) which diffuses out of the cell where it may lead to protein tyrosine nitration. One protein that is nitrated is GS resulting in loss of enzyme activity and impaired removal of brain ammonia

progression of encephalopathy and the development of brain edema. Potential effects of increased extracellular glutamate include the induction of astrocyte swelling, which has been reported in vitro, and the alteration of CBF. Increased extracellular glutamate may be secondary to a reduced expression of glutamate transporters such as the excitatory amino acid transporter (EAAT)-2 (the main astrocytic glutamate transporter in rat forebrain), EAAT-1, and EAAT-3, which have been described in

39: HEPATIC ENCEPHALOPATHY

amounts and that many had their origins in pharmaceutical BZs previously prescribed to the patients. Studies on the NS site have been more fruitful. Although coupling between the NS site and the GABA recognition site is unchanged in human HE brain [86], NSs with potent agonist properties at the NS site (such as allopregnanolone) are present in increased amounts in this material [87], the levels being sufficiently high to cause “increased GABAergic tone,” and are potent neuroinhibitory agents.

ALF and portacaval-shunted rats [78, 79]. Ammonia exposure is known to stimulate astrocytic glutamate release in cultured astrocytes, offering another potential explanation [76]. Ammonia also interferes with glutamatergic synaptic transmission by affecting glutamate receptors, but these effects depend on the receptor subtype (NMDA vs AMPA/kainate), on the brain region under study and on the acuteness of the insult [77]. For example, administration of an NMDA receptor antagonist, memantine, was associated with an improvement in encephalopathy and brain edema in rats with ALF, suggesting a pathogenic role of increased NMDA receptor activity [80]. In contrast, the activity of the NMDA receptor–NO–cGMP signaling pathway has been shown to be decreased in the portacaval-shunted rat [81], and stimulation of the NMDA receptor–NO–cGMP pathway by administration of sildenafil restores the impaired learning ability in this model.

Monoamines Patients with end-stage CLD frequently manifest extrapyramidal signs and symptoms and up to 22% of these patients exhibit moderate to severe parkinsonism associated with focal dystonia [88]. Typical features of this cirrhosis-related parkinsonism include rapid progression, symmetrical akinetic rigid syndrome, postural tremor, and gait impairment. Some patients respond to l-DOPA (l-3,4-dihydroxyphenylalanine), suggesting a role for the brain dopaminergic system in its pathophysiology. Studies of autopsied brain tissue from cirrhotic patients who died in hepatic coma reveal increased dopamine degradation [89], increased expression of the dopamine-metabolizing enzyme monoamine oxidase (MAO-A isoform) [90] and reduced densities of postsynaptic D2 receptor densities measured either biochemically [91] or by PET [92]. The most likely explanation for these changes involves manganese deposition (see later in this chapter), leading to a defect in the nigrostriated dopaminergic pathway.

GABA γ-Aminobutyric acid (GABA) is the major inhibitory neurotransmitter of mammalian brain. The last three decades have witnessed intense activity focused on the GABA system in relation to HE. Studies in the 1980s suggested that GABA synthesized in the periphery was transferred into brain via a defective BBB, resulting in increased “GABAergic tone” [82]. The postsynaptic GABA-A receptor is neuronal in localization and, being a multimeric protein complex, has several associated modulatory sites (Figure 39.5). These modulatory sites form part of the so-called “GABA-A receptor complex.” Such sites include the benzodiazepine modulatory site (BZ site) and the neurosteroid modulatory site (NS site), shown in a simplified schematic form in Figure 39.5. Clinical trials in the 1990s showed that flumazenil (Ro 15-1788), a BZ site antagonist, was effective in treating HE [83, 84] and it was suggested that this action of flumazenil resulted from the drug’s action in blocking the effects of an “endogenous BZ” present in the brains of HE patients [85]. BZ-like compounds were identified, but interest in this area was tempered by the finding that these compounds were present in only trace Benzodiazepines

BZ site

607

Cerebral blood flow (CBF) CBF influences the cerebral complications of liver failure by determining the delivery of ammonia and other putative toxins to the brain (Figure 39.6). The development of brain edema and intracranial hypertension is also influenced by the cerebral blood volume and hydrostatic pressure present in brain capillaries at a given time point

GABA

Neurosteroids

GABA recognition site

NS site Cl

Postsynaptic neuronal membrane

−

Figure 39.5 Simplified schematic representation of the GABA-A receptor complex, a multimeric protein complex localized on the postsynaptic neuronal membrane that gates chloride ion (Cl− ). Associated with the GABA recognition site are a series of modulatory sites; only two are depicted here for the sake of clarity: The benzodiazepine modulatory site (BZ site) and the neurosteroid modulatory site (NS site). Agonists of these modulatory sites include the pharmaceutical benzodiazepine diazepam and the naturally occurring neurosteroid allopregnanolone. Activation of the modulatory sites by these agonists leads to amplification of the GABA signal, increased chloride influx, and neuroinhibition

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THE LIVER: PATHOGENESIS OF HEPATIC ENCEPHALOPATHY

↑ CBF

Brain parenchyma Osmotic • ↑ brain glutamine → disturbance

↑ delivery of NH3, cytokines, etc…

↑ hydrostatic pressure

↑ blood volume

• ↑ oxidative/nitrosative stress

↑ ASTROCYTE SWELLING

↑ H2O influx

↑ BRAIN EDEMA ↑ intracranial volume compromise

↑ ICP Figure 39.6 Schematic representation of the potential mechanisms by which increased cerebral blood flow (CBF) worsens brain edema and intracranial hypertension in acute liver failure [93]. First, cerebral hyperemia increases the delivery of ammonia and other toxins to the brain, resulting in the enhancement of (i) the osmotic disturbance secondary to glutamine accumulation, (ii) oxidative/nitrosative stress, and (iii) the alterations of glucose metabolism, all of which are considered primary processes responsible for astrocyte swelling. Second, cerebral hyperemia with reduction of arteriolar tone increases hydrostatic pressure in brain capillaries, which would modify the hydrostatic pressure gradient, favoring the influx of water from blood to brain. Finally, cerebral hyperemia may increase cerebral blood volume, which in the setting of a rigid skull with a brain with poor compliance may lead to direct increases in intracranial pressure (ICP)

[93]. Alterations of CBF have been described both in acute and in chronic liver failure. Compared with healthy controls, cirrhotic patients with or without HE present regional alterations of CBF consisting of a decrease in flow to cortical regions and an increase to thalamus, basal ganglia, and cerebellum, which combined generally result in a reduction in global CBF [94, 95]. Because this pattern follows the regional changes of brain glucose utilization, CBF autoregulation (the capacity of CBF to match brain activity independently of changes in systemic arterial pressure) appears to be preserved in CLD, a suggestion supported by studies where the effect of artificially increasing and/or decreasing mean arterial pressure was assessed [96]. Cerebral vascular resistance in patients with ascites, however, correlates directly with renal vascular resistance and inversely with mean arterial pressure, suggesting that CBF in cirrhosis may be affected by systemic factors [97]. Accordingly, the capacity to maintain CBF despite a decrease in mean arterial pressure, that is, the lower limit of autoregulation, has been shown to be impaired in some patients, particularly in those with worse liver function and advanced systemic circulatory disturbances [96, 98]. Alterations of CBF are particularly noticeable in ALF. In contrast to CLD, a loss of CBF autoregulation has been found in many patients with ALF. This is relevant for the clinical management, as increases or decreases in mean

arterial pressure could easily lead to episodes of cerebral hyperemia or hypoperfusion, respectively. The absolute values of CBF in patients with ALF vary from below to above normal [66], probably reflecting the complex interplay between cerebral energy demands, CBF autoregulation, ammonia neurotoxicity, systemic arterial pressure, and ICP at different stages of the disease. Based on a retrospective analysis of sequential ICP and CBF measurements in 26 patients with ALF, Aggarwal et al. recently proposed a five-phase sequence consisting of an initial phase of low values of CBF (in accordance with low neuronal activity) and normal ICP (phase I), followed by a progressive increase in CBF (phase II) and a subsequent increase in ICP (phase III) [99]. In the final stages, intracranial hypertension would lead to a decrease in CBF (phase IV) with impaired cerebral perfusion and brain death (phase V). An increase in CBF (phase II) is a common finding in patients with severe HE that is associated with more severe brain edema, higher ICP, and mortality [93]. Manipulation of CBF strongly influences the development of brain edema and high ICP in both clinical and experimental ALF, emphasizing the pathogenetic relevance of this alteration (Figure 39.6). This five-phase classification, therefore, integrates well with previously published data, has a mechanistic correlation, and is helpful for the prognostic evaluation of these patients.

39: HEPATIC ENCEPHALOPATHY

609

Altered gene expression in the brain

Infection and inflammation

Experimental liver failure is associated with alterations in the expression of multiple genes in the brain. Although not exclusively, these alterations affect genes predominantly expressed in astrocytes and/or in endothelial cells, and they include proteins involved in the transport of water molecules (aquaporin-4), glucose (GLUT-1), and neurotransmitters (EAAT-2, EAAT-1, GLYT-1), implicated in cell structure [GFAP (glial fibrillary acidic protein)], and in the synthesis of NSs (peripheral type BZ receptor), to name a few [100, 101]. Even though the alterations of these genes may influence the molecular processes underlying the cerebral complications of liver disease, further investigations are required to determine which alterations are primarily implicated in the pathogenetic processes and which are downstream responses to other pathological changes such as astrocyte swelling.

Because of its unique cellular composition and anatomical localization, the liver can be considered a major immunological organ. Systemic inflammation is common in most forms of liver failure, either as a consequence of the injury to the liver and/or as part of the processes leading to that injury. Infection, a major cause of inflammation, is a well-recognized precipitant of HE in CLD and is also present in most patients with ALF. The relevance of infection and inflammation for the development of the cerebral complications of both acute and chronic liver failure has attracted increasing attention [108]. In patients with CLD and documented infection, induction of hyperammonemia was associated with a worsening of neuropsychometric performance that was not observed when hyperammonemia was induced after the infection and its associated inflammatory response was resolved, suggesting that inflammatory mediators modulate the effect of ammonia in the brain [109]. In rats with common bile duct ligation, the administration of lipopolysaccharide induced neurological deterioration that was associated with an increase in brain water content and protein tyrosine nitration, suggesting an exacerbation of brain edema and nitrosative stress as potential operative mechanisms [110]. In patients with ALF, the presence of infection or a systemic inflammatory response is associated with the worsening of HE [108]. Modulation of CBF by circulating inflammatory mediators is a potential mechanism by which inflammation can affect the development of the cerebral complications of ALF. In this regard, changes in the circulating levels of pro-inflammatory cytokines appear to parallel the changes in CBF and ICP in some situations [111]. In anesthetized rats, administration of ammonia exacerbated the development of cerebral hyperemia and intracranial hypertension induced by lipopolysaccharide, further supporting a synergistic effect of ammonia and inflammatory mediators on the cerebral circulation [112]. In addition to systemic inflammation, recent findings in patients developing uncontrolled intracranial hypertension [113] and also in experimental models [114] suggest the intriguing possibility that inflammation can also be generated within the brain in ALF. Activation of inflammatory cascades inside the brain can be the consequence of systemic inflammation [108], but its exact trigger(s) and its contribution to the cerebral pathological process in ALF remain unknown.

Systemic factors in addition to ammonia There is overwhelming evidence for considering ammonia the major pathogenetic factor of the cerebral complications of liver failure. Notwithstanding, evidence for other factors having a synergistic and/or influencing effect for the development of HE continues to accumulate.

Manganese Manganese is normally removed from the circulation by the hepatobiliary system. Consequently, blood and brain manganese concentrations are elevated several-fold in CLD [102]. Brain manganese concentrations are selectively increased in basal ganglia structures in both experimental and human CLD, and it has been proposed that the pallidal signal hyperintensities observed on T1 -weighted MRI are due to manganese deposition [103, 104]. The intensity of pallidal hyperintensities is significantly correlated with extrapyramidal symptoms, particularly rigidity [105]. Although considered mainly a feature of CLD, pallidal hyperintensities and parkinsonian symptoms have also been noted in ALF [106]. Manganese deposition in basal ganglia structures of patients with CLD offers a plausible explanation for alterations in the dopaminergic neurotransmitter system (dopaminergic neurons are particularly vulnerable to manganese) and for the Parkinsonian symptoms in cirrhotic patients [88, 107]. Moreover, motor slowing due to the toxic effects of manganese on the dopamine systems could explain, in part, the poor performance in psychometric tests such as number connection test (NCT)-A and -B.

Hyponatremia Chronic hyponatremia is relatively common in patients with advanced liver disease, in whom it has been associated with an osmoregulatory decrease of brain myo-inositol [115]. As discussed in a previous section,

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the induction of such compensatory changes in brain organic osmolytes may impair the capacity of brain cells to respond to acute osmotic insults. In accordance with this notion, chronic hyponatremia has been shown to exacerbate the development of brain swelling in hyperammonemic rats [116]. Acute hyponatremia can also directly favor the transport of water into the brain by decreasing plasma osmolarity and altering the equilibrium of Starling forces across the BBB [93]. Therefore, any factors disturbing the normal osmotic homeostasis of the blood may, through these mechanisms, contribute to the cerebral complications of both acute and chronic liver failure.

THERAPEUTIC APPROACHES TO HE Improving the treatment of the cerebral complications of liver disease remains a major challenge. Efficient targeting of the major pathogenetic factor, hyperammonemia, has proved to be difficult due to the essential roles of nitrogen metabolism in multiple organs. In CLD, treatment of the precipitating factor(s) and maneuvers that decrease ammonia production remain the mainstay of therapy. Precipitating factors of HE may act by increasing ammonia production (constipation, gastrointestinal hemorrhage) or altering its metabolism (renal failure), and also by affecting pathogenetic factors different than ammonia (hyponatremia, infections and inflammation, use of sedatives, etc.). The main specific targets in the therapy of HE are as follows: 1. Ammonia production: Several agents have been shown to decrease ammonia production by the gut, although the lack of conclusive clinical trials demonstrating clinical benefits against placebo has recently been acknowledged [117]. The most commonly used agents are the non-absorbable disaccharides lactulose and lactitol, which lower plasma ammonia by increasing ammonia fecal nitrogen excretion. In addition to their cathartic effect, their mechanism of action involves their catabolism to lactic and acetic acid, respectively. The resulting acidification of colonic pH favors the formation of the less absorbable NH4 + from NH3 . Several poorly absorbable antibiotics (neomycin, metronidazole, and vancomycin) lower plasma ammonia by decreasing ammonia production by the colonic bacterial flora, but they are of limited use due to side-effects and to the risk of developing multi-resistant bacteria and Clostridium difficile diarrhea. Neomycin, whose mechanism of action includes a reduction in intestinal glutaminase activity [118], has been the most commonly used [119]. The new antibiotic rifaximine may be a safer alternative that has been associated with reductions of blood ammonia and encephalopathy improvements similar

to lactitol in patients with type C HE [120]. Finally, recent reports support the use of lactic acid-producing probiotics and symbiotic preparations (combinations of probiotics and fermentable fiber), particularly for treating minimal HE [121]. In addition to acidification of colonic pH and promotion of saccharolytic over ammonia-producing bacterial flora, probiotics may have other beneficial effects such as the reduction of intestinal permeability and endotoxemia [122]. Restriction of protein in the diet for reducing ammonia load is no longer indicated as a means of preventing episodes of HE [123]. Current guidelines recommend a normal protein intake (1–2 g kg−1 per day) for patients with cirrhosis in order to maintain an adequate nitrogen balance and avoid malnutrition [124]. This is important given the role of muscle mass as a major ammonia-detoxifying organ in hyperammonemia. Oral supplementation with branched-chain amino acids may benefit selected patients, particularly those with severe protein intolerance [125]. In patients with type C HE, the administration of acarbose, an oral hypoglycemic agent, has recently been associated with improvement of HE and reductions in plasma ammonia, presumably achieved by the promotion of intestinal saccharolytic bacterial flora [126]. 2. Ammonia removal: In addition to decreasing ammonia production, a second possibility to lower plasma ammonia consists in increasing ammonia removal. Phenyl acetate (given orally as its precursor phenyl butyrate) and sodium benzoate are agents used to decrease plasma ammonia in children with congenital urea cycle disorders [127]. Phenyl acetate binds to glutamine and benzoate binds to glycine to form, respectively, phenylacetylglutamine and hippuric acid, which can then be excreted in the urine. The ammonia-lowering action theoretically resides in the reduction of glutamine and glycine, which results both in a decrease of ammoniagenic substrates and in the utilization of ammonia for synthesis and replenishment of the respective amino acid pools. Both agents have occasionally been used in patients with HE, but their safety and the possibility of competition with a non-disrupted urea cycle for the elimination of ammonia represent important concerns [128, 129]. The combination of L-ornithine with L-aspartate has been shown to reduce blood ammonia and to improve neuropsychometric performance in patients with type C HE [130], and its use is currently being tested in patients with type A HE. Its mechanism of action involves the stimulation of the synthesis of urea and glutamine in the liver and of glutamine in the muscle, the latter being most relevant in ALF [131]. L-Carnitine is an endogenous quaternary ammonium compound with antioxidant properties that is involved in the transport of fatty acids into the mitochondria. Administration of l-carnitine reduced blood ammonia in clinical and experimental HE, although its operative

39: HEPATIC ENCEPHALOPATHY

mechanism is still undefined [132]. Even though complex and more invasive, extracorporeal blood depuration methods represent another means for removing ammonia and circulating toxins from the blood. Albumin dialysis using the molecular adsorbent recirculating system (MARS), for example, has been associated with the reduction of blood ammonia and the improvement of HE in patients with advanced cirrhosis and severe HE [133]. Importantly, experimental studies in pigs with ALF suggest that depuration of toxins other than ammonia are important for the cerebro-protective effects of MARS [134]. 3. Neurotransmission: Medications acting on the GABA-receptor complex [flumazenil [83], BZ partial inverse agonists [135]), dopamine receptor agonists (l-DOPA, bromocriptine) [136], histamine H1 receptor antagonists (hydroxyzine) [137], glutamate receptor antagonists (memantine) [80], phosphodiesterase inhibitors (sildenafil) [81], and others] have been used in clinical and/or experimental studies to ameliorate selective alterations found on the different neurotransmitter systems in liver failure, usually resulting in the improvement of their related symptoms of HE. In addition to improvement of the symptomatology, some of these interventions may act by mechanisms shown to be implicated in the pathogenesis of the cerebral complications of liver failure. For example, administration of memantine (an NMDA receptor antagonist) attenuated brain edema in rats with ALF [80], and BZ receptor antagonists prevented astrocyte swelling in cultured astrocytes exposed to ammonia [138]. 4. Inflammation: Agents that modulate inflammatory responses may have a greater impact on the pathogenesis of HE and brain edema than the antagonism of specific cytokines, given the redundant roles of the latter. Administration of dexamethasone, however, did not prevent the development of intracranial hypertension in patients with ALF [139]. The utilization of non-steroidal anti-inflammatory drugs in patients with liver disease is limited due to the renal and hematological toxicity. Notwithstanding, administration of indomethacin effectively reduced brain edema and high ICP in patients with ALF and in hyperammonemic rats, effects likely related to the reduction of cerebral hyperemia [52, 140]. Ibuprofen also attenuated the impairment in learning ability in portacaval-shunted rats, a model of chronic HE [141]. Anti-inflammatory strategies remain a potential therapeutic approach that is still largely unexplored. 5. Oxidative/nitrosative stress: Attenuation of oxidative/nitrosative stress using diverse antioxidants and inhibitors of NOS decreases cell swelling in cultured astrocytes exposed to ammonia [75]. It has been proposed that the beneficial effects of N -acetylcysteine observed in ALF could involve central antioxidant mechanisms [142]. Evidence for a role of oxidative/

611

nitrosative stress in human HE awaits additional studies. Treatment of HE in ALF is mostly directed towards diminishing brain edema and intracranial hypertension. Whereas some of the previous strategies may be valuable, in practice this goal is usually achieved using approaches similar to those used for intracranial hypertension of other etiologies. The main targets of such approaches are as follows: 1. Osmotic equilibrium: Administration of intravenous (i.v.) boluses of mannitol or the infusion of hypertonic saline attenuate intracranial hypertension in patients with ALF [139, 143]. The increase in blood osmolarity induced by these agents is thought to influence the osmotic gradient across an intact BBB, favoring the retention of water in the intravascular compartment. Their exact mechanism of action, however, is not known [144]. 2. Cerebral blood flow: Maneuvers that reduce CBF and/or prevent cerebral hyperemia significantly attenuate the development of brain edema and high ICP in ALF. Hyperventilation reduces CBF by inducing hypocapnic cerebral vasoconstriction, and it has been associated with the restoration of CBF autoregulation in patients with ALF [145]. Despite the theoretical risk of developing excessive vasoconstriction leading to cerebral hypoxia, institution of hyperventilation for short-term periods has been shown to be safe in these patients. The rapid reduction of intracranial hypertension by indomethacin in clinical and experimental studies of ALF was also intimately associated with the reduction of CBF [52, 140]. A decrease in CBF is also a major mechanism explaining the cerebroprotective effects of sedative agents such as propofol or barbiturates [111]. 3. Multiple mechanisms: Evidence from clinical [146] and experimental [147, 148] studies point to the induction of mild hypothermia as an efficient therapeutic maneuver to control brain edema and high ICP in ALF. Reductions of the blood–brain transfer of ammonia, of arterial ammonia levels, and of CBF are major mechanisms explaining the remarkable ability of hypothermia to attenuate these complications, but direct effects of hypothermia on other factors such as inflammation or oxidative/nitrosative stress may also be important (see [149, 150] for reviews). Assessment of the therapeutic potential of mild hypothermia in well-designed clinical trials of ALF is essential to confirm its benefits, to identify which patients can benefit from it, and to provide the guidelines for its clinical use [151].

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140. Tofteng, F. and Larsen, F.S. (2004) The effect of indomethacin on intracranial pressure, cerebral perfusion and extracellular lactate and glutamate concentrations in patients with fulminant hepatic failure. J Cereb Blood Flow Metab, 24, 798–804. 141. Cauli, O., Rodrigo, R., Piedrafita, B., Boix, J. and Felipo, V. (2007) Inflammation and hepatic encephalopathy: ibuprofen restores learning ability in rats with portacaval shunts. Hepatology, 46, 514–19. 142. B´emeur, C., Vaquero, J., Desjardins, P. and Butterworth, R.F. (2007) Protein microarray study reveals toxin-selective cytokine profiles in experimental acute liver failure: beneficial effects of mild hypothermia and N -acetylcysteine. Hepatology, 46, 611A. 143. Murphy, N., Auzinger, G., Bernel, W. and Wendon, J. (2004) The effect of hypertonic sodium chloride on intracranial pressure in patients with acute liver failure. Hepatology, 39, 464–70. 144. Saraswat, V.A., Saksena, S., Nath, K., Mandal, P., Singh, J., Thomas, M.A., Rathore, R.S. and Gupta, R.K. (2008) Evaluation of mannitol effect in patients with acute hepatic failure and acute-on-chronic liver failure using conventional MRI, diffusion tensor imaging and in vivo proton MR spectroscopy. World J Gastroenterol , 14, 4168–78.

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40

The Kidney in Liver Disease Moshe Levi Division of Renal Diseases and Hypertension, University of Colorado, Denver, CO, USA

IMPAIRED RENAL FUNCTION IN LIVER DISEASE Impaired renal function is common in liver diseases, either as part of multi-organ involvement in acute illness or secondary to advanced liver disease.

HEPATORENAL SYNDROME

ACUTE KIDNEY INJURY IN LIVER DISEASE Acute kidney injury (AKI, previously known as ARF or ATN) occurs in approximately 20% of hospitalized patients with cirrhosis [1]. In contrast, the incidence of acute kidney injury in acute liver failure varies from 40 to 80%, especially in the setting of infections such as viral hemorrhagic fever, leptospirosis, bacterial peritonitis, and toxin-induced injuries such as acetaminophen poisoning, administration of nephrotoxic antibiotics (such as aminoglycoside antibiotics), or high doses of non-steroidal anti inflammatory drugs [2]. The mechanisms leading to acute renal injury are multiple and additive and include (1) changes in the systemic arterial circulation, including systemic arterial vasodilatation and sometimes in certain alcoholic patients the concomitant presence of cardiomyopathy, (2) portal hypertension, (3) activation of renal vasoconstrictive hormones, including the renin–angiotensin system, the sympathetic nervous system (SNS), arginine vasopressin, endothelin (ET), thromboxane A2, and leukotrienes, which all additively impair renal blood flow, and (4) suppression of

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

renal vasodilatory factors including prostacyclins which impair the kidney’s ability to counterbalance the effects of vasoconstrictors on the renal circulation (Figure 40.1). These hemodynamic factors can accelerate kidney injury in the presence of intrinsic kidney disease caused by various disorders outlined in Table 40.1.

Hepatorenal syndrome (HRS) is a unique form of functional renal failure that often complicates advanced liver disease, hepatic failure, or portal hypertension. The International Ascites Club has recently updated the diagnostic criteria for HRS [3, 4], which are outlined in Table 40.2. In cirrhotic patients with ascites, most of the cases of acute renal failure are mediated by (1) pre-renal failure and (2) acute kidney injury (acute tubular necrosis). HRS accounts for approximately 20% of renal failure. HRS is characterized by (1) an extreme expression of the profound circulatory dysfunction in cirrhosis, with marked splanchnic arterial and systemic vasodilatation, insufficient cardiac output, severe reduction of effective blood volume, homeostatic activation of vasoactive systems, and intense renal vasoconstriction, resulting in a critical decrease in renal blood flow and glomerular filtration rate, (2) absence of pathological changes in renal tissue (vasculitis, glomerulonephritis, or tubulointerstitial fibrosis), and (3) preserved renal tubular function including FENa 1.5 mg dl−1 3. No improvement in serum creatinine (decrease to C polymorphism in the bile salt export pump. World J Gastroenterol , 14 (1), 38–45. M¨ullenbach, R., Bennett, A., Tetlow, N., Patel, N., Hamilton, G., Cheng, F., Chambers, J., Howard, R., Taylor-Robinson, S.D. and Williamson, C. (2005) ATP8B1 mutations in British cases with intrahepatic cholestasis of pregnancy. Gut , 54 (6), 829–34. Lammert, F., Marschall, H.U., Glantz, A. and Matern, S. (2000) Intrahepatic cholestasis of pregnancy: molecular pathogenesis, diagnosis and management. J Hepatol , 33 (6), 1012–21. Milkiewicz, P., Gallagher, R., Chambers, J., Eggington, E., Weaver, J. and Elias, E. (2003) Obstetric cholestasis with elevated gamma glutamyl transpeptidase: incidence, presentation and treatment. J Gastroenterol Hepatol , 18 (11), 1283–86. Clayton, R.J., Iber, F.L., Ruebner, B.H. and McKusick, V.A. (1969) Byler disease. Fatal familial intrahepatic cholestasis in an Amish kindred. Am J Dis Child , 117 (1), 112–24. Bull, L.N., van Eijk, M.J., Pawlikowska, L., DeYoung, J.A., Juijn, J.A., Liao, M., Klomp, L.W., Lomri, N., Berger, R., Scharschmidt, B.F., Knisely, A.S., Houwen, R.H. and Freimer, N.B. (1998) A gene encoding a P-type ATPase mutated in two forms of hereditary cholestasis. Nat Genet , 18 (3), 219–24. Eppens, E.F., van Mil, S.W., de Vree, J.M., Mok, K.S., Juijn, J.A., Oude Elferink, R.P., Berger, R., Houwen, R.H. and Klomp, L.W. (2001) FIC1, the protein affected in two forms of hereditary cholestasis, is localized in the cholangiocyte and the canalicular membrane of the hepatocyte. J Hepatol , 35 (4), 436–43. Ujhazy, P., Ortiz, D., Misra, S., Li, S., Moseley, J., Jones, H. and Arias, I.M. (2001) Familial intrahepatic cholestasis 1: studies of localization and function. Hepatology, 34 (4Pt 1), 768–75. van Mil, S.W., Houwen, R.H. and Klomp, L.W. (2005) Genetics of familial intrahepatic cholestasis syndromes. J Med Genet , 42 (6), 449–63. Paulusma, C.C., Folmer, D.E., Ho-Mok, K.S., de Waart, D.R., Hilarius, P.M., Verhoeven, A.J. and Oude Elferink, R.P. (2008) ATP8B1 requires an accessory protein for endoplasmic reticulum exit and plasma membrane lipid flippase activity. Hepatology, 47 (1), 268–78.

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43

Adaptive Regulation of Hepatocyte Transporters in Cholestasis James L. Boyer Department of Internal Medicine and Liver Center, Yale University School of Medicine, New Haven, CT, USA

INTRODUCTION Bile secretion is a unique and vital function of the liver. This secretion delivers bile acids to the intestine for the solubilization and absorption of dietary lipids and serves as an excretory pathway for endogenous metabolic products including cholesterol, bilirubin, and porphyrins, in addition to many other foreign compounds and xenobiotics. Many different liver disorders can impair this secretion, leading to the accumulation of bile and the syndrome of cholestasis. In response to cholestatic liver injury there are a number of adaptive responses in hepatic enzymes and membrane transport proteins that synthesize and excrete bile acid, bilirubin, and other solutes in order to reduce the accumulation of these products and attenuate liver tissue injury. However, over time, these adaptive responses are not sufficient and chronic cholestatic disorders ultimately lead to the development of biliary cirrhosis and progressive liver failure. Hence there is considerable interest in devising therapeutic strategies that might augment these intrinsic adaptations. This chapter reviews current knowledge of the mechanism of these adaptive responses in order to provide a background for new strategies for therapeutic interventions.

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

The major membrane transport systems that result in the formation of bile are illustrated in Figure 43.1. Their gene designations, membrane locations, and primary functions are briefly summarized in Table 43.1. Hepatic enzymes that are involved in the regulation of bile salt synthesis and metabolism are summarized in Table 43.2. The molecular basis of bile formation is discussed in greater detail in Chapter 23. The current topic has also been the subject of several recent reviews [1–5].

Steps in bile formation The overall process of the formation of bile from the hepatocyte can be functionally divided into four different phases based on a modification of Phase I and II drug metabolizing enzymatic reactions. Phase 0 consists of hepatic uptake mechanisms. Phase I are hydroxylation enzyme reactions. Phase II represents conjugation enzyme reactions, for example, sulfation, glucuronidation, and amidation, and Phase III represents hepatic efflux mechanisms. Each of these steps is regulated by a series of transporters (Phase 0 and III) or enzyme reactions (Phase I and II).

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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THE LIVER: GENERAL OVERVIEW – ACQUIRED DEFECTS IN BILE TRANSPORT PROTEINS

BSEP Na+ BA−

NTCP

BA−, OA− OC+

OATPs

BA

FIC1

PS

MRP4 BA-S−

MRP2 GSH HCO3−

OA− GSH

OC+ PL

BCRP OA−S H+,

OCT1

OC+

MDR1

Na+

MDR3 OSTα-β

Chol OATs

OA−

ABCG5/8

?

MRP3 Bil-G−

OC+ MATE-1

Figure 43.1 Membrane transporters that determine the uptake and excretion of bile acids and other organic solutes in hepatocytes (see tables for full terminology)

This chapter focuses on the molecular adaptations in these hepatocyte membrane transporters and enzymatic reactions that serve to attenuate cholestatic liver injury and its systemic effects. Cellular events that impair signal transduction pathways, cytoskeleton structures, tight junction and gap junctional proteins, and the targeting of intracellular vesicles to the apical canalicular domain that maintains the secretory polarity of the hepatocyte all contribute to the cholestatic phenotype [1], but are not considered here. Much information has been obtained from animal models of cholestasis and from discoveries of the genetic basis of several hereditary and acquired human cholestatic disorders. Studies in human hepatocyte cell lines and cholestatic liver disorders, and also mouse knockout models, have increasingly become available and have contributed significantly to advances in this field.

GENERAL OVERVIEW – ACQUIRED DEFECTS IN BILE TRANSPORT PROTEINS The molecular characterization of transport systems that determine the formation of bile has made it possible to assess their response to cholestatic liver injury at the genetic level. Although the picture is still evolving, a common pattern of expression has begun to emerge that has resulted in the following paradigm: The basic determinants of bile formation adapt to cholestatic liver injury in a manner that tends to minimize hepatic injury by (i) diminishing the hepatic uptake of bile salts and other solutes from portal blood, (ii) reducing bile acid synthesis, (iii) augmenting bile acid detoxification mechanisms, and (iv) up-regulating mechanisms that facilitate the exit of bile salts and other toxic substances from hepatocytes [6]. Thus, transport proteins on the basolateral sinusoidal membrane which normally function to remove

bile salts and other cholephiles selectively from portal blood are usually down-regulated during cholestatic liver injury by both transcriptional and post-transcriptional mechanisms. In contrast, some canalicular transport proteins, particularly the multidrug resistance protein (MDR) homologs, are either not severely impaired or may actually be up-regulated; these include MDR1/Mdr1, BSEP/Bsep (bile salt export pump), and MDR3/Mdr2 [7–9]. These later findings suggest that the cholestatic hepatocyte attempts to maintain canalicular export function. Perhaps more importantly, transporters such as MRP3/Mrp3 [10–12], MRP4/Mrp4 [13–17], and OSTα-OSTβ (organic solute transporter) [18] that are expressed at the basolateral sinusoidal membrane at low levels in normal liver are substantially up-regulated in cholestatic hepatocytes. Here they facilitate the removal of hydrophobic bile salts and other products from the liver via these alternative pathways back into the systemic circulation where they may be cleared in part by the kidney [19, 20]. Thus, cholestasis results in a partial reversal of the bile secretory process from the apical to the basolateral plasma membrane of the hepatocyte. Less is known concerning molecular responses in cholangiocytes during cholestasis. However, bile duct proliferation is characteristic of many cholestatic disorders, particularly those that result in both intrahepatic and extrahepatic bile duct obstruction. The apical sodium-dependent bile salt transporter (ASBT), originally identified in the ileum, is also located on the luminal membranes of cholangiocytes and the proximal tubules of the kidney where it functions to remove bile salts from bile and glomerular filtrate, respectively [21–23]. Because the cholestatic hepatocyte continues to excrete bile salts, albeit at a reduced rate, ASBT may function to remove bile salts from an obstructed cholestatic biliary tree. MRP3, MRP4, and OSTα/β are also located on the blood side of the cholangiocyte and are up-regulated in the cholestatic liver [10, 11, 18, 24]. Both MRP3/Mrp3 and MRP4/Mrp4 and also OSTα-OSTβ are capable of transporting bile salts and presumably function to return

43: ADAPTIVE REGULATION OF HEPATOCYTE TRANSPORTERS IN CHOLESTASIS

683

Table 43.1 Nomenclature, location and function of the major hepatocyte plasma membrane bile acid and organic solute transporters involved in bile secretion (Phase 0 and III) Name

Abbreviation (gene)

Sodium-taurocholate cotransporter

NTCP (SLC10A1)

Organic anion-transporting polypeptides

Location

Function

0

Basolateral membrane of hepatocytes

OATPs (SLCO1B1 and 1B3)

0

Basolateral membrane of hepatocytes

Organic solute transporter alpha/beta

OSTα/β

III

Basolateral membrane of hepatocytes, cholangiocytes, ileum and proximal tubule of kidney

Organic cation transporter-1

OCT-1 (SLC22A1)

0

Basolateral membrane of hepatocytes

Organic anion transporter 2

OAT-2 (SLC22A7)

0

Basolateral membrane of hepatocytes

Multidrug-resistance-1 P-glycoproteina

MDR1 (ABCB1)

III

Canalicular and cholangiocyte apical membrane

Multidrug-resistance-3 P-glycoprotein (phospholipid transporter)a Bile salt export pumpa

MDR3 (ABCB4)

III

Canalicular membrane

BSEP (ABCB11)

III

Canalicular membrane

MRP2 (ABCC2)

III

Canalicular membrane

MRP3 (ABCC3)

III

Basolateral membrane of hepatocytes and cholangiocytes

MRP4 (ABCC4)

III

Basolateral membrane of hepatocyte; apical membrane of proximal tubule of kidney

Primary carrier for conjugated bile salt uptake from portal blood Broad substrate carriers for sodium-independent uptake of bile salts, organic anions, and other amphipathic organic solutes from portal blood Heteromeric solute carrier for facilitated transport of bile acids across basolateral membrane of ileum. Expression induced in liver in cholestasis Facilitates sodium-independent hepatic uptake of small organic cations Facilitates sodium-independent hepatic uptake of drugs and prostaglandins ATP-dependent excretion of various organic cations, xenobiotics, and cytotoxins into bile; barrier function in cholangiocytes ATP-dependent translocation of phosphatidylcholine from inner to outer leaflet of membrane bilayer ATP-dependent bile salt transport into bile; stimulates bile salt-dependent bile flow Mediates ATP-dependent multispecific organic anion transport (e.g. bilirubin diglucuronide) into bile; contributes to bile salt-independent bile flow by GSH transport Expression induced in cholestasis. Transports bilirubin and bile acid glucuronide conjugates Expression induced in cholestasis – transports sulfated bile acid conjugates and cyclic nucleotides (continued overleaf )

Multidrug-resistanceassociated protein 2 (canalicular multispecific organic anion transporter)a Multidrug-resistanceassociated protein 3a Multidrug-resistanceassociated protein 4a

Phase

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Table 43.1 (continued) Name

Abbreviation (gene)

Phase

Location

Function ATP-dependent transport of organic anions and small peptides. Mutations of MRP6 gene result in pseudoxanthoma elasticum ATP-dependent multispecific drug transporter, particularly sulfate conjugates; protoporphyrins are endogenous substrate. Substrate overlap with MRP2 Heteromeric ATP dependent transporter for cholesterol and plant sterols (stilbestrol) Organic cation/H+ exchanger extrudes cationic xenobiotics

Multidrug-resistanceassociated protein-6a

MRP6 (ABCC6)

III

Basolateral membrane of hepatocyte

Breast cancer resistance proteina

BRCP (ABCG2)

III

Canalicular membrane, proximal tubule of kidney

Sterolin-1 and 2a

ABCG5/G8

III

Multidrug and toxin extrusion protein 1

MATE-1 (SLC47A1)

III

Canalicular membrane and apical membrane of intestine Canalicular membrane and brush border of kidney

a These transporters are members of the ATP-binding cassette superfamily.

bile salts to the systemic circulation during cholestasis [20, 24, 25]. MRP3/Mrp3 has a particularly high affinity for glucuronidated conjugates and may provide the means by which bilirubin glucuronide is excreted into the blood. MRP4/Mrp4 may provide the same role for sulfated conjugates [26, 27]. Sulfated bile salt conjugates in particular are formed in the cholestatic human liver and to a less extent in mouse liver [20, 28]. More recent studies suggest that OSTα-OSTβ, which mirrors the tissue expression of ABST, also plays a major role in bile acid efflux from the cholestatic hepatocyte in human liver [18]. Renal bile salt transporters also undergo adaptive regulation in animal models of cholestasis. Down-regulation of Asbt in the renal proximal tubule results in diminished absorption of bile acids from the glomerular filtrate, thereby facilitating an extrahepatic pathway for disposal of bile salts in the urine [29] Additionally, Mrp2 and Mrp4 (which are both expressed on the apical luminal membrane of the proximal tubule in the kidney) may facilitate the tubular excretion of bile acid and other divalent conjugates. Less is known about the response of intestinal bile acid transport proteins in cholestasis, although ABST is generally down-regulated on the luminal brush border of the ileum, reducing the return of bile acids to the liver in the enterohepatic circulation [30, 31], and MRP2/Mrp2 is diminished in both humans and rats with obstructive cholestasis [32, 33]. Adaptations also occur in hepatic enzymes that determine the synthesis and metabolism of bile acids. In general, these changes result in a smaller and less hydrophobic bile salt pool, although

significant species differences exist in composition and response. For example, in cholestatic mice, bile acid pools are enriched in the highly hydrophilic bile acid, muricholic acid, often requiring feeding of cholic acid (CA) in the diet to elicit significant tissue injury. Enzymes that regulate bile acid and bilirubin conjugation are also significantly up-regulated in cholestasis. The major enzymes and their function and adaptive responses are summarized in Table 43.2. Thus, a complex pattern of adaptive regulatory responses in transporter expression and metabolism is emerging in several different tissues, including the liver hepatocyte, biliary epithelial cells, kidney, and intestine, that can be interpreted as an attempt to mitigate tissue damage from the retention of bile salts and other toxic substrates. Much of this adaptive regulatory response is mediated by transcriptional events that are regulated by liver-enriched transcription factors [hepatocyte nuclear factors (HNFs)] and nuclear receptors (NRs). Members of the HNF family tend to regulate constitutive expression of genes whereas NRs induce adaptive responses in gene expression and are activated by specific ligands. In cholestasis, these ligands are bile acids, bilirubin, oxysterols, and drugs or xenobiotics. Chief among these are bile acids which regulate gene expression primarily via the farnesoid X receptor (FXR) [34–36]. When bile acids accumulate in the cholestatic liver, the expression of genes that are regulated by FXR are usually enhanced. Other NRs that play an important role in regulating gene expression (based largely on studies of nuclear receptor knockout animal models of cholestasis) are the pregnane X receptor (PXR),

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Table 43.2 Hepatic enzymes involved in the regulation of bile salt synthesis and bile salt and bilirubin metabolism (Phase I and II) and their adaptive responses to cholestasis Name

Abbreviation (gene)

Cytochrome P450 7A1

(CYP7A1)

Cytochrome P450 8B1

(CYP8B1)

Cytochrome P450 27A1

(CYP27A1)

Cytochrome P450 3A4

(CYP3A4)

Sulfotransferase

SULT2A1

Mediates Phase II conjugation with sulfate

Uridine glucuronyl transferase

(UGT1A1/B4/B7)

Mediates Phase II conjugation with glucuronic acid

Bile acid CoA synthetase Bile acid CoA:amino acid N -acyltransferase

BACS

Forms bile acid CoA-thioesters, substrates for BAAT Mediates taurine and glycine conjugation of bile acids (amidation)

BAAT/Bat

Adaptive response in cholestasisa

Function Cholesterol 7α-hydroxylase, the rate-limiting step in bile acid synthesis from cholesterol Sterol 12α-hydroxylase, the pathway to cholic acid synthesis. Controls ratio of cholic acid to chenodeoxycholic acid Sterol 27-hydroxylase, the mitochondrial “acidic” alternative pathway of bile acid side-chain oxidation. Favors chenodeoxycholic acid formation Mediates Phase I bile acid hydroxylation

Down-regulation limits the synthesis of bile acids More favored pathway resulting in increased % of bile acids as cholic acid

Unchanged

Unchanged or stimulated; may facilitate ability of bile acids to undergo Phase II conjugations Unchanged or stimulated; facilitates bile acid conjugation to substrates for MRP2 and MRP4 Unchanged or stimulated: mediates bile acid and bilirubin conjugation to substrates for MRP2 and MRP3 ?? ??; reduced in sepsis

a ??, The adaptive response in cholestasis is not known.

the constitutive androstane receptor (CAR), the liver X receptor (LXR) [37], the vitamin D receptor (VDR), and peroxisome proliferator-activated receptor alpha (PPARα) [38]. Knockout animals for FXR, PXR, CAR, and LXR are each more susceptible to cholestatic injury than their respective wild-type [37, 39–43]. Other ligand stimulated NRs involved in the adaptive response to cholestasis include the retinoic X receptor alpha (RXRα) and the glucocorticoid receptor (GR). RXRα plays a particularly important role in these responses since it is the obligate heterodimeric partner for the class II low-affinity NRs that include FXR, PXR, LXR, CAR, and PPARα. Cholestasis also can affect the expression of NRs. Studies of inflammatory and fibrotic models of cholestasis indicate that RXRα expression is reduced [44, 45] and may be translocated out of the nucleus into the cytoplasm, where it may be degraded, thus influencing the expression of genes dependent on its NR partners [45]. Reductions in mRNA levels of FXR, and its target gene SHP (short heterodimeric partner), have also been reported in patients with cholestasis from PFIC1 and 2 (progressive familiar intrahepatic cholestasis type 1 and type 2) and biliary atresia [46, 47]. Reductions in PXR and CAR have also been

reported in late stages of biliary atresia [47]. Short-term administration of the PPARα agonist, fenofibrate, has beneficial effects on hepatocellular damage and apoptosis in common bile duct ligated (CBDL) rats [48], suggesting that this nuclear receptor may also be involved in the cholestatic response. However, the benefits of fibrate therapy in cholestatic patients with primary biliary cirrhosis has yet to be established [49, 50]. Several other regulators of transcription for which no specific ligands are known, but that also influence the expression of bile transporters and enzymes, include the SHP-1, liver receptor homologue-1 (LRH-1), and the HNF-4α. HNF-4α is a primary determinant of HNF-1α expression, which determines the transcription of genes that encode for cytochrome P-450 7A (CYP7A), Ntcp, and Oatps (organic anion transporting polypeptides) as examples [51]. Table 43.3 lists these NRs and their activating ligands and key transcription factors, and also illustrates their major target genes and summarizes the anticipated regulatory responses in cholestasis. Further details of these interactions are provided below. However, because of differences between species, it is difficult to generalize. For more detailed information on adaptive

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Table 43.3 Nuclear receptors, important ligands, transcription factors and target genes regulating the adaptive response in cholestasis (? = from rodents only)a Nuclear receptors

Ligands

Major target genes

Anticipated function in cholestasis

FXR (farnesoid X receptor) – NR1H4

Bile acids, GW4064, 6α-ethyl-CDCA, fexaramines

PXR (pregnane X receptor) – NRII

Xenobiotics, LCA, ursodeoxycholic acid, rifampacin

BSEP, OATP1B3, MRP2, MDR3, OSTα/β, CYP3A4, UGT2B4 and B7, SULT2A1, BACS and BAAT, SHP, I-BABP, FGF-15/19, PXR CYP3A4, OATP1B1?, SULT2A1, UGT1A1 MRP2, MRP3, MDR1, GST

CAR (constitutive androstane receptor) – NR3

Xenobiotics, bilirubin, phenobarbital, Yin Chin, TCPOBOP

CYP2B, CYP3A4, OATP1B1, MRP2, MRP4, UGT1A1, SULT2A1, GSTs)

LXRα (liver X receptor) – NR1H3

Oxysterols (metabolites of cholesterol) All-trans-retinoic acid

CYP7A1, CYP8B, UGT1A3 ABC5/8?, SHP, OSTα/β?, LRH-1 NTCP?, MRP2,? ASBT, MRP3

Inhibition of bile acid synthesis and ileal bile acid uptake via SHP and FGF-15/19. Up-regulation of Phase I and II bile acid hydroxylation and conjugation, induction of bile acid canalicular and basolateral transporters Induction of Phase I and II bile acid and bilirubin conjugation reactions and bile acid and bilirubin alternative export pumps Induction of Phase I and II bile acid and bilirubin conjugation reactions and bile acid and bilirubin alternative export pumps Inhibits bile acid synthesis while stimulating Phase II and III steps

Vitamin D, lithocholic acid Corticosteroids

CYP3A4, SULTs? MRP3? NTCP, ASBT, MRP2, BSEP, AE2 Obligate heterodimeric partner for all class II NRs

RARα (retinoic acid receptor) – NR1B1

VDR (vitamin D receptor) – NRIII GR (glucocorticoid receptor) RXRα (retinoic X receptor) – NR2B1 Others SHP-1 (short heterodimeric partner) – NR0B2

9-cis-Retinoic acid

None; Up-regulated by FXR

LRH-1(FTF) – liver receptor homologue-1; fetal transcription factor – NR5A2 HNF-4α (hepatocyte nuclear factor-4) – NR2A1

? Blocked by SHP

PPARα (peroxisome proliferator-activated receptor-alpha) – NR1C1

Fatty acids, eicosanoids, fibrates, statins

None

? Induction of RXR partners via metabolism to 9-cis-retinoic acid; loss in cholestasis up-regulates MRP3 Induction of Phase I and II bile acid hydroxylation ? Induction of AE2 (together with ursodeoxycholate) Reduction of RXR in cholestasis has variable effects depending on the gene

Interacts with LRH-1 to suppress CYP7A1, CYP8B1, CYP27A1. Also inhibits NTCP?, and ASBT?, OATP1B1 CYP7A1, CYP8B1. MRP3, ASBT?

Suppresses bile acid synthesis, and hepatic and ileal bile acid uptake

HNF-1, CYP7A1, CYP8B1, CYP27A1 OATP1B1, NTCP? BACS and BAAT SULT2A1, UGT2B4, MDR2/3

Inhibition by SHP via FXR of bile acid synthesis and bile acid uptake

Inhibition of bile acid synthesis (CYP8B1) and ileal uptake (ABST?) via SHP; up-regulates MRP3

Induce Phase II bile acid and bilirubin conjugation reactions

a Transporter and enzyme gene abbreviations include: ABCG5 and G8 (sterolin 1 and 2); ASBT (apical sodium-dependent bile acid transporter, SLC10A2);

BAAT (bile acid CoA:amino acid N -acyltransferase); BACS (bile acid CoA-synthase); BSEP (bile salt export pump, ABCB11); CYP7A (cytochrome P-450 7A); CYP8B (cytochrome P-450 8B); CYP27A1(cytochrome P-450 27A1); FGF-15/19 (fibroblast growth factor 15 or 19); GSTs (glutathione S -transferases); I-BABP (ileal bile acid binding protein); MDR1 or 2/3 (multidrug resistance protein 1 or 2/3, ABCB1or 4); MRP2, 3 and 4 (multidrug resistance associated protein 2, 3 or 4, ABCC2,3 or 4); NTCP (sodium taurocholate cotransporting polypeptide, SLC10A1); OATP-1B1, 1B3 (organic anion transporting polypeptide C or 8, SLC01B1 or 1B3); OSTα/OSTβ (organic solute transport protein alpha and beta); SHP-1 (short heteromeric partner 1); SULTs (sulfa transferases); UGTs (uridine 5′ -glucuronosyl-transferases).

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responses in animal models of cholestasis and the role of NRs, the reader is referred to recent comprehensive reviews [4, 52].

Regulation of transporter genes in the cholestatic hepatocyte Ion transporters: (Na+ K+ -ATPase and Na+ /H+ exchange) Several major ion transport proteins are located on the basolateral membrane and regulate important homeostatic housekeeping functions in the hepatocyte, including maintenance of the electrical potential, cell volume, and intracellular pH. These transporters include Na+ /K+ -ATPase and the Na+ /H+ exchanger-isoform 1 (NHE-1). Although many cholestatic agents have been shown to inhibit the sodium pump in vitro, the molecular expression of the sodium pump is either not significantly affected or somewhat up-regulated, as discerned from studies in several different animal models of cholestasis [53–57]. This adaptive response could result from an attempt to counteract increased sodium entry and cell swelling that may result from the detergent properties of retained bile salts. Na+ /H+ exchange is also up-regulated at both transcriptional and post-transcriptional levels following common bile duct ligation in the rat, resulting in an increase in intracellular pH. This response may also contribute to sodium entry and cell swelling in the cholestatic liver [58].

Transporters on the basolateral membrane of the hepatocyte (Phase 0) Sodium taurocholate co-transporting polypeptide, NTCP/Ntcp (SLC10A1/Slc10a1) Sodium taurocholate cotransporting polypeptide (Ntcp) and its human homolog NTCP are the major determinant of the selective hepatic uptake of conjugated bile salts from the portal circulation and are substantially down-regulated in both human and animal models of cholestasis [12, 14, 53, 55–57, 59–61]. NTCP mRNA has been evaluated in patients with biliary atresia and is diminished [47]. However, NTCP mRNA increases towards normal values if biliary drainage is restored by a successful portoenterostomy (Kasai procedure) [62]. NTCP mRNA correlates inversely with the level of the total serum bilirubin in these patients. NTCP mRNA is also reduced in patients with inflammatory diseases (alcoholic hepatitis), although levels are inversely correlated with serum bile acid levels [59]. NTCP mRNA and protein are also down-regulated in human (and rat) liver

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slices after exposure for 24 hours to lipopolysaccharides (LPSs), and correlate inversely with the levels of interleukin (IL)-1β, suggesting mediation by cytokines [63]. Other clinical studies found reduction of NTCP protein in advance stages of primary biliary cirrhosis [12]. Transcription of Ntcp is impaired in the bile duct obstructed rat, and the protein disappears rapidly from the sinusoidal membrane throughout the hepatic lobule [53]. Levels of hepatic Ntcp mRNA also correlate inversely with the level of the total serum bile acids in the bile duct obstructed rat model [64], where retention of biliary constituents, but not the depletion of bile, regulates mRNA levels of Ntcp [64]. The transcriptional regulation of NTCP/Ntcp differs between species [65]. In human tissue, the mechanism is complex as bile acids induce SHP via FXR, which reduces HNF4α binding to bile acid response elements (BAREs) in the NTCP promoter which, in turn, inhibits its transactivating effects on HNF1α [51]. HNF1α expression is highly dependent on activation by HNF4α, which is a main regulator of NTCP expression [51, 66]. Bile acids also have SHP-independent effects on HNF4α binding. However, SHP has no direct effect on NTCP/Ntcp promoter activity, and bile acid-induced signaling pathways via c-Jun N-terminal kinase may be involved. More recent studies suggest that the human NTCP gene is also activated by the glucocorticoid receptor and PPARγ coactivator-1α, and can be suppressed by bile acids via a small heterodimer partner-dependent mechanism [67]. In contrast, the promoter region of Ntcp in the rat contains several regulatory response elements including binding regions for the transactivators, HNF1, and retinoid receptors (RARα/RXRα) [68, 69], and also the signal transducer and transactivator 5 (Stat 5) [70]. Bile acids down-regulate rat Ntcp via FXR-dependent Shp expression and subsequent inhibition of retinoid activation of RARα/RXRα [71]. Several additional upstream regions are involved in cytokine responses. Cholestasis produced by administration of endotoxin (LPS) results in loss of HNF1 and the RXRα : RARα heterodimer [69], which decreases Ntcp expression that parallels the reductions in bile flow [61]. Induction of Shp-1 precedes down-regulation of Ntcp in bile duct-ligated mice [72]. LPS also result in release of cytokines [tumor necrosis factor alpha (TNF-α) and IL-1β] that may contribute to the diminished Ntcp expression [57]. In mice, Ntcp repression by CBDL and CA or taurocholate feeding is mediated by FXR and does not depend on cytokines, whereas Ntcp repression by LPS is independent of FXR. Reduced levels of HNF-1α, RXR-α, and RAR-α in CBDL FXR–/–mice and reduced DNA binding in CA fed FXR–/–mice, despite unchanged Ntcp levels, indicate that these factors may have a minor role in regulation of mouse Ntcp during cholestasis [73]. Although impairment of Ntcp in rodents and NTCP in humans can explain the reduced Na+ -dependent uptake of conjugated bile salts, sodium-independent mechanisms

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for hepatic bile salt uptake persist due to continued expression of several other sinusoidal membrane organic anion transporters [74], such as Oatp2/Oatp1a4 and Oatp4/Oatp1b2 in mice [75] and possibly OATP1B3 in humans [76, 77].

Organic anion transporting polypeptides, OATPs/Oatps (SLCO/Slco) The OATP/Oatps are members of a large super family of transporters [solute carrier organic anion transporter family (SLCO)] with broad substrate specificity that are capable of translocating a wide range of organic anions, including unconjugated and conjugated bile salts, bulky organic cations, and even certain uncharged organic substrates [78–87]. These proteins appear to function as anion exchangers, exchanging the extracellular anion with either intracellular bicarbonate or glutathione [88, 89], although the exact driving forces are still not known [87]. OATPs in human liver consist of OATP1B1 (SLCO1B1, formally OATP-C), OATP1B3 (SLCO1B3, formally OATP8), OATP1A2 (SLCO1A2, formally OATP-A), and OATP2B1 (SLCO2B1, formally OATP-B). Human OATP1B1 is the most widely studied and the major sodium-independent uptake mechanism for bile acids. Its expression is reduced in inflammatory liver disease (alcoholic hepatitis) [59] and in primary biliary cirrhosis (PBC), particularly in later stages of the disease [12]. Although transcriptional mechanisms are not known for OATPS in cholestatic patients, in vitro studies indicate that OATP1B1, like NTCP, is regulated by both FXR/SHP-dependent and -independent mechanisms where HNF1α is also a primary transcription factor [51]. In contrast to OATP1B1, earlier studies in patients with sclerosing cholangitis suggested that OATP1A2, (formally OATP-A), mRNA expression is actually increased [76]. However OATP1A2 is expressed in cholangiocytes in human liver and not hepatocytes [90]. Bile acids also stimulate expression of OATP1B3 via FXR, raising the possibility that this OATP might function in reverse and extrude organic anions from the cholestatic human hepatocyte. This concept has been confirmed in studies of bile duct obstruction in the rat [75, 91]. Several cholestatic animal models have been used to evaluate the expression of Oatp1al (Slco1a1, Oatp1), Oatp1a4 (Slc1a4, Oatp2), Oatp1b2 (Slco1b2), and Oatp 4 [75, 83, 91]. CBDL, ethinylestradiol (EE), and LPS all result in down-regulation of Oatp1al, although mRNA levels remain unchanged after EE treatment, suggesting that the mechanism of EE cholestasis is mainly post-transcriptional [7, 56, 92, 93]. Estrogen-induced cholestasis results in down-regulation of all basolateral Oatps. The moderate decline in expression of Oatp1al, Oatp1a4, and Oatp1b2 may explain the unchanged sodium-independent transport of bile acids due to overlapping substrate specificity. Reduction in transporter

gene expression seems to be mediated by a diminished nuclear binding activity of transactivators such as HNF1, C/EBP, and PXR by estrogens [94]. Cytokines, particularly IL-1β and TNF-α, play a major role in the regulation of OATP/Oatps and also other bile acid transporters in cholestasis [5, 75, 95]. Basolateral and canalicular transporter systems are down-regulated by both cytokines. Etanercept pretreatment reverses the effects of CBDL on Oatp1a1 down-regulation consistent with involvement of TNF-α [75]. Changes in transporter expression are preceded by a reduction in binding activities at inverted repeat (IR)-1, everted repeat (ER)-8, direct repeat (DR)-5, and HNF-1α sites after 4 hours by either cytokine. Nuclear protein levels of RXR-α are significantly decreased by TNF-α but only transiently affected by IL-1β. Minor reductions of RXR, FXR, PXR, and CAR nuclear proteins are restricted to 4 hours after cytokine application and are paralleled by a decrease in mRNA levels. Activity of HNF-1α as regulator of mouse Ntcp is also suppressed by both cytokines. Decreased binding activities of nuclear receptor heterodimers may also be explained in part by a reduction of the ubiquitous heterodimerization partner, RXR-α [5].

Organic cation transporter (OCT-1, SLC22a1) OCT-1 (organic cation transporter) is the major hepatic uptake transporter for small organic cations. OCT-1 expression in human cholestatic liver has not been assessed but animal models of cholestasis (CBDL and LPS) indicate down-regulation of rOct-1 at the mRNA [96, 97] and protein level and impaired uptake of Oct-1 substrates [96]. The human OCT-1 gene is transactivated by HNF-4α and, like other basolateral hepatic uptake transporters, is suppressed by bile acids via SHP [98]. Thus, hOCT-1 is also likely to be down-regulated in cholestasis.

Organic anion transporter 2 and 3 (OAT-2 and -3, Slc22a6 and Slc22a8) These two members of the OAT family are expressed in liver, and also kidney, and transport a variety of organic anions including bile acids. However, the effects of cholestasis on the expression of these transporters in liver is not well studied. LPS has no effect on Oat-2 in liver in the rat [97], although Oat1 and Oat3 are up-regulated in the kidney 21 hours after CBDL [99].

Bile acid synthesis Bile acids are formed from cholesterol in the liver by a series of enzymatic pathways consisting of the classic (neutral) pathway or the alternative (acidic) pathway

43: ADAPTIVE REGULATION OF HEPATOCYTE TRANSPORTERS IN CHOLESTASIS

[100]. The former is mediated by CYP7A1 and is rate limiting in the overall process. This pathway also involves CYP8B1, which leads to the production of CA and which determines the relative hydrophobicity of bile by determining the ratio of chenodeoxycholic acid (CDCA) to CA. which normally is approximately equal. CYP27A1 mediates the alternative pathway which leads primarily to the formation of CDCA. All bile acid species are conjugated (amidated) with either glycine or taurine by the enzymes bile acid CoA-synthase (BACS) and bile acid CoA:amino acid N -acyltransferase (BAAT). These bile acid conjugate products are then excreted into bile via the BSEP and can undergo both deamidation and 7α-dehyroxylation reactions in the intestine by bacteria, producing deoxycholic acid and lithocholic acid. Unconjugated bile acids are reamidated when they recycle back to the liver in the enterohepatic circulation [100]. During cholestasis, elevated levels of bile acids inhibit bile acid synthesis by activating FXR, which induces SHP and inhibits CYP7A1 gene transcription [101]. In the intestine, bile acids also induce the expression of fibroblast growth factor (FGF-15 in rodents and FGF-19 in humans). Reductions in bile salts in the intestinal lumen result in a decrease in production of FGF-15/19 in the ileum. FGF15/19 is an FXR regulated hormone that interacts with the FGFR4 receptor in hepatocytes in conjunction with β-klotho [102] to down-regulate the production of bile acids from CYP7A1/Cyp7a1 by inhibiting Cyp7a1 expression and reducing gallbladder filling and contractions [103, 104]. FGF19 is also up-regulated in extrahepatic obstruction in human liver [105]. In addition, pro-inflammatory cytokines (TNFα and IL-1β) also play a role via c-Jun N-terminal kinase (JNK)/cJun post-transcriptional signal transduction pathways [106–108]. Thus, multiple transcriptional and posttranscriptional mechanisms exist to reduce the formation of bile acids during cholestasis. However, the master switch appears to be at the level of the intestine, where bile acid homeostasis is exquisitely regulated. Studies in Ostα knockout mice have demonstrated an elevation of bile acids in cells in the terminal ileum, resulting in an Fxr-dependent production of Fgf-15. In contrast, Asbt knockout animals had diminished ileal bile acid uptake and reduced production of Fgf-15 [109, 110]. Furthermore, intestinal selective deletion of Fxr in mice also resulted in increased bile acid pool size and bile acid excretion [111], whereas intestinal specific over-expression of Fxr has the opposite effect. Recent studies suggest that Abst is also down-regulated by FGF15/19 and β-klotho in mouse ileum and cholangiocytes and in human Mz-Cha-1 cells [112]. This regulation may be mediated by LRH-1 via an FXR/SHP pathway in mice and RXR/RAR in humans [112]. Studies in human hepatocytes and HepG2 cells indicate that bile acids and cytokines can also activate hepatic stellate cells to release TGF-β1, which in turn activates Smad3, which enters the nucleus and inhibits HNF4α binding and coactivator interactions including deacetylation

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of the CYP7A1 chromatin [113]. Other recent studies indicate that hepatocyte growth factor also can repress CYP7A1 mRNA expression and the rate of bile acid synthesis in primary human hepatocytes. HGF appears to function by induction of c-Jun and SHP, resulting in increased phosphorylation of extracellular signal-regulated kinase (ERK)1/2, JNK, and c-Jun [114]. This may serve to reduce bile acid synthesis in the early stages of cholestasis when liver regeneration may be stimulated. In rat models of CBDL and α-naphthyl isothiocyanate (ANIT) administration, Cyp8b1 expression, but not Cyp7a1, was significantly inhibited [115], whereas 17αethinylestradiol-induced cholestasis decreased Cyp7a1 but not the acidic pathway via Cyp27a [116]. In patients with primary biliary cirrhosis, CYP7A1 mRNA decreased to 10–20% of control levels with no changes in CYP27A or CYP8B1 [16]. Thus, adaptive changes that serve to reduce bile acid synthesis occur in both rodents and patients with cholestasis.

Bile acid hydroxylation (Phase I) Studies in bile duct ligated rats indicate that most Cyp450 -mediated reactions are diminished. In vitro studies indicate that hydrophobic bile acids are more potent inhibitors, whereas conjugated bile acids are less toxic then unconjugated forms [117]. However, during cholestasis, there is an attempt by the liver to decrease the hydrophobicity of the bile acid pool through hydroxylation reactions mediated predominantly by CYP3A4. These more hydrophilic compounds and their conjugates are more readily excreted in the urine, accounting for the significant levels of (poly)hydroxylated bile acids in the urine of cholestatic animals and patients [118]. CYP3A4 can be regulated by the NRs PXR, FXR [118], VDR [119, 120], and CAR([121, 122] Bile acids also induce Cyp3a11 after CBDL in mice [118] and CYP3A4 in humanized transgenic mice independent of lithocholic acid, a potent PXR ligand [119]. However, CYP3A4 mRNA is only mildly altered in patients with PBC [16].

Bile acid conjugation (Phase II) Glucuronidation, sulfation, and amidation reactions are the major Phase II pathways for diminishing the toxicity of bile acids during cholestasis. Bile acids are conjugated with glucuronide and sulfate in the cholestatic liver, facilitating their excretion back into blood via MRP3 and MRP4 and subsequent elimination by the kidney. Human uridine glucuronyl transferase 1A3 (UGT1A3) is the major enzyme responsible for the hepatic formation of acyl chenodeoxycholic acid 24-glucuronide (CDCA-24G). Cytosolic sulfotransferase 2A (SULT2A) also plays a significant role, particularly in females. In vitro studies in the nuclear receptor Car knockout animal indicate

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that this enzyme may be up-regulated by Car-dependent mechanisms [120]. In contrast, SULT2A is negatively regulated through CDCA-mediated FXR activation in mice and humans [121]. Amidation reactions for bile acids are determined by both BACS and BAAT and both are regulated by FXR. Few studies have examined these enzymes during cholestatic liver injury. However, a rat model of polymicrobial sepsis demonstrated a suppression of transcription with a decrease in rBat expression and binding activity of FXR/RXR. Dexamethasone is able to restore this binding activity and also bile acid-CoA:amino acid N -acyltransferase (BAAT) rBAT protein expression [122]. Mutations in BAAT have also been described in hypercholanemia in childhood [123].

Hepatic efflux mechanisms (Phase III) Basolateral membrane of the hepatocyte (the multidrug resistance associated proteins MRPS/Mrps (ABCC) and the organic solute transporter, OSTα-β/Ostα-β) Cholestatic liver injury results in reversal of the secretory polarity of the hepatocyte. This phenomenon results in the adaptive up-regulation of transporters on the basolateral membrane that now function to facilitate the extrusion of bile salts and other organic solutes by an alternate pathway back into the hepatic sinusoids where they can be eliminated in the urine. Mrp1, -3, and -4 and OSTα/β-Ostα/β are normally expressed at low levels at the basolateral membrane in normal liver [124, 125]. However, these transporters are generally up-regulated during cholestasis. Mrp1, -3, and -4 are all up-regulated by LPS and bile duct ligation in the rat [7, 91]. Mrp1, -5, and -6 do not appear to be of much functional importance in the cholestatic liver, but Mrp3 and -4 are capable of preferentially transporting glucuronidated and sulfated bile acids, respectively [26], and their induction during cholestasis may account for the appearance of these bile acid conjugates in the urine [126, 127]. Induction of Mrp3 after BDL in mice is due to TNF-α dependent up-regulation of Lrh-1 with increased binding of Lrh-1 to the Mrp3 promoter [128]. Hepatic injury is more severe in bile duct ligated TNF-receptor knockout mice [128]. Studies in HepG2 cells indicate that human MRP3 expression is repressed by RXR α:RARα which occupies Sp1 activator sites in the MRP3 promoter. Because RXRα:RARα expression is diminished by cholestatic liver injury, loss of RXRα:RARα may lead to up-regulation of MRP3/Mrp3 expression by de-repressing Sp1 activation in cholestatic disorders [129]. More recent studies in Mrp4

and Mrp3 knockout mice suggest that Mrp4, rather than Mrp3, may be more important in protecting the hepatocytes from bile acid toxicity [15, 130, 131]. In contrast to Mrp3, Mrp4 does not seem to be affected by LPS-mediated inflammation in mice [132]. MRP4 protein, but not mRNA, levels are significantly increased in stage III and IV disease in patients with primary biliary cirrhosis suggesting post-transcriptional regulation [16]. Similar increases in MRP4 have been described in patients with PFIC1 [14] and in late-stage biliary atresia [47]. To date, the mechanism of up-regulation of Mrp4/MRP4 in the cholestatic liver remains unclear, although both CAR and aryl hydrocarbon receptor (AHR) response elements are present in the MRP4 promoter. Bilirubin may be a CAR activator, as suggested by studies in Car knockout mice [133]. There is no evidence that other basolateral Mrps expressed in the liver, including Mrp5 and -6, play any protective role in the adaptive response in cholestatic liver injury [134]. OSTα-β/Ostα-β is the third major hepatic exporter of bile acids and other sterols on the basolateral membrane that is up-regulated in cholestasis [18]. OSTα-β/Ostα-β is a facilitated heteromeric OST where the direction of transport is dependent on the electrochemical gradient between the cell and blood. A considerable difference exists between species and tissues in levels of expression. For example, there is little expression in the normal liver in mouse or rat [18], in contrast to the human hepatocyte. However, Ostα-β is up-regulated by bile duct ligation in mice and rats [18]. In humans, OSTα-β is up-regulated in patients with stage III and IV primary biliary cirrhosis at both mRNA and protein levels [18] . OSTα-β protein is also increased in the liver in biliary atresia.[47]. OSTα-β/Ostα-β is highly regulated by FXR in both human and mice [18, 135, 136] and also in the mouse by Lxr, which shares a DR-1 binding site with Fxr in the mouse promoter [137]. Paradoxically hepatic injury is reduced after bile duct obstruction in Ostα knockout mice (unpublished observations).

Canalicular membrane of the hepatocyte. (MDR1/Mdr1a,b; MDR3/Mdr2; MRP2/Mrp2; BSEP/Bsep; BCRP/Bcrp; ABCG5/G8) While the down-regulation of transporters that function as hepatic uptake mechanisms in the hepatocyte and the up-regulation of basolateral efflux transporters can each be viewed as adaptive responses that retard the accumulation of bile acids and other potential toxic substrates, hepatic levels of bile acids and other constituents of bile still continue to accumulate in the cholestatic liver and are primary substrates for the apical canalicular membrane efflux pumps. All of these transporters are members of the ABC superfamily and the inability of these rate-limiting transporters to excrete their substrates into bile becomes the major determinant of the molecular response to cholestatic liver injury and the cholestatic phenotype.

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Canalicular organic solute transporters Multidrug resistance protein, MDR1/Mdr1a,b (ABCB1/Abcb1) MDR1 encodes for the drug efflux pump known as P-glycoprotein 170 (Pgp-170). In contrast to most of the transport systems for hepatic uptake of organic substrates, most forms of cholestasis result in significant up-regulation of Pgp-170 in both animal models and humans. CBDL [8, 138, 139], LPS [7, 138], and ANIT [8] all increase the expression of Mdr1a/b mRNA, and the level of expression correlates with the severity of the cholestasis as reflected by the levels of plasma bilirubin and alkaline phosphatase [8]. Although the endogenous substrates for Mdr1 are not certain, Mdr1 is capable of transporting bile acids, albeit with a much lower affinity than Bsep [140]. MDR1 protein is increased in late-stage PBC and in biliary atresia [12, 47]. Molecular regulation of Mdr1/MDR1 is thought to be mediated by NF-κB transcriptional mechanisms [141]. Both CAR and PXR activators can also stimulate MDR1 expression [142, 143]. To date, no mutations have been described in MDR1 that cause human disease. However, polymorphisms have significant effects on drug absorption, excretion, and toxicity [144].

MDR3/Mdr2 (ABCB4/Abcb4) The importance of this phospholipid export pump in the pathogenesis of cholestasis is dramatically demonstrated by mutations in the MDR3/Mdr2 gene that result in PFIC-3 in children [145] and biliary cirrhosis in the mouse knockout model, Mdr2(–/–) [146]. In the absence of the canalicular Mdr2/MDR3 gene product, a phospholipid flippase, phosphatidylcholine cannot be excreted into bile, and bile salts, which continue to be excreted normally, cannot form mixed micelles. The consequence of this genetic defect in both animals and humans is progressive injury to the bile duct epithelium [147–152]. The function of MDR3/Mdr2 was established in the Mdr2 knockout mouse, which has a complete deficiency of phospholipid in the bile and which develops portal inflammation and bile ductular proliferation, characteristic of non-suppurative sclerosing cholangitis [148]. With time, fibrosis occurs and these animals develop a biliary-type cirrhosis and, in some cases, hepatocellular carcinoma [152]. Children with PFIC-3 also have defective phospholipid excretion in bile, bile duct proliferation by histological examination, and elevated γ-glutamyl transferase, thus distinguishing these patients from PFIC-1 and -2, where bile duct proliferation is absent and γ-glutamyl transferase is normal [153]. Heterozygotes for Mdr2 or MDR3 demonstrate partial deficiencies in phospholipid excretion but under normal conditions do not have a cholestatic phenotype [145, 150,

691

154]. However, mothers of PFIC-3 patients are obligate heterozygotes (MDR3+/–) and are at risk to develop cholestasis during the third trimester of pregnancy when high levels of estrogens are present [155–157]. Polymorphisms and mutations in MDR3 can predispose patients to cholestatic liver injury when exposed to other potential cholestatic agents, including drugs and environmental toxins [158]. A missense mutation in ABCB4 has been described in cases of ductopenic cholestatic liver disease in adults. These patients present a wide spectrum of disease from intrahepatic cholestasis of pregnancy through fibrosis to cirrhosis and death in childhood and adulthood [159]. Mdr2/MDR3 is up-regulated in most forms of cholestasis, including bile duct obstruction [139], and ANIT [8, 115] and in patients with biliary atresia [47]. Mdr2/MDR3 is regulated in large part by Fxr/FXR-mediated mechanisms [160], but is also regulated by Pparα in mice [161]. Trials with fibrates in patients with primary biliary cirrhosis are based in part on this finding [49, 51].

MRP2/Mrp2 (ABCC2) MRP2 encodes for the conjugate drug export pump, also known as the multidrug resistance protein 2 or the canalicular multidrug organic anion transporter (cMOAT) [124]. This ABC transporter is the major export pump for bilirubin diglucuronides, glutathione conjugates, and divalent bile acid conjugates with sulfates and glucuronides. It is a major determinant of bile acid independent bile flow and drug conjugate excretion. Levels of expression in human liver show considerable variability [162] and single nucleotide polymorphisms (SNPs) correlate with drug clearance [163]. Mutations in the Mrp2 gene result in a stop codon and premature termination of protein translocation in the mutant TR–/GY [164] and EHBR rats [165]. Mutations in the MRP2 gene in man result in the Dubin–Johnson syndrome [166–168]. These mutations are a genetically determined cause of conjugated hyperbilirubinemia. Although not a cholestatic disorder by definition, since bile salt excretion is normal, this mutation results in impaired excretion of a variety of amphipathic organic anions, including leukotrienes, conjugated bilirubin, divalent bile acids, and corproporphyrin isomer series 1, in addition to a variety of other compounds including bromosulfthalein (BSP), Indocyanine Green, and oral cholecystographic agents [124]. Antibiotics, such as ampicillin and cephtriaxone, and heavy metals are also excreted by Mrp2. Although it is not known if bile secretion is impaired in the Dubin–Johnson syndrome, bile salt independent bile flow is reduced in the rat model as a result of impaired glutathione excretion [169]. Current evidence suggests that glutathione is a low-affinity substrate for Mrp2 and that Mrp2 is the major pathway for excretion of glutathione and also oxidized GSSG and glutathione

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conjugates [170]. Because the excretion of glutathione is a major determinant of bile salt independent bile flow [171], polymorphisms in the MRP2 transporter which result in diminished glutathione excretion may contribute to or predispose to other forms of toxic and cholestatic liver injury [162–173]. Therefore, it is noteworthy that the expression of Mrp2, at both the mRNA and protein level, is markedly down-regulated in several experimental animal models of cholestasis, including CBDL, EE, and LPS [7, 138, 174]. This impairment is particularly noted following administration of LPS, providing an experimental explanation for the increase in conjugated bilirubin in serum that is characteristic of sepsis-induced jaundice [7, 138, 175]. Retrieval of Mrp2 from the canalicular membrane to a submembranous localization may precede changes in mRNA expression after LPS [176, 177] and phalloidin [178] treatment and represents early post-transcriptional events that may contribute to cholestasis. In LPS-treated human liver slices, mRNA is preserved whereas MRP2 protein is essentially absent [63], consistent with a post-transcriptional event. MRP2 protein is also markedly reduced in liver biopsies from patients with inflammatory cholestasis [59]. The promoter region of Mrp2 is activated by the nuclear receptor heterodimer, RXRα:RARα, whereas IL-1β represses promoter activity via this element [69]. Following bile duct ligation in rats, Mrp2 is down-regulated and IL-1β is up-regulated associated with down-regulation of liver RXRα:RARα nuclear protein levels and binding to the Mrp2 promoter cis element. Thus, Mrp2 expression in obstructive cholestasis is associated with cytokine-dependent alterations in the RXRα:RARα NRs. However, response elements for CAR, PXR, FXR, and Ahr also are present in the Mrp2/MRP2 promoter so that the mechanism of regulation of this transporter in cholestasis remains complex [52, 179]. As previously mentioned, down-regulation of Mrp2 in cholestasis also explains the impairment in the biliary excretion of a variety of substrates including glutathione, bilirubin diglucuronide, cysteinyl leukotrienes, antibiotics, bile acid sulfates, and glucuronides. Because bilirubin and glutathione are antioxidants, their hepatic retention in cholestasis might have some cytoprotective effects [180].

The bile salt export pump (BSEP/Bsep, ABCB11/Abcb11) These genes encode for the “sister of P-glycoprotein” the canalicular membrane bile salt export pump and the major, if not the sole, determinant of bile salt dependent bile flow. Mutations in BSEP result in PFIC-2 [181–183], and also benign recurrent cholestasis and intrahepatic cholestasis of pregnancy in adults [182, 184]. More then 100 different mutations have been described in families with these

disorders [183]. PFIC-2 resembles the Byler’s disease (PFIC-1) phenotype, with absence of bile duct proliferation and normal γ-glutamyl transferase levels in serum. Bsep knockout mice also demonstrate impaired bile salt transport into bile, although these mice do not become cholestatic unless infused with CA [185]. Together these findings provide compelling evidence that BSEP/Bsep is the canalicular transport protein for determining bile salt excretion and bile salt dependent bile formation. Therefore, it is surprising to find that the expression of Bsep in several rat models of cholestasis is only moderately impaired [7, 175]. Although a variety of cholestatic agents profoundly inhibit ATP dependent bile salt transport in vitro in isolated rat liver canalicular membrane vesicles, including LPS [55, 186], EE [187], cyclosporin A, and rifamycin [188, 189], down-regulation in vivo is less pronounced. Although 3 days of CBDL in the rat results in inhibition of Bsep mRNA and protein expression by ∼30 and 50%, respectively, after 7–14 days, mRNA and protein expression recover to ∼60 and 80% of control values, respectively and Immunofluorescence studies indicate that the transporter remains at the canalicular membrane [175]. Furthermore, bile salt excretion continues in the face of complete obstruction, albeit at a reduced rate [175]. More recent studies suggest that Bsep down-regulation during bile duct ligation in the rat is limited to periportal regions of the hepatocytes and is mediated by TNF-α and Il-1β [91]. LPS and EE administration in vivo to rats also results in only partial inhibition of Bsep expression [175]. Although studies in human liver slices exposed to LPS show loss of BSEP protein [63] and reduced staining of protein in patients with inflammatory cholestasis [59], BSEP expression is maintained in patients with primary biliary cirrhosis [12] and is reduced in early but not late stages of biliary atresia [7, 47], where it is maintained in its normal amount and location [190]. Together these experimental observations suggest that Bsep/BSEP is variably preserved during cholestatic liver injury. Bsep/BSEP is strongly regulated by Fxr/FXR in human, rat, and mice [191–193] and Bsep induction by bile acids is absent in the FXR knockout mouse [194]. Thus differences in levels of Bsep/BSEP expression in cholestasis may be related in part to differences in the levels of expression of this nuclear receptor. For example, while initially reduced, FXR and BSEP levels return to normal in late-stage cases of biliary atresia [47]. Recent studies indicate that LRH-1 also can regulate BSEP expression and thus may play a supporting role to FXR in maintaining hepatic bile acid levels and in coordinating the expression of both CYP7A1 and BSEP to determine bile acid synthesis and excretion [195]. Taurocholate transport is competitively cis-inhibited by various cholestatic drugs, including cyclosporin A, rifamycin, rifamycin SV, and glibenclamide in Sf9 cells expressing rat Bsep [189]. Altogether, these findings suggest

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that the cholestatic effects of these compounds may be determined in part by the extent to which the canalicular export pumps continue to function as export pumps. Interestingly, the cholestatic metabolite estradiol-17β-glucuronide inhibited ATP-dependent taurocholate transport only when Mrp2 was co-expressed in Sf9 cells, suggesting that this compound results in trans-inhibition of Bsep-mediated bile salt transport only after excretion into the bile canaliculi by Mrp2 [189]. Thus, some drugs may produce cholestasis by inhibiting the BSEP only after excretion into bile. Polymorphisms in BSEP also affect the level of expression of BSEP in human liver [162] and may predispose to some forms of drug-induced cholestasis [158]. ABCB11 1331T > C polymorphism is a susceptibility factor for cholestasis of pregnancy [196].

Other canalicular solute and lipid transporters/flippases (BCRP, ABCG2) and ABCG5/8, MATE-1 and FIC1) The breast cancer resistance protein (ABCG2) is also expressed on the canalicular membrane [197]. BRCP shares broad substrate specificity with MRP2 and excretes sulfated drug conjugates [198], but its role in cholestasis is unclear. BCRP is down-regulated in the duodenum of patients with obstructive cholestasis [199] but not in bile duct ligated rats [33]. Polymorphisms in BRCP are speculated to play a role in triglitazone sulfate excretion and could possibly result in cholestasis induced by this metabolite [200]. Sterolin 1 and 2 (ABCG5/8) are two ABC transporters that form a heterodimer at the canalicular membrane and account in large part for the excretion of cholesterol and plant sterols [201]. Little is know about its role in cholestasis, although estrogen-induced cholestasis results in diminished mRNA expression [202]. Reduced expression of ABCG5/8 would be expected to contribute to the hypercholesterolemia that develops during cholestasis. The multidrug and toxic compound extrusion protein-1 (MATE-1) is also expressed in the liver on the apical canalicular membrane [203]. Members of the MATE family are organic cation exporters that excrete metabolic or xenobiotic organic cations from the body by way of a H+ or Na+ exchange mechanism [203–205]. However, little is known about its specific role in the liver and whether it is regulated during cholestasis. The gene product of familial intrahepatic cholestasis-1 (FIC1, ATP8B1) is a P-type ATPase which functions as a phosphatidylserine flippase at the canalicular membrane [206–208]. Mutations in FIC1 result in PFIC-1, also called Byler’s disease [209]. PFIC-1 is phenotypically similar to PFIC-2. Whether FIC1’s function is impaired in other forms of cholestasis and contributes further to cell injury is not clear.

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Canalicular ion transporters (AE2, also SLC4A2) Anion Exchanger-2 (AE2) This transporter encodes for the canalicular Cl− /HCO− 3 exchanger that regulates the excretion of bicarbonate, a partial determinant of canalicular bile salt independent bile formation [210, 211]. Anion exchanger-2 (AE2) is also expressed on the luminal membrane of the cholangiocyte and is a determinant of bicarbonate excretion from this epithelium [212]. Thus, impairment of AE2 would be expected to reduce bile flow and thus might predispose to other cholestatic insults. Reduced levels of liver AE2 mRNA expression and immunoreactivity at the canaliculus and in bile ducts have been described in patients with PBC but not in other cholestatic and non-cholestatic liver diseases, suggesting that they might be primary rather then secondary changes [213–215]. Reduced expression of AE2 mRNA has also been observed in salivary glands in patients with PBC and the sicca syndrome, suggesting that there may be a generalized deficiency in this transporter in this disease [216]. One study examined AE2 activity in a cholestatic animal model after EE treatment but found activity to be normal [217]. Treatment of patients with ursodeoxycholic acid (UDCA) has normalized the expression of AE2 mRNA and partially restored protein expression [213]. In cholangiocytes from needle biopsies, basal AE2 activity was significantly decreased in PBC compared with normal livers or disease controls. In addition, cAMP increased AE2 activity in cholangiocytes from both normal and non-PBC livers, but not in PBC cholangiocytes, further suggesting that this was a primary defect in this secretory mechanism [218] More recent studies demonstrate that Ae2a,b-deficient mice develop altered immunological changes, mitochondrial antibodies, and histological findings that resemble PBC [214, 219]. Further, the combination of UDCA and dexamethasone was found to increase the expression of AE2b1 and AE2b2 and enhance AE2 activity in human hepatobiliary cell lines [220]. These studies suggest one of several mechanisms by which UDCA may improve outcomes in PBC patients.

CONCLUSION This chapter has focused on the adaptive response that membrane transporters in hepatocytes play in inherited and acquired forms of cholestasis and the role of NRs in this process. Information about the mechanisms of these transcriptional events is rapidly expanding, revealing the complexity and the interrelatedness of these responses in the form of extended networks [4]. Adaptative mechanisms also occur in cholangiocytes, kidney, and intestine

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that contribute to the ability to modulate this disorder but are beyond the scope of this review. In the future, progress will come from new therapeutic strategies, most likely novel nuclear receptor ligands that stimulate these protective pathways. The recent literature can be consulted for further information on future therapeutic opportunities and potential problems [35, 221, 222].

ACKNOWLEDGMENT Work cited from this laboratory is supported by USPHS DK 25636 and DK P30-34989.

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44

Pathogenesis of Portal Hypertension Roberto J. Groszmann1,2 and Juan G. Abraldes3 1 Veterans

Administration Medical Center, West Haven, CT, USA University School of Medicine, New Haven, CT, USA 3 Hepatic Hemodynamic Laboratory, Liver Unit, Hospital Clinic University of Barcelona, IDIBAPS and CiberEHD, Barcelona, Spain 2 Yale

Portal hypertension is the hemodynamic abnormality most frequently associated with cirrhosis of the liver, although it is also recognized less commonly in a variety of hepatic and extrahepatic diseases. Many of the most lethal complications of liver cirrhosis are related to the presence of portal hypertension, including hemorrhage from gastro-esophageal varices, hepatic encephalopathy, ascites and functional renal failure, bacterial infections, and hepatopulmonary syndrome. A thorough knowledge of the pathogenesis leading to portal hypertension sets the framework for a rational approach to treatment, and is therefore, central in the management of the patient with liver cirrhosis and in devising rational investigational strategies. This chapter provides an overview of the basic pathophysiological mechanisms of the intrahepatic, splanchnic, and systemic circulatory derangements involved in the genesis of portal hypertension.

RESISTANCE AND FLOW AS THE DETERMINANTS OF PORTAL PRESSURE Ohm’s law states that changes in pressure (P1 − P2 ) along a blood vessel are a function of the interplay between blood flow (Q) and vascular resistance (R): P 1 − P2 = Q × R

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

It follows that portal hypertension may develop due to an increase in portal blood flow, to an increase in resistance to portal blood flow, or both. The pathophysiology of portal hypertension is best approached by analyzing these components separately, although mathematical equations necessarily oversimplify the complex and dynamic interactions that exist in biological systems. Resistance to the flow in vessels can be expressed by Poiseuille’s law: 8nL R= πr 4 where n is the coefficient of viscosity, L the length of the vessel, and r its radius. Under physiological conditions, resistance is mainly a function of changes in r, which have a dramatic influence because these are taken to the fourth power. By contrast, L and n are basically constant because neither the length of a vessel nor the viscosity of blood varies greatly under usual circumstances. The liver is the main site of resistance to portal blood flow but the liver itself has no active role in regulating portal inflow; this function is provided by resistance vessels at the splanchnic arteriolar level. Hence the normal liver is a passive recipient of fluctuating amounts of blood flow that, due to its large and distensible vascular network, can encompass a wide range of portal blood flow with minimal effect on pressure in the portal system [1]. Hence an increase in portal venous inflow, per se, does not induce portal hypertension. This means that the primary and

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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necessary factor for the development of portal hypertension is an increased resistance to portal blood flow. However, once portal hypertension develops, a series of mechanisms (not yet fully characterized) lead to an increase in portal venous inflow that contributes to perpetuating and aggravating portal hypertension.

ABNORMALITIES IN VASCULAR RESISTANCE TO PORTAL BLOOD FLOW Hepatic vascular resistance In liver cirrhosis, the most frequent cause of portal hypertension in Western countries, marked morphological aberrations, characterized by fibrous tissue and regenerative nodules, results in vascular obliteration that leads to increased resistance to blood flow. In the early stages of liver fibrosis, sinusoidal capillarization is probably the initial factor that induces portal hypertension. Capillarized sinusoids [2] are characterized by accumulation of extracellular matrix in the space of Disse and loss of fenestrations of sinusoidal endothelial cells. This change in the phenotype is not limited to morphological changes, since it results also in functional changes of the sinusoidal endothelial cells that might lead to activation of adjacent hepatic stellate cells (HSCs) [3]. In viral hepatitis, these changes occur initially in the periportal area, whereas in alcoholic liver disease it is predominant at the pericentral area. Later, the development of fibrous septa and regenerative nodules markedly alter the hepatic angioarchitecture [4]. Recent work showed that increased thickness of fibrous septa and decreasing size of regenerative nodules were independently associated with the severity of portal hypertension [5], stressing the relevance of these architectural disturbances in the genesis of portal hypertension. Additionally, thrombosis of the small portal and hepatic venules [6] also contributes to increased hepatic resistance and could be an important factor in the progression of cirrhosis. Although these morphological changes are undoubtedly the most important factor, functional factors leading to an increase in intrahepatic vascular tone also contribute to increased intrahepatic resistance in cirrhosis. The initial demonstration that vasodilators could decrease hepatic resistance in cirrhosis was originally shown by Bhathal and Grossman [7], where the use of the isolated and perfused liver model allowed the changes in hepatic resistance to be evaluated independently of the changes in systemic hemodynamics. It was suggested that up to 30% of the increase in hepatic resistance in cirrhosis is due to increased vascular tone. There have been no convincing studies that quantified the magnitude of this functional component in patients with cirrhosis, and it is not well known whether its importance changes with the natural history of the

disease. A recent study in humans evaluating the relation between liver stiffness (with transient elastography) and portal pressure confirmed that architectural changes are the major factors contributing to portal hypertension in early phases of the syndrome, while dynamic alterations (at both the hepatic and splanchnic levels) become especially relevant in advanced portal hypertension [8]. The dynamic component in the increase in intrahepatic resistance reflects the existence of contractile structures in the liver that modulate hepatic resistance in response to endogenous or pharmacological vasoactive substances. In the normal liver, it has been demonstrated that the hepatic sinusoids can contract, and that changes in sinusoidal caliber colocalize with HSCs [9]. These cells, located in the space of Disse, normally act as a deposit for retinoids and regulate the extracellular matrix turnover, but their contractile properties allow HSCs to behave as tissue pericytes, which regulate microcirculation through capillary contraction [10]. The contractile capacity of HSCs, probably present in the normal liver [9], is particularly important after liver injury, when HSCs acquire an activated phenotype, with increased proliferative, synthetic and contractile capacity, behaving as myofibroblasts [10]. Apart from HSCs, other vascular structures modulate the hepatic vascular resistance in the liver. In the normal liver, it has been demonstrated that portal venules contract in response to vasoactive substances [11], but it is unknown whether this contributes to the increased hepatic resistance in cirrhosis. In that regard, data from a rat model of cirrhosis suggest that pre-sinusoidal resistance is decreased in cirrhosis [12]. The same study suggested that the contraction of the post-sinusoidal vascular bed could contribute to the increase in hepatic resistance and to ascites formation in cirrhosis. It is uncertain whether these results could be generalized to human cirrhosis. All these contractile structures are responsible to several vasoconstrictors and vasodilators that modulate hepatic resistance (Figure 44.1). These substances can be of hepatic origin and act in a paracrine fashion [nitric oxide (NO), prostacyclin, hydrogen sulfide (H2 S), carbon monoxide (CO), endothelin, locally produced angiotensin II, thromboxane, leukotrienes], can arrive at the liver from the systemic circulation (circulating angiotensin II, vasopressin, or norepinephrine) or can be of neural origin (norepinephrine). It is not well known which of these systems is more relevant but it is clear that in cirrhosis there is an imbalance between vasoconstrictive and vasodilating forces, characterized by an abundance of vasoconstrictors and a deficient production and deficient response to vasodilators. These abnormalities are amplified by the fact that, compared with the normal liver, the hepatic vascular bed of the cirrhotic liver exhibits an increased response to vasconstrictors, and a deficient response to vasodilators. In addition, most vasoconstrictors have profibrogenic actions, whereas vasodilators have antifibrogenic properties, so the effects of this imbalance go beyond the increase in intrahepatic vascular tone.

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Figure 44.1 Vasoactive molecules known to be involved in the regulation of vascular tone in cirrhosis. In the arterial splanchnic circulation (c), agonists such as adrenomedullin, vascular endothelial growth factor (VEGF), and tumor necrosis factor alpha (TNFα) or physical stimuli such as shear stress stimulate Akt, which directly phosphorylates and activates endothelial nitric oxide synthase (eNOS). eNOS requires co-factors such as tetrahydrobiopterin (BH4 ) for its activity. Heat shock protein 90 (Hsp90) is one of positive regulators of eNOS. Like NO, carbon monoxide (CO) produced by hemeoxygenase-1 (HO-1) causes vasodilatation by activating soluble guanylate cyclase (sGC) to generate cyclic guanosine monophosphate (cGMP) in vascular smooth muscle cells. Prostacyclin (PGI2) is synthesized by cyclooxygenase (COX) and elicits smooth muscle relaxation by stimulating adenylate cyclase (AC) and generation of cyclic adenosine monophosphate (cAMP). In the intrahepatic circulation (b), decreased NO and increased thromboxane A2 (TXA2) production in SECs results in a net reduction of vasorelaxation in the intrahepatic circulation. Endothelin-1 (ET-1) has dual vasoactive effects, mediating vasoconstriction through binding to endothelin A (ETA) receptors located on HSCs and causing HSC contraction. Binding of ET-1 to ETB receptor (ETBR) mediates vasodilatation through Akt phosphorylation and eNOS phosphorylation in normal liver. In cirrhosis, G-protein-coupled receptor kinase-2 (GRK2), an inhibitor of G-protein-coupled receptor signaling, is up-regulated in SECs, leading to the impairment of Akt phosphorylation and a reduction in NO production. An increased production of COX-1-derived vasoconstrictor prostanoid TXA2 is also an example of endothelial dysfunction in cirrhosis. Modified from Iwakiri and Groszmann [13] with permission

Deficit and hyporesponse to vasodilators Nitric Oxide (NO) In the early 1990s, it was demonstrated that in the normal liver sinusoidal endothelial cells respond to increases in shear stress with an increase in NO production [14], thus suggesting that NO was a major factor regulating intrahepatic resistance. This mechanism allows the normal liver to accommodate physiological changes in portal

blood flow, such as those occurring after the ingestion of a meal, with minimal changes in portal pressure. In contrast, the cirrhotic liver exhibits endothelial dysfunction, characterized by an insufficient production of NO, that contributes to an increase in hepatic vascular tone (and, thus, to the development and progression of portal hypertension), and in a decreased capacity to accommodate increases in flow [15–17] (Figure 44.2). This makes physiological increases in portal blood flow result in marked increases in portal pressure in cirrhotic patients [18]. Recent studies have validated hepatic NO deficiency

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THE LIVER: ABNORMALITIES IN VASCULAR RESISTANCE TO PORTAL BLOOD FLOW

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Figure 44.2 Impaired NOx production, pressure regulation, and NOS activity in cirrhotic liver. (a) NOx in the perfused liver was significantly lower in cirrhotic than in control animals. Addition of l-NMMA to the perfusate significantly inhibited NOx production in both groups. (b) Flow-induced increases in perfusion pressure in both experimental groups. The increase in perfusion pressure was greater in cirrhotic animals, as demonstrated by a significantly greater perfusion pressure at each flow rate and a significantly greater pressure–flow slope (k ). (c) NOS activity was significantly reduced in liver tissue from cirrhotic animals compared with parallel control animals when normalized per milligram of protein (a) and per milligram of liver tissue. Reproduced from Shah et al. [17] with permission

as a useful therapeutic target in cirrhosis, showing that it is possible to improve portal hemodynamics by selectively increasing NO bioavailability in the liver circulation, either by transfection of the liver with adenovirus encoding nitric oxide synthase (NOS) [19], by the administration of a liver-selective NO donor [20], or by enhancing NOS activity with drugs or gene therapy [21–23].

The molecular mechanisms leading to the insufficient production of NO in cirrhosis have attracted great attention. Endothelial nitric oxide synthase (eNOS) expression has been found to be normal [16, 17] or decreased, whereas eNOS activity has been found consistently to be decreased. This is due to abnormalities in the complex post-translational regulation of the enzyme that include protein–protein interactions, phosphorylation, and intracellular localization [13]. Among other mechanisms, interaction with caveolin-1 reduces the activity of eNOS [17], whereas Akt-dependent eNOS phosphorylation increases eNOS activity [22]. In the cirrhotic liver, an increase in caveolin-1 expression has been found and increased interaction of this protein with eNOS, with ensuing decreased activity of eNOS [17]. Also, a decrease in Akt-dependent eNOS phosphorylation has been found in the cirrhotic rat liver [22]. This might be due, in part, to the up-regulation of G-protein-coupled receptor kinase-2 (GRK2), an inhibitor of G-protein-coupled receptor signaling. Indeed, GRK2 knockdown restores Akt phosphorylation and NO production, improving portal hypertension [24]. Another mechanism that might interfere with Akt-dependent eNOS phosphorylation is an increase in the Rho-kinase signaling, that prevents the interaction between Akt and eNOS [25]. The up-regulation of Akt-dependent eNOS phosphorylation by drugs [21] or gene therapy [22] improves NO bioavailability and decreases hepatic resistance in cirrhosis. Further studies have shown that a deficiency in the eNOS essential cofactor tetrahydrobiopterin (BH4 ) [26] and increased levels of the eNOS endogenous inhibitor ADMA [27] might also contribute to the decrease in eNOS activity in cirrhosis. In the cirrhotic liver circulation, not only is NO production deficient, but also the dilating response to NO is impaired [28]. One of the mechanisms that account for this finding is the destruction of NO before it reaches its targets by increased oxidative stress and superoxide production [29]. Thus, administration of antioxidants might improve liver NO availability in cirrhosis [30]. Additionally, the downstream signal pathways of NO (the cyclic guanosine monophosphate (cGMP) pathway being the most relevant) are also impaired in cirrhosis [31]. A recent study showed that phosphodiesterase-5 (PD5), the enzyme that degrades cGMP, the second messenger of NO, is overexpressed in the cirrhotic liver. PD5 inhibition with sildenafil restores a normal response of hepatic vascular bed in the isolated perfused cirrhotic liver to NO [31].

Carbon Monoxide (CO) This is another endogenous vasodilator that has been shown to modulate hepatic resistance in the normal liver [32]. In the cirrhotic liver, an increased expression of hemo-oxygenase-1 (HO-1) (the enzyme responsible for CO synthesis) has been demonstrated [33], but the relevance of this finding in the pathogenesis of portal hypertension is unknown.

44: PATHOGENESIS OF PORTAL HYPERTENSION

Hydrogen Sulfide (H2 S) H2 S, which has been called the “third gas” (after NO and CO), is a gaseous transmitter that has been recently implicated in the regulation of vascular tone [34]. H2 S results from the metabolism of homocysteine, and in vascular tissues it is mainly produced by the enzyme cystathionine γ-lyase (CSE). A recent study has shown that H2 S might regulate hepatic vascular tone in the normal liver in an NO-independent pathway, since it relaxed HSCs and attenuated the vasoconstrictive response to an alpha-adrenergic agonist. In contrast, in the liver from rats with experimental cirrhosis there was decreased H2 S production [35] due to a decreased expression of CSE in HSC, leading to an increased response to vasoconstrictors of the liver vascular bed.

Increased production and response to vasoconstrictors As discussed previously, both the normal and the cirrhotic liver vasculatures have contractile capacity. Consequently, infusion of vasoconstrictors increases intrahepatic resistance in the isolated liver [36, 37]. In the cirrhotic liver there is an increase both in local vasoconstrictors produced in the liver itself and in circulating levels of vasoconstrictors [38]. Further, the cirrhotic liver exhibits an enhanced response to certain vasoconstrictors with respect to the normal liver [12, 36, 37, 39]. This hyper-response has been attributed to different abnormalities, namely an increase in the amount of “contractile machinery” within the liver (proliferation of myofibroblast and activation of HSC), a deficient production of vasodilators, changes in expression and sensitivity of the receptors, and amplification of the vasoconstrictive response by secondary production of vasoconstrictors. While the first two mechanisms would be non-specific and of concern to all vasoconstrictors, the other two would be associated with changes in the signaling pathways of specific vasoconstrictors.

Endothelin This is the vasoconstrictor that has been most thoroughly studied in hepatic circulation, but its role in portal hypertension remains unclear. There is evidence that circulating and intrahepatic levels of endothelin are increased in cirrhosis [40], and that endothelin can increase intrahepatic resistance in the isolated liver [41]. Endothelin exerts dual vasoactive effects. Binding of endothelin to ET-A receptors, located in smooth muscle cells and HSC, mediates constriction, whereas binding to ET-B receptors mediates both constriction (at the HSC) and dilation (through endothelial ET-B receptors, via Akt/eNOS activation) [42]. The vasoconstrictive response of the cirrhotic liver to endothelin has been found to be both increased [41] and

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decreased in comparison with the normal liver, whereas the vasoconstrictive response to selective ET-B agonists has been shown to be uniformly increased in cirrhosis. On the other hand, activation of HSC is associated with a change in endothelin receptor pattern, from a predominance of ET-A receptors in normal subjects to a predominance of ET-B receptors in cirrhosis, which could suggest a change in the sensitivity of HSC to endothelin. In vivo studies have not contributed to clarification of the role of endothelin in portal hypertension. Acute administration of the mixed ET-A/ET-B receptor blocker bosentan decreased portal pressure [43, 44], whereas chronic administration of RO 48-5695, a second-generation ET-receptor mixed blocker, did not modify portal pressure and even increased liver fibrosis [45]. In contrast, chronic selective blockade of ET-A receptor with LU 135252 dramatically decreased collagen accumulation in a secondary biliary cirrhosis model [46], whereas acute administration of another ET-A blocker (FR 139317) to cirrhotic rats did not lower portal pressure [47]. Results in humans have shown that neither ET-A nor ET-B blockers are able to decrease portal pressure in cirrhotic patients, and that ET-A blockers induce marked hypotension [48]. These results led to a decrease in the interest in the role of endothelin in portal hypertension.

Angiotensin II In patients with advanced cirrhosis, there is a marked activation of the renin–angiotensin system (RAS), that correlates with the severity of portal hypertension [38]. In experimental studies in the isolated liver, angiotensin II infusion increases intrahepatic resistance [36], and infusion of angiotensin II through the portal vein of cirrhotic patients increases portal pressure and decreases hepatic blood flow. These effects are probably mediated by the contraction of HSCs, which, once activated, express angiotensin II type I receptors, and contract in response to angiotensin II [49]. Further, HSCs express all the components of the RAS [50], which suggests that not only circulating angiotensin, but also locally produced angiotensin II has a role in the increased resistance in cirrhosis. Systemic activation of RAS, however, is a homeostatic process that contributes to maintaining arterial pressure in advanced cirrhosis [51], and therefore RAS blockade induces marked hypotension in these patients without a significant decrease in portal pressure [52].

Cysteinyl-leukotrienes These are potent vasoconstrictors that are derived from arachidonic acid through the 5-lipoxygenase (5-LO) pathway. Cysteinyl-leukotriene production is increased in the livers from cirrhotic rats, which show an increase in 5-LO expression [39]. The isolated cirrhotic liver responds to leukotrienes with a marked increase in intrahepatic

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THE LIVER: THE INCREASE IN PORTAL BLOOD INFLOW AND THE HYPERDYNAMIC CIRCULATORY SYNDROME

resistance, and 5-LO blockade decreases hepatic resistance in cirrhotic livers [39]. There is no information on the in vivo hemodynamic effects of 5-LO blockers.

Alpha-adrenergic stimulus Alpha-adrenergic agonists increase hepatic resistance [36] and this effect is higher in the cirrhotic than the normal liver [12, 15, 37]. This finding cannot be fully explained by a decreased production of NO in the cirrhotic liver [37]. Recent studies have shown that this hyper-reactivity might be mediated, in part, by an increased activity of the RhoA/Rho-kinase pathway in the cirrhotic liver, probably at the HSC level [53]. In addition, and distinct from the effect of alpha-adrenergic agonists on the contractile machinery of the liver, the cirrhotic liver responds to alpha-adrenergic agonists with an increased endothelial production of cyclooxygenase (COX)-1-derived vasoconstrictive prostanoids (mainly thromboxane A2 ). This secondary production of vasoconstrictors amplifies the vasoconstrictive effect of alpha-adrenergic agonists [54, 55]. Hence these findings expand the concept of endothelial dysfunction in cirrhosis, characterized not only by an insufficient production of NO, but also by an increased production of vasoconstrictive prostanoids. In vivo studies have shown that alpha-adrenergic blockade decreases intrahepatic resistance in patients with cirrhosis [56], but also induces severe hypotension. In summary, the cirrhotic liver exhibits a dysregulation in the production of and response to a number of vasoconstrictors and vasodilators. This situation might be acutely exacerbated by intercurrent factors such as infections [57], that might result in abrupt increases in portal pressure.

Collateral resistance The development of collaterals in portal hypertension is the key event that leads to severe complications such as variceal bleeding and hepatic encephalopathy. Collaterals develop as a consequence of the pressure increase in the portal system, allowing the decompression of the portal territory to vascular beds of low pressure. However, this decompression is ineffective, not because these collaterals have a very high resistance, but because in parallel with the development of collaterals an increase in portal blood inflow maintains portal hypertension [58–60]. Collateral formation results in part from the opening and dilation of pre-formed channels but also from an active, vascular endothelial growth factor (VEGF)-dependent angiogenesis process. In that regard, recent studies have shown that VEGF expression increases in the intestine and mesentery of rats with pre-hepatic portal hypertension and early VEGF blockade reduces by 50% the development of collaterals in this model [61, 62]. Collateral formation has also been shown

to be NO dependent [63, 64], raising the possibility that VEGF acts upstream of NO in the collateralization process. Additional studies have shown that local NADPH-dependent oxidative stress at the splanchnic circulation [65] and platelet derived growth factor (PDGF) [66] also mediate in the development and stabilization of collaterals. How all these mediators interact in the development and maintenance of collaterals, and their relative relevance in the process, require further clarification. Since in advanced portal hypertension as much as 90% of portal blood flow could be shunted through porto-systemic collaterals, changes in collateral resistance can modify portal pressure. A number of studies performed in a model in which the collateral bed is perfused in situ have demonstrated that these vessels have functional receptors for vasopressin, endothelin, serotonin, and alpha- and beta-adrenergic receptors, and respond to NO with vasodilatation [63, 67].

THE INCREASE IN PORTAL BLOOD INFLOW AND THE HYPERDYNAMIC CIRCULATORY SYNDROME In 1953, Kowalski and Abelman first demonstrated that cirrhosis is associated with a hyperdynamic circulatory syndrome, characterized by a marked decrease in systemic vascular resistance, arterial hypotension, and increased cardiac output [68]. These profound alterations in peripheral circulation in liver cirrhosis contribute to the development of complications such as ascites, hepato-renal syndrome, and hepato-pulmonary syndrome [69]. Additionally, in the 1970s, a series of studies in patients with liver cirrhosis suggested that the splanchnic circulation was also hyperdynamic (Figure 44.3), and that not only an increase in hepatic resistance, but also an increase in portal blood flow contribute to portal hypertension in cirrhosis [70–72]. This was unequivocally demonstrated in subsequent studies in rodent models of portal hypertension [60], and set the rationale for the use of vasoconstrictors in the treatment of portal hypertension. The hyperdynamic circulatory state in portal hypertension is the consequence of two pathophysiological phenomena: arterial vasodilatation and plasma volume expansion. The presence of both is required for the expression of the hyperdynamic state [51, 71].

The hyperdynamic syndrome and the hepatic artery The hepatic artery, not part of the portal system, also seems to play an important role in the development and maintenance of portal hypertension [73, 74].

44: PATHOGENESIS OF PORTAL HYPERTENSION NORMAL SUBJECT SMA HV

Inj. PATENT WITH LIVER DISEASE SMA HV

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Figure 44.3 Appearance time and mean transit time of [131 I]albumin from the superior mesenteric artery to the hepatic vein (calculated from the isotope dilution curves) in patients with alcoholic liver disease and in normal subjects. Both transit time and appearance time were shorter in alcoholic patients. Based on these data, the existence of an abnormally high flow in the mesenteric bed of patients with alcoholic liver disease was suggested. Reproduced from Kotelanski, Groszmann and Cohn [71] with permission

A hyperdynamic syndrome also develops in the hepatic arterial circulatory bed. This hyperdynamic syndrome is accentuated with the progression of chronic liver diseases. In response to the progressive loss in portal flow, the hepatic artery vasodilates and increases its flow. This response is initiated and perpetuated by a complex interaction of anatomical and functional factors and, contrary to what is observed in the intrahepatic portal system, an excess production of NO seems to play a role in the vasodilatation observed [73].

Increased vasodilatation in portal hypertension At least three mechanisms are thought to contribute to vasodilatation in portal hypertension: (i) increased concentration of systemic vasodilators, (ii) increased endothelial production of local vasodilators, and (iii) decreased vascular responsiveness to endogenous vasoconstrictors. The last mechanism is probably due to the effect of the first two components.

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evidence for a significant role in promoting splanchnic hyperemia in portal hypertension [76]. Many studies have demonstrated that plasma glucagon levels are elevated in cirrhosis. Hyperglucagonism results, in part, from decreased hepatic clearance but, more importantly, from an increased secretion by pancreatic alpha-cells [77]. Support for a role of glucagon in modulating the splanchnic blood flow comes from studies showing that the normalization of the circulating levels of glucagon partially reverses the increased splanchnic blood flow, and this can be prevented by a concomitant glucagon infusion [78]. However, some studies showed no correlation between glucagon levels and splanchnic blood flow, thus questioning a major role of hyperglucagonism in portal hypertension. Glucagon release is clearly implicated in postprandial hyperemia, which, in patients with cirrhosis, is associated with marked increases in portal pressure [79]. Collectively, these data provided the rationale for the use of somatostatin and octreotide in the treatment of patients with portal hypertension [80], although it was recently demonstrated that these drugs promote vasoconstriction by mechanisms independent of those of glucagon inhibition [81].

Endocannabinoids In rats and in patients with advanced cirrhosis, there is an increase in the production of the endogenous cannabinoid anandamide by monocytes [82, 83], and the specific blockade of the peripheral cannabinoid receptor CB1 attenuates the hyperdynamic circulation [82, 83] and decreases portal pressure [82]. In addition, resistance mesenteric arteries from cirrhotic rats exhibit an increased vasodilatory response to anandamide related to an overexpression of CB1 receptor. This was a local mesenteric phenomenon, since it does not occur in other peripheral vessels. It has been postulated that cannabinoids would act through an increase in NO production [82], but recent data do not support this contention [83]. The mechanisms that would induce anandamide production are not clear, but could be related to the frequent endotoxemia observed in cirrhosis [84]. Several other circulating vasodilators, such as calcitonin gene related peptide (CGRP) [85], adrenomedullin [86, 87], and urotensin have also been linked to the pathogenesis of vasodilatation in portal hypertension, but evidence is still scarce.

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Nitric Oxide The implication of NO in portal hypertension was initially suggested by Vallance and Moncada [88]. Several lines of evidence confirmed the central role of NO in

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the development of the hyperdynamic circulation. On the one hand, patients with cirrhosis have increased levels of nitrites and nitrates [89], the degradation products of NO. In experimental animals, it was demonstrated that NO production is increased in the splanchnic vascular bed of portal hypertensive rats, and this accounts for the hyporesponse to vasoconstrictors characteristic of portal hypertension [90]. Furthermore, inhibition of NO production reduces portal pressure and porto-systemic shunting, and prevents (although not completely) the development of the hyperdynamic circulation [91–94]. This latter finding, together with the fact that a double eNOS/inducible nitric oxide synthase (iNOS) knockout mouse still develops the hyperdynamic circulation after the induction of portal hypertension [95], suggests that NO is the principal, but not the only, mediator of vasodilatation. A number of studies have characterized at the molecular level the mechanisms leading to increased NO production in portal hypertension (Figure 44.1). At odds with the original hypothesis, which suggested that endotoxemia present in cirrhosis would up-regulate iNOS [88], overwhelming data suggest that increased NO in portal hypertension is mediated by eNOS [96]. Preliminary data suggest that neuronal nitric oxide synthase (nNOS) activation could also have a role in the increased NO production that occurs in portal hypertension [97], but this role would be far outweighed by that of eNOS [98]. The most powerful stimulus for eNOS up-regulation is shear stress. Indeed, shear stress is increased in portal hypertension once the hyperdynamic circulation is established. Furthermore, the superior mesenteric vascular bed from portal hypertensive rats shows enhanced production of NO in response to shear stress [90] (Figure 44.4). Bacterial translocation also contributes to increased NO production in advanced cirrhosis, but the mechanism involves an up-regulation of eNOS, not iNOS [99, 100]. Finally, porto-systemic shunt, per se, can induce NO-mediated vasodilatation [101]. However, sequential studies in the portal vein ligated model have shown that eNOS activation occurs before any of these three mechanisms are present [102, 103]. This indicates, on the one hand, that increased eNOS production is a primary factor in the development of vasodilatation and, on the other, that different mechanisms from the above-mentioned activate eNOS in the very early phases of portal hypertension. Recent data indicate that the initial eNOS up-regulation occurs at the microcirculation of the intestinal mucosa, and that it is secondary to VEGF up-regulation [104, 105], raising the possibility that the first stimulus that up-regulates eNOS is intestinal hypoxia, secondary to congestion or to superior mesenteric artery reflex vasoconstriction in response to increased portal pressure [103]. In keeping with these findings, it was recently demonstrated that blocking VEGF action from the onset of portal hypertension markedly attenuates the development of the hyperdynamic circulation and decreases portal blood inflow by 50% [62]. Whether

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Figure 44.4 Splanchnic circulation from portal hypertensive rats exhibit increased nitric oxide production in response to shear stress, and this occurs early after the induction of portal hypertension, before the hyperdynamic circulation develops. (a) Pressure response to flow changes in in vitro perfused mesenteric vessel beds of portal vein-ligated (PVL) and sham rats 3 days after PVL. Perfusion pressure in response to increasing flow is significantly smaller in PVL rats. (b) Relationship between amount of nitric oxide (NO) metabolites (NOx ) production and index of shear stress induced by changes in flow rate. Slopes of NOx production rate vs shear stress index were significantly higher in PVL rats than in sham rats. (c) After N ω -nitro-l-arginine (l-NNA) incubation, the pressure response to changes in flow rate in PVL and sham rats was no longer significantly different, demonstrating an NO dependence of observed hyporesponsiveness in PVL rats. Reproduced from Wiest et al. [102] used with permission of The American Physiological Society

Carbon Monoxide CO is an end product of the HO pathway, which seems to play an important role in the regulation of vascular

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(mmHg*min*ml−1*100gbw−1)

these mechanisms account for the development of the hyperdynamic circulation in human cirrhosis needs to be confirmed. When looking at the signaling level, molecular studies have shown that, in the early stages of portal hypertension, increased eNOS activity is detected before the increase in eNOS expression. This is due to activation of eNOS at the post-translational level, mediated by increased Akt-dependent eNOS phosphorylation [103, 106]. In more advanced stages of portal hypertension, NO production increases due both to an increase in eNOS expression [99] and to an increase in eNOS activity related to changes at the post-translational level. This latter mechanism involves an increased interaction of eNOS with the molecular chaperone heat shock protein 90 (Hsp90) [107], and a markedly abnormal subcellular localization of the eNOS molecule [108]. Also, it has been shown that bacterial translocation activates eNOS through a tumor necrosis factor (TNF)-alpha dependent increase in BH4 [99, 100], an essential cofactor of eNOS. Taken together, these studies show that different mechanisms up-regulate eNOS in portal hypertension, and that the relative importance of these mechanisms varies with the evolution of the syndrome. In summary, in striking contrast to what occurs in the intrahepatic circulation, in which there is a deficit in NO production, in the splanchnic circulation there is an increase in NO production [96]. This paradox poses enormous difficulties in developing NO-based therapies for portal hypertension. However, it must be taken into account that the primary defect is the increase in intrahepatic resistance, and splanchnic vasodilatation is a secondary alteration. Furthermore, the severity of the hyperdynamic circulation closely correlates with the resistance to portal blood flow [104] (Figure 44.5). Therefore, it is likely that by improving the increase in hepatic resistance, the hyperkinetic syndrome might be, at least in part, reverted [22].

PP (mmHg)

44: PATHOGENESIS OF PORTAL HYPERTENSION

Figure 44.5 Portal vein ligation surgery performed over different needles of increasing caliber [16-gauge (16G), 18G, and 20G] yields rats with different degrees of portal pressure (PP). The degree of portal hypertension is proportional to the degree of systemic circulatory abnormalities: mean arterial pressure (MAP), cardiac index (CI), and systemic vascular resistance (SVR). Modified from Abraldes et al. [104] used with permission of The American Physiological Society

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resistance in several vascular beds, including mesenteric arteries [112]. CO is generated in endothelial and smooth muscle layers of arterial vessels and, similarly to NO, induces vasodilatation through stimulation of soluble guanylyl cyclase (sGC) in vascular smooth muscle cells. Several investigations have demonstrated that the inducible isoform of HO (HO-1) is up-regulated in the systemic and splanchnic circulation of portal hypertensive animals, contributing to vasodilatation and the hyperdynamic circulatory state [113, 114].

Plasma volume expansion For many years, plasma volume expansion has been recognized in a wide variety of portal hypertensive liver diseases. In conditions of constant peripheral vascular resistance, an increase in circulatory blood volume results in increased venous return and cardiac output. However, expansion of blood volume leads to stress relaxation of the vasculature, and after this initial increase cardiac output returns to normal. This demonstrates that blood volume expansion alone is not sufficient in itself to maintain a hyperkinetic circulatory state. It is the combination of arterial vasodilatation and blood volume expansion that produces optimal conditions for maintaining the hyperdynamic circulatory state in portal hypertension. Sodium retention is the earliest and most frequent abnormality of renal function associated with portal hypertension [115]. Sodium restriction markedly attenuates the development of the hyperdynamic circulation in portal hypertensive rats, normalizing cardiac output and portal venous inflow [116]. On the other hand, it has been demonstrated that diuretics such as spironolactone decrease (although mildly) portal pressure [117]. The initial stimulus for sodium retention is arterial vasodilatation [118, 119], which leads to arterial underfilling and subsequent activation of baroreceptors. This leads to the activation of the sympathetic nervous system, RAS, and vasopressin secretion. The final result is sodium and water retention by the kidneys, and plasma volume expansion (Figure 44.6). This has been called the peripheral arterial vasodilatation hypothesis [51], and explains not only the pathophysiology of the hyperdynamic circulation in portal hypertension, but also many aspects of other complications of portal hypertension such as ascites and hepatorenal syndrome [69].

CONCLUSION The primary event leading to portal hypertension in liver cirrhosis is increased hepatic resistance. This is due not only to the architectural disturbances in liver vasculature associated with the cirrhotic process, but also to an increased intrahepatic vascular tone that results from an imbalance between excessive vasoconstrictors and deficient

Functional

Structural Varices

Increased resistance Porto-systemic collaterals PORTAL HYPERTENSION

Vasodilatation

Hypotension Central hypovolemia

Activation NE, VP, A-II

Increased flow

Increase in CO

Na and water retention increase venous return to the heart

Figure 44.6 Summary of the pathophysiology of portal hypertension. The increase in hepatic resistance leads to an increase in portal pressure. This leads to a cascade of disturbances in the splanchnic and systemic circulation characterized by vasodilatation, sodium and water retention, and plasma volume expansion, which are major players in the pathogenesis of ascites and hepato-renal syndrome. Additionally, these alterations lead to an increase in portal blood inflow that contributes to maintaining and aggravating portal hypertension. Another characteristic feature is the development of porto-systemic collaterals, which are responsible for complications such as variceal bleeding and hepatic encephalopathy. CO, cardiac output; NE, norepinephrine; VP, vasopressin; A-II, angiotensin II; Na, sodium

vasodilators. Further, portal hypertension induces marked alterations in the systemic and splanchnic circulation, characterized by a decrease in systemic vascular resistance, arterial hypotension, increased cardiac output and plasma volume expansion, and known as the hyperdynamic circulatory state. This leads to an increase in portal blood inflow that contributes to maintaining and aggravating portal hypertension (Figure 44.6).

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80. Abraldes, J.G. and Bosch, J. (2002) Somatostatin and analogues in portal hypertension. Hepatology, 35 (6), 1305–12. 81. Wiest, R., Tsai, M.H. and Groszmann, R.J. (2001) Octreotide potentiates PKC-dependent vasoconstrictors in portal-hypertensive and control rats. Gastroenterology, 120 (4), 975–83. 82. Batkai, S., Jarai, Z., Wagner, J.A., Goparaju, S.K., Varga, K., Liu, J. et al. (2001) Endocannabinoids acting at vascular CB1 receptors mediate the vasodilated state in advanced liver cirrhosis. Nat Med , 7 (7), 827–32. 83. Ros, J., Claria, J., To-Figueras, J., Planaguma, A., Cejudo-Martin, P., Fernandez-Varo, G. et al. (2002) Endogenous cannabinoids: a new system involved in the homeostasis of arterial pressure in experimental cirrhosis in the rat. Gastroenterology, 122 (1), 85–93. 84. Varga, K., Wagner, J.A., Bridgen, D.T. and Kunos, G. (1998) Platelet- and macrophage-derived endogenous cannabinoids are involved in endotoxin-induced hypotension. FASEB J , 12 (11), 1035–44. 85. Hori, N., Okanoue, T., Sawa, Y. and Kashima, K. (1997) Role of calcitonin gene-related peptide in the vascular system on the development of the hyperdynamic circulation in conscious cirrhotic rats. J Hepatol , 26, 1111–19. 86. Fernandez-Rodriguez, C.M., Prada, I.R., Prieto, J., Montuenga, L.M., Elssasser, T., Quiroga, J. et al. (1998) Circulating adrenomedullin in cirrhosis: relationship to hyperdynamic circulation. J Hepatol , 29 (2), 250–56. 87. Genesca, J., Gonzalez, A., Catalan, R., Segura, R., Martinez, M., Esteban, R. et al. (1999) Adrenomedullin, a vasodilator peptide implicated in hemodynamic alterations of liver cirrhosis: relationship to nitric oxide. Dig Dis Sci , 44 (2), 372–76. 88. Vallance, P. and Moncada, S. (1991) Hyperdynamic circulation in cirrhosis: a role for nitric oxide? Lancet , 337, 776. 89. Guarner, C., Soriano, G., Tomas, A., Bulbena, O., Novella, M.T., Balanzo, J. et al. (1993) Increased serum nitrite and nitrate levels in patients with cirrhosis: relationship to endotoxemia. Hepatology, 18, 1139–43. 90. Hori, N., Wiest, R. and Groszmann, R.J. (1998) Enhanced release of nitric oxide in response to changes in flow and shear stress in the superior mesenteric arteries of portal hypertensive rats. Hepatology, 28, 1467–73. 91. Lee, F.Y., Colombato, L.A., Albillos, A. and Groszmann, R.J. (1993) Administration of N -omeganitro-L-arginine ameliorates portal–systemic shunting in portal-hypertensive rats. Gastroenterology, 105 (5), 1464–70. 92. Pizcueta, P., Piqu´e, J.M., Fern´andez, M., Bosch, J., Rod´es, J., Whittle, B.J.R. et al. (1992) Modulation of the hyperdynamic circulation of cirrhotic rats by nitric oxide inhibition. Gastroenterology, 103, 1909–15. 93. Pizcueta, M.P., Piqu´e, J.M., Bosch, J., Whittle, B.J.R. and Moncada, S. (1992) Effects of inhibiting nitric oxide biosynthesis on the systemic and splanchnic circulation of rats with portal hypertension. Br J Pharmacol , 105, 105–184.

94. Garc´ıa-Pag´an, J.C., Fernandez, M., Bernadich, C., Pizcueta, P., Piqu´e, J.M., Bosch, J. et al. (1994) Effects of continued nitric oxide inhibition on the development of the portal hypertensive syndrome following portal vein stenosis in the rat. Am J Physiol , 30, 984–90. 95. Iwakiri, Y., Cadelina, G., Sessa, W.C. and Groszmann, R.J. (2002) Mice with targeted deletion of eNOS develop hyperdynamic circulation associated with portal hypertension. Am J Physiol Gastrointest Liver Physiol , 283 (5), G1074–81. 96. Wiest, R. and Groszmann, R.J. (2002) The paradox of nitric oxide in cirrhosis and portal hypertension: too much, not enough. Hepatology, 35 (2), 478–91. 97. Jurzik, L., Froh, M., Straub, R.H., Scholmerich, J. and Wiest, R. (2005) Up-regulation of nNOS and associated increase in nitrergic vasodilation in superior mesenteric arteries in pre-hepatic portal hypertension. J Hepatol , 43 (2), 258–65. 98. Kwon, S.Y., Groszmann, R.J. and Iwakiri, Y. (2007) Increased neuronal nitric oxide synthase interaction with soluble guanylate cyclase contributes to the splanchnic arterial vasodilation in portal hypertensive rats. Hepatol Res, 37 (1), 58–67. 99. Wiest, R., Das, S., Cadelina, G., Garcia-Tsao, G., Milstien, S. and Groszmann, R.J. (1999) Bacterial translocation in cirrhotic rats stimulates eNOS-derived NO production and impairs mesenteric vascular contractility. J Clin Invest , 104 (9), 1223–33. 100. Wiest, R., Cadelina, G., Milstien, S., McCuskey, R.S., Garcia-Tsao, G. and Groszmann, R.J. (2003) Bacterial translocation up-regulates GTP-cyclohydrolase I in mesenteric vasculature of cirrhotic rats. Hepatology, 38 (6), 1508–15. 101. Bernadich, C., Bandi, J.C., Piera, C., Bosch, J. and Rodes, J. (1997) Circulatory effects of graded diversion of portal blood flow to the systemic circulation in rats: role of nitric oxide. Hepatology, 26 (2), 262–67. 102. Wiest, R., Shah, V., Sessa, W.C. and Groszmann, R.J. (1999) NO overproduction by eNOS precedes hyperdynamic splanchnic circulation in portal hypertensive rats. Am J Physiol , 276 (4 Pt 1), G1043–51. 103. Tsai, M.H., Iwakiri, Y., Cadelina, G., Sessa, W.C. and Groszmann, R.J. (2003) Mesenteric vasoconstriction triggers nitric oxide overproduction in the superior mesenteric artery of portal hypertensive rats. Gastroenterology, 125 (5), 1452–61. 104. Abraldes, J.G., Iwakiri, Y., Loureiro-Silva, M., Haq, O., Sessa, W.C. and Groszmann, R.J. (2006) Mild increases in portal pressure up-regulate vascular endothelial growth factor and endothelial nitric oxide synthase in the intestinal microcirculatory bed, leading to a hyperdynamic state. Am J Physiol Gastrointest Liver Physiol , 290 (5), G980–87. 105. Iwakiri, Y., Abraldes, J.G., Haq, O., Supatsri, S., Loureiro-Silva, M.R. and Groszmann, R.J. (2007) Portal hypertension triggers peripheral signals that induce vascular endothelial growth factor (VEGF) and endothelial nitric oxide synthase (eNOS) expression at the intestinal microcirculation in cirrhotic rats. Hepatology, 46, 608A.

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106. Iwakiri, Y., Tsai, M.H., McCabe, T.J., Gratton, J.P., Fulton, D., Groszmann, R.J. et al. (2002) Phosphorylation of eNOS initiates excessive NO production in early phases of portal hypertension. Am J Physiol Heart Circ Physiol , 282 (6), H2084–90. 107. Shah, V., Wiest, R., Garcia-Cardena, G., Cadelina, G., Groszmann, R.J. and Sessa, W.C. (1999) Hsp90 regulation of endothelial nitric oxide synthase contributes to vascular control in portal hypertension. Am J Physiol , 277 (2 Pt 1), G463–68. 108. Iwakiri, Y., Murata, T., Gao, H., Loureiro-Silva, M.R., Sessa, W.C. and Groszmann, R.J. (2006) Altered intracellular eNOS localization leads to an excessive NO production in the splanchnic arterial circulation in cirrhotic rats with portal hypertension. Hepatology, 44 (Suppl 1), 354A. 109. Guarner, C., Soriano, G., Such, J., Teixido, M., Ramis, I., Bulbena, O. et al. (1992) Systemic prostacyclin in cirrhotic patients. Relationship with portal hypertension and changes after intestinal decontamination. Gastroenterology, 102 (1), 303–9. 110. Potenza, M.A., Botrugno, O.A., De Salvia, M.A., Lerro, G., Nacci, C., Marasciulo, F.L. et al. (2002) Endothelial COX-1 and -2 differentially affect reactivity of MVB in portal hypertensive rats. Am J Physiol Gastrointest Liver Physiol , 283 (3), G587–94. 111. Fernandez, M., Garcia-Pagan, J.C., Casadevall, M., Mourelle, M., Pique, J.M., Bosch, J. et al. (1996) Acute and chronic cyclooxygenase blockade in portal hypertensive rats. Influence on nitric oxide biosynthesis. Gastroenterology, 110, 1529–35. 112. Naik, J.S., O’ Donaughy, T.L. and Walker, B.R. (2003) Endogenous carbon monoxide is an endothelial-derived

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45

Non-alcoholic Fatty Liver Disease: A Pathophysiological Perspective Michael Fuchs and Arun J. Sanyal Division of Gastroenterology, Hepatology and Nutrition, Department of Internal Medicine, Virginia Commonwealth University School of Medicine, Richmond, VA, USA

INTRODUCTION

NAFLD AND OBESITY: AN EVOLUTIONARY PERSPECTIVE

Non-alcoholic fatty liver disease (NAFLD) is the most common cause of chronic liver disease in North America. The hepatic lesions associated with this condition extend from isolated fatty liver to steatohepatitis. Whereas full-blown steatohepatitis includes evidence of cytological ballooning, inflammation, and pericellular fibrosis, variants are commonly seen with only some of the features of this lesion. This spectrum of disease was originally felt to represent alcoholic liver disease. It is now appreciated, however, that it often occurs in those who consume little or no alcohol; in such cases, the prefix non-alcoholic is added. NAFLD affects about one-third of the general population and is an important risk factor for the development of diabetes and vascular disease. Non-alcoholic steatohepatitis (NASH) can also progress to cirrhosis and liver cancer. It is therefore a major public health problem in the Western world. In this chapter, we discuss the clinical spectrum and metabolic, cellular, and molecular basis for the disease and present a pathophysiology-driven approach to the management of the condition.

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

Reproduction, physical activity, and food supply are some of the basic necessities to ensure the survival of human beings. Prior to the development of agriculture, human ancestors were primarily hunter–gatherers. Since the outcome of a hunt was never guaranteed, there were periods of abundance of food interspersed with periods of lack of food. In order to ensure survival of the species, there was therefore a need for an “energy bank” where excess calories could be stored during periods of food abundance and from which calories could be withdrawn. This function was served by adipose tissue, since fat provided the most efficient way to store excess energy. As the human body adapted for a situation in which food was limited yet physical exertion was required for survival, “thrifty” genes evolved to promote caloric utilization and storage in the form of adipose tissue for energy needs and survival during extended periods of starvation [1]. Insulin resistance may have evolved as an important survival mechanism to preserve protein stores in times of starvation. Under such conditions, fatty acids increased

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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due to lipolysis. Although this increase in fatty acids resulted in reduced insulin-stimulated glucose uptake in muscle, other insulin signal pathways that regulate protein metabolism and other insulin-regulated processes could remain unaffected. In this way, the body preserved circulating glucose for the central nervous system and other obligate glucose-requiring organs while preserving protein stores, both of which are essential for survival. In modern times, industrialization and technological advances have led to a state of over-abundance of food, especially in the economically developed nations. This, coupled with a sedentary lifestyle, has led to an imbalance between long-term caloric intake versus expenditure. Of note, although the absolute caloric intake of modern humans is lower compared with our ancient ancestors [2], it is nevertheless high relative to the corresponding larger decrease in caloric expenditure via physical activity. As a consequence, the “thrifty” genotype that evolved for function, selective advantage, and survival in the Late-Paleolithic era is now being exposed to sedentary lifestyles, fat-rich and fiber-poor diets, positive caloric imbalance, and an extended life span, all of which have resulted in a disruption of the homeostatic mechanisms optimized during evolution. Therefore, unabated storage of fuel without the stimulus for its utilization has led to discordance in gene–environmental interactions predisposing the paleolithically programmed genome to misexpress its genes in multiple organ systems, ultimately resulting in our epidemics of obesity, diabetes mellitus, and fatty liver disease.

THE CLINICAL SPECTRUM OF NAFLD NAFLD affects up to 60 million Americans [3]. A recent study in adults employing magnetic resonance spectroscopy showed a prevalence of fatty liver in 34% [4]. This is several-fold higher than other common chronic liver diseases such as hepatitis C and alcohol-related liver disease. Importantly, 13% of children have fatty liver disease [5]. NAFLD is more common in those of Hispanic origin and African Americans appear to be relatively protected [6, 7]. Although the majority of subjects with NAFLD are obese, the condition can occur in the absence of obesity or other features of the metabolic syndrome. In patients with diabetes and morbid obesity, the prevalence of NAFLD has been shown to be as high as 62 and 96%, respectively [8, 9]. Because of the rapidly increasing prevalence of obesity and diabetes, not only in adults but also in children, NAFLD will reach epidemic proportions, contributing substantially to the burden of chronic liver disease in the coming decades [10].

NASH represents an independent cardiovascular risk factor [11], and in patients with diabetes it is an independent predictor of cardiovascular mortality [12]. Over a period of 10–15 years, 15% of patients with NASH will progress to liver cirrhosis [13]. The development of cirrhosis is associated with a loss of hepatic fat and other pathological features of NASH [14]; such patients are often considered to have cryptogenic cirrhosis. Once cirrhosis has developed, hepatic decompensation occurs at a rate of 4% annually and the 10 year risk of developing liver cancer is 10% [13, 15]. It is therefore not surprising that NAFLD is accounting for an increasing number of liver transplantations. Like other causes of chronic liver disease, NASH recurs following liver transplantation almost universally [16]. NASH is also associated with an excess of cardiovascular morbidity and mortality and also extrahepatic cancer-related deaths [17].

CELL AND MOLECULAR BIOLOGY OF NAFLD A hallmark that drives development of fatty liver is insulin resistance [18, 19]. The fundamental defect in insulin resistance is impaired insulin-mediated suppression of lipolysis, resulting in increased fatty acid release into the circulation. Before one can appreciate the mechanism of insulin resistance and how it leads to NAFLD, it is important to understand the normal effects of insulin. Insulin influences essential biochemical processes involved in carbohydrate and fat metabolism. By increasing the expression of glucose transporters, insulin promotes glucose uptake by cells. It also inhibits glycogenolysis, promotes glycogen synthesis, and inhibits gluconeogenesis. By promoting the synthesis of triglycerides and inhibiting lipolysis, insulin is also an important modulator of the formation and storage of lipids [20]. Insulin resistance is characterized by impaired glucose uptake into skeletal muscle and adipose tissue [21]. Also, insulin-mediated lipogenesis results in a net increase in free fatty acid release from adipose tissue. In the liver, hyperinsulinemia manifests with unrestrained hepatic glucose production resulting from impaired glycogen synthesis and failure to suppress gluconeogenesis. Initially, it was proposed that once fatty liver has developed, an additional cellular event promotes progression to inflammation, apoptosis, and fibrosis [22]. It is now clear that multiple mechanisms are concurrently operative to produce cell injury, apoptosis, activation of inflammation, and fibrosis to produce the phenotype of steatohepatitis. Below, we summarize our current understanding of NAFLD and highlight the achievements made since the first description of this disease [23].

45: NON-ALCOHOLIC FATTY LIVER DISEASE: A PATHOPHYSIOLOGICAL PERSPECTIVE

Molecular events leading to hepatic fat accumulation The role of gut microbiota The gut microbiota is acquired at birth and throughout life it undergoes a process of re-colonization dependent on several factors, among which are diet, antibiotic treatment, hygiene, and infections. Interestingly, the major part of the microbiota is present in the large bowel, where food products escaped digestion in the upper part of the gastrointestinal tract. The human intestinal microbiota is composed of 1013 –1014 microorganisms whose collective genome contains at least 100 times more genes than our own genome [24]. Microorganisms are composed of 400–500 phylotypes largely belonging to 30–40 dominant bacterial species. From an evolutionary standpoint, selection pressure likely altered the microbiota composition, favoring the evolution of new strains with different biological properties mainly to promote more efficient extraction and storage of energy from a given diet [25]. Bacteroidetes and Firmicutes may influence energy balance and fat storage [25–27]. In addition, changing dietary habits to a high-fat and low-fiber diet also appears to modulate the Adipose tissue

TG storage

721

gut microbiota. This appears to affect the fecal contents of lipids, further supporting a role of gut microbiota for energy harvesting [28]. Fat storage and development of fatty liver may be promoted by gut microbiota both by promoting insulin resistance and by insulin resistance-independent mechanisms (Figure 45.1): first, microbial fermentation of non-digestible polysaccharides to monosaccharides and short-chain fatty acids and their subsequent absorption stimulate de novo hepatic triglyceride synthesis through their effects on two basic helix–loop–helix/leucine zipper transcription factors – carbohydrate-responsive element binding protein (ChREBP) and sterol regulatory element binding protein 1 (SREBP1) [29, 30]; second, selective suppression of intestinal epithelial expression and secretion of angiopoietin-like protein 4, also known as fasting-induced adipose factor [29], results in increased lipoprotein lipase activity in adipose tissue [31]. This in turn facilitates triglyceride lipolysis and release of free fatty acids that are taken up by adipose tissue, thereby promoting storage of liver-derived triglycerides and the development of insulin resistance. This suppression of angiopoietin-like protein 4 also appears to occur in response to a high-fat diet [32]; third, during a high-fat diet, the gut microbial metabolism may convert choline Liver

TG synthesis ChREBP/SREBP

VLDL PC

Lipogenic substrates

Choline

Kidney LPL

ANGPTL4 Gut epithelium

Gut microbiota Choline

Methylamines

Figure 45.1 Energy harvest and storage: the role of the intestine and gut microbiota. The microbiota may act to stimulate hepatic lipogenesis through effects mediated by transcription factors such as carbohydrate-responsive element-binding protein (ChREBP) and sterol regulatory element binding protein 1 (SREBP1) and to promote lipoprotein lipase (LPL)-directed incorporation of these triglycerides into adipocytes through suppression of intestinal epithelial production of the circulating LPL inhibitor angiopoietin-like protein 4 (ANGPTL4). The gut microbial metabolism may convert choline into methylamines with subsequent urinary excretion, thereby reducing the availability of choline to synthesize enough phosphatidylcholine necessary for the assembly and secretion of very low density lipoprotein (VLDL)

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into methylamines with subsequent urinary excretion [33], thereby mimicking a choline-deficient diet known to cause NAFLD. The reduced availability of choline likely results in an inability to synthesize enough phosphatidylcholine [34] necessary for the assembly and secretion of very low density lipoprotein (VLDL), with subsequent hepatic accumulation of triglycerides [35]. The intestinal microbiota has also been implicated in the genesis of the insulin-resistant state. Recently, plasma levels of endotoxin were shown to increase two- to threefold in response to alterations in the gut microbiota and/or a high-fat diet [36, 37]. This is believed to be due to a local intestinal inflammatory response to ingress of bacterial products from the lumen due to altered intestinal permeability. Intestinal permeability is increased due to decreased expression of intercellular tight junction proteins in the epithelium [28]. The activation of the Toll like receptor 4 (TLR4) plays a central role in this process. This receptor is expressed on many cell types and represents a part of the innate immune system. It recognizes endotoxin and is also activated by fatty acids [38]. Activation of TLR4 directly activates IκB-kinase (IKK), leading to serine phosphorylation of insulin receptor substrate (IRS) proteins and impaired insulin signaling and action (Figure 45.2). Signaling through TLR4 and IKK activates the nuclear factor κB (NF-κB) pathway, leading to the local production of inflammatory cytokines such

as interleukin 6 (IL-6) and tumor necrosis factor alpha (TNF-α) [39, 40] that indirectly attenuate insulin action. IL-6-induced expression of suppressor of cytokine signaling 3 (SOCS-3) impairs insulin signaling directly by competing for phosphorylation sites on IRS proteins or by their ubiquitin-mediated degradation [41]. TNF-α is a potent activator of c-jun N-terminal kinase (JNK) that promotes the phosphorylation of IRS, thereby negatively regulating normal insulin receptor signaling [42]. Activation of these pathways in adipose tissue and liver is in line with increased gene expression of TNF-α and its receptor in patients with NAFLD [43]. Collectively, these findings implicate TRL4 as a gatekeeper by which endotoxin and fatty acids induce inflammation, leading to insulin resistance and the development of NAFLD under conditions of nutrient excess. This concept is supported by the observation that high-fat diet-induced insulin resistance and subsequent development of NAFLD may be prevented or reversed by either reducing circulating endotoxin levels [44] or by modulation of TLR4 [45].

Adipose tissue response to energy excess Dietary fat is absorbed from the intestine in the form of chylomicrons. Endothelial lipoprotein lipase in adipose tissue hydrolyzes triglycerides, releasing free fatty acids

FA/LPS

IL-6 IR

TLR4

IL-6R

TNF TNF-R

IRS

IRS

Ser-P

Ser-P

STAT3 PI3K

IR

JNK JNK

IKK IKK SOCS3 SOCS3 IL-6 TNF

AKT2 NF- B

Figure 45.2 Mechanisms by which the intestine promotes insulin resistance. Binding of dietary fatty acids and endotoxin to the Toll-like receptor 4 (TLR4) activates IκB-kinase (IKK), which causes degradation of IκBα and stimulates nuclear translocation of nuclear factor κB (NF-κB). This activates the expression of interleukin 6 (IL-6) and tumor necrosis factor α (TNF-α). Activated IKK directly increases serine phosphorylation of insulin receptor substrate (IRS) that down-regulates insulin signaling associated with IRS tyrosine phosphorylation, phosphatidylinositol-3,4,5-triphosphate kinase (PI3K) and AKT2, a member of the serine/threonine-specific protein kinase family, resulting in insulin resistance. Production of IL-6 and TNF-α indirectly causes down-regulation of insulin signaling via suppressor of cytokine signaling 3 (SOCS3) and c-jun N-terminal kinase (JNK). IR, insulin receptor; IL-6R, interleukin 6 receptor; TNF-R, tumor necrosis factor α receptor

45: NON-ALCOHOLIC FATTY LIVER DISEASE: A PATHOPHYSIOLOGICAL PERSPECTIVE

hepatic inflammation and fibrosis in patients with NAFLD [53], likely due to the greater amount of TNF-α and IL-6 produced compared with subcutaneous fat and expression of 11β-hydroxylase the rate-limiting step in steroidogenesis. Both adipocyte size and total body weight are strong predictors of the number of mature macrophages found within adipose tissue and the amount of TNF-α and IL-6 produced by macrophages [50, 54]. This inflammatory response then appears to promote insulin resistance (Figure 45.2). The major consequence of macrophage infiltration in adipose tissue is the activation of adipocytes that then produce a pro-inflammatory mix of cytokines characterized by high TNF-α and low adiponectin levels. These are released in to the circulation and sensed by the liver, where additional cytokines, for example C-reactive protein, is released as part of the acute phase reaction. These cytokines promote endothelial dysfunction, promote atherogenesis and thrombosis, impair fibrinolysis, and impair the metabolic clearance of glucose (insulin resistance). The increasing size of adipocytes during weight gain is closely related to a decrease in adiponectin expression [55]. Thus adiponectin is the only protein specific to adipose tissue the level of which is reduced in patients with NAFLD [56]. Moreover, serum adiponectin levels are inversely related to hepatic fat stores and correlate positively with insulin resistance [56]. The fact that insulin, TNF-α and IL-6 modulate gene expression and circulating levels of adiponectin [57] is in agreement with the concept of adipose tissue energy status and production of reactive

for uptake, re-esterification to triglycerides, and storage as fat (Figure 45.3). Excess carbohydrates and hyperinsulinemia also activate ChREBP, promoting de novo lipogenesis from carbohydrates in adipocytes. Activation of hormone-sensitive lipase and adipose tissue triglyceride lipase activate lipolysis and release of free fatty acids [46, 47]. This process is also promoted by activation of the β3 adrenergic sympathetic neurons [48]. Weight gain and lipid storage are first associated with expansion of the size and number of adipocytes as adipose tissue has a much greater innate lipid storage capacity than other tissues such as muscle and liver. There is an organ-specific hierarchy for safe lipid storage; peripheral/subcutaneous adipose tissue is preferable to central/visceral adipose tissue that in turn is preferable to liver. When lipid accumulation continues and finally exceeds the storage capacity, cellular and ultimately adipocyte dysfunction ensues. A key consequence of adipose tissue expansion is the infiltration of adipose tissue by macrophages [49, 50]. This is believed to be promoted by hypoxia inducible factor-1 and expression of the C-C motif chemokine ligand 2 (CCL-2) within adipose tissue. Elevated CCL-2 serum levels and increased hepatic expression of its receptor found in patients with NAFLD support their role in the disease process [51, 52]. Leptin inhibits this process. These changes are notably most marked within visceral fat [53], the fat deposit with the greatest metabolic risk, and seem to precede other features of the metabolic syndrome. It is noteworthy in this context that visceral fat is an independent predictor of s.c. adipose tissue

visceral adipose tissue

TG TG TG

TG

723

TG

TG

Lipolysis

TG TG

Liver TG TG

Lipogenesis

Esterification FA VLDL

β-oxidation

CM remnants

Gut

Figure 45.3 Sources of free fatty acids reaching the liver and their hepatic processing. Free fatty acids (FAs) derived from subcutaneous and visceral triglycerides (TGs) stores by lipolysis contribute to the hepatic free fatty acid pool. Dietary fatty acids reach the liver via intestinal processing and formation of chylomicron (CM) remnants. The size of the hepatic free fatty acid pool is modified by lipogenesis, degradation of fatty acids via β-oxidation, and esterification of fatty acids for storage as triglycerides or secretion as very low density lipoprotein (VLDL)

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oxygen species (ROS) in particular determining the extent of adiponectin release [58]. Concomitant with lower circulating adiponectin levels, TNF-α levels are significantly higher in patients with NASH compared with controls [56, 59], a finding that may be related to the ability of adiponectin to reduce TNF-α production in adipose tissue. During weight gain and visceral fat accumulation, increased secretion of retinol binding protein 4 (RBP4) directly impairs insulin signaling in adipocytes and induces hepatic expression of phosphoenolpyruvate carboxykinase, thereby reducing insulin action to suppress glucose production [60]. Increased serum levels of RBP4 have been found in patients with NAFLD [61] and in subjects with subclinically increased liver fat [62]. The close inverse association between RBP4 and adiponectin in patients with NAFLD further favors the concept that the degree of insulin sensitivity depends on the balance between TNF-α, IL-6, and RBP4 on the one hand and adiponectin on the other. More recently, decreased expression of six-transmembrane protein of prostate 2 (STAMP2), present in high abundance in adipocytes but also at lower at levels in liver, has been linked to the development of NAFLD [63]. This protein likely integrates nutrient and inflammatory signals with metabolic pathways as it is induced by fatty acids and TNF-α and appears to be required for normal insulin signaling, including tyrosine phosphorylation of both the insulin receptor and Akt

kinase. In fact, a reduced expression of STAMP2 leads to increased production of inflammatory cytokines such as IL-6 and CCL-2 [63]. The findings of both protection against excessive inflammation and up-regulation under inflammatory and nutrient-rich conditions may appear paradoxical. However, STAMP2 may act in a regulatory role, not to block activation of inflammatory pathways, but to restrict the degree of their activity. This is exemplified in the regulation of STAMP2 by TNF-α that induces both STAMP2 and inflammatory cytokine expression; however, in the absence of STAMP2, the ability of TNF-α to promote IL-6 expression is more potent [63]. Future studies will demonstrate whether such a mechanism is operational in fatty liver disease in humans.

Energy homeostasis in the liver and genesis of hepatic steatosis A key function of the liver is to integrate carbohydrate and lipid metabolic pathways to maintain energy homeostasis (Figure 45.4). Glucose is derived from uptake from portal blood, glycogenolysis, and gluconeogenesis. It undergoes glycolysis that provides glyceraldehyde 3-phosphate, a key intermediate required for triglyceride synthesis. Pyruvate generated from the glycolytic pathway enters the Krebs cycle. Under conditions of carbohydrate excess,

Citrate Acetyl-CoA SREBP ChREBP

CPT1

ACC Malonyl-CoA FAS LCE SCD1 FFA GPAT DAG

Glucose

Pyruvate Krebs cycle

ACL

L-PK ChREBP

-oxidation

TG

Figure 45.4 Metabolic pathways contributing to hepatic triglyceride synthesis. Triglyceride synthesis is nutritionally regulated and involves induction of several key metabolic pathways including glycolysis, lipogenesis, and β-oxidation. Several of the participating key enzymes are induced by sterol regulatory element binding protein 1 (SREBP1) and carbohydrate-responsive element-binding protein (ChREBP), respectively. The final step whereby glucose is converted to pyruvate involves liver-type pyruvate kinase (LPK). Pyruvate then enters the Krebs cycle to generate citrate, the principal source of acetyl-CoA used for fatty acid synthesis. Fatty acids then undergo elongation and desaturation steps before triglycerides are formed. Increased fatty acid synthesis results in elevated levels of malonyl-CoA, which inhibit carnitine palmitoyltransferase 1 (CPT1), representing the rate-limiting step of β-oxidation by facilitating the transport of fatty acids into mitochondria. ACL, ATP citrate lyase; ACC, acetyl-CoA carboxylase; FAS, fatty acid synthase; LCE, long-chain fatty acyl elongase; SCD1, stearoyl-CoA desaturase 1; GPAT, glycerol-3-phosphate acyltransferase; DGAT, diacylglycerol acyltransferase

45: NON-ALCOHOLIC FATTY LIVER DISEASE: A PATHOPHYSIOLOGICAL PERSPECTIVE

acetyl-CoA generated from pyruvate and citrate, a Krebs cycle intermediate, is used for fatty acid and eventually triglyceride synthesis. Similarly, excess dietary fatty acids or plasma fatty acids derived from lipolysis are taken up and converted to triglycerides. These are then packaged in VLDLs and secreted from the liver, a process that required microsomal transfer protein to bring the triglyceride molecules into a lipoprotein particle with apolipoprotein B 100 [64]. Fatty acids may also be used for oxidation in mitochondria and peroxisomes. Several nuclear receptors and transcription factors acting in a complex crosstalk have been identified that are in control of lipid and glucose homeostasis (Figure 45.5). The expression of key enzymes of lipogenesis is stimulated by insulin and glucose at the transcriptional level via SREBP1 and ChREBP, respectively [65]. Insulin activates the liver X receptor (LXR), which in turn increases the transcription of SREBP1 [66]. Interestingly, ChREBP was recently identified as direct target for LXR [67, 68], establishing LXR as master lipogenic transcription factor enhancing hepatic fatty acid synthesis. As shown for oxysterols, glucose effectively may bind and activate LXR [68]. The integration of glucose sensing and control of lipogenesis in a single protein may provide an explanation for the observation that a low-fat, high-carbohydrate diet induces hypertriglyceridemia [69]; LXR can sense surplus glucose, can induce fatty acid synthesis, and prompt hepatic export of VLDL [70]. Continued activity of fatty acid transport protein 5, highly expressed in liver, is required to sustain caloric uptake and fatty acid flux into the liver [71]. Adiponectin

FA

725

When hepatic fatty acid levels increase, peroxisome proliferator-activated receptor α (PPARα) and peroxisome proliferator-activated receptor γ co-activator 1 (PPARGC1) are activated and promote fatty acid oxidation and production of VLDL, the net effect being catabolism and clearance of fatty acids [72]. PPARα also directly interacts with PPARGC1. This interaction in conjunction with forkhead box protein a2 (Foxa2), also known as hepatic nuclear factor 3α, cooperatively stimulates fatty acid oxidation and VLDL production [72]. Because insulin resistance results in disrupted phosphorylation of Foxa2 with dissociation from PPARGC 1 [73], this pathway may contribute to the development of NAFLD. Transient commitment of cells to either lipid synthesis or lipid disposal is further regulated by the interaction of PPARα with LXR in a reciprocally inhibitory manner, so that the two factors can execute their opposite functions on lipid metabolism. Moreover, PPARGC1 increases SREBP1 and LXR-dependent transcriptional activity, thereby coupling lipid synthesis and lipoprotein secretion [69]. The key lipogenic enzymes and SREBP appear to be under additional control by microRNAs, for example, mir 122 [74]. Fatty acid oxidation is also promoted by adiponectin. This effect of adiponectin can be mediated either via activation of PPARα [75] or AMPK, adenosine 5′ -monophosphate-activated protein kinase [57]. AMPK decreases gluconeogenesis and fatty acid synthesis while increasing fatty acid oxidation. This cumulative result is achieved through three independent mechanisms that include the suppression of SREBP1 and ChREBP,

Cholesterol/Glucose

FA

AMPK AMPK

PPAR PPARGC1

LXR PPARGC1

SREBP1

Insulin

PPARGC1

Foxa2

ChREBP

FA oxidation

FA synthesis

AMPK VLDL

Glucose

Figure 45.5 Balancing oxidation and synthesis of fatty acids. Fatty acid (FA) oxidation is promoted by peroxisome proliferator-activated receptor α (PPARα) and adenosine 5′ -monophosphate-activated protein kinase (AMPK), both being activated by adiponectin. Fatty acid synthesis is induced by insulin, glucose, and cholesterol via the transcription factors sterol regulatory element binding protein 1 (SREBP1), carbohydrate regulatory element binding protein (ChREBP), and liver X receptor (LXR), respectively. PPARα and LXR form heterodimers with the same retinoid X receptor, allowing the hepatocyte to commit transiently to either fatty acid oxidation or fatty acid synthesis. This may require co-activators such as peroxisome proliferator-activated receptor γ co-activator 1 (PPARGC1) or forkhead box protein a2 (Foxa2)

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respectively [76–78], and inhibition of acetyl-CoA carboxylase that in turn increases fatty acid oxidation via decreased malonyl-CoA [79]. AMPK may also be activated in the liver via PPARγ [80]. Recently, SOCS have emerged as important regulators of insulin signaling. SOCS3 inhibits janus kinase (JAK)-signal transducer and activator of transcription (STAT) signaling and activates JNK signaling, which further worsens insulin signaling and activates inflammatory signaling pathways. SOCS-3 is virtually absent in basal conditions, but is rapidly and robustly induced in response to cytokines and appears to promote the development of fatty liver, likely as a consequence of insulin resistance-mediated activation of SREBP1 [81]. In patients with NAFLD, 60% of triglycerides stored in the liver arise from circulating fatty acids whereas dietary fat supply and lipogenesis account for 15 and 25%, respectively [82]. The increase in lipogenesis appears to be paradox but may be explained by partial insulin resistance to the gluco-regulatory actions of insulin, whereas the lipogenic effects of insulin are preserved [83]. The degree to which a particular pathway becomes resistant may vary from tissue to tissue. Although β-oxidation of fatty acids in the liver may be increased in NAFLD [19], this apparently is not sufficient enough to overcome elevated triglyceride synthesis. The impaired apolipoprotein production in patients with NAFLD is likely to decrease the

production of VLDL, further contributing to hepatocellular accumulation of fat in the liver [84]. Overall, these findings support the concept that fatty acid metabolism in hepatocytes is compartmentalized in such a way that fatty acids from different sources have selective access to different parts of the cell. Newly synthesized and dietary fatty acids have access to parts of the cell housing pools of PPARα. Some of these new fatty acids are transferred into the cytosolic pool of triglyceride, the bulk of which is formed from free fatty acids originating in adipose tissue. Fatty acids derived by lipolysis of this triglyceride pool may not be able to access the PPARα activating pools, but are either re-esterified and incorporated into VLDL for export or returned to the cytoplasmic triglyceride pool [85].

How hepatic steatosis worsens insulin resistance Increased circulating levels of fatty acids [86] resulting from enhanced lipolysis secondary to adipose tissue insulin resistance alter hepatic fatty acid metabolism and promote the development of fatty liver. A product of lipogenesis is diacylglycerol, which in turn activates a serine/threonine kinase cascade resulting in activation of phosphokinase C-epsilon (Figure 45.6). Activated phosphokinase C-epsilon binds to the insulin receptor Glucose

FA

FATP5

IR

SLC2A2

IRS

JNK

Tyr-P

PI3K PKC

P FOXO

DAG AKT2

PEPCK

GSK3

G6Pase

FOXO

Lipogenesis

FA oxidation Glycogen synthesis

Mitochondrium

Nucleus

Figure 45.6 Mechanisms of fatty acid-induced insulin resistance. Hepatic diacylglycerol (DAG) content is determined by fatty acid (FA) influx via the fatty acid transport protein 5 (FATP5) and the amount of lipogenesis and FA oxidation. Increasing amounts of DAG activate phosphokinase C-epsilon (PKCε) and impair insulin receptor substrate (IRS) tyrosine phosphorylation. This results in reduced activation of phosphatidylinositol-3-kinase (PI3K) and AKT2, ultimately decreasing glycogen synthesis. This pathway can also be activated by fatty acid-induced c-jun N-terminal kinase (JNK) activation. Decreased phosphorylation of forkhead box protein O (FOXO) facilitates the transcription of the rate-limiting enzymes of gluconeogenesis, phosphoenolpyruvate carboxykinase (PEPCK), and glucose-6-phosphate phosphatase (G6Pase), respectively. Export of glucose occurs via the solute carrier 2A2 (SLC2A2). GSK3, glycogen synthase kinase-3

45: NON-ALCOHOLIC FATTY LIVER DISEASE: A PATHOPHYSIOLOGICAL PERSPECTIVE

and inhibits its tyrosine kinase activity and also interferes with the ability of insulin to phosphorylate IRS 2 on tyrosine residues. This then results in reduced activation of phosphatidylinositol-3-kinase and Akt 2, as shown in patients with NAFLD [87]. Lower phosphorylation of glycogen synthase kinase-3 results in reduced insulin-stimulated glycogen synthase activity causing decreased insulin-stimulated hepatic glucose uptake and reduced insulin stimulation of hepatic glucose production. Furthermore, decreased phosphorylation of forkhead box protein O (FoxO) allows it to enter the nucleus to activate transcription of the rate-limiting enzymes of gluconeogenesis, phosphoenolpyruvate carboxykinase, and glucose-6-phosphate phosphatase, respectively [87]. Increased gluconeogenesis with activation of translocation of the glucose transporter 2 promotes hyperglycemia aggravating insulin resistance [88, 89]. In insulin resistance, FoxO not only activates the gluconeogenetic pathway but also promotes production of VLDL by activating microsomal triglyceride transfer protein transcription [90]. Fatty acids may also cause insulin resistance independent of TNF-α by activating JNK [91]. Elevated JNK activity (Figure 45.7) in NAFLD [87, 92] impairs the signaling capacity of the insulin receptor at least in part through its effects on IRS phosphorylation [93]. Insulin resistance is further promoted by steatosis-mediated activation of IKKβ and NF-κB that in turn increases IL-6 production and secretion [94].

727

Molecular events that promote progression from steatosis to steatohepatitis Steatohepatitis is differentiated from a fatty liver by the presence of increased cell injury, apoptosis, inflammation, and development of fibrosis. Whereas most pathophysiological mechanisms of cell injury are present in both fatty liver and NASH, structural mitochondrial changes with development of paracrystalline inclusions and activation of JNK have been uniquely associated with NASH.

Oxidative stress A key mechanism for the genesis of cell injury in NAFLD is oxidative stress. Oxidative stress results from an imbalance between the production of ROS and antioxidant defenses. The mitochondria, cytochrome P450 system, and peroxisomes represent important sources of ROS. Both mitochondrial and cytochrome P450-related ROS generation has been implicated in the generation of oxidative stress associated with fatty liver and steatohepatitis. The structural defects in mitochondria in NASH are associated with impaired mitochondrial respiratory chain activity, thereby producing a state of uncoupled oxidation and phosphorylation that leads to ROS production [95].

IR IRS

Metabolic stress

Tyr-P

JNK ER IRE1

Protein misfolding

Insulin resistance

XBP1

UPR

Nucleus

Figure 45.7 Mechanism of endoplasmic reticulum (ER) stress-induced disturbance of insulin sensitivity. Metabolic stress such as increased fatty acid delivery to hepatocytes and lipid overload disrupt the smooth operation of the ER and causes protein misfolding. To cope with this stress, the ER activates the unfolded protein response (UPR) via activation of the transcriptional regulator XBP1. If homeostasis is not restored, stress-induced inositol-requiring protein 1 (IRE1) activates c-jun N-terminal kinase (JNK). This in turn impairs insulin signaling, resulting in insulin resistance. IR, insulin receptor; IRS, insulin receptor substrate

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Moreover, mitochondrial free cholesterol content appears to be critical in precipitating steatohepatitis by sensitizing hepatocytes to TNF-α and Fas through depletion of mitochondrial glutathione [96]. One mechanism to increase fatty acid oxidation is the proliferation and enlargement of hepatic peroxisomes as shown in patients with NAFLD [97]. Another adaptive response is increased microsomal oxidation of fatty acids [19, 98] evident by activation of cytochrome P450 enzyme 2E1 (CYP2E1). The change in CYP2E1 is not part of a generalized increase in hepatic cytochrome P450 proteins as cytochrome P450 enzyme 3A is decreased, consistent with earlier observation that documented a decrease in antipyrine metabolism in patients with NASH [98, 99]. Dehydroepiandrosterone increases superoxide dismutase, a key antioxidant [100]. Low circulating dehydroepiandrosterone levels [101] thus explain the reduced expression of superoxide dismutase in these patients [102] and also contribute to TNF-α-induced increased NF-κB-dependent transcription in the liver [103]. Persistent generation of ROS can also diminish insulin sensitivity, as recently demonstrated [104], and reduce the capacity of the liver to export triglycerides as VLDL [105]. The redox imbalance in NASH is characterized by a reduced antioxidant potential, as evidenced by glutathione depletion and reduced superoxide dismutase activity [102, 106], increased 3-nitrotyrosine activity, high 8-hydroxydeoxyguanosine levels, and low catalase activity [107–109]. Increased production of ROS promotes lipid peroxidation within hepatocytes, resulting in the formation of malondialdehyde and trans4-hydroxy-2-nonenal [110]. These molecules are formed through the peroxidation of polyunsaturated fatty acids that are preferentially oxidized owing to decreased carbon–hydrogen bond strength in methylene groups between unsaturated carbon pairs [111]. Lipid peroxidation has the potential to induce apoptosis, inflammation, and liver fibrosis by impairing nucleotide and protein synthesis, promoting production of inflammatory cytokines leading to neutrophil chemotaxis, and activation of hepatic stellate cells (HSCs).

Activation of lysosomal and mitochondrial pathways of apoptosis In patients with NASH, lysosomal involvement in apoptosis is an early event that is activated by ROS and fatty acids [112, 113]. This pathway results in lysosomal membrane permeabilization and release of cathepsin B into cytosol. This process is preceded by cytosol-to-lysosome translocation of B-cell lymphoma 2-associated X protein (Bax), a proapoptotic member of the B-cell lymphoma 2 (Bcl-2) protein family that induces membrane channel formation [113, 114]. Lysosomal permeabilization may be facilitated by a decreased expression of Bcl-XL , an antiapoptotic protein and Bax antagonist [115]. Once released,

cathepsin B increases TNF-α via activation of the NF-κB pathway. Fatty acids also induce JNK-dependent apoptosis by up-regulation of proapoptotic B-cell lymphoma 2interacting mediator of cell death (Bim) and Bax leading to mitochondrial permeabilization, cytochrome c release, and caspase-3 and -7 activation [116]. Measurement of caspase-3 endproducts such as cytokeratin-18 fragments may serve as a serum marker to distinguish NASH from fatty liver [117]. Expression of Bim depends on activation of FoxO3A [118]. Interestingly the inherent toxic potential of individual fatty acids varies in this regard, suggesting that fatty acid profiles of patients with NAFLD may also allow prediction who will develop more severe disease. Because fatty acid stimulation of JNK and FoxO3A appears to work together to induce apoptosis, one may speculate that JNK facilitates Bax dissociation from cytosolic binding partners or its translocation to mitochondria [119]. Once this has been achieved, Bim may in turn directly bind and activate Bax or it may de-repress an antiapoptotic Bcl-2 protein to help activate Bax [120]. Upon full activation, Bax induces mitochondrial dysfunction, release of cytochrome c, activation of caspases, and ultimately apoptosis. Under physiological conditions, hepatocytes are resistant to tumor necrosis factor-related apoptosis-inducing ligand (TRAIL) cytotoxicity [121]. Under pathophysiological conditions such as fatty liver, fatty acids appear to sensitize hepatocytes to Fas- and TRAIL-mediated apoptosis [122, 123]. This is supported by a JNK-mediated enhanced hepatic expression of the TRAIL receptor DR5 observed in patients with NASH [122].

Endoplasmic reticulum stress and the unfolded protein response A delicate balance between a cell’s synthetic needs and the ability of the endoplasmic reticulum (ER) to meet these demands is an important determinant of whether a given cell lives or dies. Perturbation of this balance triggers the unfolded protein response (UPR) (Figure 45.8) that can correct the imbalance and help the cell to adapt or condemns the cell to death [124]. The membrane of the ER contains three stress sensors that include inositol-requiring protein1 (IRE1), activating transcription factor 6 (ATF6), and protein kinase RNA-like endoplasmic reticulum kinase (PERK). Activation of these sensors through increasing levels of unfolded proteins is mediated by the dissociation of the chaperone GRP78 following ER stress. Both IRE1 and PERK are oligomerized and autophosphorylated. IRE1 catalyzes the splicing of X-box protein 1 (XBP1) to generate a more potent transcription factor that promotes the endoplasmic reticulum-associated degradation (ERAD) pathway to remove terminally misfolded proteins. PERK phosphorylates eukaryotic initiation factor 2 (eIF2), leading to global translation attenuation. Under conditions of sustained ER stress with

45: NON-ALCOHOLIC FATTY LIVER DISEASE: A PATHOPHYSIOLOGICAL PERSPECTIVE

729

GRP78

ER

GRP78

ATF6

GRP78

GRP78

IRE1

IRE1

PERK

eIF2 XBP1

Bax/BAK

eIF2

Calcium

JNK ATF4

Translation arrest

ATF6

sXBP1

Caspases CHOP

GADD34

Apoptosis ERAD ER chaperones

Figure 45.8 The endoplasmic reticulum (ER) stress pathway in NAFLD. Three stress inducers (ATF6, IRE1, PERK) are located in the ER membrane that remain inactivate as long as they are bound to the chaperone GRP78. Upon activation, the endoribonuclease domain of IRE1 cleaves XBP1 to sXBP1, leading to transcriptional activation of adaptive molecules to overcome ER stress. ATF6 also activates this pathway. Translation arrest is achieved via PERK-facilitated phosphorylation of eIF2α. If these responses fail to correct the ER stress, PERK leads to selective translation of ATF4, CHOP, and activation of the apoptotic machinery. Increased phosphorylation of IRE-1also can facilitate recruitment of TNF-associated factor 2 protein and activation of JNK. GADD34 promotes dephosphorylation of eIF2α, leading to attenuation of the ER stress response. Finally, proapoptotic Bak and Bax undergo conformational alteration and permit calcium release, which leads to activation of the caspase cascade and apoptosis. ATF6, activating transcription factor 6; ATF4, activating transcription factor 4; PERK, protein kinase RNA-like ER kinase; XBP1, X-box protein 1; sXBP1, spliced X-box protein 1; JNK, c-jun N-terminal kinase; eIF2α, eukaryotic initiation factor 2α; CHOP, C/EBP homologous protein; GADD34, growth arrest and DNA damage-34; ERAD, ER-associated degradation

prosurvival efforts being exhausted, PERK leads to selective translation of ATF4, transcription of CHOP (C/EBP homologous protein), and activation of the apoptotic machinery. Increased phosphorylation of IRE-1 also can facilitate recruitment of TNF-associated factor 2 protein and activation of JNK [125]. Finally, Bak and Bax in the ER membrane undergo conformational alteration and permit calcium release, which leads to activation of the caspase cascade. The calcium efflux also leads to the activation of mitochondria-dependent apoptosis. CHOP, one of the UPR downstream effectors, inhibits the expression of Bcl-2 and thus promotes apoptosis. In patients with NAFLD, the UPR is activated [92]. It has been shown that IRE-1 activation appears to play an important role in the genesis of cell injury in NASH via activation of JNK phosphorylation. Interestingly, there seems to be a close association of IRE-1 activation with the histological activity of the disease. Failure to generate ER degradation-enhancing α-mannosidase-like protein (EDEM) in response to spliced X-box protein (sXBP) is another key observation supporting the notion that low EDEM is not critical for the development of NASH. One may speculate that patients with the lowest EDEM levels are at particular risk to progress to cirrhosis due to

insufficient degradation of unfolded proteins perturbating ER stress. Despite increased phosphorylated eIF-2α, patients with NASH are apparently unable to up-regulate ATF4, CHOP, and growth arrest and DNA damage-34 (GADD34), which contributes to the failure to recover from ER stress [126].

Activation of inflammatory signaling in NASH It seems plausible that the inflammatory response in the liver is initiated in adipocytes, as they are the first cells affected by the development of NAFLD. One such adipocyte-derived factor is adiponectin, which exerts several anti-inflammatory effects, including inhibition of NF-κB activation [127], suppression of macrophage function [128], and suppression of TNF-α synthesis and release from macrophages [129]. Lower adiponectin levels in patients with NASH compared with steatosis alone that are independent of insulin resistance contribute to increased circulating serum levels of TNF-α and the hepatic expression of TNF-α receptor in these patients [43, 56, 130]. A gradual rise in serum levels of CCL-2 from

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healthy controls to steatosis, reaching the highest levels in NASH, suggests that CCL-2 may play a role in the progression from steatosis to steatohepatitis partly by promoting infiltration of leukocytes into the liver [51]. NASH has also recently been shown to be associated with activation of the innate immune system and also lipoxygenase pathways leading to increased systemic levels of inflammatory eicosanoids [131].

Activation of fibrogenesis NASH is characterized by perisinusoidal deposition of a collagenous matrix by activated HSCs. In NASH, HSC activation correlates with portal and lobular inflammation, but not with steatosis alone, suggesting that the mechanisms implicated in fibrogenesis are likely related to signals that not only lead to hepatic fat deposition but are also pro-inflammatory and pro-fibrotic in their own right [132]. One such signal could be adiponectin, known to inhibit HSC proliferation and migration and the expression of the classic fibrogenic cytokine transforming growth factor β (TGFβ), respectively [133, 134]. Adiponectin-mediated activation of HSCs may occur via activation of AMPK [135], NF-κB [136], or Kruppel-like factors [137]. In patients with NASH, adiponectin and adiponectin receptor 2 expressions are significantly reduced compared with steatosis alone [59, 130]. Another emerging picture is the potential fibrogenic role of insulin and glucose being capable to induce the synthesis of connective tissue growth factor by HSCs [138]. Endocannabinoids have recently emerged as a potent mediator of hepatic steatosis, HSC activation, and fibrosis [139, 140]. Cannabinoid (CB) receptor 1 is expressed at low density on hepatocytes, quiescent HSCs, and endothelial cells and its expression is strongly induced in fibrosis and cirrhosis [141–143]. While CB1 receptors on hepatocytes promote lipogenesis, expression on HSCs accelerates a fibrogenic response associated with chronic liver injury, independently of the offending agent [141].

The role of genes in development of NASH and disease progression Clinical experience indicates that NAFLD and other features of the metabolic syndrome often cluster within families. Also, those of Hispanic origin appear to have more aggressive disease than those of African American origin, who seem to be protected from NASH. To identify genes contributing to NAFLD, three different genetic approaches may be chosen, among which are linkage analysis, single nucleotide polymorphism scanning, and candidate gene studies. A number of studies (Table 45.1) have reported gene polymorphisms that may be linked to the development of fatty liver or progression to NASH

and cirrhosis, respectively. In pursuing the search for genetic factors involved in NAFLD, one has to choose carefully the appropriate study design to avoid potential pitfalls. Interpretation of published studies so far is hampered, however, by small cohort sizes, inadequate statistical power, poor phenotypic criteria, and suboptimal control population selection. Nevertheless, with regard to the underlying molecular biology of NAFLD, variations in genes affecting lipid metabolism, insulin resistance, adipocytokines, and profibrogenic mediators appear to be of special interest.

TREATMENT STRATEGIES AND DRUG TARGETS To date, no single therapy has been approved for treating NAFLD. Based on our current understanding of the natural history of this disease, one may propose to treat only patients with NASH or those who are at risk of developing liver cirrhosis. This approach may change based on the recent finding that hepatic inflammation is detectable by gene expression analysis and immunochemistry in simple fatty livers in the absence of histologically detectable inflammatory changes [52]. Irrespective, treatment approaches may aim either at preventing disease progression or at reversing already established fatty liver or NASH.

Weight loss: diet, exercise, and bariatrics Energy intake has been shown to be significantly higher in patients with NAFLD than in subjects without evidence for fatty liver [144]. Saturated fatty acids constitute a significant part of the Western diet and subjects with NASH may consume a diet richer in saturated fatty acids and poorer in polyunsaturated fatty acids, fiber, and antioxidant vitamins C and E [145–147]. A 10% diet-induced body weight reduction has been shown to improve not only metabolic parameters associated with NAFLD but also serum aminotransferases, hepatic steatosis, and necroinflammatory activity [148, 149]. Bariatric surgery can lead to a reduction in serum aminotransferases and histological improvement [150], but is reserved for those individuals with a body mass index (BMI) >35 kg m−2 along with end organ disease, for example, diabetes. Although lifestyle change and exercise to reduce body weight and treat concomitant diabetes and dyslipidemia are accepted first-line therapy, they have not been shown convincingly to reduce the risk of disease progression. As our understanding of the pathological consequences of poor dietary choices and lack of exercise on adipose tissue and liver continues to improve, we should be able to develop specific guidelines for a “fatty liver diet” and exercise regimens that ultimately will decrease the severity of NAFLD.

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731

Table 45.1 Gene polymorphisms studied in patients with NAFLD Gene

Gene namea

Protein function

ADIPOQ ADRB3 ADRB2 APOE LEPR LIPC MTTP PEMT PPARA SREBF1 CLOCK PPARGC1A MTHFR SOD2 CYP2E1 TNFA IL1B CD14 KLF6 TGFB1 AT

Adiponectin1

Lipid metabolism Lipid metabolism Lipid metabolism Lipid metabolism Lipid metabolism Lipid metabolism Lipid metabolism Lipid metabolism Lipid metabolism Lipid metabolism Lipid metabolism Insulin resistance Endoplasmic reticulum stress Oxidative stress Oxidative stress Inflammation Inflammation Inflammation Fibrogenesis Fibrogenesis Fibrogenesis

β3-Adrenergic receptor2,3 β2-Adrenergic receptor4 Apolipoprotein E5–7 Leptin receptor8 Hepatic lipase9 Microsomal triglyceride transfer protein10,11 Phosphatidylethanolamine N -methyltransferase12,13 Peroxisome proliferator-activated receptor14 Sterol regulatory element binding transcription factor 115 Circadian locomoter output cycles protein kaput16 Peroxisome proliferator activated receptor γ co-activator 1A17,18 Methylenetetrahydrofolate19 Superoxide dismutase 210 Cytochrome P450 2E120 Tumor necrosis factor α21−23 Interleukin 1β2 CD1424 Kruppel-like factor 625 Transforming growth factor β26 Angiotensinogen26

a References: 1 Musso, Hepatology 2008, 47, 1167–77; 2 Nozaki, Alcohol Clin Exp Res 2004, 28, 106S–110S; 3 Shima, Clin Chim Acta 1998, 274, 167–76; 4 Iwamoto, Clin Chim Acta 2001, 314, 85–91; 5 Yang, Hepatogastroenterology, 2005, 52, 132–83; 6 Sazci, Dig Dis Sci 2008, 53, 3218–24; 7 Demirag, Dig Dis Sci , 2007, 52, 3399–403; 8 Chen, Zhonghua Gan Zang Bing Za Zhi 2006, 14, 453–55; 9 Zhan, Zhonghua Gan Zang Bing Za Zhi , 2008, 16, 375–78; 10 Namikawa, J Hepatol , 2004, 40, 781–86; 11 Gambino, Hepatology, 2007, 45, 1097–107; 12 Dong, J Hepatol , 2007, 46, 915–20; 13 Song, FASEB J , 2005, 19, 1266–71; 14 Cheng, Zhonghua Gan Zang Bing Za Zhi , 2007, 15, 65; 15 Zhuang, Zhonghua Gan Zang Bing Za Zhi , 2008, 16, 138–39; 16 Sookoian, World J Gastroenterol , 2007, 13, 4242–48; 17 Yoneda, BMC Gastroenterol, 2008, 8, 27; 18 Hui, Liver Int, 2008, 28, 385–92; 19 Sazci, Cell Biochem Function, 2008, 26, 291–96; 20 Valenti, Gastroenterology 2002, 122, 274–80; 21 Piao, World J Gastroenterol , 2003, 9, 2612–15; 22 Huang, Zhonghua Gan Zang Bing Za Zhi , 2006, 14, 613–15; 23 Tokushige, J Hepatol , 2007, 46, 1104–10; 24 Brun, Gut, 2006, 55, 1212; 25 Miele, Gastroenterology, 2008, 135, 282–91; 26 Dixon, J Hepatol , 2003, 39, 967–71.

Targeting gut microbiota The role of gut flora in the genesis of insulin resistance suggests that one could target the gut flora for the treatment of insulin resistance and its consequences, for example, NAFLD. In general, one can envision two strategies to be taken for gut microbiota-targeted therapies. One is to eliminate a specific microorganism or certain species of bacteria. However, antibiotic treatment also destroys beneficial microorganisms and drug resistance may also emerge over time. A better approach would be to alter the composition of the gut flora by employing pro- and prebiotics [151]. These approaches await development and formal testing.

Targeting insulin signaling pathways Impaired insulin signaling is a key defect in the pathogenesis of NAFLD. Targeting either individual insulin signaling steps or mimicking insulin action at the transcriptional level, bypassing defects in insulin signaling with the potential to remedy many of the defects seen in insulin resistant states, may thus represent an attractive target for treatment. Inhibition of protein kinase C epsilon may represent such a target to treat insulin resistance in fatty liver disease

[88]. Another target may be FoxO1, which plays a pivotal role in the control of gluconeogenesis by linking insulin signaling via Akt2 to decreased transcription of key gluconeogenic enzymes. Specifically, insulin-mediated Akt2 phosphorylation of FoxO1 leads to nuclear exclusion, ubiquitination, and degradation [152] that decreases the expression of gluconeogenic enzymes. Employing antisense oligonucleotides indeed decreased the expression of FoxO1 and this resulted in a lower glucose production and also a reduced hepatic triglyceride content. Hepatic and peripheral insulin action also improved [153]. The UPR can also trigger both insulin resistance and liver injury. It is therefore another attractive therapeutic target. Indeed, 4-phenylbutyric acid, known to stabilize protein conformation and improve ER folding capacity [154], has been shown to restore glucose homeostasis and to be capable of obtaining resolution of NAFLD [155]. The taurine conjugate of the endogenous bile acid ursodeoxycholic acid appears to modulate ER function by suppression of JNK activation in the liver, thereby restoring insulin signaling. Unfortunately, ursodeoxycholic acid may not show beneficial effects in patients with NAFLD [156], but this may not hold true for other compounds to be tested in the future. JNK activation and downstream activation of the IKKβ/NF-κB are additional attractive targets for the treatment of insulin resistance and NAFLD. NF-κB

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activation in the liver triggers not only low-grade inflammation but also insulin resistance and fatty liver [94], while deletion of IKKβ preserves the sensitivity to insulin [157]. Indeed, specific blocking of IKKβ has been shown to prevent the development of NAFLD and to attenuate the appearance of NASH progression [158]. Blockade of IKKβ is accompanied by enhanced PPARα signaling and thus lipid degradation through mitochondrial fatty acid oxidation [158]. On the other hand, IKKγ inhibition causes steatohepatitis, fibrosis, and liver cancer [159, 160]. This phenomenon may be related to an entire block of NF-κB signaling in the case of IKKγ, whereas inactivation of IKKβ leads only to a partial block [161]. SOCS-3, which regulates JNK activity, offers yet another potential target for therapy.

Targeting triglyceride accumulation To prevent or reduce hepatic triglyceride storage, targeting fatty acid uptake into liver, the triglyceride synthetic pathway, or fatty acid β-oxidation may represent targets for drug development. Several such targets are currently under study. Of these, the CB1 antagonists are most developed. These agents improve insulin sensitivity and promote weight loss. Another such target is the fatty acid binding protein 5; its inhibition may reverse NAFLD despite increased de novo synthesis of preferentially monounsaturated fatty acids to compensate for the disturbed fatty acid supply from the diet [162]. Diacylglycerol acyltransferase (DGAT), catalyzing the final step in triglyceride synthesis, is another such target. It exists as two isozymes; however, inhibition of DGAT2 in a mouse model of steatosis led to the development of cell injury despite decreased steatosis, suggesting that triglyceride formation may serve a protective mechanism under conditions of insulin resistance-mediated increases in delivery of fatty acids to the liver [163]. Acetyl-CoA carboxylase is another attractive candidate because it catalyzes the formation of malonyl-CoA, which leads to fatty acid synthesis and decreases the activity of carnitine palmitoyltransferase, the rate-limiting step in mitochondrial fatty acid oxidation [164]. This enzyme is modulated by AMPK, that is, downstream of adiponectin signaling; metformin targets this pathway. Although carnitine palmitoyltransferase 1a-promoted stimulation of fatty acid oxidation may be limited to more downstream steps over which carnitine palmitoyltransferase 1a exerts very low control [165], pharmacological targeting to increase hepatic fat oxidation has potential as a novel approach for the treatment of NAFLD. This assumption is based on recent studies that demonstrate a reduction in triglyceride accumulation and secretion in the presence of unchanged de novo lipid synthesis absence can be achieved with overexpression of carnitine palmitoyltransferase 1a [166].

Nuclear receptors and transcription factors as therapeutic targets Nuclear receptors are at the center of a network for metabolic control of transcription. As transcriptional regulators they are unique, because in the presence or absence of a ligand they can function as a switch, turning on or off transcription in response to small molecules and to restore dysregulated metabolic pathways in human disease. At the cellular level these receptors exert their function in mutual interaction with accessory proteins and the pathways they control are intertwined with each other. To reduce drug toxicity, minimize unwanted activities, and obtain more tissue-selective biological responses, isotype-specific modulators will be available in the future. The nuclear receptors that could be targeted include the retinoid X receptors (RXRs), farnesoid X receptors (FXRs), PPARα, PPARγ, sirtuins, and CHREBP. While RXRs have been largely neglected for drug discovery, co-administration of an RXR-selective ligand with a PPARγ agonist may achieve synergistic activity and may provide a potential route to safer, more efficacious therapies for NAFLD [167]. The principal predicted risk of this approach is the risk of teratogenecity. FXR is a metabolic nuclear receptor expressed in the liver and other organs such as intestine and adipocytes. Activation of the FXR may be beneficial by (i) reducing hepatic fatty acid synthesis via inhibition of SREBP1 [168], (ii) increasing insulin sensitivity in liver and adipose tissue [169], and (iii) enhancing hepatic fatty acid oxidation via induction of PPARα [170]. PPARγ agonists such as thiazolidinediones improve insulin sensitivity in adipose tissue and reduce hepatic inflammation [171]. As PPARγ agonists may lower RBP4 levels [60], this may contribute to the insulin-sensitizing effects of these drugs. Another mechanism that may contribute to the beneficial effects of these receptor agonists is due to the increase in adiponectin synthesis [172]. The rise in adiponectin then in turn may act via PPARGC1 to activate PPARα [173]. Unfortunately, these agents may have limited utility due to weight gain, a direct consequence of the shift of fat distribution from visceral to subcutaneous depots [174]. In general, these receptor agonists favor fatty acid oxidation in non-adipose tissues and promote fat storage in adipose tissue; these effects enable appropriate fuel partitioning and lipid storage within the body. With this in mind, dual PPARα/γ agonists appear attractive for therapy as they have the potential to improve insulin resistance, reduce circulating fatty acids, and avoid the weight gain associated with pure PPARγ agonists [175, 176]. PPARδ is another receptor that when activated in adipose tissue counteracts high-fat diet-induced weight gain and prevents NAFLD [177]. The recent discovery that PPARδ regulates the expression of multiple SREBP1

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target genes in liver may have therapeutic potential in the treatment of NAFLD [178]. Sirtuin 1 (Sirt1) is a protein deacetylase/mono-adenosine diphosphate (ADP) -ribosyltransferase that depends on nicotinamide adenine dinucleotide (NAD+ ), linking its activity to the cellular metabolic status in response to nutrient availability. Because Sirt1 can deacetylate IRS, PPARγ, PPARGC1, NF-κB, LXR, and SREBP1 [179–184], all of which are potential targets involved in either insulin resistance and/or development of fatty liver, it becomes an attractive target for drug development. Additional support comes from the observation that Sirt1 is decreased in NAFLD [185] and represents a major regulator of adiponectin expression and secretion [186, 187]. Indeed, insulin resistance can be improved through activation of Sirt1 by the natural compound resveratrol, present in juice and red wine, or by small molecules [188, 189]. In addition, Sirt1 may be capable of increasing the number of mitochondria in cells and stimulate AMPK [190], thereby increasing the cell’s capacity for fatty acid oxidation.

Targeting apoptosis Apoptosis in NAFLD appears to involve several pathways. It is at present unknown which pathway dominates and whether they occur at the same time or in a defined sequence of events. Several targets for decreasing apoptotic activity have emerged in the past. It appears possible to reduce fatty acid-induced lipotoxicity by stabilizing lysosomal and mitochondrial membrane integrity with 18β-glycyrrhetinic acid [191]. Another target for treatment may be inhibition of JNK [192]. This approach may not only ameliorate JNK-induced Bax-dependent apoptosis but also ameliorate the consequences of insulin resistance. Alternatively, targeting Bax or its natural inhibitor would appear to be a viable strategy to prevent fatty acid-induced cytotoxicity [115].

Targeting fibrosis The translation of advances in basic research into improved therapeutics for the management of liver fibrosis is still poor and proof of efficacy of most potential antifibrogenic molecules in a clinical setting is currently lacking [193]. Most of the molecules studied so far have been directed against activation of HSCs or at inflammatory signals, or aim to reduce the oxidative stress. In patients with NASH, blocking the angiotensin II type 1 receptor with losartan appears to reduce plasma TGFβ levels and to decrease the activation of HSCs, thereby improving hepatic necroinflammation and fibrosis [194, 195]. Vitamin E also may have some beneficial effect on fibrogenesis by reversing elevated plasma TGFβ levels [134].

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Activation of the endocannabinoid system in liver promotes fibrogenesis; therefore blocking this system appears attractive for the treatment of NAFLD. Although CB receptor 1 and 2 are only marginally expressed in normal liver, they undergo marked up-regulation during fibrogenesis [196, 197]. Selective inactivation of the CB receptor 1 reduces hepatic expression of TGFβ and the accumulation of fibrogenic cells in the liver after apoptosis and growth inhibition of hepatic myofibroblasts [141], molecular mechanisms that have already been shown to be important for fibrosis resolution. Another beneficial effect of selective inactivation of the CB receptor 1 is the increase in adiponectin levels [198]. Interestingly, activation of the CB receptor 2 appears to limit the progression of fibrosis [196], suggesting that the combination of a selective CB receptor 1 antagonist and a CB receptor 2 agonist might open new therapeutic avenues for the treatment of liver fibrosis. In addition to their accelerating effects on fibrogenesis [139, 141], endocannabinoids also stimulate fatty acid synthesis via activation of SREBP1 as the CB receptor 1 is also expressed on hepatocytes and adipocytes [143, 199]. Blocking the CB receptor 1 therefore may also prevent the development of NAFLD [200].

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Pathophysiology of Alcoholic Liver Disease Natalia Nieto1 and Marcos Rojkind2 1

Mount Sinai School of Medicine, Department of Medicine/Division of Liver Diseases, New York, NY, USA 2 Department of Biochemistry and Molecular Biology, The George Washington University Medical Center, Washington, DC, USA

INTRODUCTION Alcoholism is a disease that affects approximately 11 million men and 5 million women in the United States. Of these, one in five will develop alcoholic hepatitis and one-third will develop cirrhosis. There are multiple factors that determine the outcome of the disease. These include among others, the intensity and duration of alcohol intake, gender, and genetic and epigenetic factors. In this chapter we review important aspects of alcohol metabolism and the molecular mechanisms whereby ethanol and/or its metabolites induce liver damage, fibrosis, and cirrhosis. Alcohol metabolism occurs largely in the liver, where it is metabolized by three enzymatic pathways. The bulk of ethanol oxidation occurs in the liver via alcohol dehydrogenase (ADH), which is predominantly expressed in hepatocytes, although other tissues such as the gastric mucosa express ADH and contribute to the overall oxidation of ethanol [1]. The difference in the expression of gastric ADH plays a key role in gender sensitivity to alcohol toxicity, with women being more prone to develop liver disease under lower alcohol concentrations [1]. The second major pathway for ethanol elimination is the microsomal ethanol oxidizing system (MEOS) catalyzed by cytochrome P450 2E1 (CYP2E1). CYP2E1 The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

activity pathway is elevated in chronic alcoholics and induction of this pathway contributes to the metabolic tolerance to ethanol observed in alcoholics [1]. A large number of xenobiotics and drugs are substrates for CYP2E1; therefore, induction of CYP2E1 can have a major impact on the production of highly toxic metabolites in alcoholics, promoting liver injury [1] (see Figure 46.1). Whereas ADH generates mainly acetaldehyde from ethanol, CYP2E1 generates acetaldehyde, reactive oxygen species (ROS), and free radicals that induce lipid peroxidation and form adducts with several macromolecules. As a consequence of lipid peroxidation, there is formation of several reactive aliphatic aldehydes that are fibrogenic (Chapter 28) (Figure 46.1) [1]. As in the case of ADH, CYP2E1-dependent oxidation of ethanol takes place primarily in the hepatocyte [1]; however, Kupffer cells also express CYP2E1. The third pathway for ethanol metabolism is a non-oxidative pathway catalyzed by fatty acid ethyl ester (FAEE) synthase, leading to the formation of FAEEs [2]. The concentration of FAEEs are highest in organs susceptible to the toxic effects of ethanol such as liver and pancreas [2]. They accumulate in the plasma membrane, and also in the mitochondria and lysosome membranes [2], having a negative impact on the activity of these organelles.

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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THE LIVER: PATHOGENESIS AND RISK FACTORS OF ALD Acetaldehyde

H2O2

P Smad 3

TGF-β

P Smad 4

Smad 3 P

ad

Sm

P

The histological manifestations of chronic alcoholism include, among others, fatty liver, formation of Mallory bodies [12], hepatocyte apoptosis and necrosis, inflammation, formation of regenerating nodules, fibrosis, and eventually cirrhosis [13, 14]. Fibrosis initiates in response to hepatocellular damage, with inflammatory infiltrates contributing to the progression of the disease [15]. Many of the events involved in the development of fibrosis are similar to those observed in other tissue responses to injury, such as wound healing in the skin and soft tissues.

4

Sp1+Sp3 COL1A2

Figure 46.1 Schematic representation of the proximal promoter of the COLIA2 gene containing the acetaldehyde responsive element and the main transcription factors that acetaldehyde activates in HSC. The early events (up to 6 hours) are acetaldehyde dependent and result in phosphorylation of Smad 3 and formation of Smad3/4 complexes. The late events are TGF-β-dependent and result in up-regulation of Smads 3 and 4 and phosphorylation of Smad 3. Note the key role that H2 O2 plays in acetaldehyde-induced COLIA2 gene up-regulation.

Ethanol induces liver injury by multiple mechanisms. (1) It can act as a solvent and induces changes in membrane fluidity and function [3]. (2) Ethanol metabolism causes a significant change in the redox state of hepatocytes by influencing the NAD+ /NADH ratio. This could result in mitochondrial dysfunction, generation of H2 O2 , and accumulation of lactic acid [1]. (3) Ethanol metabolism leads to accumulation of reactive aldehydes (acetaldehyde, 4-hydroxynonenal (HNE), and malonaldehyde) that are fibrogenic when added to cultured hepatic stellate cells, the main type I collagen producing cells of the liver (Chapter 28). (4) Alcohol metabolism by CYP2E1 generates free radicals that form adducts with macromolecules and induces an immunological response to the new antigens (see [4–6]). (5) Hepatic stellate cells express leptin and adiponectin, chemokines known to control fat metabolism in adipose tissue. Their expression is increased during fibrogenesis [7–9] (see Section “Generation of ROS in ALD”). Therefore, changes in their concentrations and/or regulation could be responsible for liver fat accumulation observed in alcoholics. (6) Different liver cell types produce chemoattractants for inflammatory cells and these could further contribute to liver disease by producing TGF-β and other cytokines involved in the fibrogenic cascade [10]. Finally, chronic alcohol consumption is accompanied by a leaky intestine that allows translocation of bacteria (peritonitis) and/or endotoxin, a significant contributor to liver disease via Kupffer cell activation [11].

PATHOGENESIS AND RISK FACTORS OF ALD Alcohol-related lipid alterations A key feature of alcoholic liver disease (ALD) is accumulation of neutral lipids in hepatocytes leading to microand macro-vesicular steatosis and ballooning degeneration. Hypercaloric diets and the subsequent obesity also lead to similar changes as it occurs in non-alcoholic fatty liver disease (NAFLD). Hence, accumulation of lipids in hepatocytes is a pathological “footprint” of ALD and NAFLD. Dietary fat content and composition play a major role in the pathogenesis of ALD. In fact, diets rich in saturated fatty acids or medium chain triglycerides protect from alcoholic liver injury, whereas diets containing polyunsaturated fatty acids increase liver injury [16–18]. A key role in hepatic lipid metabolism is mediated, among others, by transcription factors such as peroxisome proliferator-activated receptor alpha (PPARα) and sterol regulatory element binding protein-1 (SREBP-1). PPARα is a receptor for free fatty acids, and is known to induce genes involved in transport, oxidation, and export of free fatty acids. Moreover, fatty acids serve as ligands for PPARα, and when their levels increase, activation of PPARα induces a series of fatty acid-metabolizing enzymes to restore normal fatty acid levels. Hepatic fatty acid levels are elevated during ethanol consumption. However, in vitro studies have shown that ethanol metabolism inhibits the capacity of PPARα to bind DNA and activate reporter genes. Although chronic ethanol treatment in rodents inhibits PPARα, it activates SREBPs [19–24]. SREBPs act as sensors of hepatic cholesterol levels, and activate genes involved in the synthesis of cholesterol and free fatty acids. SREBP-1a and -1c regulate genes required for hepatic triglyceride synthesis such as acetyl-coenzyme A acetyl-(CoA) carboxylase, fatty acid synthase, stearoyl-CoA desaturase 1, malic enzyme, and ATP citrate lyase. SREBP-2 modulates genes required for cholesterol synthesis, including HMG-CoA synthase, HMG-CoA reductase, and low-density lipoprotein receptor [23, 24]. In addition, ethanol inhibits 5′ -AMP-activated protein kinase

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(AMPK) which controls fatty acid metabolism by inhibiting acetyl-CoA carboxylase, reducing malonyl-CoA, and thereby allowing fatty acid transport into the mitochondria for their oxidation. The adipose tissue secretes adiponectin, which plays a crucial role in the development of ALD [25, 26]. Two adiponectin receptors (AdipoR1 and AdipoR2) have been cloned [25]. The liver expresses mainly AdipoR2, where it operates as the transducer of adiponectin-mediated activation of AMPK and PPARγ, leading to augmented fatty acid oxidation and reduced fat accumulation [25, 27]. In humans, adiposity is inversely related to adiponectin levels [27]. Chronic ethanol consumption significantly decreases circulating concentrations of adiponectin in mice [28]. Delivery of recombinant fulllength adiponectin into these mice reverses liver steatosis and injury [28]. Moreover, pioglitazone, a PPARγ agonist, prevents alcohol-induced liver injury in rats and mice [29–31]. Up-regulation of adiponectin by pioglitazone through activation of PPARγ could contribute to its hepatic protective effects [32]. Regulation of adiponectin by ethanol was further suggested in human studies, demonstrating that modest alcohol use significantly elevated plasma adiponectin in both healthy and insulin-resistant adult men [33, 34]. Consequently, the effect of ethanol on adiponectin physiology appears to depend on the quantity of alcohol consumed, the dietary milieu, and the nutritional status.

Obesity As suggested, liver and adipose tissue interact to regulate fat homeostasis. Adipose tissue is both a source of fatty acids that are delivered to the liver and a repository for triglycerides that are produced by the liver and released into the blood stream. In addition, fat plays a prominent role in regulating both intermediary metabolism and immunity by producing a variety of pluripotent soluble factors, including adipokines (leptin, resistin, and adiponectin), neurotransmitters (norepinephrine, neuropeptide Y, and angiotensin II), and cytokines [tumor necrosis factor α (TNFα) and interleukin 6 (IL-6)] [35, 36]. Thus, adipose tissue is an important neuroendocrine organ, as well as a component of the immune system. Abdominal fat produces less adiponectin and leptin, is more responsive to norepinephrine (i.e. exhibits greater lipolysis and fatty acid release), and produces significantly more proinflammatory cytokines [37]. Insulin resistance causes fatty liver, and on the other hand, fatty liver causes insulin resistance. Targeted disruption of insulin signaling in hepatocytes leads to steatosis associated with activation of SREBP-1 and induction of fatty acid synthesis [38]. Increased rates of hepatic fatty acid synthesis relative to fatty acid oxidation seem to play an important role in the pathogenesis of alcoholic fatty liver disease because hepatic steatosis is reversed

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by treatment with a PPARα agonist that up-regulates enzymes involved in fatty acid β-oxidation [21]. An increase in fatty acids in hepatocytes causes hepatic insulin resistance [39]. Even though the precise cellular mechanisms involved remain to be elucidated, differences in lipid partitioning appear to be involved because targeted overexpression of malonyl-CoA decarboxylase in hepatocytes from mice fed a high-fat diet induced hepatic steatosis (which inhibits de novo lipid synthesis and stimulates mitochondrial fatty acid oxidation), decreases hepatic lipid accumulation, and reverses fatty liver-associated hepatic, muscle, and systemic insulin resistance [40]. Insulin resistance is both a consequence and a cause of augmented TNFα activity. This notion supports the idea that hepatocyte accumulation of lipids and exposure to TNFα trigger comparable mechanisms that disrupt signaling downstream of insulin receptors that are expressed on the hepatocyte plasma membrane. A better understanding of the link between TNFα and insulin resistance has significant implications for the pathogenesis of ALD and steatohepatitis. However, despite the general agreement that TNFα is a critical mediator of damage to the hepatocyte and therefore alcoholic steatohepatitis (ASH) [41], the possibility that insulin resistance might play a role in the pathogenesis of ASH has scarcely been elucidated. On the other hand, there is evidence that the severity of insulin resistance correlates well with the severity of non-alcoholic steatohepatitis (NASH) in humans [42]. It is also suggested that obesity may promote cirrhosis. Although the absence of leptin promotes visceral adiposity and adipokine–cytokine abnormalities that cause hepatic injury and inflammation, a normal fibrogenic response to liver injury does not occur when leptin is not present. The mechanisms by which leptin promotes hepatic fibrosis are not fully understood, but they seem to involve direct interactions with hepatic stellate cells that lead to increased cell proliferation, activation, and transactivation of the COL1A1 promoter, leading to collagen I deposition [43–45]. Leptin may also regulate liver fibrosis through indirect mechanisms that involve other leptin sensitive mediators of stellate cell viability and maybe apoptosis.

Insulin resistance Chronic ethanol consumption leads to hepatic insulin resistance associated with steatosis, down-regulation of insulin-responsive genes, and increased lipid peroxidation and DNA damage. The mechanisms for ethanol-mediated insulin resistance are not completely clear; however, there is a strong correlation between insulin resistance and liver damage. NASH and fatty liver are associated with obesity, insulin resistance, and type 2 diabetes [46], suggesting that there may also be an important link between inflammatory cytokines, insulin resistance, and fatty liver during the progression of ALD. Leptin and TNFα appear to modulate differentially insulin-dependent signaling.

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Administration of metformin, known to re-establish insulin sensitivity, prevents the progression of obesitycoupled fatty liver [47]. It appears that leptin is not required to overcome hepatic insulin resistance. Rather, inhibition of TNFα-regulated genes appears to be relevant for fatty liver disease to progress [48]. Effective ligand binding is critical for intracellular signaling and many of the reported downstream adverse effects of ethanol on insulin signaling, including decreased cell survival, through PI3K/Akt/GSK3β and proliferation via the insulin receptor substrate-1 (IRS1)-dependent cascades could be mediated by impaired insulin binding to its receptors. Equilibrium binding assays confirmed that chronic ethanol consumption significantly impaired binding to the insulin receptor, but did not significantly alter binding of the insulin-like growth factor 1 (IGF1) or IGF2 receptors, despite their increased levels of gene expression [49–53]. The altered insulin signaling and downstream activation of growth and survival mechanisms suggest that chronic ethanol consumption leads to hepatic insulin resistance [31, 54, 55] and thus could serve as an vital mediator of ethanol-impaired liver regeneration. Ethanol also blocks nuclear SREBP-1c via interference of insulin signaling, which may be a possible connection between alcohol and insulin resistance. He et al. [56] demonstrated that chronic ethanol feeding to rats alters insulin signaling and results in hepatic insulin resistance by inducing TRB3, which effectively prevents Akt–Thr308 phosphorylation and the following Akt-mediated signaling. The ethanol-mediated blunting of nSREBP accumulation is translated into up-regulation of class 1 ADH transcription with the associated increase in ethanol metabolism.

Adipokines Triglycerides are stored in the white adipose tissue as excess energy and they are mobilized when energy intake is scarce. Adipose tissue is hormonally active and produces cytokines, chemokines, and adipokines [57]. Mature fat cells and infiltrating macrophages are a recognized source of adipokines within the adipose tissue. As indicated above, obesity and insulin resistance closely correlate with severe fibrogenic progression in ALD [58]. A broad range of adipokines has been associated as regulators of liver pathophysiology in ALD. Among them, leptin prevents lipotoxicity by restricting over-nutrition and promoting energy expenditure, and via induction of insulin action. The role of leptin in hepatic fibrosis has been characterized and serum leptin levels have been shown to be increased in patients with alcoholic cirrhosis [59]. Ablation of leptin or of the leptin receptor signaling as occurs in the ob/ob mice and in the fa/fa rats, respectively, triggers significant extracellular matrix remodeling and reduction of fibrosis [43, 60, 61]. The

decreased fibrogenic response in ob/ob mice regressed by supplementation with recombinant leptin [62]. Injection of leptin during acute and chronic liver injury causes significant up-regulation of collagen type I and TGF-β [63]. As for the likely mechanisms for the pro-fibrogenic role of leptin, it may increase the phagocytic activity, ROS generation, and cytokine secretion by Kupffer cells and macrophages [64], along with stimulation of endothelial cells to proliferate and to produce ROS and reactive nitrogen species (RNS) [65]. Leptin acts directly on hepatic stellate cells as they express functionally active leptin receptors, and incubation of stellate cells with leptin induces collagen I and synergizes with the pro-fibrogenic effects mediated by TGF-β [44, 66]. Leptin up-regulates tissue inhibitor of metalloproteinase 1 (TIMP1), playing a role in the insufficient degradation of fibrillar collagen in liver fibrosis [67] and in sustaining the survival of myofibroblasts [68]. Leptin can affect the activated phenotype of stellate cells as it works as a mitogen and a survival factor for stellate cells by means of activating the ERK and PI3K/Akt signaling pathways [9]. Exposure of stellate cells to leptin results in NFκB (nuclear factor kappa B)-mediated up-regulation of monocyte chemotactic protein-1 (MCP-1), a chemotactic factor for monocytes and T lymphocytes, therefore modulating the inflammatory response [69]. The highest concentration of adipokines in plasma is that of adiponectin. Whereas full-length adiponectin binds AdipoR1 with intermediate affinity, the globular domain binds AdipoR2 with intermediate affinity, but it is a high-affinity ligand for AdipoR1. AdipoR1 is highly expressed in the skeletal muscle whereas AdipoR2 is mainly expressed in the liver [25]. Apart from its role in glucose metabolism, adiponectin is hepatoprotective and antifibrogenic. Administration of adiponectin improves alcohol-induced liver injury and obesity via induction of hepatic fatty acid oxidation and fatty acid synthesis, and this effect is partially mediated by antagonizing TNFα [28]. Adiponectin prevents LPS-mediated liver injury in obese mice by interfering with TNFα [70]. In a model of ASH, serum adiponectin levels were reduced, indicating that deficiency of this adipokine may to some extent mediate alcohol-induced liver injury [28]. Adiponectin knockout mice develop more widespread liver fibrosis after CCl4 injection than controls [71]. Administration of adiponectin protects against the progression of NASH to cirrhosis and tumor formation [72]. The central mechanism of action of adiponectin in these protective effects appears to be by modulating the activated phenotype of hepatic stellate cells, which express both AdipoR1 and AdipoR2 [73, 74]. It is known that adiponectin suppresses proliferation and migration of stellate cells stimulated with platelet-derived growth factor BB (PDGF-BB) [71] and decreases the effect of TGF-β [71]. Adiponectin also reduces stellate

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cell proliferation and increases apoptosis [73]. Activated stellate cells have less adiponectin and more leptin while in quiescent stellate cells the opposite happens [73]. The adiponectin–AMPK axis is functional in stellate cells and it is partially responsible for the hepatoprotective effects of this adipokine [75]. Lastly, resistin expression in human liver increases in ALD, hepatitis C virus (HCV)-induced hepatitis, and NASH, and positively correlates with inflammation [76]. Resistin co-localizes with CD43 in livers of patients with alcoholic hepatitis [76]. Serum resistin levels are elevated in patients with cirrhosis compared to healthy individuals and are associated to the clinical and inflammatory stage of the disease [77, 78]. Expression of resistin in stellate cells has been documented only in quiescent but not in activated cells [77, 79]. However, cultured human stellate cells respond to incubation in the presence of resistin increasing pro-inflammatory chemokines such as MCP-1 and IL-8 [78].

Acetaldehyde Acetaldehyde is the first product of ethanol metabolism and together with the aldehydes generated via lipid peroxidation it induces the expression of the collagen I genes in hepatic stellate cells (HSCs) [80–84]. Although the molecular mechanisms whereby acetaldehyde up-regulates type I collagen genes remain unclear, it has been established that acetaldehyde, by itself, is a weak pro-fibrogenic factor that recruits TGF-β to exert and enhance its fibrogenic actions [82]. Early acetaldehyde-dependent events are directly induced by acetaldehyde in a protein synthesis-independent manner. However, the late events occurring after 12 hours of HSC exposure to acetaldehyde are TGF-β and protein synthesis dependent. H2 O2 is a second messenger of acetaldehyde and many of its actions are dependent on the accumulation of this ROS [81, 82]. Acetaldehyde-responsive elements that are functional in HSC have been localized to the proximal promoters of the rat α1(I) and human α2(I) collagen promoters [81, 82]. These responsive elements have been mapped to the sequences corresponding to the previously identified TGF-β and TNF α-responsible elements [85–87]. However, the differences in the transcription factors that bind to the promoters determine whether collagen gene expression is up- or down-regulated. In the rat col1a1 gene, the responsive element is localized to the −370 to −345 nucleotide sequence of the promoter [81]. This element binds p35-C/EBP-β and this transcription factor is critical for basal and acetaldehyde-dependent collagen gene transcription. However, when the dominant negative form of p35-C/EBP-β, namely p20-C/EBPβ and C/EBPδ bind to the promoter, as occurs after TNFα treatment, expression of the gene is repressed [87]. The acetaldehyde-responsive

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EtOH ADH

Acetaldehyde H2O2 TGF-β

CYP2E1

Fibrosis

Free Radicals ROS (H2O2, O2.−, etc.) LPO (4-HNE, MDA)

Adducts

Figure 46.2 Schematic representation of the pleiotropic actions of ethanol. It illustrates some alcohol metabolites resulting from alcohol metabolism. As shown, acetaldehyde metabolism via ADH generates H2 O2 and this, in turn, up-regulates TGFβ. The cytokine potentiates the fibrogenic actions of acetaldehyde. Ethanol metabolized by CYP2E1 generates acetaldehyde, and free radicals, such as ROS and lipid peroxidation end products, and induces lipid peroxidation. This, in turn, generates several aliphatic aldehydes that are also fibrogenic. The aldehydes form adducts with macromolecules and elicit immunological responses. Overall, these products and alcohol metabolism-derived species are responsible for many of the alterations observed in chronic alcoholics

element of the COL1A2 gene has been localized to the −378 to −183 nucleotide sequence of the promoter [82]. SP1 and SP3 bind to this acetaldehyde-responsive element. Moreover, complexes containing Smad 3 and Smad 4 also bind to the acetaldehyde-responsive element and are required for collagen gene transcription (see Figure 46.2). In contrast to TGF-β, acetaldehyde induces phosphorylation of Smad 3 but has no significant effect on the expression of these transcription factors. Overall, acetaldehyde and TGF-β up-regulate collagen gene expression by similar but not identical mechanisms. Gel shift assays performed to identify nuclear proteins that bind to the acetaldehyde-responsive element of the α2(I) human collagen promoter revealed the existence of a repressor complex whose composition remains unknown. This repressor complex is down-regulated upon incubation of HSC with acetaldehyde. In addition, acetaldehyde promotes the degradation of the two oncogenes Ski/Sno, known to be repressors of many TGF-β-dependent genes (M. Rojkind, unpublished observations). Acetaldehyde activates several signal transduction pathways that include, among others, PKC [88–90], JNK [91], and PI3K [92, 93]. The early response to acetaldehyde is PI3K dependent. However, the late events induced by TGF-β are PI3K independent [94]. PKC appears to be one of the upstream elements involved in the expression of collagen and the activation of ERK1/2 and also the phosphorylation of p70S6K, a member of the AKT survival pathway [92]. Overall, alcohol/acetaldehyde have additional effects. Although these may not directly modify collagen gene expression, they are important in the trans-differentiation of

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HSC into myofibroblasts, the main type I collagen producing cells in the liver. Acetaldehyde inhibits the expression of PPARγ, a receptor involved sustaining the adipogenic phenotype in HSC (see Chapter 28) [94]. This effect is also mediated by the increased formation of H2 O2 induced by acetaldehyde. Moreover, alcohol metabolism generates, in addition to acetaldehyde, ROS and alters the NAD+/NADH ratio [1]. Hence the formation of protein adducts and redox imbalance appear to play an important role in the endoplasmic reticulum (ER) stress (see Section “The Unfolded Protein Response in ALD”). The formation of free radicals and lipid peroxidation generates additional reactive aldehydes such as 4-HNE and malondialdehyde (MDA) with profibrogenic potential [83, 84]. Hence the fibrogenic actions of ethanol/acetaldehyde are complex and multifactorial and involve various liver cell types in addition to inflammatory cells.

Cytokines, chemokines and growth factors Steatosis is one of the first manifestations of injury in ALD, and in a large majority of patients it is followed by necroinflammatory activity. In some of them, fatty liver is eventually replaced by a fibrous collagenous collagen scar with the subsequent development of liver fibrosis and cirrhosis. Although steatosis per se is usually harmless and typically reversible, it is cell ballooning and the development of inflammation (steatohepatitis), what determines whether a patient progresses to irreversible liver damage and fibrosis. Over the years, the pathophysiological concept of ALD has evolved into a dual-hit hypothesis whereby the accumulation of fat in the liver constitutes the “first hit”, and pro-inflammatory challenge to the liver such as ROS and LPS constitutes the “second hit.” Cytokines and particularly chemokines are crucial in initiating and perpetuating the inflammatory infiltrate in steatohepatitis. A better understanding of this process might allow therapeutic intervention to switch off the inflammatory response before irreversible damage occurs in both ALD and NAFLD. Persistent inflammation is the result of accumulation of lymphocytes, which become organized into a stable inflammatory infiltrate [95]. Lymphocytes are recruited from the circulation via communication with the endothelium. However, increased recruitment itself is not sufficient to generate chronic inflammation as powerful mechanisms have emerged to warrant that in most cases acute inflammation resolves and liver architecture reinstates once the original offense is dealt with [96]. In chronic inflammation, the resolution and healing mechanisms fail, triggering stable lymphocyte infiltration, tissue damage, and fibrosis. The severity and distribution of the lymphocytic infiltrate are therefore critical determinants for the outcome [97, 98].

Studies with mice deficient in B and T cells showing that liver fibrosis was absent suggest a critical role for lymphocytes in this pathology [99, 100]. Lymphocyte adhesion to endothelial cells is a requirement for recruitment into the liver and it entails confinement by the action of immunoglobulins [101], activation by chemokines present on the endothelial glycocalix [102], arrest on the vessel wall as a consequence of the high-affinity integrin interactions [103], and migration all the way through the endothelium into the tissue [97]. Lymphocytes can enter the liver through endothelial cells in the portal tract, the sinusoids, or the terminal hepatic veins [104]. The fenestrated sinusoidal endothelial cells do not form tight junctions and are deficient in basement membrane, being separated from the hepatocytes by the space of Disse, the scarce amount of loose connective tissue and HSCs that are adhered to the surface of endothelial cells. Endothelial cells express low levels of adhesion molecules such as selectins and CD31 [105]. Therefore, lymphocyte recruitment into the parenchyma through the sinusoids may be modulated by other molecules, that is, vascular adhesion protein-1 (VAP-1), intercellular adhesion molecule-1 (ICAM-1), and vascular cell adhesion molecule-1 (VCAM-1), compared with other vascular beds. VAP-1 is a source of oxidants in both NASH and ALD [106]. Recruitment via portal vascular endothelium could possibly also involve P-selectin. Neutrophils are recruited into inflamed liver simply if ICAM-1 and E-selectin are expressed [107]. Alcoholic hepatitis is linked with an increase in the expression of E-selectin and ICAM-1 on portal and hepatic venous endothelium and of ICAM-1, VCAM-1, and VAP-1 on sinusoidal endothelium as an effect of local proinflammatory cytokines such as TNFα [108–111]. In alcoholic cirrhosis, augmented expression of endothelial adhesion molecules including ICAM-1, VCAM-1, and P-selectin is largely restricted to portal and septal vessels [108]. Recruitment of leukocytes to the parenchyma is firmly regulated and depends on the correct combination of endothelial adhesion molecules and chemokines. Chemokines are secreted by leukocytes and endothelial cells with a significant contribution from hepatocytes, cholangiocytes, and fibroblasts [112–116]. T cells are decisive for the development of liver damage and fibrosis [117]. Both CD8+ T and CD4+ T cells are found in expanded portal tracts and within sinusoids and lobules in areas of parenchymal inflammation and fibrosis in steatohepatitis [118]. CD8 T cells are also found in the fibrous bands between nodules and at the lining plate of hepatocytes. ICAM-1 is up-regulated on the membrane of ballooned hepatocytes in areas of parenchymal inflammation related with elevated levels of lymphocyte function-associated antigen-1 (LFA-1) on infiltrating lymphocytes and confined TNFα secretion [108, 119], suggesting that lymphocytes are implicated in hepatocyte damage, liver wound, and fibrosis. Among the mechanisms thought to play an important role are the

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Fas-dependent apoptotic pathway [120] and the TRAIL (tumor necrosis factor-related apoptosis-inducing ligand) pathway that arbitrate hepatic apoptosis, steatosis and liver damage in response to alcohol [121], and these effects are increased by inhibiting NFκB activation [121]. The complex process of liver damage and repair associated with inflammation and scarring involves both autocrine and paracrine communication between stromal cells, extracellular matrix, leukocytes, and the secreted mediators (i.e. ROS, RNS, growth factors, cytokines, and chemokines). Although stellate cells are at center stage in the pathogenesis of liver fibrosis, macrophages, Kupffer cells, and lymphocytes are highly involved [122, 123]. Macrophages and Kupffer cells enhance the fibrogenic response by secreting pro-fibrogenic mediators such as ROS [122, 123], IL-6 [123], and TGF-β [124]. Neutrophils could act as effector cells in amplifying cell damage and scar formation [125]. The liver contains large levels of CXC chemokines which can attract, activate, and promote neutrophils extravasation along with an induction of ROS production in all liver cells. Alcoholic hepatitis is typified by augmented expression of numerous CXC and CC chemokines such as CXCL1, CXCL5, CXCL8, CCL2, CCL3, CCL4, and CCL5 [102]. The expression of chemokines and their receptors is not well characterized in NAFLD and NASH. CXCL8 (IL-8), which is increased in alcoholic hepatitis, is also elevated in a number of studies of patients with NASH [126–128], and LPS-induced induction of IL-8 and IL-6 from blood-derived monocytes was higher in NASH patients than in controls, suggesting that their monocytes were prepared for chemokine secretion [128]. Serum concentrations of CCL2 and the CCR7 ligand CCL19 are elevated in steatosis [129], and possibly more elevated in NASH patients [128]. Alcoholic cirrhosis involves increased expression of several chemokines in portal tracts and fibrous septa, where CCL2, CCL4, and CCL5 are all identified. CCL2 may be relevant in the development of fibrogenesis. It is secreted by both Kupffer cells and stellate cells and helps the recruitment of macrophages and lymphocytes to areas of active matrix deposition and collagen accumulation [106, 109]. CCL2-ablated mice do not develop liver injury and fibrosis, and CCL2 secretion by stellate cells is up-regulated by adipokines, indicating a close connection between steatosis and CCL2 [69, 76, 130, 131]. In chronic inflammatory liver disease, the extension of lymphocyte infiltration from the portal tracts into the parenchyma is associated with expression of the IFNγ-dependent CXCR3 ligands CXCL9, CXCL10, and CXCL11 in the sinusoids and hepatocytes [132]. These promote the recruitment of CXCR3hi effector cells through the sinusoidal endothelium [133]. Increased expression of CXCL10 is associated with accelerated progression of inflammatory liver disease in HCV infection, suggesting that CXCR3 is an important receptor in regulating chronic liver inflammation [134, 135].

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Translocation of gram negative bacteria, endotoxin, and TNFα Among the cytokines involved in the inflammatory response in ASH and NASH, TNFα is considered the most relevant and is probably the best studied to date. Elevated serum TNFα may be due to portal vein endotoxin from bacterial translocation due to alcohol-induced gut permeability, the so-called “leaky gut” [136]. Evidence points at Kupffer cells and infiltrating monocytes/macrophages as the major sources of TNFα [137, 138]. TNFα messenger RNA (mRNA) is increased in both the liver and the adipose tissue in NASH [139]. Soluble TNFα receptor is increased in steatosis and steatohepatitis [126, 127] and circulating monocytes from patients with NASH show enhanced TNFα secretion on stimulation when compared with healthy controls [140]. Along the same lines, TNFα receptor knockout mice present less severe steatosis than wild-type littermates [141], and anti-TNFα therapy results in a significant reduction in liver injury in steatohepatitis [41, 142, 143]. Even though chronic ethanol ingestion amplifies the Kupffer cell response to endotoxin [144], and makes hepatocytes vulnerable to TNFα [145], Kupffer cell activation by endotoxin may not be the sole mediator for the sustained hepatic inflammatory state, since constant exposure to endotoxin triggers tolerance [146], while chronic administration of ethanol along with endotoxin fails to elevate alcohol hepatotoxicity [147]. Furthermore, rats chronically fed alcohol show liver injury, inflammation, and increased TNFα mRNA even in the absence of substantial plasma endotoxin. TNFα and endotoxin may also drive the production of IL-12 and IL-15 in Kupffer cells, resulting in a local Th1 environment and the expansion of NKT cells [148]. TNFα is a potent inducer of chemokine secretion and adhesion molecule expression by endothelial cells, suggesting that high local levels of TNFα promote the recruitment of lymphocytes to the liver in steatohepatitis [149]. There are two types of plasma membrane receptors, TNFR1 and TNFR2. In the liver, TNFR1 prevails. Studies using TNFα receptor knockout mice indicate that TNFR1 is necessary for ethanol-induced liver injury [150]. TNFR1 is the prototype of a family of receptors to which the Fas receptor and the nerve growth factor receptor belong. During alcohol-induced liver injury, binding of TNFα to TNFR1 leads to receptor trimerization and activation, mediated by association of the receptor with a variety of signaling proteins that interact with specific protein domains on its intracellular tail [151]. So far, the best characterized signaling pathway activated by TNFα involves proteins that bind through the death domain, a protein domain found in a variety of signaling proteins involved in activation of the pro-apoptotic machinery. Proteins such as TNFα receptor-associated death domain protein (TRADD) and Fas-activated death

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domain protein (FADD) activate caspase 8, an upstream signaling caspase protein that begins the proteolytic activation of a variety of effector proteins, including other caspases, in addition to the proapoptotic Bcl-2 family protein Bid [152]. The active, truncated form of Bid (t-Bid) acts on mitochondria to cause mitochondrial outer membrane permeabilization. This allows the release of many proapoptotic factors located in the mitochondrial intermembrane space, which, along with the effector caspases, lead to nuclear chromatin condensation and degradation typical of apoptosis [153]. Binding of TNFα to TNFR1 also activates other signaling pathways that may advance cell death. Binding of a complex of receptor-interacting protein (RIP), a death domain serine–threonine kinase, and TNF receptorassociated factor 2 (TRAF2), an adaptor protein that interacts with the activated receptor, causes activation of sphingomyelinases and ceramides synthesis which act on mitochondria to promote mitochondrial permeability transition (MPT), a process that involves opening of a large pore across both inner and outer membranes [154]. This is related to loss of mitochondrial energization, matrix swelling, and rupture of the outer membrane, along with the release of numerous proapoptotic factors. TNFα activates stress-activated protein kinase cascades in liver cells, resulting in the stimulation of p38 mitogen-activated protein kinase (MAPK) and Jun N-terminal kinase (JNK) [155]. Both affect the response of mitochondria to proapoptotic stimuli: JNK by phosphorylation of antiapoptotic proteins such as Bcl-2 and Bcl-XL, and p38 MAPK by increasing the effects of the pro-apoptotic Bcl-2 family protein Bax. Simultaneously, TNFR1 activation begins cytoprotective responses in hepatocytes via other signaling cascades. Of relevance is the activation of NFκB, through a pathway that involves TRAF2 and RIP, activating a protein kinase cascade that ends in the phosphorylation and degradation of the NFκB inhibitory protein IκB [155]. This enables NFκB to translocate into the nucleus and transactivate selected genes, such as inhibitor of apoptosis (IAP) 1 and 2, whose products block the activation of caspases through the death pathway. Thus, these protective actions are prevented when gene transcription or protein synthesis is inhibited. Additional protection occurs through a TNFR1stimulated cascade that triggers activation of PI3K, which activates Akt/PKB. The Akt/PKB phosphorylates and inactivates Bad, another proapoptotic protein of the Bcl-2 family that acts through its interaction with mitochondria [156]. Hence, the initiation of the apoptotic program depends on the appropriate balance of a variety of signaling cascades, and the balance of downstream actions of these cascades seems to target the mitochondria, which acts as a master switch for the apoptotic machinery. Ethanol preconditions hepatocytes to alter the equilibrium of cytoprotective and cytotoxic actions set off by TNFα (or other cytokines), thus impairing the suitable

response of the cell to toxic challenges. The pathways involved in TNFα-mediated cell death that are active in ethanol-treated hepatocytes seem to circumvent the classical death domain signaling cascade [157]. This is compatible with a role for other TNFα-induced cytotoxic signaling pathways (e.g. TNFα-induced ceramide) [158]. The cytotoxicity associated with TNFα treatment of hepatocytes obtained from ethanol-fed animals involves the early loss of mitochondrial function, as confirmed by mitochondrial depolarization and the release of cytochrome c from the mitochondria into the cytosol. The decline in the mitochondrial membrane potential is owed to the MPT, and inhibition of this process with cyclosporin A prevents the TNFα-induced cell death [157]. The biological effect of endotoxin is mediated through specific receptors in Kupffer cells, and stellate cells. Endotoxin binds to serum lipopolysaccharide binding protein (LBP), which interacts with the CD14 receptor [159], resulting in the translocation of NFκB to the nucleus, which in turn, transactivates the promoter of several cytokines and chemokines. The fact that endotoxin activates NFκB in the absence of CD14 implies a CD14-independent mechanism for Kupffer cell activation [159]. This is mediated through a CD11c/CD18 adhesion molecule that also serves as a transmembrane signaling receptor for endotoxin [160]. This notion is supported by studies showing that anti-CD18 antibody stimulates Kupffer cells to enhance free radical generation [161]. In addition to the deleterious effects of alcohol on the liver microcirculation, endotoxin disturbs hepatic microcirculation regulation [162]. The microcirculation disruption seen under both stress conditions is characterized by a hypersensitivity to the constrictor effects of endothelin-1, a powerful vasoconstrictor expressed by endotoxin exposure and also chronic alcohol consumption [163, 164]. Intravital microscopic assessment of the liver microcirculation of alcohol-fed rats treated with LPS or saline suggests that chronic alcohol consumption can sensitize the liver microcirculation to the effects of LPS, leading to synergistic exacerbation of the disruption [165, 166]. A study conducted by Horie and co-workers [165] indicated that alcohol-mediated blood flow disruption and sensitization to endotoxin were dependent on the effects of endothelin-1.

GENERATION OF ROS IN ALD Sources of ROS Oxidative stress plays a critical role in the pathogenesis of ALD. In hepatocytes, ethanol-induced ROS generation occurs at different sub cellular levels such as the mitochondria, the ER where cytochrome P450 is located, and the cytosol where free iron accumulates and enzymes such as xanthine oxidase and aldehyde oxidases are present. In

46: PATHOPHYSIOLOGY OF ALCOHOLIC LIVER DISEASE

addition, active enzymatic systems in neighboring cells such as Kupffer cell and stellate cell NADPH oxidase and the neutrophil myeloperoxidase also contribute to oxidative injury in ALD. The formation of key ROS such as O2• − and H2 O2 is an important source of oxidative injury in many diseases associated with free radical formation. In the presence of trace amounts of transition metals (most frequently iron), O2• − and H2 O2 will gen• erate highly-reactive hydroxyl radicals (HO ), which are then responsible of further oxidation reactions and will also enhance lipid peroxidation end products. Moreover, in hepatocytes, lipid rafts, when modified by oxidative reactions, are involved in the development of oxidative stress due to ethanol, as characterized by both ROS production and lipid peroxidation [166]. The participation of membrane fluidization for lipid raft clustering could clarify how membrane fluidity can increase ethanol-induced oxidant stress. Sergent et al. [167] postulated that the contribution of membrane fluidity in ethanol-induced oxidative stress takes place via a mechanism whereby ethanol metabolism, by triggering early ROS generation, quickly increases membrane fluidity, which eventually enhances ROS production and the subsequent lipid peroxidation.

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proteins interacting with mitochondria could prevent the TNFα-induced onset of the MPT in hepatocytes obtained from ethanol-fed rats. Hence the changes in susceptibility to TNFα-cytotoxic signals probably reflect multiple levels of adaptive (or non-adaptive) effects of chronic ethanol treatment.

NADPH oxidase NADPH oxidase is a complex enzyme composed of two membrane-bound subunits, three subunits in the cytosol, plus Rac 1 or Rac 2. Activation of the oxidase involves the phosphorylation of one of the cytosolic subunits. Crystallographic data indicate that the tail of this cytosolic subunit is localized in a groove between two Src homology 3 domains and, when phosphorylated, the tail leaves the groove and is substituted by the tail of one of the membrane-bound subunits. NADPH oxidase generates O2• − by transferring electrons from NADPH within the cell across the membrane, and coupling these to O2 to make O2• − and eventually generates H2 O2 and hypochlorous acid: NADPH + 2O2 ⇋ NADP+ + 2O2• − + H+

Mitochondrial damage Deterioration of liver mitochondrial function after chronic ethanol treatment has been well documented [168]. Mitochondria obtained from ethanol-fed rats are more susceptible to increase the permeability transition in response to a wide variety of challenges [169]. Thus, the effects of ethanol may be, at least in part, the result of its effects on mitochondria. Under chronic alcohol consumption, oxidative phosphorylation is impaired because of a defect in the synthesis of proteins encoded in the mitochondrial DNA. Even if this deficiency is not usually translated into limiting the supply of energy in the liver, and ATP levels are sustained under normal conditions, it may be harder for the cell to respond appropriately to stress conditions that place high demands on the mitochondrial bioenergetic capacity. Ethanol also provokes changes in mitochondrial membrane structure [170] which are associated with ineffective mitochondrial uptake of glutathione (GSH) via the mitochondrial GSH carrier, and the subsequent decline in the mitochondrial antioxidant defense [171]. This not only contributes to liver injury but could also further enhance the onset of the MPT under oxidative challenge [172]. The effects of chronic ethanol consumption on the mitochondria also comprise changes in the upstream signaling pathways that arbitrate the defensive or harmful effects of TNFα. The MPT is coordinated by a spectrum of pro-apoptotic and anti-apoptotic messengers that influence the permeability transition pore complex. Studies indicate that inhibition of MAPK cascades that control the equilibrium of pro-apoptotic and anti-apoptotic

Oxidant stress in Kupffer cells is one mechanism by which free radicals, formed as a result of alcohol consumption, leads to liver damage [173]. Kono et al. [174] demonstrated that reduced NADPH oxidase is a major source of free radicals in Kupffer cells after ethanol administration using P47Phos knockout mice, which lack the critical subunit for NADPH oxidase. In mice treated with ethanol for 4 weeks, fatty liver, necrosis, and inflammation appeared in association with the generation of electron spin resonance-detectable free radicals, activation of NFκB, and higher TNFα levels. In NADPH oxidase-deficient mice, pathological changes in the liver due to ethanol were abrogated [174]. The improvement of these pathological events was accompanied by a down-regulation in free radical generation, NFκB activation, and TNFα synthesis. These results confirmed a role for Kupffer cell-derived free radicals in the pathogenesis of alcoholic liver injury. This information is consistent with previous work indicating that LPS activates oxidant production in vitro via NADPH oxidase [175–177], which leads to NFκB activation [178] and TNFα production [179]. Notably, mice deficient in the endotoxin receptor CD14, which is primarily expressed on Kupffer cells in liver [180, 181], were also resistant to ethanol-induced liver damage [182]. Mice with a mutation in the Toll-like receptor-4, which makes them resistant to endotoxin, also did not display ethanol-derived pathology [183]. Both activation of NFκB and synthesis of TNFα by ethanol were ameliorated under these conditions. These data are in agreement with the hypothesis that endotoxin activates Kupffer cells through the endotoxin receptor CD14–Toll-like receptor 4 complex to produce toxic free radicals via NADPH oxidase [176].

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Oxidants activate NFκB, increasing the production of pro-inflammatory cytokines, leading to events involving neutrophil adhesion, mitochondrial dysfunction, and eventually tissue damage [178, 183, 184]. Both NADPH oxidase and iNOS knockout mice are protected completely against alcohol-induced oxidative stress, suggesting that O2• − and NO• are involved in alcohol-induced liver injury [173, 185]. Ethanol increases Ca2+ entry and modifies membrane phosphatidylinositol turnover and arachidonic acid metabolism leading to the release of secondary messengers that, as a result, fuel the translocation and activation of protein kinase C [186, 187], which in turn induces the translocation of NADPH oxidase to the plasma membrane to generate O2• − . ROS, particularly H2 O2 , generated through the dismutation of O2• − by superoxide dismutase (SOD) and other reactions, serve as secondary messengers for the translocation and activation of nuclear transcription factors leading to enhanced gene expression for a number of cytokines and chemokines. In hepatic stellate cells, NADPH oxidase appears to mediate the actions of angiotensin II on collagen accumulation [188], plays a crucial role in PDGF-induced proliferation of hepatic stellate cells [189], and phagocytosis of apoptotic bodies by hepatic stellate cells induces NADPH oxidase and is associated with liver fibrosis in vivo [190].

Xanthine oxidase Xanthine oxidoreductase is an enzyme that has the unusual property of existing both in a dehydrogenase form, which uses NAD+ , and in an oxidase form, which uses O2 as electron acceptor. Both forms have a high affinity for hypoxanthine and xanthine as substrates. In addition, conversion of one form to the other may occur under selected conditions. This enzyme seems to play a role in purine catabolism, detoxification of xenobiotics, and antioxidant capacity by producing urate. The oxidase form produces ROS and, therefore, the enzyme is involved in various pathological processes such as tissue injury due to ischemia followed by reperfusion, and in ALD. hypoxanthine + O2 + H2 O ⇋ xanthine + H2 O2 xanthine + O2 + H2 O ⇋ uric acid + H2 O2 Xanthine oxidase is widely distributed enzyme, and it is involved in the metabolism of purines which generates O2• − and it is also involved in free radical-generated tissue damage. It is present at high concentration in the liver, from where it may be released during liver injury into the circulation, binding to vascular endothelium and causing vascular dysfunction. Xanthine oxidase is present in hepatocytes and in bile duct epithelial cells, where it is concentrated towards the luminal surface [191]. Moreover, in liver disease, such as ALD, proliferating

bile ducts are also strongly positive for xanthine oxidase suggesting that the enzyme is secreted into bile [191]. Xanthine oxidase activity was 10–20-fold higher in liver tissue obtained from patients with liver disease than in healthy liver [191]. Its role in bile is unknown, but it may be involved in innate immunity of the bowel mucosa. Administration of enteral ethanol for 4 weeks increased serum transaminases, caused severe fatty infiltration, mild inflammation, necrosis, and increased NFκB binding. Allopurinol, a xanthine oxidase inhibitor and scavenger of free radicals, blunted these effects [192]. Furthermore, enteral ethanol caused free radical adduct formation, values that were reduced by approximately 40% with allopurinol [192]. These results indicate that allopurinol prevents early alcohol-induced liver injury, most likely by preventing oxidant-dependent activation of NFκB [192].

Alcohol metabolism by CYP2E1 Hepatocytes metabolize ethanol through three enzymes localized in different subcellular compartments: the cytosolic ADH, the microsomal CYP2E1, and the peroxysomal catalase. Metabolism of ethanol by any of these systems generates acetaldehyde, which can cause liver injury by impairment of the mitochondrial β-oxidation of fatty acids, GSH depletion, lipid peroxidation, ROS burst, and generation of acetaldehyde adducts [193]. Acetaldehyde is quickly metabolized to acetate by aldehyde dehydrogenase 2. While most ethanol is oxidized by ADH, CYP2E1 plays a major role at elevated concentrations of ethanol when ADH reaches saturation and after chronic alcohol consumption [193]. Metabolism of alcohol through CYP2E1 causes a hypermetabolic state in hepatocytes, triggering injury in the low-oxygen perivenular zone of the hepatic lobule [194]. Induction of CYP2E1 by ethanol additionally amplifies oxidative stress, which is stressed by acetaldehyde, which consumes antioxidants and generates extra ROS when oxidized [194]. Thus, the redox shift related to ethanol oxidation by ADH and the activation of CYP2E1 by ethanol led to increased ROS [193, 194]. In humans, CYP2E1 induction occurs not only in alcohol abusers [195], but also in moderate alcohol consumers [196]. CYP2E1-dependent monooxygenase activity is amplified several-fold in alcohol-fed mice. CYP2E1 has a high rate of NADPH oxidase activity, leading to the generation of large amounts O2• − and H2 O2 . In liver microsomes from both alcoholics and alcohol-fed rodents the CYP2E1 content correlates with NADPH oxidase activity and formation of lipid peroxidation end products. Increased generation of ROS occurs in HepG2 cells stably transfected with CYP2E1 compared with control cells [197]. Thus, the high effectiveness of CYP2E1 in reducing O2 to O2• − and H2 O2 could be considered as a key factor contributing to oxidative stress during chronic alcohol consumption.

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Experiments in rats fed alcohol [198–200] have confirmed that the induction of CYP2E1 by ethanol is related to the appearance of lipid peroxidation-derived products. However, addition of CYP2E1 inhibitors decreases oxidative stress and liver injury. These findings are in partial disparity with the observation that CYP2E1 ablation in mice does not protect from alcohol toxicity [201]. This discrepancy can be clarified, since in ethanol-fed mice CYP2E1 represents 50 mg of alcohol per day in men and 40 mg per day in women) in patients with CHC is strongly associated with progression to cirrhosis and an increased risk of HCC in both cohort- and population-based studies [118, 165, 166]. A recent meta-analysis of 20 studies of more than 15 000 patients estimated a pooled RR of cirrhosis associated with heavy alcohol intake (defined as in the range of at least 210–560 g per week) to be 2.33 (95% CI, 1.67 to 3.26) [167]. The evidence overwhelmingly showed a worsened outcome for those with chronic HCV and concurrent alcohol use. The studies examined varied widely in their definition of significant alcohol intake, and so the true threshold above which alcohol accelerates HCV disease progression remains uncertain. Whether consumption of lower amounts of alcohol is also associated with faster rates of fibrosis progression has not been adequately studied and the issue remains unresolved.

Cannabis/cigarette smoking Daily cannabis use has been shown to be independently associated with severe fibrosis in patients with CHC [168]. This effect could be mediated via activation of the CB1 receptor, the receptor for marijuana, since up-regulation of CB1 receptor has been demonstrated in experimental models of fibrogenesis [169]. Several epidemiological studies have suggested a link between cigarette smoking and severity of liver disease in patients with CHC [170, 171]. In a cross-sectional study of patients with CHC admitted for a first liver biopsy, smoking (>15 packs per year), was found to be independently associated with increased fibrosis and necroinflam-

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mation [172]. Smoking was strongly related to other risk factors, such as gender, alcohol consumption, and history of intravenous drug use. The relationship between fibrosis and smoking disappeared when the Knodell activity score was included in the multivariate analysis, suggesting that the pro-fibrotic effect of smoking may be related to increased activity of liver disease.

Herbal preparations There is no evidence that herbal preparations are associated with a worse outcome of HCV infection or that they are protective. Caffeine was demonstrated in population-based studies to be associated with lower serum ALT levels [173]. It is unknown whether caffeine may have a protective role in persons with CHC.

HEPATITIS C AND HEPATOCELLULAR CARCINOMA Worldwide, ∼25% of all HCCs are related to CHC [174]. In the United States, ∼9% of all HCCs are related to HCV infection, although this is predicted to double in the next two decades [175]. Consistent with this, as many as 42% of HCC tissues were found to contain HCV nucleic acid in one recent US series [176]. In persons with HCC, rates of anti-HCV positivity are highest in Japan, Spain, and Italy, intermediate in the United States, but low in countries where HBV is endemic [177]. The geographic variation reflects the prevalence of the two main risk factors for HCC: HBV and HCV infection. There is strong epidemiological and experimental data linking HCV with the development of HCC [140, 178, 179]. Markers of HCV infection, either anti-HCV or HCV RNA, can be detected in the serum and liver tissue from patients with HCC. In a meta-analysis of 21 case–control studies, the risk for HCC was increased 17.3-fold in anti-HCV positive compared with anti-HCV negative controls [180]. Several HCV proteins may be directly oncogenic. At least four HCV proteins, core, NS3, NS4B, and NS5A, have been shown to exhibit transformation potential in vitro. Transgenic mouse models expressing HCV core protein under control of HBV transcriptional elements or the entire viral polyprotein under control of the murine albumin promoter have been associated with HCV steatosis and the development of HCC in male mice [181, 182]. These tumors developed in the absence of inflammation, suggesting a direct oncogenic effect of the HCV transgene. Several HCV proteins interact with and may disable the function of important tumor suppressor proteins such as p53 or the retinoblastoma protein [183]. HCV may also promote the development of HCC via the induction of reactive oxygen species and oxidative stress [184].

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Risk factors that have been identified for HCC in HCV-positive individuals include male sex, older age, HBV co-infection, heavy alcohol use, and diabetes [140]. In one study that examined the contribution of HBV and HCV to the development of cancer, co-infection with HBV significantly increased the risk for HCC. For anti-HCV-positive persons, the RR of HCC was 21.3 (95% CI, 8.8 to 51.5) and for HBsAg-positive persons the RR of HCC was 13.3 (95% CI 5.5 to 32.2). However, when both anti-HCV and HBsAg were present, the RR increased to 77.0 [185]. HCC is usually the final stage in the orderly sequence of the natural history of CHC. In a natural history study of transfusion-related CHC, the period between transfusion and the diagnosis of HCC was 33–45 years [125]. Rates of HCC are also significantly higher in patients with cirrhosis than without. In fact, HCC has never been reported to occur in the absence of fibrosis in patients with CHC. However, once cirrhosis is established, the risk of developing HCC is approximately 1–4% per year [128, 178]. It is presumed that chronic liver injury, leading to repeated bouts of regeneration and proliferation, results in the accumulation of mutations which ultimately lead to malignant transformation. Whether IFN therapy can prevent the development of HCC in patients with CHC remains an unanswered question. A small prospective study in Japan randomized 90 patients with compensated HCV-related cirrhosis to either IFN or no therapy for 24 weeks and followed the patients for 2–7 years [186]. During a mean follow-up of ∼5 years, the incidence of HCC in the treated group was 4% and in untreated controls 38%. This provocative trial suggested an undeniable benefit from IFN in the prevention of HCC. However, the rate of HCC in the untreated controls was unexpectedly high and questions were raised about how such a short course of IFN could lead to such a dramatic reduction in HCC rate in the absence of a sustained viral response (SVR). In another prospective study in Japan involving 25 clinical centers, 271 patients received (6–9) ×106 U of IFN three times weekly for 26–88 weeks and 74 received no treatment [187]. All were monitored with α-fetoprotein (AFP) and abdominal ultrasonography to detect HCC. HCC was detected in 119 patients during a mean follow-up period of 6.8 years: 84 (31%) in the IFN-treated group and 35 (47%) in the untreated group. The cumulative incidence of HCC among IFN-treated patients was significantly lower than in untreated patients [Cox model: age-adjusted HR, 0.65 (95% CI, 0.43 to 0.97); p = 0.03], especially in sustained virological responders. However, this was not a randomized, controlled study, which may have biased the results. Patients enrolled in the control group had declined to receive IFN treatment even though they were eligible for treatment. Other large studies in Japan reported similar findings [188–190]. However, studies in Europe and the United States have not found a significant difference in the rate of HCC development in non-responder

patients and untreated controls [191–193]. A metaanalysis in Italy demonstrated minimal benefit of IFN therapy in HCV-related cirrhosis and prevention of HCC [194]. The wide variation in methodology among studies precluded a definite conclusion as to the benefits of IFN. However, results of the recent Hepatitis C Antiviral Long-term Treatment against Cirrhosis (HALT-C) study suggest that continued IFN therapy is unable to prevent the development of HCC in non-responder patients with CHC [195]. Successful eradication of HCV in patients with cirrhosis significantly reduces but does not completely eliminate the risk of developing HCC [196]. In a retrospective analysis, 920 Italian patients with cirrhosis who received IFN monotherapy were assessed for long-term outcomes of HCV infection [197]. An SVR was achieved in 124 patients. A total of 129 HCCs were detected during a mean follow-up period of 8 years, seven in patients with an SVR and 122 in non-responders. The incidence rates of HCC per 100 person-years of follow-up were 0.66 (95% CI, 0.27 to 1.37) for SVR and 2.10 (95% CI, 1.75 to 2.51) for non-responder patients. Non-responders had a 2.59-fold higher rate of HCC than patients with an SVR. Similar reductions in the rate of HCC in cirrhotic patients following viral clearance have been reported [198]. In summary, chronic HCV infection results in a chronic inflammatory process in the liver with the deposition of collagen within the extracelluar matrix. Over a variable period of time, the accumulation of collagen leads to distortion of the hepatic architecture with impairment of the hepatic microcirculation and hepatocyte dysfunction. Progression of hepatic fibrosis thus closely mimics the natural history of CHC. In general, this progression is slow and takes decades to develop, although it may be accelerated by host and external factors, the most significant of which include older age, male sex, and alcohol consumption. Patients with CHC should be counseled to modify known risk factors and monitored for progression of disease. Those persons identified to have progressive liver disease (bridging fibrosis on liver biopsy) should be considered for therapy. Achieving an SVR is associated with significantly lower rates of clinical outcomes (hepatic decompensation, HCC, and death) even in those with cirrhosis.

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(2007) The natural history of hepatitis C with severe hepatic fibrosis. J Hepatol , 47, 37–45. Planas, R., Balleste, B., Alvarez, M.A., Rivera, M., Montoliu, S., Galeras, J.A., Santos, J., Coll, S., Morillas, R.M. and Sola, R. (2004) Natural history of decompensated hepatitis C virus-related cirrhosis. A study of 200 patients. J Hepatol , 40, 823–30. Shimizu, I. (2003) Impact of oestrogens on the progression of liver disease. Liver Int , 23, 63–69. Di Martino, V., Lebray, P., Myers, R.P., Pannier, E., Paradis, V., Charlotte, F., Moussalli, J., Thabut, D., Buffet, C. and Poynard, T. (2004) Progression of liver fibrosis in women infected with hepatitis C: long-term benefit of estrogen exposure. Hepatology, 40, 1426–33. Codes, L., Asselah, T., Cazals-Hatem, D., Tubach, F., Vidaud, D., Parana, R., Bedossa, P., Valla, D. and Marcellin, P. (2007) Liver fibrosis in women with chronic hepatitis C: evidence for the negative role of the menopause and steatosis and the potential benefit of hormone replacement therapy. Gut , 56, 390–95. Wiley, T.E., Brown, J. and Chan, J. (2002) Hepatitis C infection in African Americans: its natural history and histological progression. Am J Gastroenterol , 97, 700–6. Crosse, K., Umeadi, O.G., Anania, F.A., Laurin, J., Papadimitriou, J., Drachenberg, C. and Howell, C.D. (2004) Racial differences in liver inflammation and fibrosis related to chronic hepatitis C. Clin Gastroenterol Hepatol , 2, 463–68. Sterling, R.K., Stravitz, R.T., Luketic, V.A., Sanyal, A.J., Contos, M.J., Mills, A.S. and Shiffman, M.L. (2004) A comparison of the spectrum of chronic hepatitis C virus between Caucasians and African Americans. Clin Gastroenterol Hepatol , 2, 469–73. El-Serag, H.B. (2002) Hepatocellular carcinoma and hepatitis C in the United States. Hepatology, 36, S74–83. Adinolfi, L.E., Gambardella, M., Andreana, A., Tripodi, M.F., Utili, R. and Ruggiero, G. (2001) Steatosis accelerates the progression of liver damage of chronic hepatitis C patients and correlates with specific HCV genotype and visceral obesity. Hepatology, 33, 1358–64. Leandro, G., Mangia, A., Hui, J., Fabris, P., RubbiaBrandt, L., Colloredo, G., Adinolfi, L.E., Asselah, T., Jonsson, J.R., Smedile, A., Terrault, N., Pazienza, V., Giordani, M.T., Giostra, E., Sonzogni, A., Ruggiero, G., Marcellin, P., Powell, E.E., George, J. and Negro, F. (2006) Relationship between steatosis, inflammation, and fibrosis in chronic hepatitis C: a meta-analysis of individual patient data. Gastroenterology, 130, 1636–42. Ramesh, S. and Sanyal, A.J. (2004) Hepatitis C and nonalcoholic fatty liver disease. Semin Liver Dis, 24, 399–413. Fartoux, L., Chazouilleres, O., Wendum, D., Poupon, R. and Serfaty, L. (2005) Impact of steatosis on progression of fibrosis in patients with mild hepatitis C. Hepatology, 41, 82–87. Castera, L., Hezode, C., Roudot-Thoraval, F., Bastie, A., Zafrani, E.S., Pawlotsky, J.M. and Dhumeaux, D. (2003) Worsening of steatosis is an independent factor

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54

Current and Future Therapy for Hepatitis B and C Gary L. Davis1 and Jean-Michel Pawlotsky2 1

Department of Hepatology, Baylor University Medical Center, Dallas, TX, USA 2 Department of Virology and INSERM U955, Henri Mondor Hospital, University of Paris 12, Cr´eteil, France

HEPATITIS B VIRUS INFECTION General principles of treatment The immediate aims of antiviral treatment of chronic hepatitis B are to achieve sustained suppression of hepatitis B virus (HBV) replication and remission of liver disease. The markers of such a treatment response include normalization of the serum alanine aminotransferase (ALT) level, decrease of HBV DNA, loss of HBeAg in patients in whom it was initially present, and improvement of hepatic histology. The long-term goal is to prevent, or at least reduce, the likelihood of progressing to complications of liver disease such as cirrhosis, hepatic failure, or hepatocellular carcinoma. Thus, the initial assessment of the patient with chronic hepatitis B must include serum biochemistry testing (ALT, albumin, bilirubin, prothrombin time), virological testing [hepatitis B surface antigen (HBsAg), hepatitis B e antigen (HBeAg), antibody to HBeAg (anti-HBe or HBeAb)], HBV DNA [by a sensitive molecular assay with a wide dynamic range, such as real-time polymerase chain reaction), antibody to hepatitis A virus (anti-HAV), and antibody to hepatitis delta virus (anti-HDV)]. A history and physical examination should be performed with special attention to risk factors for HBV infection and family history of infection or The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

hepatocellular carcinoma. In patients with relevant risk factors, testing should then also include antibody to hepatitis C virus (anti-HCV) and antibody to the human immunodeficiency virus (anti-HIV). Finally, surveillance for hepatocellular carcinoma should be made at the time of diagnosis and serially when appropriate [1]. Several practice guidelines have been published to assist the evaluation and treatment of patients with chronic hepatitis B (Table 54.1) [2–4]. Nonetheless, interpretation of testing can be confusing. For example, while low levels of HBV DNA [2 × ULN

Normal

1–2 × ULN

>2 × ULN

HBV DNA (IU ml−1 ) Liver biopsy

Any level

Any level

Any level

20 000

Consider if borderline ALT or age >40 years

Consider if elevated ALT persists or age >40 years Q3–6m Treatment should be initiated if moderate inflammation or fibrosis is present

Optional

Consider if borderline ALT or age >40 years

Optional

Q3–6m Treat if ALT elevation persists or any evidence of decompensation

Q3–6m Treatment not required unless liver biopsy demonstrates active disease

Consider if elevated ALT persists or age >40 years Q3–6m Treatment should be initiated if moderate inflammation or fibrosis is present

Monitoring Q3–6m Considerations Treatment not required unless liver biopsy demonstrates active disease

Q3-6m Treat if ALT elevation persists or any evidence of decompensation

a ULN = upper limit of the normal range, defined as ALT of 30 IU ml−1 for men and 19 IU ml−1 for women. b Adapted from the American Association for the Study of Liver Diseases Practice Guidelines [1].

availability, and cost. The following discussion describes the available drugs and considerations for their use in detail.

This genome encodes four overlapping reading frames and four RNA species whose sequences encodes for seven viral proteins, including three envelope proteins that comprise the heterogeneous HBsAg, the secretory HBeAg, the X protein, and a multifunctional viral enzyme that has reverse transcriptase, DNA polymerase, and ribonuclease (RNase) activities. Replication of HBV occurs mostly in hepatocytes, although some have suggested that other cells may also support replication in vivo. HBV replication is summarized in Figure 54.1. The virus attaches via a yet

Molecular and pathophysiological basis for therapy The HBV is a small enveloped encapsidated DNA virus with a 3.2 kb partially double-stranded genome [6, 7].

Cell binding and entry

HBsAg export

New virion export

Uncoating Assembly

Translation

DNA repair (DNA polymerase)

Pre-S1 Pre-S2 S

2.4 & 2.1 kb mRNA

Positive DNA strand synthesis (DNA polymerase)

cccDNA 3.5 kb mRNA

Pregenomic RNA degraded (RNAse H)

Translation Pregenomic RNA HBV polymerase Core

Negative DNA strand synthesis (reverse transcriptase)

Encapsidation

Figure 54.1 Schematic of the hepatitis B virus replication cycle in the hepatocyte. Adapted from Ghany M, Liang TJ. Drug targets and molecular mechanisms of drug resistance in chronic hepatitis B. Gastroenterology, 2007, 132,1574–1585.

54: CURRENT AND FUTURE THERAPY FOR HEPATITIS B AND C

unidentified receptor complex at the cell surface, but the precise mechanism of binding is not known. Upon entry and uncoating, the circular DNA genome homes to the nucleus where host and viral polymerases repair (close) the open portion of the double-stranded genome to form the fully double-stranded covalent closed circular deoxyribonucleic acid (cccDNA) that serves as the template for transcription of three (maybe four) viral messenger ribonucleic acids (mRNAs). These RNAs are transported to the cytoplasm where they serve as templates for translation of the HBV proteins, including polymerase, core/HBe protein, the envelope proteins, and the X protein, respectively. The pregenomic RNA, which is 3.5 kb long, serves as a template for genomic DNA synthesis through reverse transcription. Assembly of the nucleocapsid involves polymerase-dependent binding of the pregenomic RNA with the core protein which forms the structure within which the pregenomic RNA serves as the template for first negative strand synthesis and then partial positive strand synthesis and circularization. In the meantime, nucleocapsids bud into the endoplasmic reticulum membrane that contains newly translated envelope proteins and form new virions that are exported out of the cell. Finally, newly synthesized DNA-containing nucleocapsids may also move back to the nucleus in order to maintain the small cccDNA pool required to sustain replication in the infected cell (5–50 copies per infected cell). It is apparent from the previous discussion that the essential and multifunctional viral polymerase is the most obvious and arguably the most efficient target of antiviral therapies. Nucleoside and nucleotide analogs that target the reverse transcriptase function of the polymerase (Figure 54.1) are currently the major class of agents licensed and under development for the treatment of HBV infection. Specific targets, mechanisms of action, and issues with these agents are discussed in detail below. Several other potential targets for therapy are possible, including cell uptake or binding inhibitors, cell assembly and glycosylation inhibitors, immune-enhancing drugs or vaccines, and inflammation modulators. In general, these classes of agents are either theoretical or early in preclinical development and so will not be discussed in this chapter.

Current agents HBV polymerase/reverse transcriptase inhibitors The HBV polymerase carries out the primary enzymatic functions required for viral replication, including the RNA-dependent DNA polymerase (i.e. reverse transcriptase) activity, DNA polymerase activity, and RNase H activity (see above). Given its critical role in replication, it is not surprising that it has been the major target for the development of antiviral agents. The nucleoside/nucleotide

901

analogs are synthetic compounds that are similar enough to endogenous nucleosides to be incorporated into growing DNA strands, but different enough to ensure that the resultant DNA is non-functional, usually as a result of reverse transcriptase inhibition and termination of chain elongation. Nucleoside/nucleotide analogs require sequential phosphorylation by cellular kinases to their nucleoside triphosphates (three phosphorylations in the case of nucleoside analogs, two in the case of nucleotide analogs such as adefovir and tenofovir) before being capable of binding to and inhibiting the viral polymerase [7]. The activated forms (dNTPs) bind to active sites in palm portion of the polymerase near the 3′ terminus of the primer strand and thereby inhibit normal enzyme function. The drugs then inhibit the HBV polymerase and prevent chain elongation by one of several mechanisms, including impairing base priming, impairing reverse transcription of the negative strand from the pregenomic RNA, or inhibiting synthesis of the positive strand of HBV DNA (Figure 54.1). The agents currently approved for clinical use and most under development are nucleoside/nucleotide analog polymerase inhibitors that fall into three general classes. (i) The HBV polymerase has a preference for l-nucleoside, in contrast to most others that prefer the d-configuration. Thus, many of the HBV polymerase inhibitors are l-nucleoside analogs, including lamivudine, its 5-fluoro derivative emtricitabine, telbivudine (l-thymidine), and clevudine (l-FMAU). These are all pyrimidine nucleoside analogs. Lamivudine and emtricitabine are obligate chain terminators. (ii) The acyclic nucleoside phosphonates adefovir and tenofovir have a stable phosphate-mimetic group which allows them to bypass the rate-limiting initial phosphorylation step (nucleotide analogs). They have high affinity for viral DNA polymerases and impair chain elongation. (iii) Finally, the cyclopentanes are deoxyguanosine analogs in the natural d-configuration. Entecavir falls into this group. The characteristics of these agents and the clinical outcomes of treatment are listed in Table 54.2 and discussed below. Generally, the HBV polymerase inhibitors must be used for a prolonged and indefinite period of time in order to suppress chronically virus replication below levels associated with progressive liver disease. Although clinically well tolerated, the requirement for long-term administration makes viral resistance more likely, particularly when virus suppression is incomplete or delayed. The hepatitis B virus is prone to develop drug resistance since it has a high rate of replication (1012 virions per day) and the HBV reverse transcriptase lacks proof-reading capability [7]. The genesis of viable mutations may be somewhat constrained because of the overlapping reading frames of the genome. Nonetheless, drug-resistant viral variants likely exist before initiation of drug therapy. The probability of a mutation being selected during therapy relates to the ability of the drug to suppress virus, the degree of drug resistance that a particular mutation confers, the fitness of the resistant variant, and the susceptibility of the

l-Nucleoside l-Nucleoside l-Nucleoside Acyclic nucleotide Acyclic nucleotide d-Cyclopentane

Lamivudine Emtricitabine Telbivudine Adefovir

0.5–1.0

300

100 —b 600 10

67

76

39 39 60 21

HBV DNA undetectable (%)

22

21

22 14 26 24

HBe seroconversion (%)

90

93

72 79 88 51

HBV DNA undetectable (%)

0

0

24 13 10 0

HBeAg negative 1 year (%)

1.2

No data

67 No data No data 29

Drug resistance (treatment na¨ıve) 5 year (%)

Yes

Yes

Yes Yes Yes Yes

Renal dose adjustment

C

B

B B B C

Use in pregnancy (categorya )

a Category A indicates no risk of fetal harm, category B indicates no proven risk in animal studies but no human data is available, category C indicates animal data showing fetal risk, but no human data. b Not used as a single agent; available in combination with tenofovir, but does not have FDA approval for use in hepatitis B.

Entecavir

Tenofovir

Class

Agent

Dose (mg qd)

HBeAg positive

Table 54.2 Characteristics of and clinical responses to nucleoside/nucleotide analogs that inhibit hepatitis B viral polymerase function [8, 9]

902 THE LIVER: HEPATITIS B VIRUS INFECTION

54: CURRENT AND FUTURE THERAPY FOR HEPATITIS B AND C

903

Table 54.3 Drug-resistant mutations by major drug and therapeutic alternatives [10, 14] Agent

Primary resistance mutations

Secondary “fitness” mutations

Lamivudine

rtM204V/I/S, rtA181V/T

Telbivudine Adefovir

M204I rtN236T, rtA181V/T

rtL80I/V, rtL180M/C, rtV173L rtL80I/V and rtL180M —

Tenofovir

Unknown in patients

—

Entecavir

rtM204V/I

rtM250V, rtI169T, rtT184G, rtS202I/G

drug’s site of action on the polymerase to mutational pressure. Resistance is usually related to steric hindrance of drug binding, but associated mutations may compensate for this inhibition to a degree. Since nucleos(t)ide analogs inhibit replication but do not eliminate existing virus or significantly decrease cccDNA, eventual drug resistance is an expected outcome in a subset of patients treated with any of these drugs. Clinical resistance is most easily recognized by less than a 1 log decline in HBV DNA after 3 months or a 1 log rise in HBV DNA level above nadir in patients who initially responded. Drug resistance should be suspected in such cases and can be confirmed by genotype testing. A detailed discussion of the specific dominant and minor drug-resistant mutations is beyond the scope of this chapter, but excellent reviews are available [7, 9]. The major mutations for each drug and the therapeutic alternative after emergence of drug resistance are listed in Table 54.3 and discussed under each drug below.

Lamivudine Lamivudine (3TC) is a cytosine analog which inhibits HBV polymerase by competing with natural triphosphates to terminate RNA chain elongation [12]. It is a potent inhibitor of HBV replication and was the first nucleoside analog approved for treatment of chronic hepatitis B. However, its use has been limited by a high rate of viral resistance ranging from 10 to 20% per year, resulting in loss of clinical response. Approximately 70% of patients develop resistance after 4–5 years of continuous treatment [13]. Resistance to lamivudine increases the likelihood of developing resistance to other agents, particularly entecavir (see below) [14]. Nevertheless, the high potency and low cost of lamivudine still make it an attractive drug in certain clinical situations where only short-term treatment is anticipated, such as the third trimester of pregnancy in highly viremic pregnant women, severe acute hepatitis, or as prophylaxis for reactivation in patients receiving immunosuppressive or chemotherapeutic agents [11].

Rescue therapy Add adefovir or tenofovir Add adefovir or tenofovir Add lamivudine Add telbivudine Add entecavir Add lamivudine Add telbivudine Add entecavir Add adefovir or tenofovir

Emtricitabine (FTC) Emtricitabine is the 5-fluoro derivative of lamivudine and, as such, has a potency and resistance profile similar to its parent drug [15]. It is not approved for use as a single agent for hepatitis B but is available in combination with tenofovir as Truvada.

Telbivudine Telbivudine is a thymidine analog that derives its high potency against HBV from its reported ability to inhibit the viral polymerase through all three of the aforementioned mechanisms [16]. It suppresses HBV and has similar HBeAg seroconversion rates to lamivudine, adefovir, and entecavir [8]. Resistance occurs in 5–10% per year and selects the rtM204I mutation, which is also resistant to lamivudine and entecavir [8, 9, 11].

Adefovir Adefovir is administered as its pro-drug of adefovir dipivoxil, and has the advantage of having activity against both wild-type and lamivudine-resistant strains of the virus. However, it has limited potency at the administered dose of 10 mg per day and up to one-third of patients fail to have a clinically significant virus reduction on therapy, particularly HBeAg-positive patients with high HBV DNA levels [17]. Hence it has been suggested that the primary applications of the drug might be in HBeAg-negative patients who tend to have lower baseline HBV DNA levels or in patients who have developed lamivudine resistance [11]. Resistance is observed in approximately 30% of cases after 5 years [18].

Tenofovir Like adefovir, tenofovir is an acyclic adenine nucleotide, but unlike adefovir, it has less renal toxicity and can be

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THE LIVER: HEPATITIS C VIRUS INFECTION

used at higher doses, thereby resulting in greater potency. It is approved in Europe and the United States. Approximately 75% of HBeAg-positive patients lose detectable HBV DNA after 1 year of therapy and HBeAg seroconversion is similar to that seen with the nucleoside analogs [19]. Resistance is extremely low but long-term experience with the drug is still limited.

a finite duration of treatment, IFN-α has not maintained favor in practice because of the subcutaneous route of administration and frequent side effects. Its use accounts for less than 10% of all prescriptions for HBV treatment in the United States [11].

Combination therapy Entecavir Entecavir is an acyclic guanosine derivative with potent activity against HBV. HBV DNA suppression is rapid, but the HBeAg seroconversion rate is not different to that with the other nucleoside analogs [20]. Viral resistance is rare in patients who have never been treated with lamivudine [21], but may emerge in more than 40% of lamivudine-resistant patients after 5 years [14].

Interferon alpha Interferon (IFN) refers to a family of natural glycoproteins produced by the cells of most vertebrates in response to challenge by foreign agents, such as infectious organisms (viruses, bacteria, fungi, and parasites), and by tumor cells. IFN can be produced by cells of the innate and adaptive immune systems and by non-immune cells such as fibroblasts and epithelial cells. IFNs can be commercially manufactured by cell culture or recombinant technology, and have been commercially available for the treatment of chronic hepatitis for more than 15 years. IFNs inhibit the replication of many viruses, including hepatitis viruses, through induction of intracellular genes that initiate a variety of antiviral mechanisms, including direct virus inhibition (inhibition of virus attachment and uncoating, induction of intracellular proteins and RNases), and by amplification of specific [cytotoxic T-lymphocyte (CTL)] and non-specific [natural killer (NK) cell] immune responses. These effects are discussed in more detail below as they relate to hepatitis C treatment. The specific mechanism(s) of action for IFN-α in chronic hepatitis B relate to both of these pathways although IFNs induce a smaller decline in HBV DNA than the nucleoside analogs. Rather, it appears to work predominantly through augmentation of the cell-mediated immune response to the virus [22]. Standard IFN-α administered daily or three times per week was first approved for use in chronic hepatitis B in 1992 [23], but its use has been largely supplanted by once per week pegylated IFN-α for 48 weeks [24–26]. Treatment with pegylated IFN-α achieves an HBeAg seroconversion rate of 33%, which is comparable to the best results with nucleoside analogs [8], and HBsAg clearance occurs in 3–5%, which appears to be much higher than that observed with polymerase inhibitors to date. Despite the excellent virological response rates and

The potential advantages of the combination of agents active against HBV include more rapid viral reduction, increased rate of viral response (undetectable HBV DNA or HBeAg seroconversion) and reduced drug resistance incidence in the long term. However, while the combination of antiviral medications is of clear benefit in the treatment of HIV and HCV infection, there remains no proven benefit in patients with HBV infection. The combination of either standard or pegylated IFN-α and lamivudine results in more HBV DNA suppression than with lamivudine alone, but the benefit following discontinuation of treatment was not different from that with IFN alone [27–31]. This combination also resulted in a lower rate of lamivudine resistance than with lamivudine alone; however, it is not known whether the same phenomenon would be observed if nucleoside analogs with a greater barrier to resistance, such as entecavir, were studied. Combinations of nucleos(t)ide polymerase inhibitors have been limited. Lamivudine and adefovir showed no greater likelihood of HBV DNA suppression, HBeAg loss, or ALT normalization after 52 weeks, although drug resistance was less than with lamivudine monotherapy (15% versus 43%) [32]. Similarly, the combination of lamivudine and telbivudine was not better than telbivudine alone [33]. Add-on therapy has been shown to be superior to switching to another drug with no cross-resistance in the case of lamivudine resistance. Indeed, addition of a second agent significantly reduces the chance of resistance selection and a clinical flare when treating drug resistance [34] and has been suggested as part of management of these cases (Table 54.3) [10]. In na¨ıve patients, current recommendations are to treat first line with potent drugs with a high genetic barrier to viral resistance, that is, entecavir or tenofovir, and add a second drug if HBV DNA is still detectable after 1 year in order to reduce the long-term potential for drug resistance.

HEPATITIS C VIRUS INFECTION General principles of treatment The treatment of chronic hepatitis C is based on the use of a combination of pegylated IFN-α and ribavirin. Chronic HCV infection is curable. Therefore, the goal of therapy is a cure, characterized by the sustained virological response (SVR), defined as an undetectable HCV RNA

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24 weeks after the end of treatment. Cured patients do not develop the complications of chronic HCV infection, except cirrhotic patients, who may develop hepatocellular carcinoma and need appropriate monitoring. Numerous new anti-HCV approaches are in development. The most advanced classes of drugs are specific inhibitors of HCV enzymatic functions. These drugs potently inhibit HCV replication but, when used in monotherapy, select for frequent and early HCV resistance. In the mid-term, the new standard-of-care will most likely be a triple combination of pegylated IFN-α, ribavirin, and a specific HCV inhibitor. In the longer term, short-term oral therapies based on combinations of HCV inhibitors may be indicated.

those that are infected [38]. IFNs can stimulate the effector function of NK cells, cytotoxic T lymphocytes, and macrophages, up-regulate the expression of major histocompatibility complex (MHC) class I and class II molecules, induce immunoglobulin synthesis by B cells, and stimulate the proliferation of memory T cells [39]. IFN-α acts through activation and maturation of dendritic cells, leading to MHC up-regulation. IFN-α can also up-regulate various chemokines, chemokine receptors, and costimulatory molecules that, in turn, stimulate CD4- and CD8-positive T cell responses and modulate T lymphocyte responses through the promotion of T-helper 1 (Th1) differentiation [40].

Molecular and pathophysiological basis for therapy

Pegylation of IFN-α

Properties of type I IFNs IFNs are natural glycoproteins produced by the cells of most vertebrates in response to challenge by foreign agents, such as infectious organisms (viruses, bacteria, fungi, and parasites), and by tumor cells. IFNs can be produced by cells of the innate and adaptive immune systems and by non-immune cells such as fibroblasts and epithelial cells. Type I IFNs form a superfamily of innate cytokines that comprise IFN-α, with 13 human subtypes, IFN-β, IFN-ω, IFN-τ, IFN-κ, IFN-ε, IFN-λ, and IFN-ζ. Only IFN-α, IFN-β, IFN-ω, IFN-κ, and IFN-ε are expressed in humans. Type I IFNs bind to their specific receptor complex, a heterodimer composed of two chains, interferon alpha receptor-1 (IFNAR-1) and IFNAR-2. IFNAR-1 and IFNAR-2 are intimately associated with two Janus family tyrosine kinases, Tyk2 and Jak1, both located on the cytoplasmic side of the cell membrane [35]. Type I IFN interaction with the specific receptor complex activates a transcription factor, interferon-stimulated gene factor 3 (ISGF3), which induces the expression of interferon-stimulated genes (ISGs) that mediate their cellular actions [36]. ISG expression results in two types of effects: antiviral and immunomodulatory. The best-known antiviral effectors produced as a result of IFN cascade induction include 2′ ,5′ -oligoadenylate synthetase (2′ ,5′ -OAS), double-stranded ribonucleic acid activated protein kinase (PKR) and myxovirus (Mx) proteins. Other, minor effectors include ribonucleic acid-specific adenosine deaminase 1 (ADAR1), 20 kDa ISG product (ISG20), ISG54, and ISG56. IFN-induced micro RNAs might be important in IFN antiviral properties [37]. IFN antiviral properties confer a global antiviral state in the target cells. In addition to its direct antiviral properties, IFN-α exhibits potent immunomodulatory properties which contribute to its antiviral effects by involving cells other than

Poly(ethylene glycol) (PEG) is formed by linking repeating units of ethylene glycol to form polymers that are linear or branched molecules with different masses. Pegylation is the process by which PEG chains are covalently attached to IFN molecules. Pegylation confers a number of properties on IFN-α molecules, such as sustained blood levels that enhance antiviral effectiveness and reduce adverse reactions, and also a longer half-life and improved patient convenience [41]. Two forms of pegylated IFN-α have been approved by the FDA and by the European Medicines Agency (EMEA) for the treatment of chronic hepatitis C in adults, namely pegylated IFN-α2a and pegylated IFN-α2b, in which the IFN-α molecules are linked to PEG molecules of different sizes. Pegylated IFN-α2a (Pegasys; Roche, Basel, Switzerland) is a 40 kDa branched monomethoxy PEG conjugate of IFN-α2a [42]. The pegylated IFN-α2a molecule contains a large, branched PEG moiety attached at a single point to the native protein by means of a hydrolytically stable amide bridge. The molecule is very stable. In vitro, the specific antiviral activity of the 40 kDa branched pegylated IFN-α2a molecule is only 7% of that of the non-pegylated molecule [42]. In healthy volunteers after a single subcutaneous injection, the pegylated form shows sustained absorption, reduced systemic clearance, and a longer half-life in comparison with standard IFN-α2a. High serum levels of pegylated IFN-α2a are reached 3–8 hours after subcutaneous administration, and the maximum serum concentration is reached approximately 80 hours post-injection [43]. The steady-state serum concentration is reached within 5–8 hours after weekly subcutaneous administrations, and a uniform drug concentration is present throughout the dosing interval [43]. In rats, pegylated IFN-α2a is distributed to both the liver and kidney, whereas standard IFN-α localizes principally in the kidney. Pegylated IFN-α2a requires hepatic metabolism by non-specific proteases. Active renal excretion is limited (85 kg), raising the doses of pegylated IFN-α and ribavirin significantly increased the SVR rate [103]. In a small number of cirrhotic patients infected with HCV genotype 1 who had not responded to a previous course of pegylated

54: CURRENT AND FUTURE THERAPY FOR HEPATITIS B AND C

IFN-α and ribavirin, 180 µg of pegylated IFN-α2a every 5 days, combined with ribavirin, induced an SVR in several cases [104]. Ongoing trials are assessing more frequent administration and higher weekly doses of pegylated IFN-α, and higher doses of ribavirin, in non-responder and “difficult-to-treat” patients. Patients receiving such reinforced therapy must be carefully monitored for toxicity, and the merits and drawbacks of growth factor administration should be considered.

Novel forms of IFN-α and alternatives to ribavirin Novel forms of IFN-α Albumin (Alb)–IFN-α2b (Human Genome Sciences, Rockville, MD, USA, and Novartis, Basel, Switzerland) is an 87.5 kDa recombinant protein consisting of an IFN-α2b molecule attached to a human albumin moiety. Alb–IFNα2b has a half-life of up to 159 hours, allowing dosing at intervals of 2–4 weeks [105]. Alb–IFN-α2b has been reported to induce a dose-dependent antiviral response in previously untreated patients and in non-responders to the pegylated IFN-α–ribavirin combination [105, 106]. The results of a phase II trial in untreated genotype 1-infected patients showed SVR rates to be not significantly different among four groups of patients receiving either the standard pegylated IFN-α2a–ribavirin combination or Alb–IFN-α2b at doses of 900 µg every 2 weeks, 1200 µg every 2 weeks and 1200 µg every 4 weeks [107]. End-of-treatment responses to Alb–IFN-α2b administered every 2–4 weeks in combination with ribavirin are similarly frequent in patients infected with HCV genotypes 2 and 3 [108]. The occurrence of two severe cases of pulmonary fibrosis led to a dose reduction to 900 µg every 2 weeks in the ongoing phase III trial. Multiferon (Viragen, Plantation, FL, USA) is a highly purified, multi-subtype natural human IFN-α derived from human leukocytes. It has already been approved for HCV therapy in several countries. Medusa (Flamel Technologies, Lyon, France) is a selfassembled poly(amino acid) nanoparticle system that can be used as a protein carrier for novel long-acting native protein drugs. IFN-α2bXL uses the Medusa technology to decrease the C max of IFN-α2b. In a single dose-ranging phase I study, IFN-α2bXL was shown to be safe and reduce peak serum IFN-α concentration while providing sustained release of IFN-α. In a recent phase Ib trial, a better antiviral efficacy was observed with 27 × 106 units of IFN-α2bXL than with 1.5 µg kg−1 of pegylated IFN-α2b during the second week of administration [109].

Alternatives to ribavirin Because of its adverse effects, and especially hemolytic anemia, ribavirin dose reduction or discontinuation

909

is frequently necessary, and this can lead to viral breakthroughs and to post-treatment relapse. Taribavirin (Valeant Pharmaceuticals, San Diego, CA, USA) is an amidine prodrug of ribavirin that is extensively converted into ribavirin in hepatocytes by adenosine deaminases [110]. Taribavirin is preferentially taken up by the liver, the main site of HCV replication, and does not efficiently enter red blood cells [111]. In two recent phase III trials, in combination with pegylated IFN-α2a and IFN-α2b, respectively, taribavirin at a flat dose of 0.6 g twice per day was not as effective as a weight-based ribavirin dose in patients with chronic hepatitis C of various genotypes [112, 113]. The incidence of hemolytic anemia was, however, significantly lower with taribavirin. In a more recent phase II trial, taribavirin administration at weight-based doses of 20, 25, and 35 mg kg−1 per day in combination with pegylated IFN-α2a was associated with slightly higher rates of undetectable HCV RNA, and significantly lower rates of severe anemia, than the standard-of-care at week 12 [114]. The final results of this study are awaited.

New therapies for hepatitis C In the last decade, insights into the virology of HCV have unraveled several targets for potential novel therapeutics that, unlike IFN-α and ribavirin, are specifically targeted to HCV. Many drugs are at the preclinical developmental stage and several are in clinical development, but initial trials using some of these inhibitors alone have raised concerns about their tolerability and the development of viral resistance. A number of specifically targeted therapies are now also being tested in combination with pegylated IFN-α and ribavirin.

The HCV life cycle, the target of specific HCV inhibitors The successive steps of the HCV life cycle, as described briefly here, constitute a very large number of potential targets of intervention for specific anti-HCV drugs (Figure 54.3). The reader is referred to Chapter 51 for a more detailed discussion. HCV entry involves assembly of two HCV envelope glycoproteins E1 and E2 at receptor molecules on the surface of target cells. HCV binding and internalization are mediated via a receptor complex currently considered to be comprised of glycosaminoglycans that serve as the site for HCV attachment, the tetraspanin CD81 which may facilitate post-attachment entry, along with scavenger receptor B1 (SR-B1), claudin-1 and occludin, which appear to act late in the entry process [115–118]. Several other receptor molecules have also been suggested [119–121]. After attachment, HCV entry into cells is pH dependent and related to clathrin-mediated endocytosis, which is followed by a fusion step within an acidic endosomal

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THE LIVER: HEPATITIS C VIRUS INFECTION

Figure 54.3 Schematic of the hepatitis C virus replication cycle in the hepatocyte. Every step of the life cycle offers a variety of potential targets for novel drug classes. Reproduced from Pawlotsky et al ., Gastroenterology, 132: 1979–1998, Copyright (2007), with permission from Elsevier

compartment. Decapsidation of viral nucleocapsids liberates free positive-strand genomic RNAs in the cell cytoplasm, where they serve, together with newly synthesized RNAs, as mRNAs for synthesis of the HCV polyprotein. The HCV 5′ -untranslated region (5′ UTR), the most conserved region of the HCV genome, contains the internal ribosome entry site (IRES), which controls HCV genome translation by recruiting both cellular proteins, including eukaryotic initiation factor (eIF) 2 and 3, and viral proteins [122, 123]. The large precursor polyprotein generated by HCV genome translation is targeted to the endoplasmic reticulum membrane, where post-translational processing of the HCV polyprotein results in the generation of at least 11 proteins, including three structural proteins (C or core, E1 and E2), a small protein, p7, whose function

has not yet been definitively defined, six non-structural (NS) proteins (NS2, NS3, NS4A, NS4B, NS5A, and NS5B), and the so-called “F” protein that results from a frameshift in the core coding region. At least two host cellular peptidases are required for processing of the HCV structural proteins, including host signal peptidase and signal peptide peptidase. Two viral peptidases are involved in the post-translational processing of HCV NS proteins: NS2, a zinc-dependent metalloproteinase that cleaves the site between NS2 and NS3 , and NS3/4A, a serine proteinase that catalyzes HCV polyprotein cleavage at the NS3/NS4A, NS4A/NS4B, NS4B/NS5A, and NS5A/NS5B junctions [124]. HCV replication is catalyzed by the NS5B ribonucleic acid-dependent ribonucleic acid polymerase (RdRp). The NS5A protein plays an important role in virus replication

54: CURRENT AND FUTURE THERAPY FOR HEPATITIS B AND C

through unclear mechanisms. The NS3 helicase–NTPase domain of the NS3 protein has several functions important in replication, including RNA-stimulated NTPase activity, RNA binding, and unwinding of RNA regions of extensive secondary structure [125]. Finally, NS4B is an integral membrane protein which serves as a membrane anchor for the replication complex [126]. Infection with HCV leads to rearrangements of intracellular membranes to form a replication complex that associates viral proteins, cellular components, and nascent RNA strands (Figure 54.2). By analogy with other positive-strand RNA viruses, it is believed that the positive-strand genome RNA serves as a template for the synthesis of a negative-strand intermediate of replication. Then, negative-strand RNA serves as a template to produce numerous strands of positive polarity that will subsequently be used for polyprotein translation, synthesis of new intermediates of replication, or packaging into new virus particles [127]. Finally, viral particle formation is initiated by the interaction of the core protein with genomic RNA in the endoplasmic reticulum lumen, although the details of this process and subsequent export of mature virions from the hepatocyte are poorly understood [128, 129].

Specific HCV inhibitors HCV entry inhibitors Inhibition of HCV entry can be based on the use of specific polyclonal or monoclonal antibodies that neutralize infectious particles and prevent their attachment to the receptor molecules. Polyclonal hepatitis C immune globulins have shown no efficacy on HCV replication in HCV-infected liver transplant recipients [130]. Anti-HCV monoclonal antibodies have shown moderate and transient antiviral properties in vivo [131, 132]. Studies evaluating more frequent daily dosing at higher doses and/or combinations with potent antiviral molecules using different mechanisms will need to be undertaken. Inhibition of HCV entry can also be based on the use of specific entry inhibitor molecules. Since our understanding of HCV entry mechanisms remains rudimentary, the development of specific small molecule inhibitors has been hampered. It should improve with the recent development of a productive cell culture system [133, 134]. Fusion could also become an interesting target for novel therapies when its mechanisms are better understood.

Inhibitors of HCV translation Antisense DNA or RNA oligonucleotides, the sequence of which is complementary to the target mRNA sequence, can prevent translation of viral proteins. Several oligonucleotides targeting the 5′ UTR, the most conserved region

911

of HCV genome, have been reported to inhibit HCV gene expression in cell culture systems, but their clinical development was halted due to lack of antiviral efficacy and concern about their tolerance [135]. A synthetic phosphorodiamidate morpholino oligomer (PMO) has also been shown to inhibit HCV protein translation in vitro and in animal models, but no significant reductions in HCV RNA levels have been observed in vivo. Ribozymes are RNA molecules that catalyze cleavage of a target RNA molecule based on sequence-specific recognition. A chemically modified ribozyme has been shown to cleave HCV RNA within the IRES, but early-phase clinical trials were halted because of animal toxicity. RNA interference is initiated by small interfering ribonucleic acids (siRNAs) or short hairpin ribonucleic acids (shRNAs) that associate with various proteins to form an RNA-inducing silencing complex harboring nuclease and helicase activity. siRNAs and shRNAs targeting the 5′ UTR have been shown to inhibit HCV IRES-mediated translation in various models [136]. However, because of their size and chemical composition, siRNAs and shRNAs currently are not orally bioavailable and require parenteral administration. The three-dimensional functional IRES complexed with cellular and viral proteins and the ribosomal subunits offer a promising target for small-molecule inhibitors. Future improvement of our understanding of the mechanisms of HCV polyprotein translation will boost the development of small-molecule inhibitors targeting this function.

Inhibitors of post-translational processing Peptidomimetic NS3/4A serine proteinase inhibitors Highly selective, potent peptidomimetic inhibitors of HCV NS3/4A proteinase have been derived from the viral substrate of the enzyme using structurebased drug design techniques. A number of drugs currently have reached clinical development and are promising, the most advanced of which are: telaprevir (VX-950; Vertex Pharmaceuticals), boceprevir (SCH503034; Schering-Plough), TMC435350 (Tibotec), and ITMN-191/R7227 (InterMune & Roche). Telaprevir has shown antiviral activity against HCV in vitro. In a phase Ib clinical trial, HCV genotype 1-infected patients receiving telaprevir monotherapy for 14 days all had at least a 2 log decrease in HCV RNA levels. Viral breakthroughs occurred in the patients with lower exposure to the drug, due to selection of telaprevir-resistant HCV variants [136]. Combination with pegylated IFN-α with or without ribavirin could theoretically prevent telaprevir resistance. The results of two phase II clinical trials were recently reported. The PROVE 1 trial tested the combination of telaprevir with pegylated IFN-α and ribavirin in treatment-na¨ıve US patients infected with HCV genotype 1 versus standard-of-care [137]. A sustained virological response was observed in 41% of the patients receiving the combination of pegylated IFN-α and ribavirin

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for 48 weeks, and in 61% of those receiving a triple combination of telaprevir, 750 mg every 8 hours, pegylated IFN-α, and ribavirin for 12 weeks followed by 12 weeks of pegylated IFN-α and ribavirin, and in 67% of those receiving the triple combination for 12 weeks, followed by 36 weeks of pegylated IFN-α and ribavirin (p = 0.02 and 0.001, respectively) [137]. In the PROVE 2 trial, performed in treatment-na¨ıve European patients infected with HCV genotype 1, the sustained virological response rates were 48% in the control arm, 62% in the patients receiving the triple combination for 12 weeks, and 68% in the patients receiving the triple combination for 12 weeks followed by 12 weeks of pegylated IFN-α and ribavirin (p = 0.08 and 0.01, respectively) [138]. The sustained virological response rate was 36% only in the patients receiving the combination of pegylated IFN-α and telaprevir for 12 weeks, essentially due to a high rate of relapse in relation with the selection of telaprevir-resistant variants [138]. In both trials, dermatological side effects, including rash and pruritus, were frequent (more than 50%). Other side effects included gastrointestinal symptoms and moderate anemia [137, 138]. Boceprevir is a potent inhibitor of HCV replication in vitro [139, 140]. One week of boceprevir monotherapy in patients who had previously failed to respond to pegylated IFN-α alone or in combination with ribavirin was well tolerated and induced an average HCV RNA load reduction of approximately 1.0 and 1.6 log IU ml−1 at doses of 200 and 400 mg three times per day, respectively [141]. Preliminary results of a phase II clinical trial (SPRINT-1) have been reported recently [142]. Sustained virological response rates were available in only two arms, one receiving the triple combination of boceprevir, 800 mg tid, pegylated IFN-α, and ribavirin for 28 weeks, the other receiving the same triple combination for 24 weeks after 4 weeks of a “lead-in” phase with pegylated IFN-α and ribavirin only. The sustained virological response rates were 55 and 57%, respectively (not significantly different) [142]. The main side effect was anemia and no skin events were reported. TMC435350 at a dose of 200 mg once per day induced on average a more than 3.5 log HCV RNA level reduction in patients infected with HCV genotypes 1a and 1b in a recent phase Ib trial [143]. ITMN-191 reduced HCV RNA levels by an average of 3.4 log at a dose of 200 mg every 12 hours, and 3.9 log at a dose of 200 mg every 8 hours in another phase Ib trial. A number of other peptidomimetic protease inhibitors are currently progressing to clinical development.

Inhibitors Interaction

of

NS4A–NS3

Proteinase

Other approaches have been developed to inhibit the NS3–NS4A serine proteinase function. Acylthiourea compounds inhibit binding of NS4A to the NS3 proteinase, therefore inhibiting polyprotein processing by preventing the formation of the active

proteinase complex. Administration of 300 mg twice daily for 5 days of one of these compounds resulted in an average change in HCV RNA level from baseline of −0.9 log HCV RNA IU ml−1 [144]. The development of this drug has been halted, however, because of biological signs of proximal tubular dysfunction [144].

Inhibitors of HCV replication Given the complexity of the replication process and the number of actors involved, HCV replication offers a variety of targets for antiviral intervention. So far, specific nucleoside or non-nucleoside inhibitors of HCV RdRp and cyclophilin inhibitors have undergone evaluation in clinical trials.

Nucleoside RdRp inhibitors

Nucleoside inhibitors target the catalytic site of the enzyme. Two drugs are currently being evaluated in clinical trials, R1626 (Roche) and R7128 (Pharmasset and Roche). R1626 is a 4′ -azidocytidine nucleoside analogue. In a recent phase II trial, 84% of patients receiving the combination of R1626, 1500 mg twice per day, and pegylated IFN-α and ribavirin for 4 weeks had undetectable HCV RNA at week 4 [145]. No viral resistance to R1626 was observed after 4 weeks of the triple combination. The main side effects were neutropenia, and thrombocytopenia, although to a lesser extent [145]. The drug’s development has been halted because of very severe lymphopenias in combination with IFN and ribavirin. R7128 is a nucleoside analog that demonstrated a –2.7 log mean HCV RNA level decrease with 1500 mg three times per day. The same dose of R7128 was administered in combination with pegylated IFN-α and ribavirin. The combination induced a –5.1 log HCV RNA level reduction on average and 85% of the patients had undetectable HCV RNA at week 4. Pegylated IFN-α and ribavirin without R7128 induced an average HCV RNA level reduction of –2.5 log. R7128 appeared to be safe and well tolerated over the 4 weeks [146].

Non-nucleoside RdRp Inhibitors Nonnucleoside inhibitors target allosteric sites of the RdRp. Five such sites, named A to E, have been identified so far within the RdRp structure. Various resistance profiles have been described in vitro in the replicon system for the different families of molecules targeting these sites. GS-9190 is a non-nucleoside inhibitor targeting allosteric site E. After eight days of administration, GS-9190 reduced HCV RNA levels by 1.4 and 1.7 log at doses of 40 and 120 mg twice per day, respectively. Other drugs belonging to all of the non-nucleoside inhibitor classes are at the preclinical developmental stage. Cyclophilin inhibitors

DEBIO-025 (DebioPharm) is a cyclophilin inhibitor which potently inhibits HCV

54: CURRENT AND FUTURE THERAPY FOR HEPATITIS B AND C

replication [147]. In a Phase Ib trial, DEBIO-025 was administered to treatment-na¨ıve patients coinfected with human immunodeficiency virus at a dose of 1200 mg twice daily during 15 days [148]. All treated patients experienced an HCV RNA level decrease of more than 2 log IU ml−1 upon administration, with a maximum HCV RNA level reduction of –3.6 log IU ml−1 , independent of the HCV genotype [148]. Hyperbilirubinemia was frequent, however, and led to treatment withdrawal in several cases. In a recent phase II trial, DEBIO-025 at daily doses of 600 and 1000 mg demonstrated an additive antiviral effect in combination with pegylated IFN-α. However, hyperbilirubinemia was observed at a dose of 1000 mg daily [149]. Other cyclophilin inhibitors currently are under study.

Inhibitors of HCV assembly and release Imino sugars have been suggested to be able to inhibit competitively endoplasmic reticulum-resident α-glucosidases, alter envelope proteins glycosylation, and interfere with the assembly of different viruses [150]. Celgosivir (Migenix), an imino sugar derivative, has shown a modest antiviral effect on HCV in monotherapy.

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100. Snoeck, E., Wade, J.R., Duff, F., Lamb, M. and Jorga, K. (2006) Predicting sustained virological response and anaemia in chronic hepatitis C patients treated with peginterferon alfa-2a (40KD) plus ribavirin. Br J Clin Pharmacol , 62, 699–709. 101. Jensen, D.M., Morgan, T.R., Marcellin, P., Pockros, P.J., Reddy, K.R., Hadziyannis, S.J., Ferenci, P. et al. (2006) Early identification of HCV genotype 1 patients responding to 24 weeks peginterferon alpha-2a (40kd)/ribavirin therapy. Hepatology, 43, 954–60. 102. McHutchison, J.G., Manns, M.P., Brown, R.S., Reddy, K.R., Shiffman, M.L. and Wong, J.B. Jr (2007) Strategies for managing anemia in hepatitis C patients undergoing antiviral therapy. Am J Gastroenterol , 102, 880–89. 103. Fried, M., Jensen, D., Rodriguez-Torres, M., Nyberg, L., Di Bisceglie, A., Morgan, T., Pockros, P.J. et al. (2006) Improved sustained virological response (SVR) rates with higher, fixed doses of peginterferon alfa-2a (40kD) (Pegasys) plus ribavirin (RBV) (Copegus) in patients with “difficult-to-treat” characteristics. Hepatology, 44 (Suppl 1), 314A. 104. Hezode, C., Bouvier-Alias, M., Roudot-Thoraval, F., Zafrani, S., Dhumeaux, D., Mallat, A. and Pawlotsky, J.M. (2006) Efficacy and safety of peginterferon alpha-2a administrated every five days in combination with ribavirin in HCV genotype 1-infected patients with severe fibrosis not responding to weekly administrations of peginterferon in combination with ribavirin. Hepatology, 44 (Suppl 1), 322A. 105. Balan, V., Nelson, D.R., Sulkowski, M.S., Everson, G.T., Lambiase, L.R., Wiesner, R.H., Dickson, R.C. et al. (2006) A phase I/II study evaluating escalating doses of recombinant human albumin-interferon-alpha fusion protein in chronic hepatitis C patients who have failed previous interferon-alpha-based therapy. Antivir Ther, 11, 35–45. 106. Bain, V.G., Kaita, K.D., Yoshida, E.M., Swain, M.G., Heathcote, E.J., Neumann, A.U., Fiscella, M. et al. (2006) A phase 2 study to evaluate the antiviral activity, safety, and pharmacokinetics of recombinant human albumin-interferon alfa fusion protein in genotype 1 chronic hepatitis C patients. J Hepatol , 44, 671–78. 107. Zeuzem, S., Benhamou, Y., Bain, V., Shouval, D., Pianko, S., Flisiak, R., Grigorescu, M. et al. (2007) Antiviral response at week 12 following completion of treatment with albinterferon alpha-2b plus ribavirin in genotype 1, IFN-na¨ıve, chronic hepatitis patients. J Hepatol , 46 (Suppl 1), S293. 108. Bain, V.G., Marotta, P., Kaita, K., Yoshida, E., Swain, M.G., Bailey, R., Neumann, A.U. et al. (2007) Comparable antiviral response rates with albumin interferon alpha-2b dosed at Q2W or Q4W intervals in na¨ıve subjects with genotype 2 or 3 chronic hepatitis C. J Hepatol , 46, S7. 109. Trepo, C., Guest, M., Meyrueix, R., Rouzier, R., Raffanel, C., Belhadj-Tahar, H., Maynard-Muet, M. et al. (2008) Evaluation of antiviral activity and tolerance of a novel sustained release interferon alpha-2b (IFN-alpha-2bXL) compared to pegylated

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55

Biological Principles and Clinical Issues Underlying Liver Transplantation for Virus-induced End-stage Liver Disease James R. Burton Jr1 , Hugo R. Rosen1 and Paul Martin2 1 University 2

of Colorado, Denver, CO, USA Division of Hepatology, University of Miami, Miami, FL, USA

INTRODUCTION

NATURAL HISTORY OF RECURRENT HCV

Hepatitis C virus (HCV)-related end-stage liver disease is the leading indication for liver transplantation (LT) worldwide. Recurrent HCV infection is the most significant issue facing transplant physicians today, as recurrence is immediate with a significant percentage developing severe histological recurrence. Treatment of HCV in the transplant setting can be challenging given the limited applicability and tolerability and lower efficacy than in the non-transplant setting.

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

Based on the presence of HCV RNA in the serum, recurrence is immediate and universal with viral levels 1–3 months post-LT often being 20-fold greater than in the pre-LT period [1]. The natural history of recurrent HCV after LT is accelerated compared with HCV infection in the non-transplant setting. Between 20 and 40% of patients transplanted for HCV develop allograft cirrhosis in 5 years compared with less than 5–20% at 20 years

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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THE LIVER: PRE-TRANSPLANT ANTIVIRAL THERAPY

in the non-transplant setting [2–4]. Once cirrhosis develops after LT, two-thirds will develop decompensation (variceal bleeding, ascites, hepatic encephalopathy) within 3 years, compared with ∼10% in immunocompetent HCV patients with cirrhosis [3, 5]. Once decompensation develops post-LT, outcome is very poor, with 6 g dl−1 and alkaline phosphatase levels greater than five times the upper limit of normal. The pathogenesis remains undefined but is likely immunologically different from typical HCV-induced allograft failure with a preferential Th2 cytokine production by intrahepatic lymphocytes being implicated [8]. Treatment focuses on minimizing immunosuppression and viral suppression with indefinite use of interferon-based antiviral therapy [9].

FACTORS ASSOCIATED WITH SEVERE HCV RECURRENCE A number of factors have been associated with severe HCV recurrence that affects both patient and graft survival. Table 55.1 outlines these donor, viral, and transplant factors. Given the detrimental effects of treating acute cellular rejection with corticosteroids [10–12], suspicion of rejection in HCV patients should always be confirmed with liver biopsy. Differentiating recurrent HCV alone from recurrent HCV and rejection can be challenging [13]. Mild rejection may be treated without steroids by

increasing calcineurin inhibitors or addition/increase of mycophenolate mofetil [14, 15]. The long-term effects of this approach are unknown. How best to handle maintenance immunosuppression is a hotly debated topic. Despite a vast literature on the role of immunosuppression and the effects on recurrent HCV, no clear conclusions can be drawn.

TREATMENT OF RECURRENT HCV Given the accelerated natural history of HCV recurrence, several approaches have been proposed to prevent or slow disease progression (Table 55.2). Unfortunately, many published studies investigating the role of treating recurrent HCV with standard interferon and pegylated interferon with or without ribavirin have been small, single-center, uncontrolled trials with significant variability in patient selection, timing of antiviral therapy, and study endpoints. Rates of sustained virological response (SVR) (HCV RNA undetectable 6 months after stopping therapy) are far less than those achieved in immunocompetent HCV-infected patients (on average 20–25% less). Table 55.3 outlines potential explanations for lower response rates post-LT.

PRE-TRANSPLANT ANTIVIRAL THERAPY Treatment of HCV prior to LT may eliminate the risk of developing recurrent HCV post-LT and in some cases may avoid LT altogether. Treatment should be strongly considered in patients with cirrhosis who have Child-Pugh (CP) scores ≤7 or Model for End-Stage Liver Disease (MELD) scores ≤18 {1 MELD score = 10[0.957 ln(serum creatinine) + 0.378 ln(total bilirubin) + 1.12 ln (INR) + 0.643], where INR = intestinal normalized ratio} [7]. A special group that should be strongly considered for antiviral therapy while awaiting transplantation are patients with well-compensated liver disease upgraded on the transplant

Table 55.1 Risk factors associated with severe HCV recurrence Category

Factor

Donor factors

Donor age >40 years Prolonged cold ischemia time Female sex Living donor liver transplant experience (20 cases) HCV genotype 1 in some studies High viral load at liver transplantation Cytomegalovirus infection Treatment of acute cellular rejection with: Corticosteroid boluses OKT3 use

Viral factors Transplant factors

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Table 55.2 Antiviral treatment strategies for recurrent HCV Treatment strategy

Timing of antiviral therapy

Advantages

Disadvantages Low virological response Limited tolerability and potential for serious adverse effects Maximum immunosuppression Higher risk of rejection and infection High HCV RNA levels More advanced fibrosis

Pretransplant

Prior to LT

Eliminate or reduce risk of recurrent HCV Potential to avoid LT

Preemptive

Early post-LT

Relatively low HCV RNA levels Minimum or no histological disease

Established disease

Diagnosis of acute hepatitis or established and/or severe disease

Lower immunosuppression Improved clinical status and better tolerance Lower risk of rejection

Table 55.3 Contributing factors leading to lower response rates to antiviral therapy in liver transplant patients

• High HCV RNA levels post-LT • High prevalence of genotype 1, previous non-responders to antiviral therapy

• Presence of immunosuppression • Poor clinical status, especially early post-LT leading to poor tolerability

• Cytopenias as result of immunosuppression requiring dose reduction and use of growth factors

• Renal insufficiencya limiting dose of ribavirin on account

the early transplant period. Only about 60% of LT recipients are eligible for preemptive therapy with the need for dose reduction occurring in up to 50% of treated patients [17]. Randomized trials of standard and pegylated interferon monotherapy have shown no significant differences in virological response, with high rates of discontinuation. Published experience using pegylated interferon with ribavirin is limited. In summary, the efficacy of preemptive antiviral therapy remains to be defined and should only be considered in patients undergoing retransplantation for rapidly progressive HCV recurrence [7].

of its associated risk of hemolytic anemia a Not uncommonly seen as the current model for end-stage liver disease

(MELD) score-based allocation system incorporates serum creatinine.

list solely for an indication for hepatocellular carcinoma (HCC). Few studies have examined the role of antiviral therapy in patients awaiting LT. The largest, by Everson et al., enrolled 124 patients [56 Child-Pugh class (CPC) A, 45 CPC B, 23 CPC C; mean CP score 7.4 and mean MELD score 11] [16]. Using a low accelerating dose regimen of primarily non-pegylated interferon, 24% achieved SVR (13% in genotype 1 and 50% in non-genotype 1 patients). Twelve out of 15 (80%) of patients who were HCV RNA negative pre-LT remained HCV RNA negative at least 6 months after LT.

PREEMPTIVE ANTIVIRAL THERAPY While it is tempting to consider antiviral therapy immediately after LT when viral levels are low and before the development of recurrent hepatitis, from a clinical standpoint treatment at this time is most challenging due to poor clinical status, cytopenias from maximum immunosuppression, and higher rates of rejection and infection in

TREATMENT OF ESTABLISHED DISEASE Given the lack of efficacy of preemptive therapy, many transplant centers have opted to delay treatment until significant recurrent disease is verified. This approach focuses treatment on those likely to achieve benefit with antiviral therapy, avoiding unnecessary toxicity and side effects in those without significant disease recurrence. Two general approaches have been used to treat established disease. One is to initiate antiviral therapy at diagnosis of acute recurrent hepatitis, and the other, which is followed by most transplant centers, is to initiate antiviral therapy when clinically significant evidence of recurrence exists. This latter approach utilizes protocol and/or clinically indicated liver biopsies reporting both the grade and stage of recurrent disease. The use of protocol liver biopsies is justified in patients transplanted for HCV given the potential for significant fibrosis progression, and has become standard-of-care [7]. Fibrosis progression is not linear over time, progressing more rapidly in the first post-LT year [18]. The 12 month liver biopsy has the best ability to stratify fibrosis progression with the approach that those developing severe disease recurrence within the first transplant year are at high likelihood of progressing to cirrhosis, identifying those

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THE LIVER: RISK OF ACUTE CELLULAR REJECTION

who should be considered for antiviral therapy [18, 19]. Furthermore, the absence of fibrosis 12 months post-LT is associated with excellent cirrhosis-free survival [18]. Patients with fibrosis stage ≥2 out of 4, severe inflammation (grade 3 or 4) or evidence of significant hepatic dysfunction (elevated bilirubin, prolonged prothrombin time) should be strongly considered for antiviral therapy [7]. Whether HCV patients treated for acute cellular rejection (ACR) with corticosteroids should be followed with earlier or more frequent protocol liver biopsies is not known. The measurement of hepatic venous pressure gradients (HVPGs) (wedged hepatic vein pressure minus free hepatic vein pressure) may be a useful tool in assessing disease severity in recurrent HCV. Recently, HVPG measurements ≥6 mmHg at 1 year post-LT have been shown to be extremely accurate in predicting clinical decompensation better than liver biopsy [20]. The same group has also shown a good correlation between HVPG measurements and histological response with antiviral therapy [21]. The current standard-of-care for treating established recurrent HCV is pegylated interferon and ribavirin. Berenguer et al. reported an SVR of 50% in 36 patients treated with pegylated interferon and ribavirin versus 13% in 31 patients treated with standard interferon and ribavirin [22]. There has been only one published randomized controlled trial evaluating pegylated interferon and ribavirin for established disease [21]. In this study of 81 patients, 54 patients (85% genotype 1) with mild recurrent HCV (stage 0–2/4) were randomized to either treatment or no treatment, while 27 patients with severe recurrence (stage 3–4/4 (n = 11), fibrosing cholestatic hepatitis (n = 4) or cholestatic hepatitis (lobular necroinflammatory score >2 and cholestasis, n = 12)) all revived antiviral treatment. SRV was 48% in patients with mild recurrence (versus 0% in the control group) compared with 18.5% in patients with severe recurrence. Generally SVR is associated with limiting fibrosis progression [21, 23–25]; however, one of these reports revealed fibrosis progression in 35% at 2 years and 20% at 3–5 years despite achieving SVR [23]. Viral relapse and reports of persistent hepatitis and fibrosis progression despite SVR suggest continued viral replication in the allograft [23, 24]. Intrahepatic HCV RNA detection in the liver could be useful for determining the duration of antiviral therapy, which in transplant patients is probably greater than the standard 48 weeks in the non-transplant setting. The only factor on multivariate analysis in the randomized controlled trial utilizing pegylated interferon and ribavirin to predict SVR (48% versus 0% in control group) was early virological response [≥2 log10 drop from baseline or HCV RNA negativity at week 12 from treatment initiation; odds ratio (OR) 16, 95% confidence interval (CI) 1.92 to 135; p < 0.001] [21].

MAINTENANCE THERAPY What to do with patients who fail to achieve SVR is not clear. Biochemical response [normalization of alanine aminotransferase (ALT) with treatment] has been shown to be associated with either stabilization or improvement in fibrosis in patients undergoing antiviral therapy, suggesting a possible benefit of continued therapy in this group [21]. Results from the Hepatitis C Antiviral Long-term Treatment against Cirrhosis (HALT-C) study showed a lack of efficacy of long-term maintenance pegylated interferon (half dose without ribavirin) in preventing fibrosis progression in non-responders to antiviral therapy with advanced fibrosis. Benefits of maintenance interferon in the transplant setting have not been defined. A critical problem in the treatment of recurrent HCV is the risk of severe disease progression with cessation of antiviral therapy, as evident from significant increases in HCV RNA and deterioration of allograft function even in virological non-responders [25, 26]. This finding may support the use of maintenance interferon in patients who do not achieve SVR. This issue has been looked at in a recent pilot study of 21 patients receiving maintenance therapy (interferon-α2b, 3 × 106 units three times weekly and ribavirin 600 mg daily) after completing 12 months of antiviral therapy started after development of acute HCV reinfection (i.e. low fibrosis stage). Fourteen of 21 achieved viral clearance after 12 months of antiviral therapy, with 9/14 undergoing unlimited maintenance therapy (mean 36 ± 12 months). Fibrosis scores improved in three and remained unchanged in six. None relapsed on therapy. Four virological non-responders continued on maintenance interferon and did not develop progressive fibrosis. No episodes of rejection were reported during maintenance therapy. Findings of this pilot study suggest that persistent suppression of intrahepatic HCV replication may potentially be beneficial. Figure 55.1 summarizes treatment approaches above into a recommend treatment algorithm.

RISK OF ACUTE CELLULAR REJECTION A potentially serious and controversial complication of antiviral therapy in transplant patients is rejection. Concern exists that pegylated interferon may be associated with an increased risk of rejection because of its extended half-life. Three uncontrolled trials of pegylated interferon and ribavirin yield conflicting results with no cases of rejection in two studies [27, 28] and a rate of 25% in the other [29]. Controlled trials of pegylated interferon monotherapy as prophylactic therapy and for treatment of established disease showed no difference in

55: BIOLOGICAL PRINCIPLES AND CLINICAL ISSUES UNDERLYING LIVER TRANSPLANTATION

HCV Cirrhosis Listed for LT

Pre-LT Antiviral Treatment? MELD score 107 copies ml−1 of HBV DNA, suggesting that more effective antiviral therapy initiated before transplant will further reduce the risk of HBV recurrence. These results strongly suggest that there is no need for routine i.v. HBIG administration as part of the immunoprophylaxis regimen and that the most potent oral regimen should be used pretransplant to render the patient serum HBV DNA negative. An earlier approach had been to switch from initial i.v. administration of HBIG to i.m. later post-transplant, but Gane et al.’s report makes this unnecessary. There are clearly patients who require little or no HBIG if they receive oral therapy in the absence of viral replication at the time of OLT, as shown in a series of patients who had received an HBIG-free regimen with lamivudine alone [55]. The licensing of several potent antiviral agents allows the opportunity to study HBIG-free regimens in the near future [56]. However, antiviral resistance remains a particular concern in this population because of the risk of a disease flare, although with prompt recognition of viral resistance pre-OLT and modification of antiviral therapy no adverse clinical outcomes were observed in a large group of patients [57]. An important consideration, however, is that residual HBV DNA can be detected in recipients who have no clinical evidence of recurrence [58]. Freshwater et al. using polymerase chain reaction (PCR) were able to identify HBV DNA in serum samples from two-thirds of a cohort of patients transplanted for HBV and apparently successfully protected against recurrence by immunoprophylaxis for up to 13 years [59]. This implies that it will never be possible to withdraw antiviral therapy completely.

ROLE OF IMMUNIZATION There is continued interest in vaccination as a strategy to protect transplant recipients against HBV infection either due to reinfection in an already infected candidate or de novo infection contracted from a core antibody-positive donor. Generally, patients with decompensated cirrhosis have a diminished rate of developing protective antibodies following HBV vaccination [60]. Attempts to vaccinate transplant recipients receiving immunoprophylaxis to prevent HBV recurrence have met with variable results. Endogenous anti-HB production has been described in Asian

LT recipients receiving antiviral prophylaxis, although antibody levels were not durable [61]. Standard vaccination regimens have been used as an adjunct to immunoprophylaxis in HBV-infected recipients. Generally, this approach has not resulted in substantial antibody production and has not replaced HBIG [63].

TRANSMISSION OF HBV INFECTION BY DONOR GRAFTS De novo acquisition from grafts harvested from donors with markers of prior, apparently resolved, HBV infection with absent HBsAg but positive core antibody can transmit HBV infection to the recipient, reflecting residual low-level replication after serological resolution of infection [36]. Transmission under these circumstances can be prevented by administration of antiviral prophylaxis similar to that used in infected recipients [64]. Use of these grafts from “isolated core” antibody donors helps expand the donor pool with low risk to the recipient, although ideally they should be reserved for HBsAg-positive recipients.

IMPACT OF ORAL THERAPY ON REQUIREMENT FOR LIVER TRANSPLANT As discussed earlier, oral antiviral therapy has a clear benefit in improving hepatocellular function in patients with overtly decompensated cirrhosis. A study by Kim et al. suggested that the number of patients being listed for HBV-related cirrhosis has declined in the United States in recent years, perhaps as a result of oral antiviral therapy [65].

FUTURE DIRECTIONS Clearly, preventing recurrence of HBV post-OLT has been one of the major triumphs in transplant hepatology. Future advances will likely result in regimens that do not require the use of HBIG.

REFERENCES 1. Chazouilleres, O., Kim, M., Combs, C. et al. (1994) Quantization of hepatitis C virus RNA in liver transplant recipients. Gastroenterology, 106, 994–99. 2. Gane, E.J., Portmann, B.C., Naoumov, N.V. et al. (1996) Long-term outcome of hepatitis C infection after liver transplantation. N Engl J Med , 334, 815–20.

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3. Berenguer, M., Prieto, M., Rayon, J.M. et al. (2000) Natural history of clinically compensated HCV-related graft cirrhosis following liver transplantation. Hepatology, 32, 852–58. 4. Fattovich, G., Giustina, G., Degos, F. et al. (1997) Morbidity and mortality in compensated cirrhosis type C: a retrospective study of 384 patients. Gastroenterology, 112, 463–72. 5. Charlton, M. and Wiesner, R.H. (2004) Natural history and management of hepatitis C infection after liver transplantation. Semin Liver Dis, 24, 79–88. 6. Forman, L.M., Lewis, J.D., Berlin, J.A. et al. (2002) The association between hepatitis C infection and survival after orthotopic liver transplantation. Gastroenterology, 122, 889–96. 7. Wiesner, R.H., Sorrell, M. and Villamil F. International Liver Transplant Society Expert Panel (2003) Report of the first International Liver Transplant Society expert panel consensus conference on liver transplantation and hepatic C. Liver Transpl , 9, S1–9. 8. Zekry, A., Bishop, G.A., Bowen, D.G. et al. (2002) Interhepatic cytokine profiles associated with posttransplantation hepatitis C virus-related liver injury. Liver Transpl , 8, 292–301. 9. Gopal, D.V. and Rosen, H.R. (2003) Duration of antiviral therapy for cholestatic HCV recurrence may need to be indefinite. Liver Transpl , 9, 348–53. 10. Charlton, M., Seaberg, E., Wiesner, R. et al. (1998) Predictors of patient and graft survival following liver transplantation for hepatitis C. Hepatology, 28, 823–30. 11. Prieto, M., Berenguer, M., Rayon, J.M. et al. (1999) High incidence of allograft cirrhosis in hepatitis C virus genotype 1b infection following transplantation; relationship with rejection episodes. Hepatology, 29, 250–56. 12. Neumann, U.P., Berg, T. and Bahra, M. (2004) Longterm outcome of liver transplants for chronic hepatitis C: a 10-year follow-up. Transplantation, 77, 226–31. 13. Burton, J.R. Jr. and Rosen, H.R. (2006) Acute rejection in HCV-infected liver transplant recipients: the great conundrum. Liver Transpl , 11 (Suppl 2), S38–47. 14. Klintmalm, G.B., Washburn, W.K., Rudich, S.M. et al. (2007) Corticosteroid-free immunosuppression with daclizumab in HCV(+) liver transplant recipients: 1-year interim results of the HCV-3 study. Liver Transpl , 13, 1521–31. 15. Bahra, M., Neumann, U.I., Jacob, D. et al. (2005) MMF and calcineurin taper in recurrent hepatitis C after liver transplantation; impact on histological course. Am J Transplant , 5, 406–11. 16. Everson, G.T., Trotter, J., Forman, L. et al. (2005) Treatment of advanced hepatitis C with a low accelerating dosage regimen of antiviral therapy. Hepatology, 42, 255–62. 17. Terrault, N.A. (2003) Prophylactic and preemptive therapies for hepatitis C virus-infected patients undergoing liver transplantation. Liver Transpl , 9, S95–100. 18. Neumann, U.P., Berg, T., Bahra, M. et al. (2004) Fibrosis progression after liver transplantation in patients with recurrent hepatitis C. J Hepatol , 41, 830–36.

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19. Firpi, R.J., Abdelmalek, M.F., Soldevila-Pico, C. et al. (2004) One-year protocol liver biopsy can stratify fibrosis progression in liver transplant recipients with recurrent hepatitis C infection. Liver Transpl , 10, 1240–47. 20. Blasco, A., Forns, X., Carrion, J.A. et al. (2006) Hepatic venous pressure gradient identified patients at risk of sever hepatic C recurrence after liver transplantation. Hepatology, 43, 492–99. 21. Carrion, J.A., Navasa, M., Garcia-Retortillo, M. et al. (2007) Efficacy of antiviral therapy on hepatitis C recurrence after liver transplantation: a randomized controlled trial. Gastroenterology, 132, 1746–56. 22. Berenguer, M., Palau, A., Fernandez, A. et al. (2006) Efficacy, predictors of response, and potential risks associated with antiviral therapy in liver transplant recipients with recurrent hepatitis C. Liver Transpl , 12, 1067–76. 23. Abdelmalek, M.F., Firpi, R., Soldevila-Pico, C. et al. (2004) Sustained viral response to interferon and ribavirin in liver transplant recipients with recurrent hepatitis C. Liver Transpl , 2, 199–207. 24. Bizollon, T., Ahmed, S.N., Radenne, S. et al. (2003) Long term histological improvement and clearance of intrahepatic hepatitis C virus following sustained response to interferon-ribavirin combination therapy in liver transplanted patients with hepatitis C virus recurrence. Gut , 52, 283–87. 25. Kornberg, A., Kupper, B., Tannapfel, A. et al. (2006) Antiviral treatment withdrawal in viremic HCV-positive liver transplant patients; impact on viral loads, allograft function and morphology. Liver Int , 26, 811–16. 26. Stravitz, R.T., Shiffman, M.L., Sanyal, A.J. et al. (2004) Effects of interferon treatment on liver histology and allograft rejection in patients with recurrent hepatitis C following liver transplantation. Liver Transpl , 10, 850–58. 27. Oton, E., Barcena, R., Garcia-Garzon, S. et al. (2005) Pegylated interferon and ribavirin for the recurrence of chronic hepatitis C genotype 1 in transplant patients. Transplant Proc, 37, 3963–64. 28. Biselli, M., Andreone, P., Gramenzi, A. et al. (2005) Pegylated interferon plus ribavirin for recurrent hepatitis C infection after liver transplantation in na¨ıve and non-responder patients on a stable immunosuppressive regimen. Dig Dis Sci , 38, 27–32. 29. Dumortier, J., Scoazec, J.Y., Chevallier, P. et al. (2004) Treatment of recurrent hepatitis C after liver transplantation; a pilot study of peginterferon alfa-2b and ribavirin combination. J Hepatol , 40, 669–74. 30. Chalasani, N., Manzarbeitia, C., Ferenci, P. et al. (2005) Peginterferon alfa-2a for hepatitis C after liver transplantation: two randomized, controlled trials. Hepatology, 41, 289–98. 31. Kugelmas, M., Osgood, M.J., Trotter, J.F. et al. (2003) Hepatitis C virus therapy, hepatocyte drug metabolism, and risk for acute cellular rejection. Liver Transpl , 9, 1159–65. 32. Azoulay, D., Linhares, M.M., Guguet, E. et al. (2002) Decision for retransplantation of the liver: an experienceand cost-based analysis. Ann Surg, 236, 713–21.

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33. McCashland, T., Watt, K., Lyden, E. et al. (2007) Retransplantation for hepatitis C: results of a U.S. multicenter retransplant study. Liver Transpl , 13, 1247–53. 34. Burton, J.R. Jr., Sonnenberg, A. and Rosen, H.R. (2004) Retransplantation for recurrent hepatitis C in the MELD era: maximizing utility. Liver Transpl , 10, S59–64. 35. Mutimer, D. (2006) Review article: hepatitis B and liver transplantation. Aliment Pharmacol Ther, 23 (8), 1031–41. 36. Dickson, R.C., Everhart, J.E., Lake, J.R., Wei, Y., Seaberg, E.C., Wiesner, R.H., Zetterman, R.K., Pruett, T.L., Ishitani, M.B. and Hoofnagle, J.H. (1997) Transmission of hepatitis B by transplantation of livers from donors positive for antibody to hepatitis B core antigen. The National Institute of Diabetes and Digestive and Kidney Diseases Liver Transplantation Database. Gastroenterology, 113 (5), 1668–74. 37. Davies, S.E., Portmann, B.C., O’Grady, J.G., Aldis, P.M., Chaggar, K., Alexander, G.J. and Williams, R. (1991) Hepatic histological findings after transplantation for chronic hepatitis B virus infection, including a unique pattern of fibrosing cholestatic hepatitis. Hepatology, 13 (1), 150–57. 38. Samuel, D., Muller, R., Alexander, G., Fassati, L., Ducot, B., Benhamou, J.P. and Bismuth, H. (1993) Liver transplantation in European patients with the hepatitis B surface antigen. N Engl J Med , 329 (25), 1842–47. 39. Faria, L.C., Gigou, M., Roque-Afonso, A.M., Sebagh, M., Roche, B., Fallot, G., Ferrari, T.C., Guettier, C., Dussaix, E., Castaing, D., Brechot, C. and Samuel, D. (2008) Hepatocellular carcinoma is associated with an increased risk of hepatitis B virus recurrence after liver transplantation. Gastroenterology, 134 (7), 1890–99; quiz 2155. 40. Devarbhavi, H.C., Cohen, A.J., Patel, R., Wiesner, R.H., Dickson, R.C. and Ishitani, M.B. (2002) Preliminary results: outcome of liver transplantation for hepatitis B virus varies by hepatitis B virus genotype. Liver Transpl , 8 (6), 550–55. 41. Burbach, G.J., Bienzle, U., Neuhaus, R. et al. (1997) Intravenous or intramuscular anti-HBs immunoglobulin for the prevention of hepatitis B reinfection after orthotopic liver transplantation. Transplantation, 63, 478–80. 42. Yilmaz, N., Shiffman, M.L., Stravitz, R.T. et al. (2007) Prophylaxis against recurrence of hepatitis B virus after liver transplantation: a retrospective study spanning 20 years. Liver Int , 28, 72–78. 43. Hooman, N., Rifal, K., Hadem, J. et al. (2008) Antibody to hepatitis B surface antigen trough levels and half-life do not differ after intravenous and intramuscular B immunoglobulin administration after liver transplantation. Liver Transpl , 14, 435–42. 44. Zuckerman, J.N. (2007) Review: hepatitis B immune globulin for prevention of hepatitis B infection. J Med Virol , 79, 919–21. 45. Ghany, M.G., Ayola, B., Villamil, F.G., Gish, R.G., Rojter, S., Vierling, J.M. and Lok, A.S. (1998) Hepatitis B virus S mutants in liver transplant recipients who were reinfected despite hepatitis B immune globulin prophylaxis. Hepatology, 27 (1), 213–22.

46. Perrillo, R.P., Wright, T., Rakela, J., Levy, G., Schiff, E., Gish, R., Martin, P., Dienstag, J., Adams, P., Dickson, R., Anschuetz, G., Bell, S., Condreay, L. and Brown, N. Lamivudine North American Transplant Group (2001) A multicenter United States–Canadian trial to assess lamivudine monotherapy before and after liver transplantation for chronic hepatitis B. Hepatology, 33 (2), 424–32. 47. Markowitz, J., Martin, P., Conrad, A. et al. (1998) Prophylaxis against hepatitis B recurrence following liver transplantation using combination of lamivudine plus hepatitis B immuneglobulin. Hepatology, 28, 585–89. 48. Fontana, R.J., Hann, H.W., Perrillo, R.P. et al. (2002) Determinants of early mortality in patients with decompensated chronic hepatitis B treated with antiviral therapy. Gastroenterology, 123, 719–27. 49. Marzano, A., Lampertico, P., Mazzaferro, V. et al. (2005) Prophylaxis of hepatitis B virus recurrence after liver transplantation in carriers of lamivudine-resistant mutants. Liver Transpl , 11, 532–38. 50. Rosenau, J., Bahr, M.J., Tillmann, H.L., Trautwein, C., Klempnauer, J., Manns, M.P. and B¨oker, K.H.W. (2001) Lamivudine and low-dose hepatitis B immune globulin for prophylaxis of hepatitis B reinfection after liver transplantation possible role of mutations in the YMDD motif prior to transplantation as a risk factor for reinfection. J Hepatol , 34 (6), 895–902. 51. Schiff, E.R., Lai, C.L., Hadziyannis, S. et al. (2007) Adefovir dipivoxil for pre- and post-liver transplantation patients with lamivudine-resistant mutants. Liver Transpl , 13, 349–60. 52. Han, S.B., Ofman, J., Holt, C. et al. (2000) An efficacy and cost-effectiveness analysis of combination hepatitis B immune globulin and lamivudine to prevent recurrent hepatitis B after orthotopic liver transplantation compared with hepatitis B immune globulin mono-therapy. Liver Transpl , 6, 741–48. 53. Naoumov, N.V., Lopes, A.R., Burra, P., Caccamo, L., Iemmolo, R.M., de Man, R.A., Bassendine, M., O’Grady, J.G., Portmann, B.C., Anschuetz, G., Barrett, C.A., Williams, R. and Atkins, M. (2001) Randomized trial of lamivudine versus hepatitis B immunoglobulin for long-term prophylaxis of hepatitis B recurrence after liver transplantation. J Hepatol , 34 (6), 888–94. 54. Gane, E.J., Angus, P.W., Strasser, S., Crawford, D.H., Ring, J., Jeffrey, G.P. and McCaughan, G.W. Australasian Liver Transplant Study Group (2007) Lamivudine plus low-dose hepatitis B immunoglobulin to prevent recurrent hepatitis B following liver transplantation. Gastroenterology, 132 (3), 931–37. Epub 2007 Jan 5. 55. Yoshida, H., Kato, T., Levi, D.M. et al. (2007) Lamivudine monoprophylaxis for liver transplant recipients with non-replicating hepatitis B virus infection. Clin Transpl , 21, 166–71. 56. Buti, M., Mas, A., Prieto, M. et al. (2003) A randomized study comparing lamivudine monotherapy after a short course of hepatitis B immune globulin (HBIg) and lamivudine with long-term lamivudine plus HBIg in the prevention of hepatitis B virus recurrence after liver transplantation. J Hepatol , 38, 811–17.

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57. Osborn, M.K., Han, S.H., Regev, A. et al. (2007) Outcomes of patients with hepatitis B who developed antiviral resistance while on the liver transplant waiting list. Clin Gastroenterol Hepatol , 5, 1454–61. 58. Roche, B., Feray, C., Gigou, M. et al. (2003) HBV DNA persistence 10 years after liver transplantation despite successful anti- HBs passive immunoprophylaxis. Hepatology, 38, 86–95. 59. Freshwater, D.A., Dudley, T., Cane, P. and Mutimer, D.J. (2008) Viral persistence after liver transplantation for hepatitis B virus: a cross-sectional study. Transplantation, 85 (8), 1105–11. 60. Chalasani, N., Smallwood, G., Halcomb, J., Fried, M.W. and Boyer, T.D. (1998) Is vaccination against hepatitis B infection indicated in patients waiting for or after orthotopic liver transplantation? Liver Transpl Surg, 4 (2), 128–32. 61. Lo, C.M., Fung, J.T., Lau, G.K. et al. (2003) Development of antibody to hepatitis B surface antigen after liver transplantation for chronic hepatitis B. Hepatology, 37, 36–43.

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62. S´anchez-Fueyo, A., Rimola, A., Grande, L., Costa, J., Mas, A., Navasa, M., Cirera, I., S´anchez-Tapias, J.M. and Rod´es, J. (2000) Hepatitis B immunoglobulin discontinuation followed by hepatitis B virus vaccination: a new strategy in the prophylaxis of hepatitis B virus recurrence after liver transplantation. Hepatology, 31 (2), 496–501. 63. Angelico, M., Di Paolo, D., Trinito, M.O. et al. (2002) Failure of a reinforced triple course of hepatitis B vaccination in patients transplanted for HBV-related cirrhosis. Hepatology, 35, 176–81. 64. Manzarbeitia, C., Reich, D.J., Ortiz, J.A., Rothstein, K.D., Araya, V.R. and Munoz, S.J. (2002) Safe use of livers from donors with positive hepatitis B core antibody. Liver Transpl , 8, 556–61. 65. Kim, W.R., Benson, J.T., Hinman, A. et al. (2007) Decline in need for liver transplantation for endstage liver disease secondary to hepatitis B in the United States. Hepatology, 46, 238A.

PART SIX : HORIZONS

56

Tissue Engineering of the Liver Gregory H. Underhill1, Salman R. Khetani1, Alice A. Chen1 and Sangeeta N. Bhatia1,2 1 Harvard-MIT

Division of Health Sciences and Technology/Electrical Engineering and Computer Science, Massachusetts Institute of Technology, Boston, MA, USA 2 Division of Medicine, Brigham & Women’s Hospital, Boston, MA, USA

INTRODUCTION Cell-based therapies for liver disease and failure offer the potential to augment or replace whole organ transplantation; however, the development of such therapies poses unique challenges, largely stemming from the complexity of liver structure and function. The field of liver tissue engineering encompasses several approaches collectively aimed at providing novel therapeutic options for liver disease patients and elucidating fundamental characteristics of liver biology. These approaches include the development of in vitro model systems that recapitulate normal liver function and also three-dimensional implantable therapeutic constructs. Advances in both of these areas are reviewed in this chapter within the context of current treatments for liver disease and additional clinical alternatives such as extracorporeal bioartificial liver devices and cell transplantation strategies.

CELL-BASED THERAPIES FOR LIVER DISEASE AND FAILURE Liver failure is a significant health problem, representing the cause of death of over 40 000 individuals in the United States every year [1], and can be generally separated into two major categories: fulminant hepatic failure, The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

also referred to as acute liver failure, and chronic hepatic failure resulting from chronic end-stage liver disorders. Liver transplantation is the only therapy shown to alter mortality directly, and therefore is the standard-of-care in most clinical settings. In order to expand the supply of available livers, several surgical options have been pursued, including the use of non-heart-beating donors or split liver transplants from cadaveric or living donors [2]. Partial liver transplants take advantage of the body’s role in the regulation of liver mass and the significant capacity for regeneration exhibited by the mammalian liver [3]. However, liver regeneration is difficult to control clinically, and although partial liver transplants have demonstrated some effectiveness, biliary and vascular complications are major concerns in these procedures [2]. Furthermore, in spite of these surgical advances and improvements in organ allocation, there is an increasing divergence between the number of patients awaiting transplantation and the number of available organs [4], suggesting that it is unlikely that liver transplantation procedures alone will meet the increasing demand. Consequently, alternative approaches are needed and are actively being pursued including several non-biological extracorporeal support systems such as plasma exchange, plasmapheresis, hemodialysis, and hemoperfusion over charcoal or various resins [5–7]. These systems have shown limited success, probably due to the narrow range of functions inherent to these devices. The liver exhibits a complex

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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THE LIVER: CELL-BASED THERAPIES FOR LIVER DISEASE AND FAILURE

Figure 56.1 Cell-based therapies for liver disease. Extracorporeal devices perfuse the patient’s blood or plasma through bioreactors containing hepatocytes. Hepatocytes are transplanted directly or implanted on scaffolds. Transplants with transgenic animal tissue have also been pursued in order to reduce complement-mediated damage of the endothelium [5]. Reproduced from Allen et al. (2001) with permission

array of over 500 functions, including detoxification, synthetic, and metabolic processes. Hence recapitulation of a substantial number of liver functions will be required to offer sufficient liver support. As a means to provide the multitude of known and also currently unidentified liver functions, cell-based therapies have been proposed as an alternative to both organ transplantation and the use of strictly non-biological systems [8]. These therapies encompass approaches aimed at providing temporary support such as extracorporeal bioartificial liver (BAL) devices and also more permanent adjunct interventions such as cell transplantation, transgenic xenografts, and implantable hepatocellular constructs (Figure 56.1).

Extracorporeal bioartificial liver devices One promising approach for cell-based therapies for liver failure is the development of extracorporeal support devices, which, analogous to kidney dialysis systems, would process the blood or plasma of liver failure patients. These devices are principally aimed at providing temporary support to liver failure patients to enable sufficient regeneration of the host liver tissue or to serve as a bridge to transplantation. In particular, substantial efforts have been made towards the development of extracorporeal BAL devices containing hepatic cells which would exhibit the myriad of critical liver functions and could be employed in a clinical setting. BAL devices that have been proposed and studied can be broadly categorized into four main types, and have been

reviewed extensively elsewhere [9, 10]. These include hollow-fiber devices, flat plate, and monolayer systems, perfusion bed or porous matrix devices, and suspension reactors (Figure 56.2), with each of these general designs exhibiting innate advantages and disadvantages. Overall, an effective BAL device would satisfy several important criteria, including maintenance of cell viability and hepatic functions, efficient bidirectional mass transfer, and scalability to therapeutic levels. A number of BAL devices have been tested clinically, and improvements in device and trial design continue to be implemented. Ultimately, even if clinical trials of the current BAL systems do not demonstrate sufficient efficacy, information obtained from these studies coupled with improvements in cell sourcing and functional stabilization will represent the foundation for the next generation of devices.

Cell transplantation and sourcing In addition to temporary extracorporeal support, the development of cell-based therapies for liver treatment aimed at the eventual replacement of damaged or diseased tissue is an active area of investigation. One potential cell-based approach is the transplantation of isolated mature hepatocytes. In experiments utilizing rodent models, transplanted hepatocytes were demonstrated to exhibit substantial proliferative capacity and the ability to replace diseased tissue under certain conditions [11–13]. The in vivo proliferation of transplanted hepatocytes is highly dependent on the presence of an adequate “regenerative” environment. In animal models, regenerative stimulation is provided by transgenic injury, partial hepatectomy, portacaval shunting, or the administration of hepatotoxic agents prior to cell transplantation. Similar to other cell-based approaches, the feasibility of hepatocyte transplantation is constrained by the availability of allogeneic human hepatocytes. Only a limited supply of human hepatocytes is currently available from collagenase perfusion of organs regarded as inappropriate for transplantation. Interestingly, the transplantation and subsequent proliferation of human hepatocytes in genetically altered mouse strains have been proposed as an approach not only to generate humanized mouse models, but also as a potential human hepatocyte expansion platform [14–17]. Despite the significant proliferative capacity during regenerative responses in vivo, mature hepatocyte proliferation in culture is limited, particularly for human hepatocytes [18], and as discussed in more detail later, hepatocytes exhibit a loss of liver-specific functions under many conditions in vitro. As a result, alternative cell sources for liver cell-based therapies are being investigated, such as various stem cell populations, which can retain significant proliferative ability in vitro and exhibit either pluripotency or multipotency, thereby constituting a possible source of hepatocytes in addition to other liver cell

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Hollow Fiber Devices

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Figure 56.2 Schematics of cell-based bioreactor designs. The majority of liver cell-based bioreactor designs fall into four general categories: hollow-fiber devices, flat plate systems, perfusion beds, and suspension reactors [8]. Reproduced from Allen et al. (2001) with permission

types. Through the formation of embryoid bodies and the addition of various growth factors, early studies suggested that pluripotent embryonic stem cells could be induced to differentiate the hepatocyte lineage in culture [19–21]. More recent approaches have employed co-culture configurations and refinements in the type and timing of growth factor additives with the aim of improving this differentiation efficiency, and concurrently, have more fully explored both in vitro and in vivo the functional capacity of the differentiated populations [22–24]. In addition to embryonic stem cells, a wide range of fetal or adult stem/progenitor populations have similarly been investigated. For instance, multipotent adult progenitor cells (MAPCs) derived from bone marrow [25], and also various mesenchymal stem cell preparations [26–30], have been shown to exhibit hepatocyte differentiation potential. Moreover, recent reports suggest that amniotic fluid may contain cells with the capacity for multilineage (including hepatic) differentiation [31–33]. Fetal hepatoblasts and oval cells are also intriguing possible cell types for liver cell-based therapies. Hepatic development proceeds through the differentiation of liver precursor cells, termed hepatoblasts, which exhibit bipotential differentiation capacity, defined by the ability to differentiate into both hepatocytes and bile duct epithelial cells [34]. Notably, in certain types of severe and chronic liver injury, an adult progenitor cell population termed oval cells, which share many phenotypic markers and functional properties with fetal hepatoblasts, mediates compensatory liver repair through a similar differentiation program [35, 36]. Work by Weiss and colleagues demonstrated the development of bipotential mouse embryonic liver (BMEL) cell lines that are derived from mouse E14 embryos and exhibit characteristics reminiscent of fetal hepatoblasts and oval cells [37]. These cell lines are non-transformed, proliferative, demonstrate upregulation of hepatocyte or bile duct epithelial markers under distinct culture conditions in vitro [37], and exhibit the capacity to home to the liver and undergo bipotential differentiation in vivo within a regeneration context [38]. Overall, elucidation of the mechanisms regulating the proliferation and hepatic differentiation of stem and progenitor cell populations could lay the groundwork for the development of robust cell-based therapies.

However, although diverse stem and progenitor cell types exhibit vast potential for integration into hepatic treatments, many challenges remain, including the ability to dictate differentiation completely, particularly for ex vivo applications and within multicellular systems. Furthermore, regardless of the cell source, the stabilization of hepatocyte functions outside the liver remains a primary issue. Microenvironmental signals including soluble mediators, cell–extracellular matrix interactions, and cell–cell interactions have been implicated in the regulation of hepatocyte function. Accordingly, the development of robust hepatocyte in vitro culture models which allow for the controlled reconstitution of these environmental factors is a fundamental prerequisite towards a thorough understanding of mechanisms regulating hepatocyte processes and the improved functionality of liver cell-based therapies.

IN VITRO HEPATIC CULTURE MODELS One of the major thrusts in hepatic tissue engineering is an effort towards improvements in the functionality and utility of current in vitro culture platforms. In the past, an extensive range of liver model systems have been developed, some of which include: perfused whole organs and wedge biopsies; precision cut liver slices; isolated primary hepatocytes in suspension or cultured upon extracellular matrix; immortalized liver cell lines; isolated organelles, membranes or enzymes; and recombinant systems expressing specific drug metabolism enzymes [39–41]. While perfused whole organs, wedge biopsies, and liver slices maintain many aspects of the normal in vivo microenvironment and architecture [39, 42, 43], they typically suffer from short-term viability (3 months in cultured HepG2 cells [41]. Similarly to the FVs, lentiviruses, which are gammaretroviruses, form PICs that can be translocated through the pores of intact nuclear membranes. They are, in fact, being explored as potential vectors for gene transfer into non-dividing cells. However,

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lentivirus-based gene transfer in the liver in vivo may also require the hepatocytes to be in cell cycle [42]. The retroviral LTR contains a strong enhancer that can activate protooncogenes near its site of integration, causing neoplastic transformation in some cases [43]. Promoters and enhancers within the U3 region of the 3′ LTR are copied into the 5′ LTR during reverse transcription (RT) into double-stranded DNA. Therefore, deletions introduced into this region result in the loss of functional U3 regions in both the 5′ and the 3′ LTR. These “self-inactivating” (SIN) vectors, in which the transgene of interest is driven by internal promoters, are considered to be less liable to cause insertional activation of a deleterious gene by the integrated proviral genome [44]. As a further precaution against insertional activation by the enhancer contained in the transcription unit expressing the therapeutic gene, insulator sequences are being explored for use as barriers to the potential activation of promoters of genes near the insertion site [45].

Recombinant Adeno-associated Virus Adeno-associated virus-2 (AAV-2) is a small (4.7 kb) single-stranded DNA virus of the parvovirus family that integrates preferentially on the q13.4-ter arm of human chromosome 19 [46]. When infection with a “helper virus,” such as adenovirus [47] or herpes simplex virus [48] occurs, AAV sequences are “rescued” from the integration site, resulting in productive lytic infection. In the absence of a helper virus, the infection becomes latent, and the viral genome remains integrated in the host chromosome for long periods. A permissive state for AAV replication can also be induced by the treatment of cells with a variety of genotoxic stimuli, such as heat shock, hydroxyurea, UV light, and irradiation [49–51]. Thus, it is proposed that the helper virus proteins are not directly involved in AAV replication, but provide factors, such as adenoviral proteins E1A and E1B, that maximize the synthesis of the AAV gene products and host proteins necessary for AAV replication [52]. There are 11 known serotypes of human AAV, and new serotypes continue to be added to the list. While the AAV type 2 is used most commonly for vector development, characterization of other serotypes is under way. The AAV type 2 receptor is a membrane-associated heparin sulfate proteoglycan, which is present in many cell surfaces, thus explaining the broad host infectivity of this virus. Receptor binding is required for its internalization. The wild-type AAV is not known to cause human disease and has a broad range of infectivity. The AAV genome comprises three promoters (p5, p19, and p40), a polyadenylation signal, a non-structural gene (Rep) and the structural Cap gene. At both ends of the viral genome are 145 bp inverted terminal repeats (ITRs) that form T-shaped hairpin structures. The ITRs are needed for integration of the virus into the host genome [53]. The 3′ -OH

end of the ITR primes second-strand synthesis, generating a double-stranded DNA with a covalently closed hairpin at one end. Rep mediates nicking at the terminal resolution site, thereby generating a linear duplex, which is the substrate for a new round of replication. The second-strand synthesis is needed for gene expression and integration of the wild-type virus. The ITR hairpin structures and cellular recombination pathways are required for viral integration. The viral Rep protein also directs the site specificity of AAV integration into chromosome 19. The viral genome integrates as a head-to-tail concatamer. To generate recombinant AAV vectors, all viralencoded genes, approximately 96% of the viral genome, can be replaced with foreign DNA of choice and packaged into an AAV virion. The cis-acting AAV ITRs that are retained do not appear to contain dominant enhancer/ promoter activity. Thus, the expression of the transgene is driven by the transcriptional regulatory elements inserted in the expression cassette. This ability of AAV vectors to carry regulatory elements, such as tissue-specific enhancers/promoters, and splice sites, without interference from the viral genomes allows for greater control of transferred gene expression. When toxic gene products, such as suicide genes, are expressed, a tight transcriptional control of the transgene using tissue or cell-specific promoter is critical in achieving a high therapeutic ratio. Tumor-specific expression of HSV-TK gene under the control of α-fetoprotein/albumin promoter was obtained in hepatocellular carcinoma cell lines [54]. Recombinant adenoassociated virus (rAAV) vectors are generated by cotransfecting into packaging 293 cells two plasmids, one containing the transgene, flanked by ITRs, and the other encoding Rep, Cap and adenoviral proteins (Figure 59.2). The 293 cells provide E1A in trans. Alternatively, the recombinant AAV can be generated by infection with a helper virus such as adenovirus or herpes simplex virus. When helper virus infection is utilized, the recombinant AAV needs to be purified from the helper virus. The viral Rep protein, which directs the site specificity of AAV integration into chromosome 19, is absent in the recombinant vectors, which, therefore, lose the ability to integrate and remain in the nucleus as episomes [55]. Delayed attainment of a plateau of transgene expression is characteristic of AAV vectors. Initial trials of intramuscular administration of recombinant AAV-2 vectors expressing coagulation factor IX in patients with hemophilia B resulted in low levels of protein production [56]. Portal vein infusion of higher doses of rAAV in factor IX-deficient dogs achieved 5% of normal levels of factor IX activity in plasma [57]. The effect of rAAV-mediated cancer gene therapy could be augmented when used in combination with conventional tumor therapies, such as irradiation or chemotherapy [58]. Although the greatest experience with AAV-mediated gene transfer has been based on the use of rAAV-2, it is now clear that various AAV serotypes have distinct tissue tropism,

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Figure 59.2 Production of recombinant adenoassociated virus (AAV). Two plasmids, one containing the transgene, flanked by inverted terminal repeats (ITRs) and another encoding the helper proteins, rep and cap proteins are transfected into 293 packaging cells. AAV is generated by infecting the transfected cells with a helper virus (such as, E1A-deleted adenovirus) or by transfecting a plasmid expressing the critical adenoviral protein. If a helper virus is used, the recombinant AAV is purified subsequently from the helper adenovirus

depending on their capsids. Eleven different serotypes, AAV-1 through AAV-11, and over 100 variants have been isolated from human and non-human tissues [59]. AAV serotypes 1–9 are currently being developed as gene therapy vectors. Since the AAV-2 genome is best characterized, AAV-2 genome has been cross-packaged with capsids of other serotypes. AAV-2/AAV-8 hybrids are particularly efficient for transferring genes into hepatocytes, although the pancreas, heart, and skeletal muscles are additional targets of this hybrid vector. rAAV vectors can persist for up to 1 year in hepatocytes, but they are lost eventually because of their episomal nature. The ability to readminister rAAV vectors is limited by host humoral and cellular immune responses. However, the use of pseudotyped rAAV containing capsids of alternative serotypes may permit repeated administration. Efforts are under way to generate AAV vectors containing capsids with desired characteristics by mutagenesis of the Cap gene. An alternative approach consists of “guided evolution” in which random DNA recombination is used to generate vectors with certain desired attributes, such as the ability to avoid recognition by antibodies and cytotoxic lymphocytes, or to transfer genes into specific tissues with high efficiency [60]. The small genome of AAV accommodates transcription units of limited size only. To expand the packaging

capacity of AAV, two recombinant vectors are generated. A 5′ vector includes the enhancer/promoter cassette and a half-intron-carrying a splice donor site. A complementing 3′ vector contains another half-intron-carrying a splice acceptor site linked with the cDNA encoding the target gene and a polyadenylation signal. Following coinfection into cells (e.g. 293 cells), genomes of the 5′ and 3′ vectors dimerize, generating a larger vector that expresses the target gene [61].

Simian virus 40-based vectors Recombinant simian virus 40 (SV40) is a non-enveloped DNA virus of the papova family with a 5.2 kb circular double = stranded genome. The large (Tag) and small (tag) T antigens, that are expressed by differential splicing of a single RNA transcript, are required for transcription of the viral structural genes, VP1, VP2, and VP3. Tag is the most immunogenic protein of SV40 and is able to impart an immortalizing effect on the cell. In the recombinant viral genome, the Tag gene is replaced by a target transgene (Figure 59.3). Viral particles are generated by transfecting the recombinant genome into COS-7 cells that provide Tag in trans. Because the recombinant virus lacks the Tag gene, it cannot replicate [62], and its immunogenicity is markedly reduced [63]. The SV40 is a small

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Figure 59.3 Generation of a replication-defective recombinant SV40 virus. The full-length SV40 genome is cloned into a plasmid from which the coding region of the T-antigen (Tag) is deleted and replaced by the transgene. The recombinant viral genome is excised and recircularized. The recircularized genome is transfected into COS cells, which provide T-antigen in trans, generating a helper-free replication-defective recombinant SV40

virus and the capacity of the recombinant SV40 to accommodate exogenous DNA is limited to 4.7 kb. SV40 vectors can be concentrated to 1012 infectious units per milliliter [64]. Preliminary studies suggest that the recombinant SV40 integrates into the host chromosomal DNA. The ability of these vectors to infect non-dividing cells makes them attractive for liver-directed gene therapy [65, 66].

Recombinant adenovirus Adenoviruses are large, linear, double-stranded DNA viruses. Vectors derived from human adenovirus type 5 and type 2 are commonly used for gene transfer. These vectors can be generated at high titers and can express transgenes in both dividing and quiescent cells [67, 68]. In addition, following systemic administration in animals, the virus is preferentially localized to the liver, leading to the transduction of the great majority of hepatocytes in

vivo [69]. The coxsackie adenovirus receptor, CAR, that is also involved in the internalization of the Coxsackie virus, is highly concentrated in rodent liver. Following infection, the virus exits the endocytotic vesicles and translocates to the nucleus, where it persists episomally, without integration into the host genome. It is uncertain, however, whether such efficient gene transfer occurs in the human hepatocytes in vivo. Recombinant adenoviral vectors are produced by disrupting the E1 domain which encodes transcription factors required for the expression of other adenoviral genes by cloning the transgene of interest into this region. Additional adenoviral genes, for example E3 and E4, may be deleted to increase the “stuffing space” in the vector. The recombinant virus is generated in a helper cell line that provides the viral proteins in trans [70]. Application of adenoviral vectors in gene therapy is limited by the highly immunogenic nature of the viral proteins. Host neutralizing antibodies block gene transfer by

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repeated administration of the vector. Anti-adenoviral cytotoxic lymphocytes attack the adenovirally infected host cells, causing hepatitis and rapid loss of the transgene after secondary gene transfer [71]. To prevent any de novo expression of adenoviral genes, “gutless” vectors, devoid of viral structural genes, have been developed. Although these vectors exhibit prolonged transgene expression [72], immunogenicity can be retained because of the presence of viral proteins provided in trans by the packaging cells during generation of the recombinant virus [73]. Therefore, repeated gene transfer requires the generation of vectors based on a different strain of adenovirus. Expression of immunomodulatory genes, such as adenoviral E3, by the recombinant vector is being explored for abrogating adenovirus-specific host immune response [74]. Alternative strategies seek to tolerizing the host specifically to adenoviral antigens, leaving the general immune system intact. Injection of recombinant adenovirus in utero [75], to newborn rats [69], intrathymic inoculation of adenoviral proteins in young adult rats [76], or oral administration of small doses of adenoviral proteins in adult rats [77] have been used successfully to abrogate humoral and cell-mediated immune response against adenoviral antigens in rodents. Inhibition of costimulation between antigen-presenting cells and cytotoxic lymphocytes by the administration of CTLA4-Ig and CD-40 antibodies to the host has also been explored in rodents, with limited success [78]. Although the tolerization methods permit repeated administration of adenoviral vectors in experimental animals, safety concerns remain regarding tolerization of human hosts to adenoviruses, which, in the wild form, are human pathogens.

Recombinant baculovirus The recombinant baculovirus Autographa californica nuclear polyhedrosis (AcNPV) is commonly used for generating recombinant proteins in insect cells [79]. Among mammalian cells, hepatocytes can be infected by baculoviral vectors and the transgene is expressed with the use of internal promoters that function in mammalian cells. Recombinant baculoviruses can accommodate large segments of exogenous DNA. Although the gene transfer efficiency and immunogenicity of recombinant baculoviruses in vivo have not been well characterized, recombinant vectors consisting partly of baculoviral sequences and partly of sequences from other viruses are being evaluated for in vivo application.

Herpes simplex virus-1 Herpes simplex virus-1 (HSV-1) is a 150 kb doublestranded DNA virus with a broad host range [80, 81]. These vectors efficiently infect non-dividing cells and are useful for gene transfer into neuronal cells and hepatocytes. However, long-term gene expression in the liver

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has not been achieved with the currently available HSV vectors.

NON-VIRAL VECTORS Although viral vectors have been widely utilized in gene therapy research, owing to their efficient delivery of transgenes into cells, non-viral gene transfer methods can be potentially less toxic and easier to standardize. Three major categories of synthetic non-viral delivery systems for systemic delivery of nucleic acids to tissues have been studied extensively [82]. (1) Lipid-based delivery systems encompassing both lipid-encapsulated and cationic lipid–nucleic acid complexes (lipoplex) provided the initial proof-of-principle for systemic transgene delivery [83]. (2) Polyplex systems consist of complexes formed by the addition of nucleic acid to a polycation, such as poly-l-lysine [84, 85], polyethylenimine (PEI) [86], polyglucosamines [87, 88], lipopolyamines [89], and cationic peptides [90], generating water-soluble complexes that are efficient delivery systems. Free amino groups on these agents permit the attachment of a variety of targeting ligands. (3) Lipopolyplex delivery systems are hybrid complexes that contain both polycationic polymers and lipids. Compaction of high molecular weight DNA with polycations prior to lipid encapsulation/complexation produces a DNA core surrounded by a lipid shell, reducing the final particle size and protecting the nucleic acid from nuclease degradation. The cationic lipid transfecting agents such as lipofectamine, DOTAP(1-oleoyl-2-[6-[(7-nitro-2-1,3-benzoxadiazol-4-yl) amino]hexanoyl]-3-trimethylammonium propane), and the cationic polymers, including poly-l-lysine and PEI, have improved nucleic acid transfer across the plasma membrane into the cytoplasm [91, 92]. Unfortunately, these nucleic acid–cationic complexes are cytotoxic because of their large particle size [93] and the high positive zeta potential required for uptake of the complexes via non-specific cation-mediated endocytosis [86,94–96]. Plasma proteins reduce the transfection efficiency of these particles both in vitro and in vivo by neutralizing the zeta potential and increasing the particle size to >100 nm [97–100]. Cytotoxicity of the cationic-based delivery systems is reduced by incorporation of poly(ethylene glycol) (PEG), which stabilizes, prevents aggregation, reduces binding to serum proteins, and maintains the small size required for endocytosis [101]. However, the shielding of the cationic charge by PEG reduces the transfection efficiency of the PEG-modified delivery systems. To overcome these hurdles, and to make the delivery system cell-type specific, ligands are utilized to promote receptor-mediated delivery. For example, asialoorosomucoid (ASOR) [84, 85] and galactose [89, 102, 103] have been conjugated to polylysine, lipopolyamines, or

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PEI for targeting to the asialoglycoprotein receptor (ASGPR) on hepatocytes. Lipid-based delivery systems have utilized galactocerebrosides as the targeting moiety for the ASGPR [103–105]. Receptor-mediated ligand targeting has also been exploited using transferrin, folate, and cell-specific antibodies conjugated to the polycations or liposomes [106–109]. These ligand-targeted systems have been shown to increase hepatocyte-directed gene delivery both in vitro [105, 109, 110] and in vivo [84, 111]. The ligand/receptor-mediated endocytosis obviates the need for an overall net positive charge. In fact, negatively charged particles promote more effective ligand/receptor-mediated nucleic acid delivery [107]. Additionally, ligand/receptor-mediated delivery systems are, unlike cationic lipids or polycations, able to transfect non-adherent cell lines efficiently [108]. Recently, significant reductions in size of non-viral particles targeting the ASGPR hepatocytes have been reported [112–114]; yet some off-target delivery was still observed in vivo. A promising new non-viral liver delivery system has been described [115] in the sub-50 nm range that is able to encapsulate oligonucleotides or plasmids up to 12.8 kb efficiently, exhibits a negative zeta potential, and is a true capsule with the nucleic acid cargo completely enclosed by a shell composed of a targeted receptor’s ligand. The use of hyaluronan (HA) as the capsule shell targeting the liver sinusoidal endothelial cells (LSECs) HA receptors [116] or ASOR for the hepatocyte ASGPR [117] can achieve cell-type specific liver targeting in vivo, with no apparent off-target delivery [115]. Although hepatocyte-targeted gene delivery has been achieved in vivo using ASGPR-mediated endocytosis, transgenes delivered via receptor-mediated endocytosis are naturally translocated to lysosomes, where a large fraction of it is degraded, resulting in a low level of expression. Disruption of the translocation endosomes to lysosomes, for example, by transient depolymerization of microtubules, prolongs transgene expression, but the DNA remains compartmented in cytoplasmic vesicular pools, whereby the level of transgene expression remains at a low level [118]. Destabilization of endosomal vesicles, for example, by using proton sponges (e.g. PEI) or disruptor peptides have been incorporated into the delivery vehicles to promote endosomal release of the nucleic acid into the cytoplasm [119, 120]. A novel alternative approach employs proteoliposomes containing the naturally galactose-terminated F-glycoprotein of the Sendai virus envelope [121]. In the absence of the other Sendai virus envelope protein (hemagglutinin), binding of F-protein to the cell surface is dependent on ASGPR, making it hepatocyte specific. However, the fusogenic activity of the F-protein results in deposition of the contents of the proteoliposome directly into the cytosol, bypassing the endosomal pathway, thereby enhancing transgene expression. Rapid clearance

of the gene delivery vehicle from the plasma by hepatocytes reduces the exposure of other tissues to the gene transfer vehicle, markedly reducing its immunogenicity [122]. Translocation of the transgenes from the cytosol to the nucleus can be enhanced by incorporating nuclear localization signal peptides on the plasmid constructs [123]. In one example, by including a muscle-specific transcription factor binding site in the construct, the introduced transgene demonstrated tissue-specific nuclear localization, while remaining in the cytoplasm of non-muscle cells [124]. It should be noted, however, that expression of episomal DNA is inherently transient. For long-term expression, especially for the treatment of inherited diseases, it may be desirable to use a system that permits integration of the transgene into the host genome. During the last few years, a transposon system, termed Sleeping Beauty, which was originally found in mutated form in salmonid fish [125], has been used to promote transgene integration at a TA dinucleotide site via a cut-and-paste mechanism [126]. In the current version, the system consists of a plasmid containing two transcription units, one expressing the enzyme, Sleeping Beauty transposase, and the other expressing the target DNA. The transcription unit expressing the target DNA is flanked by inverted/direct repeat sequences that are cleaved by the expressed transposase, followed by insertion of the transgene into the host genome. The Sleeping Beauty transposition system has been used recently in UGT1A1-deficient jaundiced Gunn rats, resulting in long-term amelioration of hyperbilirubinemia [127, 128]. In addition, the Sleeping Beauty system using hydrodynamic push [129] to deliver naked plasmid vectors to the liver has been used successfully for persistent expression of coagulation factor IX, [130] factor VIII [131], α1 -antitrypsin [132], β-glucuronidase [133] or α-l-iduronidase [133], and fumarylacetoacetate hydrolase (FAH) deficiency [134]. In all cases, significant to complete correction of the disease phenotype was observed. However, immune consequences resulting in loss of phenotypic correction were observed with both factor VIII [131] and α-l-iduronidase [133], in part related to the non-specific delivery of the naked DNA by hydrodynamic push to hepatocytes, LSECs, and Kupffer cells [135]. By coupling the Sleeping Beauty transposon system with targeted cell-type specific delivery, long-term persistent factor VIII levels and phenotypic correction of the bleeding diathesis were achieved [136]. These studies, coupled with the genomic Sleeping Beauty-mediated insertion profile, which although not entirely random for the TA dinucleotide selected indicate that it preferentially inserts into non-transcribed and intronic regions of the genome [137–139], suggest that clinical liver gene therapy trials using Sleeping Beauty will occur in the near future.

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TARGETED GENE MODIFICATION Homologous recombination In many experimental systems, the successful transfer of a cloned, modified gene into the genome of the host organism is now possible [140, 141]. Ideally, the introduced gene is returned only to its homologous location and is integrated at the target site. The process is a precise means for repairing mutated or damaged DNA, insuring accurate chromosomal disjunction during meiosis. Our understanding of the process of homologous pairing and DNA strand exchange is predominately derived from extensive biochemical analyses of purified RecA protein from Escherichia coli and genetic studies carried out in bacteriophage and yeast. As structural homologs of yeast recombination proteins, rad51 and/or rad52 and also many of the other proteins involved in the homologous recombination pathway have been identified in higher eukaryotes [142]. The transformation protocols studied in lower eukaryotes ultimately developed into the strategies for gene targeting in higher organisms. However, the observed genetic contradictions and the lack of biochemical data hindered identification of precise mechanisms to pursue experiments in higher organisms. Since the precise duplication of genomic DNA is vital to the survival of a species, eukaryotic evolution has produced elaborate and redundant mechanisms to limit the effects of mutagens and exogenous genetic invaders. Unfortunately, even with the explosive increase in studies designed to characterize the various genomic DNA repair pathways in cultured cells from higher eukaryotes, successful targeted modification of genomic DNA by homologous recombination has been limited [4]. Significant impediments remain to be resolved for this process to function effectively as an in vivo approach to gene therapy. First, the integration of exogenous DNA into the genome is extremely inefficient, and can occur in the absence of sequence homology between the introduced gene and genomic integration site [143]. Second, as a defensive strategy, sequences in the chromatin are sequestered, thus promoting a low efficiency of pairing between the introduced DNA and its genomic target [144]. However, the utility in vitro for targeted gene disruption to establish the functional activity of a particular gene has been fairly successful [145, 146]. With our growing understanding of how homologous recombination is regulated in mammalian cells, the process of generating only the desired integration at the target site with significant increases in the frequencies is possible. In particular, a 1% frequency of targeted homologous replacement or generation of the 3 bp deletion associated with cystic fibrosis transmembrane conductance regulator

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using a short (488 nucleotide) DNA fragment indicated potential for therapeutic use [147]. Use of single-stranded DNA produced by bacteria phage has also proved successful [148], dramatically reducing the labor-intensive nature of this approach. Moreover, using rAAV vectors as a source of the single-stranded DNA, in vivo gene repair of β-glucuronidase [149] and FAH deficiency [150] has been achieved in the livers of transgenic mice. Ironically, many of the factors involved with improving the frequencies of targeted gene replacement require inactivation of other DNA repair proteins and/or pathways [4], which is not feasible in an in vivo setting. Additionally, homologous recombination is cell cycle regulated with reduced activity outside of S phase, precluding its effective use in many quiescent cell types, such as hepatocytes [151–154]. Recently, a new approach to targeted gene replacement based on the use of zinc finger nucleases (ZFNs), resulting in improved efficiency of gene targeting by introducing DNA double-strand breaks in target genes, which then stimulate the cell’s endogenous homologous recombination machinery, has been reported [155]. Moreover, the ZFNs can be engineered to target specifically unique DNA sequences, providing the ability to target almost any region of the genome [156]. Using this approach, gene replacements using engineered ZFNs as large as 7.7 kb have been efficiently introduced into the host genome of mammalian cells [157]. In addition, the ZFNs have been used successfully for selectively targeting and degrading mutated mitochondrial DNA in cultured cells [158]. The significant enhancement in targeted gene replacement stimulated by ZFNs indicates its potential for use in quiescent cells such as hepatocytes.

Triplex DNA The use of triplex DNA to perform site-specific modification of genomic DNA is based on the formation of a three-stranded or triple-helical nucleic acid structure. In short, the exogenous third strand of nucleic acid binds in the major groove of a homopurine region of the DNA, forming Hoogsteen or reverse Hoogsteen hydrogen bonds with the purine base [159]. The triplex formation can occur at physiological pH, but the polypurine regions must be guanine rich and 12–14 nucleotides in length for adequate triplex formation to occur. Additionally, monovalent cations such as Na+ and K+ inhibit triplex formation, rendering their formation under physiological conditions difficult. Modified bases such as the thymidine purine analog 7-deaza-2′ -deoxyxanthosine can counteract the inhibitory effect of monovalent cations on triplex formation, in addition to maintaining an all-purine backbone motif [160]. The initial approach to the triple-helix site-directed modification of DNA used cross-linking agents, such as psoralen or other mutagens, to attach covalently to the

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triplex forming oligonucleotide [161]. After intercalation of the psoralen at the target 5′ ApT3′ site in the DNA, UV irradiation results in cross-linking of the thymines in the two strands. This substrate is then repaired by the endogenous DNA repair activity in the cell, producing the characteristic T:A to A:T transversions at a low frequency. This technique has been used to modify episomal DNA in mammalian cells in vitro [162], and targeted gene knock-outs of the genomic hypoxanthine phosphoribosyl transferase gene in cultured cells were successfully created [163]. However, rather than the expected T to A transversions, the majority of the knock-outs resulted predominately from small deletions and some insertions, suggesting that the endogenous repair pathway involved was not that of mismatch repair. Further investigation indicated that these triple helix-forming oligonucleotides could promote insertions and deletions at target sites in the absence of cross-linking agents, by inducing recombination via a nucleotide excision repair (NER) pathway [164, 165]. The ability of the triplex-directed DNA substrates to induce recombination was only partially dependent on functional NER, suggesting that other endogenous DNA repair pathways are also involved. However, it is now well established that triplex DNA-mediated recombination is not affected in mismatch repair-deficient cells. The problem of sequence constraint for triple helix formation has been overcome, in part, by the use of novel approaches such as bifunctional oligonucleotides [166–168]. These oligonucleotides contain regions that form triple-helical structures in addition to conventional Watson–Crick base pairs. These modified oligonucleotides have been used successfully to promote site-specific nucleotide correction and gene targeting in both cell-free systems and cultured cells [169]. Moreover, these triplex-forming oligonucleotides can be used to inhibit gene expression [170] and in such a capacity have been used successfully to modulate expression of genes involved in liver fibrosis [171, 172]. These and other exciting new advances using triple helix-forming oligonucleotides suggest that this form of genomic modification may have therapeutic potential in treating genetic disorders.

Ribozymes, Antisense, and DNA Ribonucleases, RNAi Ribozymes are RNA enzymes that bind to specific RNA substrate sequences and catalyze endoribonucleolytic cleavage [173–175]. RNA cleavage is a naturally occurring intramolecular reaction primarily involved in the processing of certain introns to form the mature RNA. However, endoribonucleolytic cleavage can be engineered to occur as a trans-acting event. Ribozymes hybridize to complementary RNA sequences in which the central portion forms a specific secondary structure where closely

located reactive groups mediate the directed cleavage of the target RNA. The domains or helices of ribozymes that base pair to substrate RNAs are functionally separable from the moieties that effect cleavage. As a result, the substrate specificity of ribozymes can be altered within certain constraints to allow catalytic, trans-cleavage of specified sites within the target RNA. Thus, they have been used to target viral RNAs in infectious diseases, dominant oncogenes in cancers, and specific somatic mutations in a variety of genetic disorders. Hairpin ribozymes require only a guanosine (G) residue immediately 3′ to the cleavage site, although a GUC sequence is optimal. The hammerhead ribozyme is even less constrained, requiring only a UN dinucleotide for cleavage where N is either A, C, or U. The resulting RNA fragments are rapidly degraded, rendering the molecule non-functional. Ribozymes can be expressed either in cells or synthesized and packaged for cellular uptake. Moreover, they have been shown to remain catalytically active for weeks during expression in an intact organ after somatic gene transfer. In one study, recombinant adenoviral vectors containing human growth hormone ribozyme expression cassettes were used to ablate growth hormone in both cultured cells and livers of mice [176]. Ribozymes have been designed to cleave the RNAs of human hepatitis viruses. Both hammerhead [177, 178] and hairpin [179] ribozymes have been used successfully to inhibit viral production of both hepatitis B and hepatitis C infection in cells. The expressed hammerhead ribozymes, individually or in combination, were efficient at reducing or eliminating the respective plus or minus strand hepatitis C virus RNAs expressed in cultured cells and primary human hepatocytes from chronic hepatitis C-infected patients [177]. In contrast, cleavage-deficient ribozymes with a point mutation in the hammerhead domain had no significant effect. It is also possible to target a variety of highly conserved hepatitis C viral RNA sequences simultaneously with multiple ribozyme genes expressed from a single vector [180]. This type of gene therapy could, in fact, result in a constant and continuous supply of multiple intracellular ribozymes, thereby decreasing the potential development of drug-resistant viral variants. HBV, a partially double-stranded DNA virus, replicates through a pregenomic RNA intermediate, providing a therapeutic target for gene therapy based on ribozyme RNA cleavage. Hairpin ribozymes have been successfully designed and formulated to disrupt HBV replication by targeting the pregenomic RNA intermediate [181]. The 5′ -non-translated region of hepatitis C virus contains important elements that control hepatitis C virus translation, and therefore is an attractive target for inhibition. Antisense oligonucleotides bound to asialoglycoprotein–polylysine complexes targeted to the ASGPR on HuH-7 cells specifically inhibited the C virus-directed protein synthesis in the cells [182]. A 21-mer phosphorothioate-linked oligonucleotide DNA complementary to the HBV polyadenylation signal and

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5′ -upstream sequences was also targeted via the ASGPR to HBV-infected cells, resulting in specific inhibition of viral protein synthesis and replication in vitro [183]. DNA ribonucleases are catalytic molecules consisting of synthetic single-stranded DNA that specifically cleave substrate RNA in a manner analogous to ribozymes with catalytic efficiencies exceeding that of comparable ribozymes [184]. They are easier to prepare and deliver to cells and less sensitive to chemical and enzymatic degradation. Similarly to the structure of ribozymes, the DNA ribonucleases have three domains. A catalytic domain consisting of 15 nucleotides is flanked by two substrate-recognition domains which bind target RNA through Watson–Crick base pairing. These DNA analogs of ribozymes could be prepared to inhibit hepatitis C. DNA ribonucleases directed against the hepatitis C viral genome can specifically cleave the targeted RNA. DNA ribonuclease with point mutations in the catalytic domain had significantly lower inhibitory effects; however, activity was not eliminated, suggesting the presence of some antisense contribution. DNA ribonucleases can be made to cleave specifically target hepatitis B viral RNA and substantially inhibit intracellular viral gene expression [185]. The most important recent discovery for gene therapy applications is RNA interference (RNAi) [186] mediated by a novel class of RNAs that modulates gene expression rather than encoding proteins. This class of small non-coding (nc) RNAs ∼22 nt in length were originally identified in plants, nematodes, and then humans from nc, imperfect complementary, stem-loop RNA precursors [187]. MicroRNA (miRNA) genes are transcribed primarily from intronic regions by RNA polymerase II and are capped and polyadenylated similarly to normal mRNAs [188]. These “primary miRNA” (pri-miRNA) transcripts exist as stem-loop configurations [189], that are processed by the nuclear Drosha RNases, digesting the pri-miRNA into the small ∼70–90 nt hairpin “precursor miRNA” (pre-miRNA). These pre-miRNA are then transported from the nucleus to the cytoplasm via Exportin-5. In the cytosol, the pre-miRNA undergo several cleavages via Dicer-1, and are shortened to ∼22 nt fragments. Mature miRNAs are paired with RISC (RNA-induced silencing complex), which aids in binding miRNA to the target mRNA, thereby acting as an active repressor of expression. siRNAs are processed much like miRNAs, in that they both require Dicer RNases for cleavage and shortening. In addition, siRNAs also require RISC to become active gene regulators. They utilize RISC to establish precise base pair targeting to mRNAs. The fate of target mRNA is primarily dependent on the extent of base pairing to the ncRNA. Perfect complementarity to the target will typically result in transcript turnover via cleavage mediated by siRNAs. The presence of multiple, partially complementary target sites filled with miRNAs will inhibit translation without significantly affecting mRNA levels [190]. The mechanism of miRNA-triggered mRNA

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degradation is not entirely known. However, it appears to be mediated in part by “Slicer” activity that is expressed by the human Argonaut (Ago2) protein. The cleavage mediated by Slicer activity is believed to mark the target mRNA for further degradation via deadenylation, decapping, and 5′ –3′ exonuclease degradation in the cytosol. In contrast, mRNAs that are translationally repressed by miRNAs are translocated to specialized processing (P) bodies located in the cell cytoplasm [191]. It should be noted, however, that miRNA-mediated translational repression occurs independently of the movement of the miRISC:mRNA to P-bodies [11, 82]. The mRNA has two fates within the P-body, which in part acts as a reservoir. It can either enter the mRNA decay pathway or be returned to the cytosol to re-engage the translational machinery [192–194]. An improved understanding of RNAi has provided powerful tools for gene therapy for many acquired liver diseases, such as hepatocellular carcinoma, hepatitis B and C, and liver fibrosis [195–199]. Their delivery via hydrodynamic push to the liver [200] has demonstrated their utility for modulation of hepatic gene expression in vivo. The rapid movement of the field suggests that RNAi based gene therapy will become a mainstay in the future for treatment of hepatic disorders resulting from deleterious protein expression.

Single nucleotide modification A novel approach to gene therapy has recently been developed which utilizes the endogenous repair pathways of the cell to correct single base pair mutations [6, 7]. The gene repair technology, called “chimeraplasty,” is based on the ability of specifically designed chimeric RNA–DNA oligonucleotides to correct point mutations in genomic DNA. The targeted correction is site-specific and permanent, thereby maintaining endogenous gene regulation. This gene alteration approach was based on studies elucidating the molecular aspects of DNA repair. A significant increase in efficiency of pairing between an oligonucleotide of ∼50 bases and a genomic DNA target occurred only if RNA replaced DNA in a portion of the targeting oligonucleotide [201, 202]. Other modifications of the hybrid oligonucleotide, or chimeraplast, were made to increase stability and improve localization to genomic target sites in mammalian cells (Figure 59.4) [6]. In the original design, two single-stranded ends, comprised of unpaired nucleotide hairpin caps, flank the double-stranded region of the chimeric molecule. The 5′ and 3′ ends are juxtaposed and sequestered; and together with 2′ -O-methylated modification of the RNA residues contributes to enhanced nuclease resistance of the chimeraplast. The length of the oligonucleotide dictates the extent of homology between the chimeraplast and its genomic target; a 68-mer is designed to include a 25 bp region of homology containing a single mismatch with the gene sequence. This engineered

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Figure 59.4 Key features of the chimeric RNA–DNA molecule. Chimeroplasts are typically 68 nt oligonucleotides, including 20 2′ -O-methyl-modified ribonucleotides in the 25 nt segment that is homologous to the target gene. The DNA and RNA residues in the homology region are shown in dark and light gray, respectively. They contain two hairpin caps of four T-residues each (black) and a 5 bp GC clamp. The 25 nt region of homology is mismatched with the genomic sequences at a single nucleotide (marked X) that is targeted for change. The numbers of nucleotides are indicated in parentheses

mismatch appears to initiate modification of the genomic target, with the chimeraplast acting as a template for the alteration of the DNA sequence. The sequestered 5′ and 3′ ends minimize end-to-end ligation while the RNA segments, the region of homology, and the “nick” are all essential for chimeraplast activity. The physical and enzymatic stability conferred by its secondary structure and the modified RNA help ensure the survival of the chimeraplast en route to and within the cell. It is postulated that the mismatch created between the chimeraplast and its target genomic DNA creates the “illusion” of a base mutation, thus activating certain endogenous DNA repair functions [7]. The process harnesses the efficient, endogenous DNA repair activity used to repair mutations caused by natural and artificial mutagens for the targeted modification of genes.

Probing the mechanism The machinery for homologous recombination and DNA repair is highly conserved throughout evolution [142]. Therefore, to investigate the process of chimeraplasty and critical aspects of their structural design, test systems in both bacteria and mammalian cell-free extracts utilizing plasmid-based selectable systems or colorimetric identification to identify chimeraplast-mediated gene conversion were developed. The plasmid-selectable systems used neomycin phosphotransferase (neo), gene conferring kanamycin (kanR), or tetracycline resistance (tetR) [203, 204]. The colorimetric assay exploited the β-galactosidase enzymatic activity of the lacZ gene to cleave the synthetic substrate, X-gal, and produce the characteristic blue color [156]. The three genes were first modified by in vitro site-directed mutagenesis to introduce single nucleotide changes resulting in inactive gene products to serve as substrates for gene repair. If site-specific nucleotide correction or insertion occurred, then the plasmids conferred appropriate antibiotic resistance or blue-colored cells or colonies following cleavage of the X-gal substrate. Using these systems, a variety of structural changes in chimeraplast design were characterized for improved

targeting and nucleotide conversion [205]. For example, chimeraplasts exhibit a significant correlation of increased homology length with increased gene repair activity, similarly to classical homologous targeting experiments [206]. As the chimeraplast is designed to be complementary to the Watson–Crick strands of the target DNA, it is expected that two pairing events could occur; an RNA–DNA hybrid and the other a DNA–DNA duplex, each containing a mismatched base pair. To establish the potential contribution to the repair process of the “hybrid” or the DNA–DNA mismatched base pair, chimeraplasts that contained only one mismatched strand with the neo gene were designed [205]. The oligonucleotide with the mismatch on the “all-DNA” strand directed the repair at a higher efficiency than that with the mismatch on the “RNA–DNA” strand, or even the original “double-mismatched” chimeraplast. The results suggest that mismatches created by the DNA strand are more efficiently repaired with the primary role of the RNA–DNA hybrid strand being to increase structural stability and enhance pairing with the target DNA. The various endogenous DNA repair pathways involved were investigated using E. coli strains deficient in specific repair proteins [205]. Gene modification by chimeraplasty was either significantly reduced or undetectable in strains containing defects in RecA, a DNA pairing protein, or in the mismatch repair binding protein MutS. However, other DNA and RNA modification enzymes such as adenine (dam) and cytosine (dcm) methylases and also dUTPase (dut) and uracil N -glycosylase (ung) involved in base excision repair were, in fact, dispensable. The requirement for both RecA and MutS proteins implies at least a two-step process, involving factors from homologous recombination and also mismatch repair pathways (Figure 59.5). It is speculated that RecA-dependent pairing of the chimeraplast with its target DNA is followed by the recognition of the mismatch and its repair, involving MutS and other proteins. The RNA portion of the oligonucleotide appears to provide an essential role in strand pairing, as control oligonucleotides comprised entirely of DNA with the identical sequence and structure exhibited no detectable activity.

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(a)

(b)

Figure 59.5 The process of gene repair by RNA–DNA chimeroplasts. (a) Chimeroplasts, designed as described in Figure 59.4, align with its target homologous genomic DNA by Watson–Crick base pairing, except for the engineered single base-pair mismatch. (b) Endogenous DNA repair pathways activated by the mispairing modify the genomic sequence complementary to that of the chimeroplast, correcting the single nucleotide mutation or deletion. Location of the gene in the chromosome is not altered and the gene expression remains under the control of the native promoter and enhancers, permitting normal physiological regulation

Mammalian studies Both recombination and mismatch repair pathways are evolutionarily conserved, suggesting that chimeraplasty would function in mammalian systems [207–209]. In fact, it was shown that HuH-7 cell-free extracts could support in vitro dose-dependent conversion of the mutant neo gene and also insertion of a deleted base pair into the tetracycline gene. Further, when a cell line lacking the mismatch repair protein hMSH2 was used to prepare the extracts, or an antibody to hMSH2 was included in the reaction, little chimeraplast repair activity was detected. Interestingly, using this same in vitro approach, rat liver mitochondrial protein extracts also supported chimeraplast conversion of a plasmid encoding a neo resistance gene [210]. Using the lacZ plasmid system, cell-free nuclear extracts from a variety of different cell lines efficiently catalyzed dose-dependent nucleotide conversion [204]. Also, the nuclear extract from a homozygous isogenic p53+ /p5+

embryonic fibroblast cell line induced 300-fold less conversion than p53 null extract. Hence wild-type p53 may inhibit the initial pairing step in chimeraplast repair, since p53 decreases the recombination activity of RecA and its human homolog Rad51 [211–213]. Overall, these data suggest a significantly conserved mechanism of nucleotide conversion that appears to be distinct from that of homologous recombination in its requirement for MutS or hMSH2 protein.

Cell culture studies It was originally reported that the chimeric RNA–DNA oligonucleotides were able to effect site-specific nucleotide conversion of episomal DNA in cultured cells. This was followed by the targeted single nucleotide conversion to correct the sickle cell genomic DNA point mutation in cultured lymphoblastoid cells [214]. Subsequently, it was shown that chimeric oligonucleotides could

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introduce a missense mutation in genomic DNA in cultured HuH-7 human hepatoma cells [215]. Interestingly, under conditions in which RNA–DNA hybrids were active in promoting nucleotide exchange, the corresponding all-DNA duplexes, despite nuclear uptake, were essentially inactive [214, 215]. These early studies, however, emphasized the need for improved delivery of chimeraplasts to hepatocytes. Moreover, therapeutic benefit requires not only targeted delivery of chimeraplasts to the liver but also efficient nuclear translocation and directed nucleotide exchange in quiescent G0 hepatocytes. Thus, the galactose sugar was used as the targeting ligand for delivery of these molecules to the liver, based on previous reports of high specificity and efficiency in targeting nucleic acids to the hepatocyte ASGPR. Additionally, anionic liposomes containing galactocerebroside for targeting to the ASGPR were formulated to encapsulate the oligonucleotides for delivery [216]. The specificity and efficiency of hepatocyte uptake and nuclear localization of fluorescein-labeled chimeraplasts was demonstrated by confocal microscopy. Chimeric oligonucleotides were designed that targeted both the transcribed and non-transcribed rat factor IX genomic DNA strands [217]. The molecules were identical in sequence with the wild-type gene except for the engineered nucleotide mismatch. The targeted nucleotide change at Ser365 would introduce a missense mutation in the rat genomic sequence resulting in the active site Ser365 to Arg365, characteristic of certain human factor IX mutations. Primary rat hepatocytes were transfected with chimeraplasts using both the PEI and liposomal delivery systems. The conversion efficiency was determined by polymerase chain reaction (PCR) amplification of a 374 nt fragment spanning the targeted nucleotide change, and also by sequence analysis. A to C conversion at Ser365 was detected in 25% of the gene pool for factor IX, irrespective of the targeted strand. Similar results were obtained with correction of the missense mutation responsible for the factor IX deficiency in isolated hepatocytes from the Chapel Hill strain of hemophilia B dogs [218]. The UGT1A1 -deficient Gunn rat animal model of Crigler–Najjar syndrome type I was tested to determine whether chimeraplasts could also correct a frameshift mutation. Insertion of a single guanosine at position 1206 in the UGT 1 coding sequence would restore the proper reading frame of the UGT1A1 mRNA, production of the UDP-glucurosyltransferase enzyme, and re-establish the wild-type BstNI restriction site [219]. The design of the correcting chimeraplast (CN3) differed from those in previous studies in that the DNA region flanked by the modified RNA bases was increased to nine nucleotides with a central mismatched base pair. Isolated Gunn rat hepatocytes were transfected with the CN3 chimeraplasts using the targeted anionic liposome delivery system. Nucleotide insertion was determined by differential hybridization analysis of the wild-type corrected sequence (1206G ) and the 1206A mutant sequence and resulted

in a dose-dependent frameshift correction frequency of about 25%.

Site-directed nucleotide conversion in vivo Success in the primary rat hepatocytes suggested utility for in vivo gene modification. Thus, a series of experiments were performed in male rats to determine if the chimeric oligonucleotides could mutate the factor IX in hepatocytes [217]. The molecules complexed to lactosylated PEI were delivered by tail vein injection and after several days liver tissue was collected and genomic DNA was isolated for analysis. The PCR-amplified products were analyzed by duplicate filter lift hybridization and indicated a dose-related genomic DNA conversion frequency of 15–40%. Similar results were obtained by RT-PCR analysis for RNA, suggesting a potential biological effect of the genomic DNA conversion. Sequence analysis of the cloned PCR products confirmed the specificity of the A to C conversion at Ser365 from both the cell culture and in vivo studies. The factor IX coagulant activity was determined by an activated partial thromboplastin time (aPTT) assay using human factor IX-deficient plasma. The results indicated a significant reduction (∼40%) in factor IX activity, which together with the percentage genomic conversion remained unchanged through 72 weeks. Moreover, direct sequence analysis of the amplicons from random samples at 18 months confirmed the polymorphism at Ser365. Hence the site-directed conversion of the factor IX gene in intact liver appears to be permanent and phenotypically stable. In addition, the replicative stability of the targeted nucleotide conversion was determined by performing a 70% partial hepatectomy after nucleotide modification. The surgical procedure induces the liver remnant to undergo compensatory regeneration, a process in which 95% of the hepatocytes replicate in two synchronous waves [220]. Factor IX activity in these animals was determined periodically by aPTT assays between 9 and 78 weeks post-injection/hepatectomy. The factor IX coagulant activity in these test animals decreased to ∼40%, while the conversion frequencies remained unchanged, indicating that the genomic mutation is stable during replication. Experiments were then performed to establish that chimeric RNA–DNA oligonucleotides could efficiently replace a deleted nucleotide. The CN3 chimeraplast designed to insert a single base pair in the mutated UGT1A1 gene in Gunn rat hepatocytes was administered intravenously using both delivery systems [221]. DNA was isolated from liver tissue after 1 week and revealed that the targeted G was inserted at a frequency of ∼20% in the animals. In contrast, no G insertion was observed in the vehicle control animals. This genomic alteration was unchanged after 6 months, and was confirmed by restriction fragment length polymorphism (RFLP) analysis and direct sequencing of the PCR-amplified region of the UG1A1

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gene [219]. Southern blot analysis of genomic DNA from these animals indicated that the BstNI restriction site was also partially restored in the CN3-treated animals, whereas the DNA isolated from the vehicle-treated controls remained resistant to BstNI digestion. Western blot analysis confirmed that the genotypic correction re-established expression of the 52 kDa UDPglucuronosyltransferase type I microsomal enzyme in the liver. In addition, serum bilirubin levels dropped below 50% of their pretreatment levels, and remained at these levels more than 1 year after treatment. Repeated administration of the correcting chimeraplasts resulted in further reductions in serum bilirubin levels whereas the levels in control animals treated with a non-relevant chimeraplast increased over the same time period. Finally, high pressure liquid chromatography(HPLC) analysis of bile from the CN3-treated animals detected both mono- and diglucuronide-conjugated bilirubin, whereas the bilirubin in control bile remained unconjugated. Taken together, these data suggest that the mismatch repair pathways in hepatocytes may be sufficiently active to make this strategy feasible for other liver-related disorders resulting from single base-pair mutations or deletions such as hemophilia B and α1 -antitrypsin deficiency. The chimeric RNA–DNA oligonucleotides are capable of promoting site-specific base conversion or insertion in intact liver resulting in sustained phenotypic changes associated with modification of the genomic DNA. However, structural aspects of the targeted nucleotide in the DNA loci may influence the conversion efficiency since repair of mismatched DNA has been shown to be bidirectional and strand specific and also depends on nucleosomal position [222–224]. There have been a number of additional successful in vivo applications of chimeric oligonucleotides for the modification of genomic sequences in mouse primary kidney tubular cells [225], the tyrosinase gene mutation responsible for melanin production in albino mouse melanocytes [226], and for correction of the mdx point mutations responsible for muscular dystrophy in both mice and dogs [227]. Collectively, these studies suggest that numerous cell types are capable of performing chimeraplast-mediated modifications of their genomic sequences.

Single-stranded oligonucleotides Interestingly, further investigation of the molecular pathways involved in mediating the targeted gene conversion directed by the chimeraplast indicated that single-stranded oligonucleotides (SSOs), if designed with a single base mismatch, would also function effectively for promoting targeted single base exchange [228]. In contrast to chimeraplasty, it is independent of the mismatch proteins MSH2 and MSH3 [228, 229]. In fact, it appears to proceed by a mechanism of strand incorporation [230],

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and does not involve the ATM/ATR homologous recombination pathway used for double-stranded break repair [231]. It exhibits some strand bias in favor of targeting the non-transcribed strand [232], but the overall rate of targeted nucleotide replacement is enhanced more than 10-fold by transcriptional activation of the targeted loci [233]. Although SSOs are significantly less costly and easier to synthesize, they are much more susceptible to intracellular degradation. Significant effort has been spent investigating the most effective designs for increasing both SSO persistence and efficiency in promoting the desired nucleotide change at the target site [234–236]. In particular, modifying the 3′ end of the SSO with at least three phosphorothioates significantly improves the rate of conversion, yet it is essential that the 3′ end nucleotide presents a 3′ -hydroxyl group, consistent with a mechanism of strand incorporation of the SSO. At the 5′ end, significantly greater conversion is observed with phosphodiester nucleotides than when phosphorothioates are used, and 5′ modification with the intercalating agent Acridine Orange can increase the targeted nucleotide exchange frequency 10-fold [237]. Coupling microarray gene expression analysis with oligonucleotide targeting studies [236], SSOs do not induce the DNA damage signaling and DNA repair genes, a potential advantage over double-stranded DNA-mediated repair. The efficiency/persistence of repair also appears to be significantly influenced by cell cycle [238]. The highest conversion was observed in S-phase in cultured cells and further improved if the rate of DNA replication was slowed [239]. Manipulation of various proteins involved in recombination and other DNA repair pathways has also been shown to modulate the gene repair activity [240–243]. In actively cycling hepatocytes, the abundant expression of mismatch repair genes inhibits the SSO-mediated conversion, with inhibition of MSH2 by RNAi resulting in a 25–30-fold increase in gene repair [244]. A novel approach to increasing the rate of repair looked at co-transfection with increasing amounts of non-specific SSOs to titrate/sequester cellular activities interfering with the targeted nucleotide conversion [245]. A 6-fold increase in gene repair frequency was observed that was independent of non-specific SSO length, but dose dependent on the molar amount of non-specific SSO co-transfected. However, without modulation of any of these parameters, 4.5% repair efficiencies can be routinely achieved in replicating cells in culture by using modified SSOs with canonical 5′ -phosphate (5′ P) and 3′ -hydroxyl (3′ OH) termini [246]. Successful in vivo application of SSOs has been reported in mice [247]. More recently, SSO-mediated gene correction using polylysine-conjugates targeted to the hepatocyte ASGPR was successful in correcting the acid α-d-glucosidase gene as assessed by both PCR and phenotypic change [248]. Using a 75-mer in transgenic mice expressing the mutant murine transthyretin (TTR) Val 30Met gene,

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targeted nucleotide replacement leading to gene repair was achieved in 9% of the adult mouse hepatocytes with phenotypic hepatic changes [249]. Moreover, we have used 45-mer SSOs modified at their 3′ end with three phosphorothioate residues and phosphorylated at their 5′ end delivered to transgenic spf ash ornithine transcarbamylase (OTC)-deficient pups, resulting in restoration of enzymatic activity to 15% of wild-type, reflected in the ∼10–15% conversion of the targeted nucleotide from the mutant A to wild-type G [250]. Thus, we enter the second decade of the 21st century with the potential for these exciting new non-viral technologies, Sleeping Beauty transposons for gene augmentation, RNAi for gene knock-down, and ZFNs and SSOs for gene repair. Coupled with the new non-viral in vivo delivery systems, the prospect of obtaining success in gene therapy for treating liver-based genetic disorders, either inherited or acquired, appears to be within reach.

ACKNOWLEDGMENT The authors acknowledge the important contributions by many investigators in the field of liver-directed gene therapy that could not be cited here because of limited space.

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215. Kren, B.T., Cole-Strauss, A., Kmiec, E.B. et al. (1997) Targeted nucleotide exchange in the alkaline phosphatase gene of HuH-7 cells mediated by a chimeric RNA/DNA oligonucleotide. Hepatology, 25, 1462–68. 216. Templeton, N.S., Lasic, D.D., Frederik, P.M. et al. (1997) Improved DNA:liposome complexes for increased systemic delivery and gene expression. Nat Biotechnol , 15, 647–52. 217. Kren, B.T., Bandyopadhyay, P. and Steer, C.J. (1998) In vivo site-directed mutagenesis of the factor IX gene by chimeric RNA/DNA oligonucleotides. Nat Med , 4, 285–90. 218. Evans, J.P., Brinkhous, K.M., Brayer, G.D. et al. (1989) Canine hemophilia B resulting from a point mutation with unusual consequences. Proc Natl Acad Sci U S A, 86, 10095–99. 219. Roy Chowdhury, J., Huang, T.J., Kesari, K. et al. (1991) Molecular basis for the lack of bilirubin-specific and 3-methylcholanthrene-inducible UDP-glucuronosyltransferase activities in Gunn rats. The two isoforms are encoded by distinct mRNA species that share an identical single base deletion. J Biol Chem, 266, 18294–98. 220. Higgins, G.M. and Anderson, R.M. (1931) Experimental pathology of the liver. I. Restoration of the liver of the white rat following partial surgical removal. Arch Pathol , 12, 186–202. 221. Kren, B.T., Parashar, B., Bandyopadhyay, P. et al. (1999) Correction of the UDP-glucuronosyl-transferase gene defect in the Gunn rat model of Crigler–Najjar syndrome type I with a chimeric oligonucleotide. Proc Natl Acad Sci U S A, 96, 10349–54. 222. Fang, W.H. and Modrich, P. (1993) Human strand-specific mismatch repair occurs by a bidirectional mechanism similar to that of the bacterial reaction. J Biol Chem, 268, 11838–44. 223. Klungland, A. and Lindahl, T. (1997) Second pathway for completion of human DNA base excision-repair: reconstitution with purified proteins and requirement for DNase IV (FEN1). EMBO J , 16, 3341–48. 224. Wellinger, R.E. and Thoma, F. (1997) Nucleosome structure and positioning modulate nucleotide excision repair in the non-transcribed strand of an active gene. EMBO J , 16, 5046–56. 225. Lai, L.-W., Chau, B. and Lien, Y.-H. (1999) In vivo gene targeting in carbonic anhydrase II deficient mice by chimeric RNA/DNA oligonucleotides. Conference Proceedings: 2nd Annual Meeting of the American Society of Gene Therapy, Washington, DC, p. 236a. 226. Alexeev, V., Igoucheva, O., Domashenko, A. et al. (2000) Localized in vivo genotypic and phenotypic correction of the albino mutation in skin by RNA–DNA oligonucleotide. Nat Biotechnol , 18, 43–47. 227. Bartlett, R.J., Denis, M.M., Kornegay, J.N. et al. (1998) Can genetic surgery be used to revert muscular dystrophy mutations in live animals? Conference Proceedings: 1st Annual Meeting of the American Society of Gene Therapy, Seattle, WA, p. 153a.

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Decoding the Liver Cancer Genome Ju-Seog Lee1 and Snorri S. Thorgeirsson2 1 Department

of Systems Biology, The University of Texas M. D. Anderson Cancer Center, Houston, TX, USA 2 Laboratory of Experimental Carcinogenesis, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA

INTRODUCTION Hepatocellular carcinoma (HCC) is one of the most common cancers in the world, accounting for an estimated 600,000 deaths annually [1]. While HCC is common in Southeast Asia and sub-Saharan Africa, the incidence of HCC has continued to increase in the United States and Western Europe over the past 25 years and the incidence and mortality rates of HCC are expected to double over the next 10–20 years [2–4]. Although much is known about both the cellular changes that lead to HCC and the etiological agents [i.e. hepatitis B virus (HBV) and hepatitis C virus (HCV) infection and alcohol] responsible for the majority of HCC, the molecular pathogenesis of HCC is not well understood [5–7]. Moreover, the severity of HCC, the lack of good diagnostic markers and treatment strategies, and the clinical heterogeneity have rendered the disease a major challenge [7, 8]. Patients with HCC have a highly variable clinical course [6, 9], indicating that HCC comprises several biologically distinctive subgroups. Despite considerable efforts to establish rational as an alternative to empirical approaches to create a classification system [6, 9–14], clinical and pathological diagnosis and classification of HCC remain unreliable in predicting patients’ survival and responses to therapy. The prognostic The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

variability likely reflects a molecular heterogeneity that has not been appreciated from methods traditionally used to characterize HCC. Improving the classification of HCC patients into groups with homogeneous prognosis in addition to a more comprehensive understanding of the underlying biology of HCC development at the molecular level would improve the application of currently available treatment modalities and offer the possibility of new treatment strategies. One of the most exciting developments in recent years has been the clinical validation of targeted drugs that inhibit the action of pathogenic gene products such as protein kinases and proteinases [15]. Treatment with these targeted drugs has proven more efficient in altering the natural history of the disease and reducing mortality. Identification of cancer-type specific oncogenes that play key roles during progression of cancer can lead to advances in classification of cancer and targeted drug therapy. However, molecular characterization of HCC aimed at identifying driver oncogenes (potential therapeutic targets) has lagged in comparison with other cancers. Therefore, in order to improve treatment options and reduce mortality, it is crucial to develop treatment strategies that can be applied in the near future while improving our understanding of hepatocarcinogenesis.

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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THE LIVER: MICROARRAY-BASED TECHNOLOGIES

COMPARATIVE GENOMIC HYBRIDIZATION: GENOME-WIDE SCREENING OF THE LIVER CANCER GENOME Since the discovery of genetic rearrangements in cancer [16], cytogenetic approaches have been extensively used to uncover the chromosomal basis for these genetic alterations. The comparative genomic hybridization (CGH) technique was developed in the early 1990s and was the first genomic tool to provide a genome-wide characterization of copy number changes in cancer [17]. With improvements in microscope and labeling technologies, CGH has become a frequently used tool to examine DNA copy number changes in cancer and to identify altered expression and function of genes residing within the affected region of the genome. Such genomic loci with decreased and increased copy numbers are believed to harbor tumor suppressor genes and oncogenes, respectively. Despite limited spatial resolution of CGH mapping, approximately 10 Mbp for low copy-number gains and losses and close to 2 Mbp for high-copy-number amplifications, this technology uncovered many candidate loci for tumor suppressor genes and oncogenes in HCC. Identification of genomic loci with copy number aberrations combined with the capacity to identify the genes residing in these loci led to a better understanding of cancer development. For example, increased copy number of the 8q24 region has been reported in many studies and the most potent oncogene residing in 8q24 is MYC [18–21]. The most frequently reported decreased copy number region is 13q14 where tumor suppressor RB1 resides [22]. In addition to 8q and 13q, CGH data revealed that gains of chromosomal material were most prevalent in 1q, 6p, and 17q, and losses were most frequently present in 8p, 16q, 4q, and 17p [18, 21]. The sensitivity to detect copy-number variations has improved with the arrival of microarray-based technology, array comparative genomic hybridization (aCGH), where arrays of genomic sequences such as BAC clones and oligonucleotides replaced metaphase chromosomes as hybridization targets. Coupled with improved annotation of genome sequence data, these technologies are facilitating the identification of new genomic loci that are associated with the progression of cancer.

across the entire genome. Although the completion of the human genome sequence was a crucial prerequisite for cataloging our genetic makeup, comprehension of the sequence data alone is not sufficient to decipher complex physiological processes during tumor development. The concomitant advances in miniaturization technology, hybridization biochemistry, and biomolecule detection have made it possible to capture a multitude of biological events during tumor development. Since the first use of cDNA microarrays in gene expression analysis [23], the technology has rapidly become a major genomic tool for cancer researchers. Microarrays have evolved from representing less than 50 genes to containing over 50 000 transcripts on whole genome arrays for complex human genomes. In some cases, over 106 individual features for every exon in human genes are represented [24]. Microarray assays allow massive parallel data acquisition and analysis. Although parallelism greatly increases the speed of data collection, massive amounts of data present daunting challenges in both processing and interpretation. Microarray-based gene expression profiling studies in a variety of cancers have discovered consistent gene expression patterns associated with pathological or clinical phenotypes, and identified subtypes of cancer previously unidentified by conventional technologies [25–27]. This new technology has been used successfully to predict clinical outcomes and survival rates and to identify potential therapeutic targets and prognostic marker genes [28–30]. Although initial applications of microarray technology have been limited to gene expression studies, this technology has also been adopted by many investigators in other fields (Figure 60.1). aCGH, in which arrays of genomic sequences are used as hybridization targets, was quickly established as a substitute for conventional CGH [31, 32]. The greatest advantage of aCGH is the ability to perform copy number analyses with much higher resolution than was ever possible using conventional

HCC

RNA

DNA

PROTEIN

MICROARRAY-BASED TECHNOLOGIES The genetic and/or epigenetic basis of complex diseases such as cancer was difficult to understand prior to completion of the human genome project and the arrival of new microarray-based technologies. These technologies have enabled investigators to describe genetic variations

Expression microarray

Array CGH

RPPA

Figure 60.1 Application of microarray-based technology

60: DECODING THE LIVER CANCER GENOME

CGH in which metaphase chromosomes were used as hybridization targets. Protein microarrays have also been developed by adopting the knowledge and technical innovations derived from DNA microarrays. The technical aspects of miniaturizing traditional methods, such as Western blotting and protein dotting on to nitrocellulose or nylon membranes, have been quickly introduced into protein microarray technology. Two approaches for producing protein microarrays exist: forward phase protein arrays (FPPAs) and reverse phase protein arrays (RPPAs). In a forward phase array, antibodies are immobilized on the surface of slides and each array is incubated with one test sample such as a tissue lysate or serum sample, and multiple protein features such as expression and phosphorylation from that sample are measured simultaneously. In contrast, the RPPA format immobilizes an individual tissue lysates in each array spot, and thus an array is comprised of hundreds of different patient samples. Each array is then incubated with one antibody, and a protein feature is measured and directly compared across multiple samples. FPPAs (antibody arrays) are particularly ill suited for tissue-based analysis since they require substantial amounts of tissue lysates for incubation, hence RPPAs (tissue lysates arrays) are a better choice of platform in cancer research [33–36].

GENE EXPRESSION PROFILING: RESHAPING THE PATHOBIOLOGY OF HCC Conventional approaches for the prognostic classification of HCC largely rely on single or multiple clinicopathological variables such as the severity of liver function and characteristics of the tumor (i.e. size, number of nodules, vascular invasion, distant metastasis, and tumor grade), and the response to therapeutic interventions. However, the utility of existing prognostic factors is limited because they largely measure tumor differentiation and bulk but do not otherwise characterize and/or measure the underlying properties that likely drive the biology of the tumor. In a previous study [27], an unbiased analytical approach applied to gene expression data from human HCC identified distinct subtypes of HCC patients with significant association with patients’ survival (Plate 60.1). These data suggest that gene expression profiling signatures well reflect biological and clinical differences between subtypes of HCC and would be highly valuable in predicting the prognosis of patients. The current clinical challenge is to identify those patients who do not derive much benefit from current therapies and to offer alternative treatments. If key (or master) regulators (genes, pathways, and/or networks) driving the biology of the tumor can be identified, they might lend themselves to therapeutic exploitation. However, in this context, it is not enough to rely entirely

993

on gene expression signatures that are indicative of prognosis, since the profiles may fall short of explaining at the molecular level what drives the prognostic difference between subtypes of tumors.

COMPARATIVE SYSTEMS GENOMICS: CROSS-SPECIES COMPARISON OF GENE EXPRESSION DATA Although differences exist, similarities in the process of cancer development between humans and mice are particularly striking [38, 39], leading many investigators to exploit the mouse as a model organism for the study of this complex disease. Recent studies provide clues on how to extend gene expression profiling studies beyond the current general practice of collecting massive data from human cancer. In an effort to identify the best-fit mouse HCC models that mimic the human condition, gene expression data from patients were integrated with those from mouse HCC [40]. Gene expression patterns of mouse HCC were obtained from seven HCC mouse models. Orthologous human and mouse genes from both data sets were selected before gene expression data were integrated. In hierarchical clustering analysis of integrated data, gene expression patterns of HCC developed in Myc, E2f1 , and Myc/E2f1 transgenic mouse models had the highest similarity with those of the better survival group of human HCC, whereas the expression patterns of HCC in the Myc/Tgfa transgenic mouse model were most similar to those of the poor survival group of human HCC (Plate 60.2a, b). These results suggest that these two classes of mouse models might more closely recapitulate the molecular patterns of the two subclasses of human HCC. The similarity of gene expression profiles between human and mouse models is in good agreement with the phenotypic characteristics of the tumors (Plate 60.2c,d). The human tumors with increased proliferation, decreased apoptosis, and worse prognosis are paired with the mouse models with the same characteristics. The gene expression-based prediction of mouse models is highly concordant with the phenotypes of mice. Myc/Tgfa mice have a typically poor prognosis phenotype, such as an earlier and higher incidence of HCC development, higher mortality, higher genomic instability, and higher expression of poor prognostic markers [37, 41]. Although the precise molecular mechanism driving hepatocarcinogenesis is yet to be discovered, the relative similarity of Myc/Tgfa mice to the human poor survival group HCC indicates a role of the EGFR, receptor for TGFA [42], or related signaling pathways in the prognosis of human HCC. These results strongly suggest that well-defined gene expression signatures from well-defined experimental conditions and/or animal models can be used to stratify human cancer patients into more homogeneous groups at the molecular

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level. In many cancers, specific drugs are effective in specific subtypes of cancer. Herceptin is only effective in the subpopulation of breast cancer patients who express C-erbB-2 oncoprotein (ERBB2) [43], and Gleevec is used for chronic myeloid leukemia patients harboring the BCR–ABL1 gene fusion [44]. We can anticipate that unique molecular identities of each subclass of HCC uncovered by comparative analysis of a genome-wide survey of gene expression from human and animal models will provide new therapeutic strategies to maximize the efficiency of treatments.

INTEGRATIVE SYSTEMS GENOMICS: INCREASING THE DIMENSIONALITY OF GENE EXPRESSION DATA Cancer cells do not invent new pathways. They evolved from normal cells by using pre-existing pathways in different ways or by combining components of these pathways in a way that effectively drives tumorigenesis. By mapping and refining pathway maps in developing or normally functioning liver, gene expression profiling studies might provide insight into the connectivity of these pathways in HCC. In a recent study [45], the gene expression signature unique to rat fetal liver progenitor cells was integrated with those from human HCC in an attempt to determine the fraction of human HCC that shares gene expression patterns with liver progenitor cells. This approach identified a novel subtype of HCC that may arise from hepatic progenitor cells. This new subtype accounts for around 20% of HCC patients examined in this study and shows extremely poor prognosis (Plate 60.3). Previous studies in diffuse large B cell lymphoma and T cell acute lymphoblastic leukemia indicated that the cellular origins of a tumor largely dictate the clinical outcome of patients [25, 46], since mitogenic, motogenic, and morphogenic responses and also the propensity for apoptosis may vary at different stages of normal differentiation. Genes involved in an invasive phenotype (MMP1 , PLAUR, TIMP1 , CD44 , and VIL2 ) were highly expressed in this newly identified subtype (with hepatic progenitor cell features) and may account for the extremely poor prognosis. This subtype showed marked activation of the AP-1 complex, which is essential for normal hepatogenesis during embryonic development and critical for initiation of HCC development in mice [47, 48]. Cancer cells arise from normal cells following accumulation of genetic alterations. One of the most important consequences of this process is the resurrection of pre-existing but dormant signaling pathways that were active during embryonic development [49]. Hence this finding supports the growing appreciation that signaling pathways that control vertebrate embryonic development are also important in human carcinogenesis.

IDENTIFICATION OF THERAPEUTIC TARGETS BY INTEGRATIVE SYSTEMS GENOMICS Previous studies clearly demonstrated that gene expression signatures can be used to classify the tumors and provide prognostic information [8, 27, 40, 45, 50–52]. The current research focus has shifted, however, more towards identifying genetic determinants that are components of specific regulatory pathways altered in cancers, potentially leading to the discovery of novel therapeutic targets [7, 53–55]. However, selection of relevant candidate genes for further studies from lengthy gene lists generated from gene expression profiling studies is a significant challenge due to the many confounding factors embedded in the gene expression profile data from human cancers. These factors include age, hospital care, treatments, non-parallel cancer progression, and unspecified environmental factors that are irrelevant to cancer development. Moreover, the gene expression profile from patients is only a “snapshot” of gene-to-gene interactions that lacks information on the interactive time-dependent changes that occur during tumorigenesis. Accordingly, it is difficult to discriminate the genes (drivers) that drive the tumorigenic process from genes (passengers) whose expression patterns simply reflect loss of organ function and/or the degree of differentiation of the cancer cells. CGH, and more recently array CGH analyses, have identified a number of recurrent regions of DNA copy number changes in many cancers. Frequent DNA copy number gains at 1q, 8q, and 20q and frequent DNA copy number losses at 1p, 4q, 8p, 13q, 16q, and 17p have been identified in HCC [7]. Some of these genomic loci contain well-characterized and/or putative oncogenes and tumor suppressor genes. Moreover, a number of genes in these regions have been linked to disease pathogenesis and clinical behavior. For example, the association of DNA copy number aberrations with prognosis has been found for a variety of tumor types, including prostate cancer, breast cancer, gastric cancer, multiple myeloma, lymphomas, and HCC [55–60]. However, some amplified or deleted regions are often large, and many of the genes residing in the recurrent regions are not expressed either in normal tissues or in tumors. Moreover, functional validation of genes residing in these loci is impractical when confronted with hundreds of candidate genomic loci. Therefore, there is inevitably a need for the development of a new strategy that can overcome the limitations of gene expression data and array CGH data. In a recent study in breast cancer [61], investigators tested the possibility of whether integrating gene expression and gene copy-number data from the same patient cohort would help identify potential driver genes. This

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study clearly demonstrated that gene copy number data provided extra prognostic relevance of genes as compared with when only gene expression data were available. Integrative analysis of gene expression and gene copy-number data also uncovered nine potential drivers which are activated by recurrent gene amplifications in breast cancer and may show an association with aggressive tumor types. Future application of this innovative approach to HCC could greatly increase the chances of identifying potential therapeutic targets. Alterations of expression patterns and genomic copy numbers of thousands of genes are fundamental properties of the cancer cells. Since the application of high-throughput microarray-based genomic technologies for the analysis of cancer inevitably generates many false-positive results, it is almost impossible to select a reasonable number of candidate genes for therapeutic targets and/or biomarkers for diagnosis and prognosis. Therefore, it is important to cross-compare and integrate two or more genomic scale datasets (i.e. coding and non-coding gene expression, and array CGH data) independently collected from the same patient cohort.

INTEGROMICS: BEYOND GENOMICS Although gene expression and copy number profiling can provide important information on somatic genetic events during tumor progression, they are unable to provide an effective recapitulation of fluctuating protein-based signaling events that are the direct executors of cellular function. RPPA is a newly developed high-throughput functional proteomic technology [33–36]. It extends the power of immunoblotting, by acting equivalently to a multiplex enzyme-linked immunosorbent assay (ELISA), to provide a quantitative analysis of the differential expression of signaling proteins. Moreover, the phosphorylation status of proteins can be detected and measured using specific anti-phosphoprotein antibodies. Through the use of these phospho-specific antibodies, it is possible to evaluate the state of the entire portions of a signaling pathway or cascade, by looking at dozens of kinase substrates at the same time through multiplexed phospho-specific antibody analysis. With RPPA, all samples are spotted at the same time and analyzed with a single antibody, making this method ideally suited for the analysis of a large number of specimens. However, their assessment to signaling pathways is limited by the number of available antibodies, which is far smaller than the number of gene probes in expression microarrays. Hence the current limitation of these data sets can be overcome by integrating data sets together during analysis (Plate 60.4). Integration of genomic and proteomic data will undoubtedly enhance our understanding

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of tumor progression by increasing the dimensionality of molecular features. Moreover, the identification of driver or contributor genes can be greatly accelerated by the integration of more than one dimension of genomic information systems.

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Genome-wide Expression Profiling of Human Hepatocellular Carcinoma Anuradha Budhu and Xin Wei Wang Liver Carcinogenesis Section, Laboratory of Human Carcinogenesis, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA

THE CURRENT STATUS OF HEPATOCELLULAR CARCINOMA DIAGNOSIS, TREATMENT, AND PROGNOSIS Well-defined and generally accepted staging systems are available for diagnosis, prognosis, and treatment stratification of almost all cancers. However, hepatocellular carcinoma (HCC) is an exception as many different staging systems have been introduced around the world and currently there is no clear consensus on which one is best [1, 2]. Prognostic assessment and choice of treatment options in HCC are complex because they dually depend on the grade of cancer spread (tumor staging) and the grade of residual liver function (liver disease stage). The lack of a consensus on the definition and staging of HCC combined with the wide heterogeneity of the disease interferes with clinical recommendations and progress. Despite many studies of HCC, information regarding phenotypic and molecular changes associated with the development of this disease is still limited [3, 4]. This emphasizes the importance of understanding the molecular mechanisms The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

underlying HCC and the development of new screening and prognosis/treatment stratification programs to refine diagnosis and improve patient outcome.

HCC diagnosis Although routine screening by ultrasonography and serum α-fetoprotein (AFP) levels of high-risk individuals has aided HCC diagnosis, most patients are diagnosed at late stages. The detection of HCC, particularly in early disease stages, still remains a challenge. Although AFP is the only widely used serum marker for HCC, elevated serum level is observed only in a subgroup of patients with small HCC (33–65%), non-specific elevation occurs in 15–58% of chronic hepatitis patients, and levels can vary significantly between ethnic groups [5]. A number of additional serum proteins have been suggested to improve HCC diagnosis, including complement C3a, des-γ-carboxyprothrombin, α-l-fucosidase, glypican-3, TGFB1, IGF-II, IGFBP-2, HCCR, Golgi protein 73, HGR, and KL-6 [6, 7]. However, these markers lack sensitivity and specificity and in some studies it is

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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unclear which circulating protein form is relevant, whether the circulating protein level is altered due to hepatic carcinogenesis or rather due to other cellular changes such as inflammation, lack confirmatory studies, or await the development of quantitative methods to evaluate their utility. It is possible that a single marker may not be sufficient to diagnose HCC, hence it may be important to test combinations of markers to improve diagnostic performance. The AFP marker remains the gold standard for HCC diagnosis and improvement of the current screening system is a major and crucial goal.

HCC treatment HCC treatment is especially difficult due to impaired liver function, raising the possibility of drug-induced liver failure. In fact, systemic chemotherapy appears to be very ineffective in HCC, due to the expression of multi-drug resistance genes [8]. As noted, most HCC patients are diagnosed at an advanced stage, excluding them from potentially curative therapies. To date, surgical resection of small tumors and liver transplantation represent the best achievements in HCC treatment. Resection is the most common in patients with relatively good liver function. However, at the time of diagnosis, only about 20% of HCC patients are eligible for this treatment option and their post-surgical survival rate is only 30–40% at 5 years [9]. Percutaneous ethanol injection, radiofrequency ablation, and transarterial chemoembolization have been employed to improve the survival rate of HCC patients [10]. Liver transplantation applying the Milan criteria increases the 5 year survival to 50–70% from 500 spotted miRNAs, including the detection of mature and precursor forms [19–21].

Protein arrays (proteome/tissue) Proteome arrays provide a better means to understand gene function since some mRNAs are transcribed, but are not translated, and thus mRNA copy number may not reflect the number of functional protein molecules in a cell. Two methods for proteome arrays exist, protein function or detecting arrays, involving immobilization of antibody probes to detect antigens in a sample, or vice versa. Each can quantify proteins, determine post-translational modifications, and correlate proteins with disease advancement or with certain treatments/environments [22]. Profiling tissues can be performed using TMAs which utilize small cylinders of formalin-fixed tissues arrayed in a single paraffin block [23]. The main limitation of protein arrays concerns the protein concentration range required for direct protein detection within a given sample, which can differ by several orders of magnitude. Current instrumentation allows for only a fraction of the proteome to be examined and measurement of low-abundance targets such as viral particles remains a challenge. High-affinity probes, such as SELEX (systematic evolution of ligands by exponential enrichment) aptamers can help to resolve this problem [24, 25].

Genomic arrays (CGH/methylation) Array comparative genomic hybridization (aCGH) permits high-resolution multi-loci mapping of small genomic regions with copy number changes, such as amplification or deletion [26]. Currently, copy number can be screened using the BAC (bacterial artificial chromosome)-based and the more recent oligonucleotide-based comparative genomic hybridization [27]. BAC aCGH is limited by costly, time-consuming, low-yield clone production, and noisy data due to non-specific hybridization of repetitive sequences. Oligonucleotide aCGH allows for flexibility in probe design, greater genomic coverage, and higher resolution (∼50 kb). New tiling BAC arrays (where each BAC overlaps with its contiguous BAC), however, can increase resolution and signal intensity and more accurately define the boundaries of genomic aberrations, but require a high concentration of high-quality BAC DNA for good array performance [28, 29]. This high-throughput data type has prompted the development of computation strategies to understand the genetic changes that occur with disease [30]. Recently, a few CGH array studies have been followed by bisulfate DNA sequencing or

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methylation-specific PCR to begin to identify the epigenetic changes that are associated with HCC.

Microarray analysis Methodologies for the analysis of array data are divided into two categories: unsupervised (to characterize the components of a dataset without a priori input or knowledge of a training set) and supervised (analysis to determine genes that fit a predetermined pattern) [31–33]. Unsupervised methods try to find internal structure or relationships in data sets by three main techniques: feature determination, which groups genes with interesting properties (principal component analysis), cluster determination, which groups genes or samples with similar patterns of gene expression (nearest-neighbor clustering, self-organizing maps, k -means clustering, and one- and two-dimensional hierarchical clustering), and network determination, which graphs gene–gene or gene–phenotype interactions (Boolean networks, Bayesian networks, and relevance networks). On the other hand, supervised methods are used to find genes with expression levels that are significantly different between groups of samples (e.g. cancer classification) and to find genes that accurately predict a characteristic of that sample (e.g. survival or metastasis). The significance found by supervised methods has been evaluated using parametric, non-parametric, and analysis of variance procedures. These methods involve permutation analyses, random partitioning of the studied dataset and false discovery limits to assess the validity of signatures associated with a tested feature and to rule out the chance of finding signatures sets by random chance. Several criteria exist for differential expression, including absolute expression level, subtractive degree of change between groups or differences in expression level across samples, fold change between groups, and ratio of expression levels across samples. These methods include the nearest-neighbor approach, decision trees, neural networks, and support vector machines. The trifold gold standard has been proposed for array studies which encompass the use of a training dataset initially to identify a signature, a test dataset to assess its predictive/classification capacity, and an independent set to validate the findings.

EMERGING CONCEPTS FROM MICROARRAY STUDIES Diagnostic signatures Chronic liver disease signatures HCC develops largely in a previously diseased liver, attributed to hepatitis viral attack, genetic/metabolic

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disorders, alcohol abuse, and/or environmental influences that are generally referred to as chronic liver disease (CLD) [14, 34]. The HCC population is thus characterized by a great heterogeneity, since both tumor and CLD may be diagnosed at different evolutionary stages, each with different therapeutic perspectives and survival probabilities. Several gene expression profiling studies have been conducted to determine the effects of CLD associated with various HCC-related etiologies (mainly viral infection) in order to identify diagnostic markers, particularly for early detection. Expression profiling studies have addressed the effects of hepatitis B and/or C infection on the liver. cDNA arrays have shown that genes associated with the TH1 immune response (including lymphocyte/monocyte activation), fibrosis, extracellular matrix remodeling, cell–cell interactions, proliferation, cell growth regulation, and apoptosis are up-regulated in hepatitis C virus (HCV)-CLD [35–37]. Candidate genes (n = 260) involved in signal transduction pathways, cell cycle control, metastasis, transcriptional regulation, immune response, and metabolism were aberrantly expressed under HBx induction by cDNA array [38]. This method has also identified several oncogenes (IGFR-2, RhoA), cell cycle regulators (p55CDC), intracellular transducers (thrombin receptor, MLK-3, MacMARCKS), stress response genes (HSP27), apoptosis related genes (FAST kinase, Bak), and transcription factors (p21 WAF) that are up-regulated in response to hepatitis B virus (HBV) infection whereas transcription factors (transcription elongation factor SII) and growth factors (monocyte chemotactic protein 1, T-lymphocyte-secreted protein I-309) were down-regulated [39]. Several of these altered genes could be correlated with chromosome regions with amplification (1q, 8q, 13q) or loss of heterozygosity (LOH) (4q, 8p, 16q, 17p) [40]. In our laboratory, we have shown that primary hepatocytes expressing HBx have altered expression of several cellular genes including oncogenes (c-myc, c-myb) and tumor suppressor genes (APC, p53) [41]. Array analysis has also been used to compare the genes altered by HBV and HCV infection. Differential gene expression was shown by cDNA array between chronic HBV and HCV hepatic lesions, with HBV affecting genes related to inflammation whereas HCV affected genes related to the anti-inflammatory process [42]. In another cDNA array study, only a slight difference between HBV and HCV host cell infection was found; however the genes that were differentially expressed were clearly regulated in a reciprocal manner and included FABP, asialoglycoprotein receptor, and thioltransferase [43]. An OLIGO array study revealed 176 genes that were altered upon HBV or HCV viral infection, including the interferon-inducible-gene IFI27 [44]. IFI27 was also shown to be highly up-regulated in HCV-HCC in an OLIGO array-based study in our laboratory in which human hepatocytes were infected with HBV- or HCV-related genes [45]. OLIGO arrays have also shown

that HCV-specific genes (NS5A) can modify gene expression to alter cell motility and adhesion, lipid transport and metabolism, calcium homeostasis, and regulation of the immune response [46]. Several of the up-regulated genes contained NFKB binding sites in their promoter region, suggesting a connection between NS5A and NFKB. Altered cellular responses to interferon have also been studied by OLIGO array, showing that 59 genes were altered in the presence of IFN [47]. The strongest effect was a down-regulation of OAS-69, an adenylate synthetase implicated in the antiviral action of IFN, and up-regulation of IL-8, which inhibits IFN antiviral activity. A proteomic array study showed that angiogenic factors, including vascular endothelial growth factor (VEGF), were up-regulated in HCV-HCC tissues [48]. Taken together, these observations suggest that a high degree of changes take place in tissue that is challenged by CLD. The identification of these premalignant changes may be useful for early cancer detection and to classify patients with CLD at risk for developing HCC, in addition to providing a window of opportunity to intervene with an effective therapy. These array-based studies have also shown that some genes are consistently altered in preneoplastic conditions and HCC, highlighting early changes that may also play a role in disease progression. However, many of these studies are relatively small, identify relatively large signatures/classifiers, do not provide sufficient follow-up data to confirm patient outcome, or are not validated in independent cohorts. Prospective studies that include a large number of patients with CLD and/or meta-analysis of existing datasets will be needed to validate the potential clinical use of these CLD-related markers as diagnostic tools.

Tumor biomarkers (tumor vs non-tumor) Microarray studies have also enhanced our understanding of how a disease process perturbs the regulatory network of genes and proteins in a way that differs from the respective normal counterpart. cDNA-based comparison between HCC and surrounding non-tumor tissue or disease-free samples have identified discriminatory expression changes involved in tumorigenesis. For example, cDNA analysis of HBV-related cell lines revealed signatures (356 genes) composed of up-regulated ribosomal-related genes including RPL6and RPS16 [49]. TIPUH1, a regulator of transcription and RNA processing of growth control genes, has also been shown to be up-regulated in HCC by cDNA array [50]. Studies have also found alterations in genes involved in protein synthesis, transcription, protein degradation, p53, Wnt–β-catenin, metabolism and tumorigenesis pathways in HCC [51]. Activators of neutrophils, antiapoptotic genes (Bcl-2), interferon response genes, and proteins related to cell differentiation or development have also been identified as differentially expressed in HCV-HCC by cDNA array [52]. Recently, cystatin B (CSTB) was

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identified as a serum biomarker for HCC by cDNA array [53]. Integrin and Akt/NFKB signaling involving SPP1, GPC3, ANXA2, VIM, and S100A10 were also up-regulated in HCC using cDNA arrays [54]. OLIGO arrays have shown that p53-related genes (n = 83) are affected by HCV infection and alter immune response, transcription, transport, signal transduction, and metabolism in tumors [55]. Several of these pathways, along with growth factor alterations, were found by Delpuech et al. in cDNA arrays comparing HBV- or HCV-positive tumor vs non-tumor tissue [56]. The authors found a clear distinction between HBV and HCV samples, where HBV affected genes involved in apoptosis, p53, and the G1/S transition whereas HCV-affected genes were more heterogeneous and included TGF-β. In a separate cDNA array study, up-regulation of mitosis-promoting genes was observed in the majority of HBV or HCV tumors vs non-tumor whereas differentially expressed genes between HBV and HCV tumors encoded enzymes that metabolize carcinogens and/or anticancer agents associated with malignant/invasive phenotype, apoptosis, or immune regulation [57]. Proteomic and TMA arrays have also been used to address the difference between HCC and non-tumor samples. A proteomic analysis of human HCV-related HCC, conducted by Yokoyama et al., found alterations in glycolysis enzymes, mitochondrial β-oxidation pathways, and cytoskeletal proteins when comparing tumor vs non-tumor tissue [58]. Other HCC-related protein classifiers include Hsp27, Hsp70, GRP78, and metabolism related enzymes [59, 60]. Proteomic studies have also shown alterations in proteins that play roles in glycolysis, fatty acid transport and trafficking, amino acid metabolism, cell cycle regulation, and cell stress, including alteration of ferritin light subunit, adenylate kinase 3 alpha-like 1, and biliverdin reductase B expression [61]. Other up-regulated genes in HCC include IGF (insulin growth factor) II, ADAM (a disintegrin and metalloproteases) 9, STAT (signal transducers and activators of transcription) 3, SOCS (suppressors of cytokine signaling) 3, and cyclin D1, whereas collagen I, SMAD 4, FHIT (fragile histidine triad), and SOCS1 were down-regulated [62]. A TMA study of HCC vs non-tumor comparisons found that the transcription repressor zinc fingers and homeoboxes 2 (ZHX2) protein expression was detected only in HCC and correlated with differentiation stage [63]. Multiple studies have aimed to determine regions of genetic loss or gain in HCC versus normal tissue. A study of 34 HCC samples found LOH at 1p, 4q, 6q, 8p, 9p, 16q, and 17p [64]. A comparison of tumor vs non-tumor HCC samples using BAC aCGH included frequent DNA copy number gains of 1q, 6p, 8q, and 20q, and losses of 4q, 8p, 13q, 16q, and 17p [65]. A study of HCV-associated HCC revealed that increases in DNA copy number were frequent at 1q, 8q, 6p, and 10p and that decreases were frequent at 17p, 16q, 4q, 13q, 10q, 1p,

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and 8p [66]. The authors found increases in copy numbers of the LAMC2, TGFB2, and AKT3 genes (located on 1q) and decreases in copy numbers of FGR/SRC2 and CYLD (located on 1p and 16q, respectively) in tumors. Another study identified narrow regions of frequent amplification on chromosome 1p and frequent deletion on 17q [67]. Paternally expressed 10 (PEG10) residing within the chromosome region 7q21 has also been implicated in HCC [68]. miRNAs have recently been utilized as potential HCC diagnostic markers. miRNAs were analyzed in human HCC by expression profiling, and defined the liver-specific miR-122 to be highly down-regulated in HCC tumors and cell lines [69]. miRNA array studies have also demonstrated that aberrant expression of miR-21 can contribute to HCC growth and spread by modulating PTEN expression, thus mediating phenotypic characteristics of cancer cells such as cell growth, migration, and invasion [70]. In other miRNA-based studies, mir-224 and a novel mRNA-like non-coding RNA named highly up-regulated in liver cancer (HULC), and a 16-miRNA set were found to be significantly up-regulated in HCC [71–73]. In another study comparing HCC samples and adjacent non-tumor, eight miRNAs were shown to be significantly altered, five of which were down-regulated in HCC and could predict HCC with 97% accuracy [74].

Tumor biomarkers (tumor vs cirrhosis) Several array studies have also compared early neoplastic stages (fibrosis/cirrhosis) with HCC in human samples. A study of 59 preneoplastic CLD (hepatitis, autoimmune hepatitis, primary biliary cirrhosis, etc.) conducted in our laboratory found genes associated with high or low risk of HCC development [75]. This 273-gene signature was validated in three independent cohorts and included 12 secretory genes in the top gene set. aCGH of 63 HCCs found etiology-dependent copy number gains and MYC overexpression in viral and alcohol-related HCCs, resulting in up-regulation of MYC target genes on 8q24 [76]. In a separate cDNA array-based study, 25 cirrhosis-specific genes were identified that were related to the inflammatory status of adjacent HCC tissue [77]. In an OLIGO array-based study of fibrotic stages, genes involved in carbohydrate metabolism were more highly expressed in HCC patients than in cases at the F3–4 fibrotic stage [78]. The use of comprehensive proteomic profiling of sera to differentiate HCC from CLD found 250 proteins that were significantly different between the HCC and CLD cases [79]. In a comparison of HCC with CLD (either HBV or HCV positive) or HCC without CLD in an OLIGO array, genes involved in transcription, metabolism and cell growth were differentially expressed [80]. A study of cirrhosis vs HCV-HCC showed that eight genes were significantly altered (GPC3, TERT, Survivin, XLKD1, and CDH1) [81]. Genome-wide miRNA arrays were used to identify 35

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miRNAs including let7 and miR-181 family members that differed between HCC and cirrhosis [82]. Two separate proteomic profiling studies revealed an 11-peak SELDI profile or four-peptide panel that could distinguish HCC from HCV-related cirrhosis and was an independent predictor of HCC [83, 84].

Tumor biomarkers (epigenetic signatures) HCC development is thought to be a multistep process, involving not only accumulation of genetic changes, but also epigenetic changes, such as methylation and histone modification, which can reversibly alter regulatory genes. A few studies have begun to address the epigenetic changes that occur in HCC. In a cDNA array and bisulfite PCR study, insulin-like growth factor binding protein was hypermethylated and down-regulated in 75% of HCCs [85]. In another cDNA–bisulfite PCR study, the demethylating agent 5-aza-dC was used to identify hepatocyte growth factor (HAI-2/PB) as a frequent hypermethylated gene in HCC [86]. An OLIGO-based analysis of human HCC cell lines showed that treatment with the demethylating agent 5-aza-dC resulted in a decrease of the tissue factor pathway inhibitor TFPI-2 [87]. A study of 60 primary HCCs using aCGH and methylation-specific PCR found no causal relationship between the methylation status of nine CpG islands, including p16, COX2, and APC, and patient outcome [88]. Thus, multiple array studies have shown that alterations occur in tumors compared with non-tumor/early disease samples or in epigenetic status. These changes have been observed using several platforms and offer potential avenues for exploration in terms of diagnostic potential and biological insight. Within platform types, however, marker sets are quite different from one another, despite similar comparison groups. This could be due to platform make-up, sample heterogeneity, or differences in etiology or ethnicity among samples. Many of these studies lack validation in most instances and are drawn from only a small dataset. Further studies will be needed to determine whether the identified changes can be useful for diagnostic or HCC classification purposes, but these studies clearly demonstrate that measurable changes occur during HCC development that may be useful for early detection.

Prognostic HCC signatures Metastasis/survival/recurrence signatures in HCC tumor tissues A main goal in HCC treatment is the prevention or inhibition of metastasis. Understanding the mechanisms involved in the process of tumor invasion and metastasis

is a major challenge, but will have the potential to improve methods to predict whether tumor cells will spread. Important questions related to metastasis involve the relationship between primary and metastatic tumors, the initiation steps leading to cell metastasis, and whether these changes are inherent to the cell or are acquired through time and/or environmental status. The current model for metastasis suggests that carcinogenesis is a multi-stage process from a benign to a malignant state that is initiated by rare genetic alterations in a single cell, followed by clonal selection and population expansion [89]. In HCC, however, such stepwise and specific progression-related genetic changes have not been illustrated [3]. Several array studies have addressed whether measurable changes occur in the transcriptome, proteome, and genome of metastatic HCC cells. A comprehensive cDNA analysis was performed in HCV-related HCCs that identified 35 genes involved in PVI [90]. The authors found that the inhibitor of deoxyribonucleic acid binding 2 (ID2), encoding a liver-rich dominant-negative helix–loop–helix protein was associated with PVI and validated this finding by quantitative reverse transcriptase polymerase chain reaction (qRT-PCR), Western blot analyses, and in an independent set. A cDNA array was also employed to profile gene expression patterns in two subtypes of HCC, solitary large hepatocellular carcinoma (SLHCC) and nodular hepatocellular carcinoma (NHCC), which differ significantly in metastatic incidence [91]. RhoC expression was significantly decreased in SLHCC compared with NHCC and strongly correlated with HCC metastasis, implicating RhoC as a potential prognosis marker and therapeutic target for HCC [92]. Another cDNA-based study found that HCC with high expression of the ubiquitin-conjugating enzyme Ube2c displayed PVI and poor disease-free survival rates [93]. The MAPK pathway (ERK1/2 and p38) has also been implicated in HCC metastasis by OLIGO array [94]. In our laboratory, we have applied cDNA arrays to show that intra-hepatic metastatic lesions are indistinguishable from their primary HCC [95]. However, primary metastasis-free HCC was distinct from primary HCC with metastasis. These data indicate that metastatic potential is an inherent quality of the primary tumor rather than a capability acquired over time through mutation. A 153-HCC metastasis gene signature, whose lead gene was osteopontin (OPN), was developed to classify metastatic HCC accurately. An OLIGO array-based study found 39 genes that were significantly correlated with metastasis, including cortactin, a cortical actin-associated protein substrate of Src [96]. TMAs and aCGH have also been used to study HCC metastasis. ZHX2, described earlier as a possible HCC diagnostic marker, was also found by TMA to be expressed significantly higher in primary lesions with metastasis than in those without this phenotype [63]. The clinical significance of FGF3 overexpression was studied by TMA in 60 pairs of primary/metastatic HCCs

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and showed that overexpression of FGF3 was significantly associated with HCC metastasis and recurrence (p < 0.01) [97]. A significant overexpression of clusterin (CLU) was found in metastatic HCC in a paired tissue study (n = 104) using TMA-containing arrays [98]. In addition, Id-1 (inhibitor of differentiation/DNA synthesis) and also Rac and VEGF, key angiogenic factors in cancer progression, were correlated with HCC metastasis [99]. Meanwhile, aCGH analysis of early and advanced components of nodule-in-nodule HCC found that genetic inactivation of the APC gene played a significant role in the progression of sporadic HCC, possibly through activation of the Wnt/β-catenin pathway [100]. aCGH was also used to examine the 7q21–q22 region for its involvement in HCC and found alterations in PFTAIRE protein kinase 1 (PFTK1), ODAG, CDK6, CAS1, PEX1, SLC25A, and PEG10 within this region [101]. The authors suggested that up-regulation of PFTK1, in particular, may confer a motile phenotype in malignant hepatocytes that correlates with metastasis. LOH has also been observed at 16q and 17q in HCC and occurred more frequently in metastatic lesions [102]. In our laboratory, we have investigated whether the expression of certain miRNAs is associated with HCC metastasis [103]. We examined the miRNA expression profiles of 482 cancerous and non-cancerous specimens from radical resection of 241 HCC patients. Using a clinically well-defined cohort of 131 cases, we built a unique 20-miRNA metastasis signature that could significantly predict (p < 0.001) primary HCC tissues with venous metastases from metastasis-free solitary tumors. A survival risk prediction analysis revealed that a majority of the metastasis-related miRNAs were associated with survival. Furthermore, the 20-miRNA tumor signature was validated in 110 additional cases as a significant independent predictor of survival (p = 0.009) and was significantly associated with both survival and relapse in 89 early-stage HCC (p = 0.022 and 0.002, respectively). These 20 miRNAs may provide a simple profiling method to assist in identifying HCC patients who are likely to develop metastases/recurrence. Functional analysis of these miRNAs may enhance our biological understanding of HCC metastasis. Tumor recurrence complicates resection in a large percentage of cases due to either true metastases or the development of de novo tumors. Vascular invasion, multinodularity, and degree of differentiation are the major predictors of recurrence. Kurokawa et al. addressed the issue of molecular prediction of early recurrence of HCC after resection and identified a 20-gene signature using a PCR-based platform in 100 HCC patients that could predict recurrence with 70% accuracy in an independent cohort of 40 patients [104]. A 12-gene OLIGO array-based signature has also been shown to predict recurrence within 1 year post-surgery with 93% accuracy [105]. Another OLIGO study identified a 57-gene signature that could predict recurrent disease at diagnosis with 84% accuracy

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and was validated in an independent test set [106]. cDNA array of HCCs identified claudin-10 expression level to be associated with disease recurrence and was validated by qRT-PCR and associated with survival in multivariate Cox regression analysis [107]. In addition, DNA microarray analysis in a training set of 33 HCCs found 46 genes linked to early intrahepatic recurrence included a down-regulation of immune response-related genes encoding MHC class II antigens (HLA–DRA, HLA–DRB1, HLA–DG, and HLA–DQA) [108]. HLA family members have also been implicated in early intrahepatic recurrence by cDNA array [109]. cDNA arrays have also been used to identify a 46-gene signature associated with extrahepatic recurrence [110].

Metastasis/survival/recurrence signatures in HCC non-tumor tissues Microarray technology can also help to resolve the factors that determine the organ distribution of metastasis. Studies have suggested that while tumor cells affect metastatic capacity, the organ can also contribute to this phenotype [111–113]. Despite considerable tumor cell dissemination, frequently observed in the hepatic venous system, metastases are rare and may be influenced by permissive target environments. A large percentage of metastasis distribution can be explained based on the circulatory anatomy; however, many metastatic sites cannot be predicted. Some mechanisms of metastasis have been set forth that include an equal chance of tumor cell dissemination in all organs or preferential growth in specific organs. These mechanisms may play a role in various degrees depending on the tumor model system. To determine the role of the hepatic microenvironment in HCC metastasis, our laboratory compared the cDNA profiles of non-cancerous surrounding hepatic tissues (n = 115) from HCC patients with venous metastases, which we termed a metastasis-inclined microenvironment (MIM) sample, with those without detectable metastases, which we termed a metastasis-averse microenvironment (MAM) sample [114]. We identified a unique change in the gene expression profiles associated with a metastatic phenotype which was refined to 17 immune-related genes. This signature was inherently different from the HCC tumor signature found in our laboratory and was validated in an independent cohort (n = 95). The non-tumor signature could successfully predict venous and extra-hepatic metastases by follow-up with >92% overall accuracy and was a superior and independent prognostic indicator compared with other available clinical parameters for determining patient survival or recurrence. Dramatic changes in cytokine responses, favoring an anti-inflammatory microenvironmental condition, occur in MIM samples, where a predominant Th2-like cytokine profile, favoring a humoral response, was associated with MIM cases. Colony-stimulating factor-1 (CSF1)

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may be one of the cytokines overexpressed in the liver milieu that is responsible for this shift. Metastasis and recurrence continue to be a problematic issue affecting HCC patient outcome. Profiling methods have identified many alterations that occur in HCC metastasis, some involving well-known metastasis-associated factors such as the angiogenesis-related VEGF and others identifying novel players related to this phenotype. These metastasis-related tumor and non-tumor signatures have increased our knowledge of the biological pathways that are affected during this process. In addition, these biomarkers may have possible clinical utility to identify HCC patients who may be prone to these outcomes and are tools that can be used to stratify patients for adjuvant therapy. However, the signatures discussed above are largely non-overlapping, suggesting a significant heterogeneity. Although some of these markers have been associated with outcome, future validation, and functional studies, including the use of large cohorts and independent validation sets, will be needed to assess their prognostic significance.

Hepatic stem-cell signatures The variability in the prognosis and the heterogenic nature of HCC suggest that this disease may comprise several distinct biological subtypes. These subtypes may result from activation of different oncogenic pathways during tumorigenesis and/or from different cell origins. Microarray analysis can aid in determining the characteristics of separate HCC subtypes that can provide insight into the cellular origin of the tumor. Recent studies suggest that HCC may arise from liver stem cells or cells with stem-cell like features which are capable of cellular plasticity, dynamic cell motility, and integral interaction with the microenvironment. It is known that certain groups of HCC patients differ in their prognosis. Array-based studies have defined HCC subgroups that have similarities to stem cells and are associated with poor outcome. Integrated gene expression data from fetal hepatoblasts and adult hepatocytes with HCC from human and mouse models found that individuals with HCC who shared a gene expression pattern with fetal hepatoblasts had a poor prognosis [115]. The gene subset included markers of hepatic oval cells, suggesting that HCC of this subtype may arise from hepatic progenitor cells. Analyses of the gene networks associated with this subtype showed activation of AP-1 transcription factors which might play key roles in tumor development. In our laboratory, we have used cDNA arrays to identify a HCC subtype with features of hepatic stem cells that expresses AFP and a cell surface hepatic stem-cell marker, EpCAM [116, 117]. EpCAM-positive cells from this subtype have self-renewal and differentiation traits and can initiate highly invasive HCC in NOD/SCID mice [118]. The Wnt/β-catenin signaling

pathway is augmented in this subtype, suggesting that therapeutic approaches geared towards Wnt/β-catenin signaling inhibitors may impact the survival of HCC patients with this stem cell-like subtype. We have also recently found that miRNAs are associated with this stem cell-like HCC subtype, suggesting that targeting miRNA pathways may alleviate the poor prognosis of HCC patients [119]. Thus, recent studies have identified subtypes of HCC that are related to stem cell-like/progenitor cell-like phenotypes and associated with poor outcome. Therefore, a clear understanding of the multiple subtypes of HCC through further biological and functional studies may identify specific factors that determine more aggressive HCC. Our ability to determine these subtypes using biomarkers may help to refine treatment options for patients based on their HCC subtype classification. Furthermore, functional follow-up studies of these prognosis-related biomarkers will aid the generation of novel therapeutic approaches to block pathways associated with poor outcome and thus help to alleviate dismal prognosis.

CONCLUSION Microarray technology has provided the unprecedented ability to study the human liver transcriptome/proteome/genome in a high-throughput fashion in relation to disease pathogenesis and clinical measures. This method, using multiple sample types, array platforms, and data analysis methods, has allowed the definition of mechanisms related to HCC carcinogenesis, cancer subtype classification, molecular diagnosis, and prognosis markers and also provided therapeutic targets (Figure 61.1). Thus, the advent of this technology has afforded the integration of descriptive characteristics of a biological system with a genomic readout and potential clinical application. Microarrays have steadily become more comprehensive and stable, not only increasing in number of elements that can be arrayed but also expanding with regard to the types of material that can be analyzed. Despite these advances, many fundamental issues still remain to be resolved. Array profiling requires the physical destruction of cells/tissues and therefore other consequential assays cannot be conducted on the same material. There are also several substantial sources of variation in arrays (among samples, within arrays, mixed cell types, etc.) and there is often a failure to account for such variations, leading to overinterpretation or spurious functional gene associations. Techniques such as laser capture microdissection have improved this problem somewhat by permitting the isolation of specific cells from samples. The overall quality and amount of starting material are a major challenge and are limited by the amount and complexity of the sample in addition to user-related handling. In addition, there

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Figure 61.1 A synopsis of molecular marker identification, validation, and clinical assessment from global gene expression profiling. In the marker discovery phase, samples are collected from a training sample set, hybridized to an array(s) and analyzed using bioinformatic algorithms to select molecular markers of interest. In the marker validation phase, the classification/prediction capacity of the identified molecular markers is tested in both a test set and in an independent set of samples using an array method or a different technology such as quantitative RT-PCR. In the clinical utility phase, the molecular markers that have passed the discovery and validation stages are assessed for their diagnostic/prognostic value. Markers which show efficacy in differentiating patient groups can be further tested for their therapeutic capacity. FFPE, formalin-fixed paraffin embedded

are numerous error-prone steps in microarray protocols which have been and continue to be improved through advances in automation. In addition, many oncogenic processes are not accounted for by array analysis since they are regulated post-transcriptionally. Therefore, elements such as protein localization and modification need to be included in HCC profiling. Another confounding issue in microarray-based studies is the use of multiple array platforms among published studies, which causes great difficulty in data comparison. The detailed names and information regarding genes of interest might not be available and this complicates the interpretation of results. Also, probe sets once thought to be unique for a particular gene might not remain unique as more genomic data are collected and may not recognize all isoforms of a gene. Resolution range is a large limitation in array analysis, whereby important changes may not be assessed or studied due to the cut-off criteria in the analysis. Improvements in hybridization, platform

composition, detection, and labeling have been used to address these issues. In addition, the software packages and analysis algorithms used in array studies differ among publications. Some data analysis issues include statistical sample size, study design, modeling, permutation, significance, and normalization, each of which may affect assessment between published data. Such problems may be alleviated by setting adherence guidelines for microarray statistical analysis and reporting, such as those established by the International Microarrays Gene Expression Data group, the REMARK guidelines or incorporation of proper study design that is suitable for array-based biostatistical analyses [120–122]. Lastly, each microarray can only provide information concerning the targets that are included on that array. Future studies may require integrative analysis of multiple platforms in order to define the exact cancer-related molecular changes on multiple biological levels and to distinguish the key players

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from their downstream effects. The utilization of a Biological Expression Network Discovery (BLEND) strategy, integrating global molecular profiling data along with mechanistic/functional studies, may improve the diagnosis, treatment, and prognosis of HCC patients. Advances in statistical methods to integrate multiple platforms will be required in order to be able to make such assessments. Recently, systems have been developed (e.g. the Illumina Genome Analyzer) that offer whole genome analysis using a massive parallel sequencing that is useful for discoveries in genomics and gene expression studies. Such systems offer an extremely high-throughput method to complete large scale global studies in an accurate manner and may allow for ease in cross-platform-type analyses since an enormous multilevel dataset can be achieved with a relatively small amount of the same starting material. Despite multiple publications on diagnostic and/or prognostic HCC markers, we are facing critical challenges in translating the findings to clinical practice. In order to improve the outcome of HCC, molecular profiling has to reach clinical applicability in terms of reproducibility and reliability. Clinical markers must be easily accessible for measurement, preferably through a method that is not invasive to a patient (blood product, urine, etc.) and be specific. In addition, since many of the identified molecular sets from array studies are large in number, they will need to be refined to a smaller number of informative biomarkers such that they can be more readily interrogated in a clinical setting. Studies will need to be performed to assess appropriate sample size for accurate diagnostics and appropriate validation cohorts that incorporate gender, race, and underlying etiological differences among HCC patients. This will require large prospective studies but have the capacity ultimately to improve the dismal outcome associated with HCC. Nonetheless, the biomarkers that have been identified through gene profiling, particularly those expressed in serum, are the first and major stepping-stones toward useful clinical application. Overall, molecular profiling studies have become powerful tools for understanding biological effects on a more global scale. Current HCC-related array studies, such as those presented in this chapter, and continuous improvements in microarray technology, experimental design, and statistical analyses will undoubtedly lead to important and major advances in our understanding of the molecular mechanisms of this disease. The molecular markers identified from such studies not only extend our biological insight but also provide the framework towards predictive personalized care for HCC patients. We are now at the horizon of implementing molecular markers from global gene expression profiling in clinical practice to guide treatment simply and accurately with the hope of improving the outcome of the many patients who suffer from HCC.

ACKNOWLEDGMENTS A multitude of global gene expression studies have been implemented in studies of HCC. The authors apologize for the many noteworthy references that could not be included in this chapter owing to space limitations.

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and Fan, S.T. (2006) Regulation of angiogenesis by Id-1 through hypoxia-inducible factor-1alpha-mediated vascular endothelial growth factor up-regulation in hepatocellular carcinoma. Clin Cancer Res, 12, 6910–19. Katoh, H., Shibata, T., Kokubu, A., Ojima, H., Kosuge, T., Kanai, Y. and Hirohashi, S. (2006) Genetic inactivation of the APC gene contributes to the malignant progression of sporadic hepatocellular carcinoma: a case report. Genes Chromosomes Cancer, 45, 1050–57. Pang, E.Y., Bai, A.H., To, K.F., Sy, S.M., Wong, N.L., Lai, P.B., Squire, J.A. and Wong, N. (2007) Identification of PFTAIRE protein kinase 1, a novel cell division cycle-2 related gene, in the motile phenotype of hepatocellular carcinoma cells. Hepatology, 46, 436–45. Nagai, H., Pineau, P., Tiollais, P., Buendia, M.A. and Dejean, A. (1997) Comprehensive allelotyping of human hepatocellular carcinoma. Oncogene, 14, 2927–33. Budhu, A., Jia, H.L., Forgues, M., Liu, C.G., Goldstein, D., Lam, A., Zanetti, K.A., Ye, Q.H., Qin, L.X., Croce, C.M. et al. (2008) Identification of metastasis-related microRNAs in hepatocellular carcinoma. Hepatology, 47, 897–907. Kurokawa, Y., Matoba, R., Takemasa, I., Nagano, H., Dono, K., Nakamori, S., Umeshita, K., Sakon, M., Ueno, N., Oba, S. et al. (2004) Molecular-based prediction of early recurrence in hepatocellular carcinoma. J Hepatol , 41, 284–91. Iizuka, N., Oka, M., Yamada-Okabe, H., Nishida, M., Maeda, Y., Mori, N., Takao, T., Tamesa, T., Tangoku, A., Tabuchi, H. et al. (2003) Oligonucleotide microarray for prediction of early intrahepatic recurrence of hepatocellular carcinoma after curative resection. Lancet , 361, 923–29. Wang, S.M., Ooi, L.L. and Hui, K.M. (2007) Identification and validation of a novel gene signature associated with the recurrence of human hepatocellular carcinoma. Clin Cancer Res, 13, 6275–83. Cheung, S.T., Leung, K.L., Ip, Y.C., Chen, X., Fong, D.Y., Ng, I.O., Fan, S.T. and So, S. (2005) Claudin-10 expression level is associated with recurrence of primary hepatocellular carcinoma. Clin Cancer Res, 11, 551–56. Matoba, K., Iizuka, N., Gondo, T., Ishihara, T., Yamada-Okabe, H., Tamesa, T., Takemoto, N., Hashimoto, K., Sakamoto, K., Miyamoto, T. et al. (2005) Tumor HLA-DR expression linked to early intrahepatic recurrence of hepatocellular carcinoma. Int J Cancer, 115, 231–40. Uchimura, S., Iizuka, N., Tamesa, T., Miyamoto, T., Hamamoto, Y. and Oka, M. (2007) Resampling based on geographic patterns of hepatitis virus infection reveals a common gene signature for early intrahepatic recurrence of hepatocellular carcinoma. Anticancer Res, 27, 3323–30. Iizuka, N., Tamesa, T., Sakamoto, K., Miyamoto, T., Hamamoto, Y. and Oka, M. (2006) Different molecular pathways determining extrahepatic and intrahepatic recurrences of hepatocellular carcinoma. Oncol Rep, 16, 1137–42.

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111. Paget, S. (1989) The distribution of secondary growths in cancer of the breast. 1889. Cancer Metastasis Rev , 8, 98–101. 112. Fidler, I.J. (2002) Critical determinants of metastasis. Semin Cancer Biol , 12, 89–96. 113. Liotta, L.A. (1985) Mechanisms of cancer invasion and metastasis. Important Adv Oncol , 28–41. 114. Budhu, A., Forgues, M., Ye, Q.H., Jia, L.H., He, P., Zanetti, K.A., Kammula, U.S., Chen, Y., Qin, L.X., Tang, Z.Y. et al. (2006) Prediction of venous metastases, recurrence and prognosis in hepatocellular carcinoma based on a unique immune response signature of the liver microenvironment. Cancer Cell , 10, 99–111. 115. Lee, J.S., Heo, J., Libbrecht, L., Chu, I.S., Kaposi-Novak, P., Calvisi, D.F., Mikaelyan, A., Roberts, L.R., Demetris, A.J., Sun, Z. et al. (2006) A novel prognostic subtype of human hepatocellular carcinoma derived from hepatic progenitor cells. Nat Med , 12, 410–16. 116. Yamashita, T., Budhu, A., Forgues, M. and Wang, X.W. (2007) Activation of hepatic stem cell marker EpCAM by Wnt–β-catenin signaling in hepatocellular carcinoma. Cancer Res, 67, 10831–39. 117. Yamashita, T., Forgues, M., Wang, W., Kim, J.W., Ye, Q., Jia, H., Budhu, A., Zanetti, K.A., Chen, Y., Qin,

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62

Cell Cycle Control in the Liver Jeffrey H. Albrecht1 and Lisa K. Mullany2 1 Division

of Gastroenterology, University of Minnesota and Hennepin County Medical Center, Minneapolis, MN, USA 2 Minneapolis Medical Research Foundation, Minneapolis, MN, USA

INTRODUCTION The control of cell division has been the focus of significant interest because of remarkable progress in defining the molecular machinery that drives cell cycle progression. This has provided substantial insight into normal cell proliferation as well as an understanding of how abnormal regulation of the cell cycle contributes to cancer and organ failure. The purpose of this chapter is to provide an overview of key concepts related to the role of cell cycle proteins in the liver. We do not highlight important work regarding growth factors, cytokines, signal transduction pathways, and transcription factors because these are addressed elsewhere in this volume (e.g. see Chapters 36 and 37. In addition, readers interested in a comprehensive discussion of the cell cycle may wish to consult more general reviews [1, 2]. Most of the literature on cell cycle proteins in the liver has focused on models of hepatic regeneration in vivo and hepatocyte proliferation in culture, although a number of studies have examined these proteins in liver diseases. In addition, because of the critical link between the cell cycle machinery and cancer [1–6], numerous investigators have examined how deregulated expression of proliferation-control proteins might contribute to the development of hepatocellular carcinoma (HCC) and other liver malignancies. At present, the field of cell cycle control in the liver might be considered a “work in progress.” Although certain basic concepts are widely accepted, a large number of mechanistic details remain to be defined, The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

and this area of research has not yet had an impact on the care of patients with liver disease. However, given the rapid pace of developments, it is likely that therapies directed at the cell cycle machinery will play a role in clinical medicine. Of particular interest to hepatologists are potential strategies to promote adaptive cell proliferation in disease states and interventions to prevent or treat HCC.

BASICS OF THE CELL CYCLE The essential function of the cell cycle is to produce two daughter cells containing normal cellular components and genetic material. Traditionally, the regulation of cell growth (increased cell size) has been viewed separately from cell proliferation (cell division) because the two processes can be regulated distinctly. However, under normal conditions, growth and proliferation are coordinately regulated so that each generation of cells is the appropriate size [3]. Compared with the understanding of cell cycle progression, less is known about the biochemical pathways regulating growth. A key determinant of cell growth is the protein synthetic apparatus, which must be substantially up-regulated to allow for normal growth and proliferation. Although a review of global protein synthesis and growth is beyond the scope of this chapter, it is becoming increasingly apparent that the pathways controlling growth and cell cycle progression are intertwined (see Chapter 37) [3, 7–9].

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

THE LIVER: BASICS OF THE CELL CYCLE

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Figure 62.1 Stages of the cell cycle. Cells respond to extracellular signals such as growth factors to exit the quiescent state (G0) and enter the cell cycle. During the G1 phase, cells remain dependent on mitogens until they reach the restriction point in late G1 phase, after which the cell cycle progresses autonomously. If growth factors are withdrawn in early G1 phase, the cells can revert to G0. After completion of mitosis, cells may continue to proliferate or return to the quiescent state

The cell cycle consists of the ordered phases Gap phase 1 (G1), S, G2, and M (Figure 62.1). Entry of quiescent cells (G0) into the initial stage of the cell cycle (G1) is regulated by extracellular stimuli such as growth factors or cytokines that activate specific signal transduction cascades (see Chapters 36 and 37). In addition, cell cycle progression requires an adequate supply of nutrients and attachment to an appropriate extracellular matrix. Cells generally remain dependent on mitogenic stimuli (particularly growth factors) for most of the G1 phase. If these stimuli are withdrawn, the cell will revert back to G0. In addition, anti-proliferative factors such as transforming growth factor-β (TGFβ) can inhibit progression through the G1 phase. However, once a key checkpoint in late G1 phase, (called the restriction point), is reached, growth factors are no longer required and cell cycle progression will proceed autonomously through mitosis – that is, the restriction point represents “the point of no return” with regard to mitogens. The G1 phase is therefore the stage of the cell cycle that is most subject to regulation by extracellular signals. As is discussed below, these signals eventually converge on the core cell cycle machinery that controls proliferation, consisting of cyclins and cyclin-dependent kinases (cdks). Importantly, a hallmark of malignant cells is the capacity to proliferate in the absence of appropriate mitogenic signals, which is mirrored by abnormal regulation of proteins involved in the G1 restriction point [2, 5]. In many systems, the G1 phase is the longest stage of the cell cycle, yet the cellular events that occur during this phase are still incompletely characterized. After stimulation by growth factors, cell surface receptors acutely

activate various signal transduction cascades, including the mitogen-activated protein kinase (MAPK) and phosphatidylinositol 3-kinase (PI3K) pathways that trigger the expression of immediate early genes including transcription factors (see Chapter 36). The initial activation of these pathways in early G1 phase is transient and then the signals are damped. Since these early events occur before the restriction point, their effects are reversible, and withdrawal of growth factors leads to regression into G0 phase. In the presence of continued mitogen stimulation, a second burst of signal transduction kinase activity occurs later in the G1 phase, which is associated with induction of the D-type cyclins and progression through the restriction point [10–13]. The critical biochemical events that occur between the early and late G1 peaks of signal transduction kinase activity have not been fully determined, but appear to include a host of metabolic adaptations to accommodate the demands of growth and proliferation. Cells enter the S phase shortly after progression through the restriction point. The S phase is characterized by the rapid synthesis of DNA to replicate the parent chromosomes. This is followed by the G2 phase (Gap phase 2 ), during which the cells prepare for mitosis. During the M phase, cells form mitotic spindles that segregate chromosomes and other cellular materials into two compartments, culminating in cytokinesis that produces two daughter cells. Of note, hepatocytes sometimes fail to undergo cytokinesis after progressing through the cell cycle, leading to the appearance of binucleated or polyploid cells in the normal liver [14, 15]. As a general rule, progression through S, G2, and M phases occurs on a fixed schedule after cells traverse the restriction point. However, each of these phases can be interrupted by additional cell cycle checkpoints that are activated in response to adverse conditions. An example of an important cell cycle checkpoint is that which occurs in the setting of DNA damage. In normal cells, DNA damage will trigger mechanisms that pause the cell cycle to allow for repair of DNA or, in more severe cases, lead to cell cycle withdrawal and apoptosis. This is mediated by the protein kinases ATR and ATM and downstream effectors including the Chk1 and Chk2 kinases and the p53 tumor suppressor protein. Chk1/2 lead to diminished activity of the cdc25 family of phosphatases that normally activate cdks, and p53 up-regulates the expression of the p21 cdk-inhibitory protein. The resulting inhibition of cdk activity leads to a cell cycle arrest. In addition to the DNA damage checkpoint, cells can be arrested by other cellular stresses including nutrient deprivation, oxidative stress, telomere dysfunction, and mitotic spindle abnormalities. A key function of these checkpoints is to prevent propagation of chromosomal abnormalities, so that genetic errors are not passed on to daughter cells. The loss of normal checkpoint function (as is seen in p53 mutations) is a feature of most cancers [2, 5].

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CYCLINS, CDKS, AND ASSOCIATED REGULATORY PROTEINS The study of cell cycle control has been greatly facilitated by the fact that the molecules that regulate proliferation have been remarkably well conserved throughout evolution. Thus, pioneering work in yeast has provided a framework for understanding the mechanisms that regulate the mammalian cell cycle [16]. The core cell cycle machinery is composed of protein kinases complexes consisting of cdks and their activating partners, the cyclins. Activation of different cyclin–cdk complexes promotes progression through discrete phases of the cell cycle (Figure 62.2). The expression of most cdks does not vary substantially during proliferation (with the exception of cdk1). However, cyclins are induced at different stages of the cell cycle and activate their appropriate cdk partner(s). During the G1 phase, one or more of the D-type cyclins is induced in mid-to-late G1 phase in response to mitogenic signaling pathways. In hepatocytes and most other cell types, cyclin D1 is the primary mitogen-sensitive cyclin during the G1 phase [13, 17–19]. Numerous signal transduction cascades converge to regulate cyclin D1 expression at the level of transcription, messenger RNA (mRNA) stability, translation, and protein stability. Cyclin D1 thus serves to integrate external signals that determine whether cells proceed through the G1 restriction point.

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Figure 62.2 Cyclin–cdk complexes in the cell cycle. Mitogens stimulate the expression of cyclin D1, which binds and activates cdk4 beginning in the G1 phase. This is followed by activation of cyclin E–cdk2, cyclin A–cdk2, cyclin A–cdk1, and cyclin B–cdk2 as cells progress through the cycle

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Cyclin D1 binds to cdk4 (or cdk6) to form active kinase complexes in late G1 phase. The best characterized target of the cyclin D1–cdk4 kinase is the retinoblastoma protein (Rb). In quiescent cells, Rb binds to and inhibits members of the E2F transcription factor family (E2F1, 2, and 3), which are key regulators of cell cycle progression (Figure 62.3). This inhibition occurs through recruitment of transcription co-repressors (e.g. histone deacetylases) and by binding of Rb to the transactivation domain of the E2Fs. Phosphorylation of Rb leads to its disassociation from E2F1–3, which then transcriptionally activate the expression of numerous genes involved in cell cycle progression. A key target gene is cyclin E, which is induced downstream of cyclin D1 and E2F. Cyclin E forms a kinase complex with cdk2, which then phosphorylates Rb at distinct sites and further promotes E2F-dependent gene expression at the G1/S phase boundary. Progression through the S and M phases is regulated by the cyclin A–cdk2 and cyclin B–cdk1 complexes, respectively. Importantly, activation of cyclin D1–cdk4, inactivation of Rb, and transcription by E2F appear to be the key molecular events during progression through the G1 restriction point. As is discussed below, deregulation of the cyclin D–Rb–E2F pathway occurs in virtually all cancers. Cyclin D1 protein expression persists as long as mitogens are present, but the expression of cyclins E, A, and B occurs periodically during proliferation. The induction of these proteins occurs through increased transcription, mRNA stability, and translational efficiency at discrete points in the cell cycle. Furthermore, each undergoes rapid proteolysis as the cell cycle progresses, a process which is controlled by a series of specific ubiquitin ligases. The combined effect of these mechanisms results in precisely timed expression of cyclins at the appropriate stages of the cell cycle. In addition to binding to the appropriate cyclin partner(s), cdk activity is regulated by its phosphorylation. After binding to its cyclin partner, full activation of each cdk requires phosphorylation of a key threonine site, which is performed by cyclin-dependent kinase-activating kinase (CAK). In mammals, the primary CAK is the cyclin H–cdk7–Mat1 complex, which is expressed constitutively in quiescent and proliferating cells. Cdk activity is negatively regulated by phosphorylation of tyrosine residues in the N-terminal portion of the protein. These are phosphorylated by the wee1 and myt1 kinases and are dephosphorylated by the cdc25 family of phosphatases. As noted above, cdc25 activity is inhibited during certain cell cycle checkpoints (e.g. DNA damage), which results in increased phosphorylation of tyrosine residues in the cdk, inhibited cyclin/cdk activity, and a cell cycle arrest. Another key mechanism by which cdk activity is regulated is binding to cyclin-dependent kinase inhibitor (CKI) proteins. The ink4 family of CKIs (p15, p16, p18, and p19) bind to cdk4 and cdk6 and prevents their association with the D-type cyclins, thereby inhibiting cyclin D–cdk4

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Figure 62.3 Regulation of the G1 restriction point. In quiescent cells, cyclins are expressed at low levels and monomeric p27 is present in high abundance. As cyclin D1 is induced in mid-G1 phase, it binds cdk4 and activates this kinase (after phosphorylation by CAK). Cyclin D1–cdk4 complexes also serve to sequester p21 and p27, thereby allowing activation of cyclin E–cdk2 and cyclin A–cdk2 acting downstream in the cell cycle. Rb is progressively phosphorylated as cells proceed through the restriction point, which allows full activation of E2F-mediated gene transcription

activity. The ink4 proteins are generally not highly expressed in proliferating cells, but are up-regulated in response to anti-proliferative signals such as TGFβ or in senescent cells. The cip/kip family of proteins (p21, p27, and p57) can bind and inhibit each of the cyclin–cdk complexes in the cell cycle. The best-studied CKI proteins are p21 and p27. p27 is abundantly expression in quiescent cells, and exists in a monomeric form capable of binding cyclin–cdk complexes. During the G1 phase, the concentration of p27 monomers is diminished by two main mechanisms. The first is its proteolytic degradation via the skp2 ubiquitin ligase. The second mechanism involves sequestration of p27 by cyclin D1–cdk4 complexes. As cyclin D1 expression increases during the G1 phase, this results in increased binding of p27 to cyclin D1–cdk4, thereby reducing the amount of monomeric p27 available to inhibit downstream cyclin–cdk complexes (Figure 62.3). Interestingly, at low concentrations, p21 and p27 can facilitate the assembly of active cyclin D1–cdk4 complexes, whereas at higher concentrations they are inhibitory [20–22]. In addition, phosphorylation of p21 and p27 can play a role in determining their inhibitory activity [23]. The expression of p21 is negligible in quiescent cells, but it is induced by a wide range of stimuli and can result in a cell cycle delay or arrest, depending on its stoichiometric concentration relative to cyclin–cdk complexes. In response to mitogenic stimulation, p21 is induced in

parallel to cyclin D1 and serves to slow progression through the G1 phase; thus, p21−/− cells enter the S phase more rapidly than wild-type cells [24–26]. In addition, elevated p21 expression arrests cell cycle progression in response to a number of growth inhibitory factors. One example is the p53 tumor suppressor protein, which activates transcription of p21 in response to DNA damage and other cellular insults. Of all the CKIs, p21 appears to be the most highly regulated in experimental conditions. Given the importance of precise regulation of the cell cycle, it is not surprising that there are overlapping mechanisms of cell cycle control that provide substantial redundancy. This has been best characterized in transgenic mice in which one or more cell cycle genes have been deleted [27–30]. In general, deletion of key cell cycle mediators leads to compensatory expression of other proteins that blunts the phenotypic effect. However, as is discussed below, the use of transgenic mice has provided important insight into the role of cell cycle proteins in normal physiology and disease states including cancer. A large number of proteins have been shown to be phosphorylated by cyclin–cdk complexes in proliferating cells, and it is highly likely that further substrates will be defined [27, 31, 32]. In addition, emerging evidence suggests that cyclins regulate cellular processes independently of cdks. For example, cyclin E mutants incapable of activating cdk2 can still promote DNA synthesis [33, 34].

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Cyclin D1 regulates the transcription of numerous genes through a mechanism that does not require cdk4, and appears to play a role in other cellular functions including metabolism and motility [35–37]. Furthermore, cyclin D1 can promote organ growth (including in the liver [38]) through mechanisms that do not appear to require Rb phosphorylation or cell cycle progression [27, 39–41]. The targets of cyclins and cdks have not been fully identified, but a substantial and growing body of literature suggests that they regulate diverse cellular processes during proliferation.

of the cyclin D1–Rb–E2F pathway, including resistance to apoptosis, telomere stabilization, and other abnormalities (see Chapter 61) [2, 5]. Cancer cells are often refractory to anti-proliferative signals and have defects that undermine other cell cycle checkpoints, including those that inhibit proliferation in response to chromosomal and DNA damage. As noted above, the loss of p53 checkpoint function is a common abnormality in malignancies. The failure to prevent propagation of cells with DNA and chromosomal abnormalities further contributes to genomic instability that accelerates cancer progression.

ABNORMAL REGULATION OF THE CELL CYCLE MACHINERY IN CANCER

CELL CYCLE REGULATION IN THE LIVER

Uncontrolled cell proliferation is a key feature of cancer [1–6]. In almost all malignant cells, this is associated with deregulation of the cyclin D–Rb–E2F pathway that regulates the G1 restriction point. Abrogation of this checkpoint can occur via overexpression of the D-type cyclins, decreased expression or activity of CKI proteins, or inactivating mutations of the Rb protein itself. The resulting loss of checkpoint functions leads to diminished mitogen dependence in proliferating cancer cells. Increased expression of cyclin D1 occurs in 40–50% of all cancers [1–6]. This can occur through constitutive activation of upstream signaling pathways (e.g. by activating mutations of Ras), gene amplification, and chromosomal rearrangements leading to constitutive expression of the cyclin D1 gene. The importance of cyclin D1 in the development of malignancy has been established in cyclin D1−/− mice, which are resistant to the development of breast cancer caused by activating Neu or Ras mutations [42]. Furthermore, targeted overexpression of cyclin D1 in transgenic models leads to cancer in a number of different organs, including the liver [28, 43]. Activation of cdk4 appears to be a key effect of cyclin D1 in tumorigenesis, because mice with replacement of cyclin D1 with a mutant incapable of activating this kinase are resistant to the development of breast cancer [44]. Another mechanism by which malignant cells bypass the G1 restriction point is though inhibition of CKI activity. A common example is the p16 protein, which is deleted or repressed by promoter methylation in a wide variety of tumors, including a substantial portion of HCCs [45]. Inhibition of p27 is another frequent finding in human cancers, which can occur via decreased gene expression, protein degradation, or cytoplasmic sequestration. A recent study also documented that p21 can serve as a tumor suppressor in the liver [46]. In addition to the abnormalities noted above, aberrant expression of many other cell cycle regulators have been described in human malignancies [1–6]. The development of cancer also requires abnormalities beyond deregulation

Although the liver is comprised of a diverse population of cell types, most studies of cell cycle control in this organ have focused on hepatocyte replication. Unlike many other differentiated cell types, mature hepatocytes retain a remarkable proliferative capacity that allows for rapid expansion of these cells in response to acute hepatic injury. This has been best characterized in the classic model of liver regeneration, that of 70% partial hepatectomy (PH) (see Chapter 36). Following PH, a majority of the remaining hepatocytes progress through the cell cycle in a relatively synchronous fashion, and this model provides a unique system to study cell cycle regulation in vivo. Hepatocyte proliferation is frequently observed in chronic liver diseases, although the relative contribution of mature hepatocytes and liver stem cells has not been clearly settled (see Chapters 36 and 38). In addition, isolated hepatocytes in culture proliferate readily in response to mitogens, which allows for a more detailed dissection of molecular pathways than do in vivo models. Following PH in rats or mice, hepatocytes undergo an initial “priming” phase characterized by the induction of immediate–early genes, including mitogenic transcription factors, which is followed by the up-regulation of cell cycle genes at later time points [47, 48]. The expression of cyclins and cdks in the regenerating liver after PH was first reported by Lu et al. [49], and these findings have been extended by numerous investigators (representative studies include [19, 24–26, 50–65]). Cyclin D1 is not significantly expressed in quiescent liver tissue but is markedly induced prior to the onset of DNA synthesis, and remains elevated for several days. This is followed by induction of cyclins E, A, and B as the population of hepatocytes progresses through the cell cycle. Cyclin D3 is present in quiescent liver and modestly induced, but is expressed at a much lower concentration than cyclin D1 in regenerating liver [19]. Cdk2, cdk4, and cdk6 are expressed in quiescent liver and only modestly induced after PH, whereas cdk1 is markedly up-regulated as cells enter S phase. The CKI p27 is expressed in quiescent liver and has been reported to increase or decrease modestly after PH [24, 66, 67]. On the other hand, p21 is absent in

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quiescent liver and markedly induced during regeneration. Among the cell cycle regulators, a role for cyclin D1 and p21 have been most clearly defined in hepatocytes. A significant body of evidence suggests that cyclin D1 plays an important role in the G1–S phase transition in hepatocytes in culture and in vivo. As noted above, it is not significantly expressed in quiescent hepatocytes, but it is induced by mitogens in late G1 phase at time points corresponding to the G1 restriction point [17, 19, 54, 68]. After its induction in late G1 phase, the cyclin D1–cdk4 kinase is activated and this complex sequesters most of the cellular p21 and p27 (Figure 62.3) [18, 19]. Transfection of cultured hepatocytes with cyclin D1 is sufficient to promote cell cycle progression in the absence of mitogens [68]. Similarly, expression of cyclin D1 is sufficient to promote both hepatocyte proliferation and liver growth in normal liver [43, 69]. More recent data indicate that siRNA-mediated knockdown of cyclin D1 significantly impairs mitogen-stimulated hepatocyte cell cycle progression in culture (Hanse, E.A. and Albrecht, J.H., in preparation). Furthermore, diminished cyclin D1 expression has been observed in many different models of impaired liver regeneration including cirrhosis, immune abnormalities, chronic ethanol ingestion, and protein deprivation [38, 51, 70–78]. The available data suggest that cyclin D1 is a key mediator of progression through the G1 restriction point in hepatocytes. The expression of cyclin D1 mRNA is governed by MAPK, PI3K, and jun N-terminal kinase (JNK) pathways acting downstream of mitogens in hepatocytes, which are necessary for full induction of this gene [13, 19, 79, 80]. Its expression requires attachment of hepatocytes to an appropriate extracellular matrix, which regulates the MAPK pathway [11, 81]. Cyclin D1 transcription is also regulated by the availability of certain amino acids [71]. In mouse liver, cyclin D1 can also be induced by activation of the constitutive active/androstane receptor (CAR) and thyroid hormone receptor, through mechanisms that may be distinct from those seen after PH or growth factor stimulation [82, 83]. Numerous transcription factors that play a role in liver regeneration also have been shown to bind and regulate the cyclin D1 promoter, including AP-1, NF-κB, β-catenin, ATF3, and CREB [35, 36]. In addition to transcriptional control, evidence suggests that cyclin D1 can be regulated at the level of mRNA stability after PH [84]. Furthermore, translation of the cyclin D1 mRNA is regulated by the target of rapamycin (TOR) pathway in hepatocytes [38, 85]. Through these and potentially other mechanisms, a wide variety of mitogenic stimuli regulate the expression of cyclin D1 at the level of mRNA and protein expression. Hepatocyte cell cycle progression can also occur in the absence of cyclin D1 induction. As opposed to adult liver, cyclins D2 and D3 appear to play a predominant role in hepatocyte proliferation in fetal animals [86]. Transient transfection of either cyclin D2 or D3 can trigger hepatocyte proliferation in culture and in vivo in the

absence of other mitogenic stimuli [87]. Treatment with the CAR activator TCPOBOP leads to delayed hepatocyte cell cycle progression in cyclin D1−/− mice as compared with wild-type mice, but proliferation and liver growth occur at later time points [88]. This is compatible with the observation that liver regeneration is driven by alternative mechanisms when one or more pathway is interrupted [89]. Studies from several systems have shown that cyclin E mRNA expression is induced in late G1 phase downstream of cyclin D1 via E2F-mediated transcription. However, the expression of cyclin E protein is also regulated independently of the cyclin D1–Rb–E2F pathway, and recent studies suggest that it regulates hepatocyte cell cycle activity in vivo. Hepatocyte-specific deletion of the ubiquitin ligase Cul3, which normally targets cyclin E for degradation, leads to increased cyclin E expression, proliferation, and polyploidy [90]. These results suggest that cyclin E levels are repressed in quiescent hepatocytes in part by Cul3-mediated proteolysis. Liver growth induced by transient expression of the oncogenic kinase Akt is associated with marked induction of cyclin E via a translational mechanism, and also hepatocyte polyploidy and limited cell division [91]. Cyclin E has also been linked to ribosome biogenesis, a key determinant of growth, in mice with liver-specific deletion of the ribosomal protein S6 (see Chapter 35) [92]. In S6-deleted livers, liver growth is markedly impaired after PH, and this is associated with a failure to induce cyclin E. Cyclin E may therefore be a sensor of the cellular ribosome capacity, thereby linking growth to the cell cycle machinery as has been proposed in Drosophila and other systems[39, 93]. In addition, a truncated form of cyclin E may play a role in inhibiting hepatocyte cell cycle progression [94]. These studies suggest that cyclin E can regulate hepatocyte proliferation or ploidy independently of cyclin D1, although the contribution of these mechanisms in the setting of PH or liver injury remains to be determined. A number of investigations have revealed that p21 plays a cell cycle inhibitory role in the liver. It is expressed at very low levels in the normal adult liver, but is induced by PH and other injuries associated with regeneration [57, 95]. Transgenic mice with constitutive expression of p21 in the liver have abnormal liver histology and markedly impaired hepatocyte proliferation after PH [96].On the other hand, p21−/− mice demonstrate a more rapid and robust progression of hepatocytes through the cell cycle during liver regeneration [24–26]. In mice with enlarged livers due to manipulation of growth-control proteins, p21 is markedly induced and suppresses further proliferation, suggesting that it may play a role in hepatic size regulation [69, 91, 98]. Altered expression of p21 appears to regulate variations in liver regeneration during the circadian rhythm [99]. Increased expression of p21 in human or rodent liver may also impair hepatocyte proliferation in the setting of fatty liver, acute liver failure, viral hepatitis, and cirrhosis [76, 78, 100–106]. Interestingly, p21

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can also regulate hepatocyte cell death [107]. These data suggest that p21 is an important determinant of hepatocyte proliferation, and that high-level expression of this protein in liver diseases prevents normal replication of these cells. An evolving area in the field of liver regeneration and repair is the concept that liver stem cells are an important source of hepatocyte replacement in certain types of liver injury (see Chapter 38). Although cell cycle protein expression has been documented in these cells, their regulation and actions have not been extensively characterized [108, 109]. Furthermore, the role of cell cycle control proteins in normal and aberrant proliferation of stellate cells, biliary epithelial cells, and other non-parenchymal liver cells is an area of potential interest [110, 111].

ABNORMAL CELL CYCLE CONTROL AND LIVER CANCER Similarly to other forms of cancer, deregulation of cell cycle proteins is commonly observed in HCC and other liver tumors [45, 112–115]. In general, HCCs display a heterogeneous pattern of molecular aberrations when the expression patterns of individual proteins are examined, although analyses of gene networks have provided a framework for understanding molecular phenotypes of this malignancy (see Chapter 61). The heterogeneity of reported cell cycle protein abnormalities in human HCC may be related to different patient populations and pathogeneses, the techniques used, and the apparently random aneuploidy that is usually observed in this malignancy. As is the case with other cancers, the large majority of HCCs have demonstrated abnormalities of the cyclin D1–Rb–E2F pathway. In addition, deregulation of upstream signaling pathways that control the cell cycle machinery is a nearly universal finding. Finally, the frequent loss of tumor suppressor proteins p53 and p16 compromise cell cycle checkpoints that normally prevent replication of cells with potentially malignant characteristics. Hepatocyte replication has been associated with improved survival in patients with liver diseases [116–118], but chronic high-level proliferation of these cells may also predispose to the development of HCC [119–121]. Although a causative role has not been clearly established, human HCCs frequently show overexpression of cyclins (including D1, E, and A) and diminished expression of the p16 and p27 CKI proteins [2, 45, 112–115]. Increased expression of cyclin D1 has been shown to occur via gene amplification in a minority of these tumors, but more often its expression is induced as a result of constitutive activation of upstream signaling molecules such as β-catenin and Ras. Diminished expression of p16 can result from deletion of this gene or, more commonly, by promoter methylation. In addition, loss or mutation of p53 is a common occurrence in HCC, which can result in defective cell cycle checkpoint function.

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Transgenic models have demonstrated that deregulation of the cyclin D–Rb–E2F pathway predisposes to HCC. For example, mice with constitutive hepatocyte-directed expression of cyclin D1 develop spontaneous HCC [43]. Even short-term expression of cyclin D1 leads to centrosome abnormalities and aneuploidy in hepatocytes, suggesting that this protein contributes to chromosomal instability that may further promote the progression to malignancy [122, 123]. Liver-specific deletion of Rb leads to increased chromosome instability and tumor formation in carcinogen-treated mice [124]. A tumor-suppressor role for p21 has recently been demonstrated in a model of liver injury due to hereditary tyrosinemia, suggesting that induction of p21 in the setting of liver injury may help to prevent replication of hepatocytes with DNA damage that are predisposed to malignant transformation [46]. These studies suggest that in experimental systems, aberrant expression of cell cycle proteins can trigger the development of HCC.

POTENTIAL RELEVANCE TO CLINICAL HEPATOLOGY The field of cell cycle control has matured substantially since the description of yeast cyclin and cdk proteins in the 1980s [16]. However, these scientific breakthroughs have not yet translated into diagnostic and therapeutic strategies with a major impact on patient care. Examples of clinical applications include the use of cell cycle protein expression profiles to classify malignancies and predict the response to treatment [125, 126]. In addition, small-molecular inhibitors of cdks are being studied as potential chemotherapeutic agents [4]. Although the impact has been modest thus far, it is likely that further insight into the cell cycle machinery will lead to the development of new therapies, and this field represents a potentially fruitful area for translational research. Perhaps the most important goal of liver regeneration research is to discover therapies that promote regeneration and restore hepatic function in patients with severe acute or chronic liver disease. Hypothetically, treatments that induce controlled growth and proliferation of normal hepatocytes (or liver stem cells) could provide substantial benefit to patients with severely diminished hepatocellular function, although this would not likely relieve symptoms related to other complications of liver disease such as portal hypertension. Despite a rapidly expanding understanding of the pathways that regulate hepatocyte proliferation and liver regeneration, we do not yet have regenerative therapies that can be applied to our patients with advanced liver disease. Hopefully, a more detailed knowledge of the key stimulatory and inhibitory proteins involved in liver regeneration may lead to rationally designed therapies that promote adaptive cell proliferation. Proof-of-principle studies suggest that transient transfection of one to two cell cycle proteins is sufficient to

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trigger abundant hepatocyte replication and substantial liver growth in mice, even under conditions that normally inhibit regeneration [38, 69, 71, 98]. However, such crude approaches are unlikely to be clinically useful for a number of reasons [127, 128], not least of which is the possibility that even transient overexpression of cell cycle proteins may promote chromosomal abnormalities [122, 129]. More likely, successful therapies will be tailored for individual patients, and will involve targeting the physiologically relevant cell cycle regulators that are disrupted in distinct liver diseases. A number of studies suggest that interventions directed at the cell cycle machinery could be used to prevent or treat cancer. For example, disruption of cyclin D1 expression prevents cancer in some animal models. Furthermore, targeted expression of CKI proteins, or the use of chemical cdk inhibitors, holds promise as potential cancer treatment agents [3, 4, 28]. In the liver, conditional deletion of the FoxM1B transcription factor prevents HCC in a mouse model of chemical carcinogenesis [130]. Since FoxM1B promotes hepatocyte proliferation in part through induction of cyclin–cdk complexes [131, 132], novel therapies to disrupt Foxm1b function may provide a means to inhibit the cell cycle machinery and hepatocarcinogenesis [130]. Furthermore, other dietary and pharmacological measures could potentially provide protection against the development of HCC by regulating cell cycle proteins [71, 133–143]. Thus the cell cycle is an attractive target for future therapies in patients with liver diseases.

ACKNOWLEDGMENTS The authors wish to acknowledge colleagues whose relevant publications were omitted from this chapter because of space considerations. We thank Emily Albrecht for help with illustrations. The authors’ research is supported by NIH grants DK54921 (J.H.A.) and F32DK074320 (L.K.M.).

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is correlated with the quiescent cell cycle status of hepatocytes in vivo. Hepatology, 44, 164–73. Serfas, M.S., Goufman, E., Feuerman, M.H., Gartel, A.L. and Tyner, A.L. (1997) p53-independent induction of p21WAF1/CIP1 expression in pericentral hepatocytes following carbon tetrachloride intoxication. Cell Growth Differ, 8, 951–61. Wu, H., Wade, M., Krall, L., Grisham, J., Xiong, Y. and Van Dyke, T. (1996) Targeted in vivo expression of the cyclin-dependent kinase inhibitor p21 halts hepatocyte cell-cycle progression, postnatal liver development and regeneration. Genes Dev , 10, 245–60. Stepniak, E., Ricci, R., Eferl, R., Sumara, G., Sumara, I., Rath, M., Hui, L. et al. (2006) c-Jun/AP-1 controls liver regeneration by repressing p53/p21 and p38 MAPK activity. Genes Dev , 20, 2306–14. Nelsen, C.J., Rickheim, D.G., Timchenko, N.A., Stanley, M.W. and Albrecht, J.H. (2001) Transient expression of cyclin D1 is sufficient to promote hepatocyte replication and liver growth in vivo. Cancer Res, 61, 8564–68. Grechez-Cassiau, A., Rayet, B., Guillaumond, F., Teboul, M. and Delaunay, F. (2008) The circadian clock component BMAL1 is a critical regulator of p21WAF1/CIP1 expression and hepatocyte proliferation. J Biol Chem, 283, 4535–42. Hui, T.T., Mizuguchi, T., Sugiyama, N., Avital, I., Rozga, J. and Demetriou, A.A. (2002) Immediate early genes and p21 regulation in liver of rats with acute hepatic failure. Am J Surg, 183, 457–63. Crary, G.S. and Albrecht, J.H. (1998) Expression of cyclin-dependent kinase inhibitor p21 in human liver. Hepatology, 28, 738–43. Brunt, E.M., Walsh, S.N., Hayashi, P.H., LaBundy, J. and Di Bisceglie, A.M. (2007) Hepatocyte senescence in end-stage chronic liver disease: a study of cyclin-dependent kinase inhibitor p21 in liver biopsies as a marker for progression to hepatocellular carcinoma. Liver Int , 27, 662. Lunz, J.G. III, Tsuji, H., Nozaki, I., Murase, N. and Demetris, A.J. (2005) An inhibitor of cyclin-dependent kinase, stress-induced p21Waf-1/Cip-1, mediates hepatocyte mito-inhibition during the evolution of cirrhosis. Hepatology, 41, 1262–71. Marshall, A., Rushbrook, S., Davies, S.E., Morris, L.S., Scott, I.S., Vowler, S.L., Coleman, N. et al. (2005) Relation between hepatocyte G1 arrest, impaired hepatic regeneration, and fibrosis in chronic hepatitis C virus infection. Gastroenterology, 128, 33–42. Morita, T., Togo, S., Kubota, T., Kamimukai, N., Nishizuka, I., Kobayashi, T., Ichikawa, Y. et al. (2002) Mechanism of postoperative liver failure after excessive hepatectomy investigated using a cDNA microarray. J Hepatobiliary Pancreat Surg, 9, 352–59. Wagayama, H., Shiraki, K., Yamanaka, T., Sugimoto, K., Ito, T., Fujikawa, K., Takase, K. et al. (2001) p21WAF1/CTP1 expression and hepatitis virus type. Dig Dis Sci , 46, 2074–79.

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121. Borzio, M., Trere, D., Borzio, F., Ferrari, A.R., Bruno, S., Roncalli, M., Colloredo, G. et al. (1998) Hepatocyte proliferation rate is a powerful parameter for predicting hepatocellular carcinoma development in liver cirrhosis. Mol Pathol , 51, 96–101. 122. Nelsen, C.J., Kuriyama, R., Hirsch, B., Negron, V.C., Lingle, W.L., Goggin, M.M., Stanley, M.W. et al. (2005) Short term cyclin D1 overexpression induces centrosome amplification, mitotic spindle abnormalities, and aneuploidy. J Biol Chem, 280, 768–76. 123. Duesberg, P., Li, R., Fabarius, A. and Hehlmann, R. (2005) The chromosomal basis of cancer. Cell Oncol , 27, 293–318. 124. Mayhew, C.N., Carter, S.L., Fox, S.R., Sexton, C.R., Reed, C.A., Srinivasan, S.V., Liu, X. et al. (2007) RB loss abrogates cell cycle control and genome integrity to promote liver tumorigenesis. Gastroenterology, 133, 976–84. 125. Liang, J., Zubovitz, J., Petrocelli, T., Kotchetkov, R., Connor, M.K., Han, K., Lee, J.H. et al. (2002) PKB/Akt phosphorylates p27, impairs nuclear import of p27 and opposes p27-mediated G1 arrest. Nat Med , 8, 1153–60. 126. Keyomarsi, K., Tucker, S.L., Buchholz, T.A., Callister, M., Ding, Y., Hortobagyi, G.N., Bedrosian, I. et al. (2002) Cyclin E and survival in patients with breast cancer. N Engl J Med , 347, 1566–75. 127. Raper, S.E., Chirmule, N., Lee, F.S., Wivel, N.A., Bagg, A., Gao, G.P., Wilson, J.M. et al. (2003) Fatal systemic inflammatory response syndrome in a ornithine transcarbamylase deficient patient following adenoviral gene transfer. Mol Genet Metab, 80, 148–58. 128. Ilan, Y., Saito, H., Thummala, N.R. and Chowdhury, N.R. (1999) Adenovirus-mediated gene therapy of liver diseases. Semin Liver Dis, 19, 49–59. 129. Spruck, C.H., Won, K.A. and Reed, S.I. (1999) Deregulated cyclin E induces chromosome instability. Nature, 401, 297–300. 130. Kalinichenko, V.V., Major, M.L., Wang, X., Petrovic, V., Kuechle, J., Yoder, H.M., Dennewitz, M.B. et al. (2004) Foxm1b transcription factor is essential for development of hepatocellular carcinomas and is negatively regulated by the p19ARF tumor suppressor. Genes Dev , 18, 830–50. 131. Kalinichenko, V.V., Gusarova, G.A., Tan, Y., Wang, I.C., Major, M.L., Wang, X., Yoder, H.M. et al. (2003) Ubiquitous expression of the forkhead box M1B transgene accelerates proliferation of distinct pulmonary cell types following lung injury. J Biol Chem, 278, 37888–94. 132. Wang, X., Kiyokawa, H., Dennewitz, M.B. and Costa, R.H. (2002) The Forkhead Box m1b transcription factor is essential for hepatocyte DNA replication and mitosis during mouse liver regeneration. Proc Natl Acad Sci U S A, 99, 16881–86. 133. Campbell, T.C. and Junshi, C. (1994) Diet and chronic degenerative diseases: perspectives from China. Am J Clin Nutr, 59, 1153S–61S. 134. Hu, J.F., Cheng, Z., Chisari, F.V., Vu, T.H., Hoffman, A.R. and Campbell, T.C. (1997) Repression of hepatitis

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Prospective study of chemoprevention of hepatocellular carcinoma with Sho-saiko-to (TJ-9). Cancer, 76, 743–49. Suzui, M., Masuda, M., Lim, J.T., Albanese, C., Pestell, R.G. and Weinstein, I.B. (2002) Growth inhibition of human hepatoma cells by acyclic retinoid is associated with induction of p21(CIP1) and inhibition of expression of cyclin D1. Cancer Res, 62, 3997–4006. Okuno, M., Kojima, S., Matsushima-Nishiwaki, R., Tsurumi, H., Muto, Y., Friedman, S.L. and Moriwaki, H. (2004) Retinoids in cancer chemoprevention. Curr Cancer Drug Targets, 4, 285–98. Koga, H. (2003) Hepatocellular carcinoma: is there a potential for chemoprevention using cyclooxygenase-2 inhibitors? Cancer, 98, 661–67. Murata, H., Tsuji, S., Tsujii, M., Sakaguchi, Y., Fu, H.Y., Kawano, S. and Hori, M. (2004) Promoter hypermethylation silences cyclooxygenase-2 (Cox-2) and regulates growth of human hepatocellular carcinoma cells. Lab Invest , 84, 1050–59.

63

miRNAs and Liver Biology Charles E. Rogler and Leslie E. Rogler Marion Bessin Liver Research Center, Division of Hepatology, Department of Medicine, Albert Einstein College of Medicine, Bronx, New York, NY, USA

INTRODUCTION The purpose of this chapter is twofold: first, to provide a historical perspective and broad overview of a newly described class of small non-coding RNA molecules, designated microRNAs (miRNAs), and second , to describe early work that has begun to understand which miRNAs are expressed in the liver and how some of them may function in liver biology. It must be said that we are in the early days of discovery for miRNA research in liver biology. Therefore, a major function of this chapter will also be to outline experimental approaches being used in the field, which will be of interest to those entering this field of research.

BRIEF HISTORICAL PERSPECTIVE For many years since their first discovery, RNAs were viewed as having two major functions that centered on mediating protein synthesis. These included mRNAs (messenger RNAs) that were the protein-coding genes that act as templates for protein synthesis, and rRNAs (ribosomal RNAs) and tRNAs (transfer RNAs), that had structural, catalytic, and information-decoding roles in protein synthesis. Small RNAs were overlooked in biochemical analyses for many years because of their size, ∼20–30 nucleotides, and because they are poor mutagenesis targets. miRNAs and short interfering RNAs (siRNAs) comprise two classes of small non-coding RNAs whose The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

primary functions are to silence gene expression at the post-transcriptional level. Our current understanding of RNA-induced silencing began with the Nobel Prize-winning work of Andrew Fire and Craig Mello and colleagues [1]. Working with the worm Caenorhabditis elegans, they were the first to show that double-stranded RNAs could act catalytically and efficiently to silence gene expression, in a phenomenon they called RNA interference (RNAi). Their studies were complemented by elegant work from the laboratories of Victor Ambros [2, 3] and Gary Ruvkun [4, 5] that characterized gene silencing in C. elegans using genetic approaches and identified the genes encoding short non-coding RNAs that mediated gene silencing. These were the first members of the new class of genetic elements now known as microRNA genes (Figure 63.1). For these studies, Ambros and Ruvkun, along with David Baulcombe, who described RNA interference in plants [6–8], have received the prestigious 2008 Albert Lasker Basic Medical Research Award. In 2000, the Ruvkun group in collaboration with H.R. Horvitz and A. Rougvie described a second miRNA gene called Let-7 [5]. Unlike the earlier C. elegans genes, Let-7 was widely conserved across animal species [9]. This discovery stimulated a frantic search for new miRNAs that led, within 2 years, to simultaneous publications by Lee and Ambros [3] and the groups of David Bartel [10] and Thomas Tuschl [11] reporting the discovery of hundreds of miRNAs that were conserved from worms to humans. There are now approximately 677 mammalian genes that produce miRNAs, and several websites devoted

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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siRNA pathway

miRNA pathway

piRNA pathway

miRNA-encoding genes

Viral infection Transcription of sense and antisense strands

miRNA precursor dsRNA

Nucleus Cytoplasm

Dicer

Long, single-stranded RNA

Drosha

Dicer

miRNA duplex

piRNAs

siRNAs

Repression of mRNA translation

miRNP

RISC

piRNP Piwi

Ago

Ago mRNA

AAAA AAAA Germ-cell development

mRNA halves

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Figure 63.1 The structure and biogenesis pathways of miRNAs, siRNAs, and piRNAs. Left column: small interfering RNAs (siRNAs) are processed from double-stranded RNAs (dsRNAs) that form by base pairing of complementary RNAs. An enzyme called Dicer cleaves dsRNA into shorter double-stranded siRNAs that are roughly 20 base pairs long. One siRNA strand then assembles into an effector complex known as an RNA-induced silencing complex (RISC). This complex uses the siRNA guide to identify mRNAs with a sequence perfectly complementary to the siRNA. RISC then cleaves the mRNA in the middle of the mRNA–siRNA duplex, and the resulting mRNA halves are degraded by other cellular enzymes. Middle column: microRNAs (miRNAs) are processed from specific genome-encoded precursors, which fold into intramolecular hairpins containing imperfectly base-paired segments. The processing generally occurs in two steps, and is catalyzed by the enzymes Drosha (in the nucleus) and Dicer (in the cytoplasm). One strand of the resulting miRNA duplex, resembling an siRNA, then incorporates into a RISC-like miRNA–ribonucleoprotein (miRNP) complex. The main components of RISC and miRNPs are proteins of the argonaute (Ago) family. Depending on the level of complementarity, miRNAs induce mRNA degradation or repress their translation. Right column: Piwi-associated RNAs (piRNAs) are generated from long, single-stranded precursors in a process independent of Drosha and Dicer. These small RNAs associate with a subfamily of argonaute proteins called Piwi proteins. Tens of thousands of piRNAs have been identified, although they are far from understood. It is known, however, that, together with their Piwi partners, they are essential for the development of germ cells. Reproduced from Großhans & Filipowicz, Nature 451: 414–416, with permission from Nature Publishing Group

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to cataloging them [12]. These studies were foundational for the new field of miRNA biology.

CLASSES OF SMALL NON-CODING RNAs Three major classes of small non-coding RNA have been described in mammalian cells: miRNAs, siRNAs, and Piwi-interacting RNAs (piRNAs) [13–17]. The use of deep sequencing technologies is leading to the discovery of additional classes of small non-coding RNAs and more members of the existing families of genes encoding them. A summary of differences in structure and biogenesis of miRNAs, siRNAs, and piRNAs is presented in Figure 63.1, and a brief description of each class is presented below.

miRNAs miRNAs are the most abundant class of endogenous small non-coding RNA, with individual miRNAs generally present in thousands of copies per cell, up to 50,000 copies per cell, in the case of miR-122 in hepatocytes [18]. Fully processed miRNAs are single-stranded RNAs of 21–24 nucleotides that are present in the RNA-induced silencing complex (RISC), located in the cytoplasm of eukaryotic cells [17] (Figure 63.1). The primary mode of gene silencing mediated by RISC is through repression of translation. It is predicted that miRNAs may eventually be shown to regulate, to some extent, at least 20–30% of all the expressed genes in mammals [17, 19]. Evidence so far is that miRNAs are generally not on/off switches, but “rheostats” that add fine tuning to gene expression [20]. The reach of biological actions of miRNAs is continuously expanding. Their initial presence was discovered as a result of their role in controlling development of C. elegans [4, 21, 22], and major roles in embryonic development from zebrafish through mouse have now been shown [23–25]. miRNA gene expression is developmentally regulated with specific miRNA expression profiles for different cell types [18, 26–28]. Individual miRNAs have been shown to control cellular differentiation [29–31], and they play multiple roles in carcinogenesis in a wide range of cancers [32–38]. Roles for miRNAs in regulating intermediary metabolism, and also roles in disease, are rapidly being discovered [39–41]. One of the goals of this chapter is to review our early understanding of the functions of some of the major miRNAs expressed in the liver.

siRNAs Endogenous siRNAs are 22–24 nucleotides in length, but they are generally rare in mammalian cells. They

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are formed by random cleavage of long double-stranded RNA molecules transcribed from transposable elements, repetitive genes, transgenes, and viruses and other genetic elements [13]. Their precursor RNAs do not contain specific hairpin structures, as do those of miRNAs. In contrast to miRNAs, siRNAs silence genes by cleavage of the target RNA at sites that have exact sequence complementarity with the siRNA (Figure 63.1). siRNAs are believed to have primary roles in the silencing of transposable elements, repetitive genes, transgenes, and some viruses. Functions of siRNAs are less understood than miRNAs and will not be discussed further in this chapter.

Synthetic siRNAs These RNAs are artificially synthesized to be 100% homologous to selected cellular mRNAs. When they are transfected into cells, they utilize the natural mechanisms of miRNAs for gene silencing. They are designed to silence specific genes and have become a major tool for gene silencing in molecular biology and genetics [42] and will not be considered further in this chapter.

piRNAs (Piwi-associated RNAs) These are small RNAs between 25 and 30 nucleotides in length that are generated from long, single-stranded precursors that generally originate from repeated DNA sequences [14, 15]. They appear to be generated by a unique “ping-pong” mechanism and are associated with transposon silencing [14]. They are expressed mainly in germline cells and are essentially absent in the liver. Their functions are poorly understood and, since they are not expressed in the liver, they will not be discussed further in this chapter.

miRNA GENETICS AND BIOLOGY Genomics The majority of miRNAs are transcribed from genomic loci that are distinct from previously annotated genes. Nearly all the miRNAs cloned from human and mouse are highly conserved [11, 17, 43]. The structure of miRNA genes suggests that they are transcribed primarily as independent transcription units by Pol II, and the primary transcripts, designated pri-miRNAs, are capped and polyadenylated. Many of these genes are located in genomic locations that are associated with regulatory genes, such as in Hox clusters [44, 45] and tumor suppressor gene locations [46, 47].

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Although most miRNA genes encode only a single miRNA, a substantial number of miRNAs occur in clusters which can include up to six miRNAs (Plate 63.1). The miRNAs within genomic clusters are sometimes closely related to each other in sequence [26]. Some conserved families of miRNAs, such as the let-7 gene family, have expanded from a few members in C. elegans to over 10 in humans [9, 48]. A sizeable minority of miRNAs are located in the introns of coding genes (Plate 63.1) [49]. In these cases, the intronic RNAs generated during splicing become substrates for miRNA biogenesis [50]. The synthesis of these miRNAs is therefore linked to the parent protein coding gene. This raises the possibility that the miRNA and protein could act cooperatively. The major liver miRNA is miR-122, which is a single miRNA gene located on human chromosome 18 (mouse syntenic chromosome 18). miR-122 is processed from a pri-miRNA that is a non-coding RNA. The transcription factors regulating miR-122 pri-miRNA expression have not been described. However, miR-122 is turned on early during murine liver development at a time (prior to e12.5) when specification first occurs. miR-122 miRNA accumulates in hepatocytes up to 50,000 copies per cell, making it the most abundant miRNA in the liver, accounting for ∼70% of all the miRNAs [18, 26] . Hence it is possible that miR-122 may play a role in hepatocyte specification.

The primary miRNA binding protein in RISC is a member of the argonaute protein family. There are four argonaute proteins in mouse and human (Ago I–IV) [62]. Argonautes I and II are the major proteins associated with RISC in eukaryotic cells [63]. Argonaute proteins contain a PAZ domain, which binds to both single-stranded and duplex RNA [64, 65] and helicase and endonuclease (Ago II only) activities [66–68]. Additional proteins, including the RNA binding protein TRBP, are part of the multiprotein RISC complex [62]. During assembly of RISC, the passenger strand of the double-stranded Dicer processed miRNA is eliminated and eventually degraded and the miRNA guide strand is stably incorporated into RISC [60, 69, 70]. The mechanism for selective retention of the guide strand in RISC is poorly understood. One model is that the strand that enters RISC is generally the one whose 5′ end is less tightly paired and that a helicase activity unwinds the duplex and incorporates the guide strand selectively into RISC [26, 71, 72]. Once present as a single-stranded RNA, tightly bound to an argonaute protein in RISC, the miRNA identifies its target mRNA initially by homology with nucleotides on its 5′ end that have been designated as the “seed” sequence [55, 62].

BIOGENESIS OF miRNAs AND THE RNA-INDUCED SILENCING COMPLEX (RISC)

Cell-free in vitro systems have demonstrated that efficient miRNA guided translational inhibition is generally dependent on a 7-methylguanine (m7G) cap and a poly(A) tail on mRNAs [73–75]. However, this is not exclusively the case [74]. Ago proteins also contain a highly conserved motif that is similar to the m7G-cap binding domain of eIF4E and experimental evidence supports the hypothesis that Ago proteins compete with eIF4E for cap binding, causing inhibition of translational initiation [74, 76]. A protein that is also part of the RISC complex is TNRC6A (trinucleotide repeat containing protein), also known as GW182. This protein may function to recruit deadenylation enzymes to mRNAs that are in RISC and thus destabilize them [77]. Studies in Drosophila have shown an inhibition of 80S complex formation by specific miRNAs and the formation of “pseudo-polysomes.” These structures appear to be large protein–mRNA aggregates that are reversible structures [78]. These structures may interact with, or aid in the formation of, structures that are designated “P bodies.” P bodies are sites in the cytoplasm of eukaryotic cells that have been linked with mRNA degradation and the inhibition of translation. Hence their interaction with RISC is of interest and the subject of much ongoing study. Another model for miRNA-mediated translational repression is the ribosome drop-off model. In this mechanism, ribosomes are induced to drop off mRNAs when miRNAs bind to the 3′ UTR [79]. This is supported by

Pri-miRNAs are initially processed in the nucleus by an RNase III endonuclease named Drosha in combination with an RNA-binding protein, designated DGCR8 [51] (Figure 63.1). Drosha creates a staggered cut at the base of stem-loop structures in pri-miRNAs, and liberates 60–70 nt stem-loop intermediates [52, 53]. These are the miRNA precursor molecules, termed pre-miRNAs, [54] that are actively transported to the cytoplasm by RanGTP and exportin-5 [55]. Once in the cytoplasm, the pre-miRNA is processed by a second RNase III endonuclease called Dicer [56, 57]. Dicer recognizes the 5′ phosphate and the 3′ overhang at the base of the stem-loop and, at about two helical turns away from the base, it makes a second staggered cut of both RNA strands. This liberates the mature miRNA that has a 5′ phosphate and a 2 nt 3′ overhang on each end of the double-stranded RNA molecule [58–60]. The mature miRNAs contains a “guide” strand that is antisense to target sites on mRNAs, and a “passenger” strand that is unstable. The next step in biogenesis of active miRNAs involves their incorporation into a ribonucleoprotein complex known as the RNA-induced silencing complex [61].

MODELS FOR TRANSLATIONAL REPRESSION BY miRNAs

63: miRNAs AND LIVER BIOLOGY

studies showing that nascent polypeptides are rapidly degraded when the they come under miRNA regulation [80]. Furthermore, the length of the 3′ UTR and the position of the miRNA target site in the 3′ UTR affect the “context” of the miRNA site and its interplay with other regulatory RNA-binding proteins. This model is also not mutually exclusive of the previous models [73].

miRNA TARGET IDENTIFICATION An important goal of miRNA research has focused on determining how miRNAs recognize their targets [81, 82]. The importance of Watson–Crick base pairing between nucleotides in the 5′ end of the miRNA and the target mRNA was an initial observation that is now well established [17, 81, 83–85]. The hybridizing region between the mRNA target and the miRNA has been termed the “seed” sequence and it is a minimum of 6 bp which are nucleotides 2–7 from the 5′ end of the miRNA. Seed sequences are characterized up to 8 bp according to additional homology at position 8 of the target or the presence of an A at target position 1. The Target Scan 4.1 website (http://www.targetscan.org/), characterizes the various types of target seed sequences as 6mer, 7mer, 7mer-A1, 7mer-m8, or 8mer. The 8mer sites are the strongest and comprise a “seed” match (nucleotides 2–7) plus a match at position 8 and the A at position 1 [83]. However, the goal of identifying authentic miRNA target sites in mRNAs is much more complicated than just identifying different seed sequences. This is confirmed by experimental evidence that shows that “seed” matches alone are generally not sufficient to produce validated miRNA target sites [17, 86]. Therefore, determining the “context” of a “seed” sequence, in the 3′ UTR or other segment of a mRNA as become an important component of bioinformatics algorithms that predict miRNA target sites. A study by Grimson et al. [86] has identified five characteristics of miRNA target sites that have been incorporated into the calculation of a “context” score (Table 63.1). These are as follows: (i) closely spaced miRNA sites often act synergistically; (ii) Watson–Crick pairing at nucleotides 12–17 from the 5′ end of the miRNA, in addition to the seed match, enhances miRNA targeting; (iii) effective miRNA target sites preferentially reside within a locally AU-rich context; (iv) effective miRNA sites preferentially reside in the 3′ UTR but not too close to the stop codon; and (v) effective miRNA sites preferentially reside near both ends of the 3′ UTR. According to the criteria above, miRNA target sites are assigned a context score of up to 100, with 100 being the strongest predicted site. The context scores of bioinformatically predicted miRNA target sites are listed on the Target Scan 4.1 website. These computational tools have greatly aided investigators in picking miRNA sites

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Table 63.1 Parameters used in determining the “context” score of computationally identified miRNA target sites [86] 1. Closely spaced sites often act synergistically 2. Additional Watson–Crick pairing at nucleotides 12–17 enhances miRNA targeting 3. Effective sites preferentially reside within a locally AU-rich context 4. Effective sites preferentially reside in the 3′ UTR but not too close to the stop codon 5. Effective sites preferentially reside near both ends of the 3′ UTR

for study. However, as is demonstrated in the next section, direct experimental approaches to target identification continue to be needed for target validation.

PROETOMICS APPROACH TO TARGET IDENTIFICATION Since miRNAs are established as post-transcriptional regulators of gene expression, the most relevant endpoint readout of their regulatory effects is on protein output. Two reports have adapted a novel quantitative mass spectrometry-based approach using SILAC (stable isotope labeling with amino acids in cell culture) to investigate the effects of miRNAs on protein levels and identify miRNA targets [20, 87] (Table 63.2). These studies have validated the bioinformatics-based “context” calculation approach for target identification. The studies revealed that hundreds of proteins were directly repressed by individual miRNAs and the 3′ UTRs of these proteins were highly enriched for bioinformatically predicted target sites for those miRNAs. This strong evidence has also to be balanced with the other finding that the bioinformatics approaches still over estimate the number of miRNA target sites and experimental validation remains a necessary component of miRNA target identification. Another important finding of the SILAC approach is that miRNAs have mainly a fine “tuning” effect on protein synthesis. The terminology “tune” was used because the Table 63.2 Principles of miRNA regulation determined from proteomic approaches [20, 87] 1. SILAC identified thousands of proteins that are regulated either directly or indirectly by miRNAs 2. MiRNA regulation is primarily as a “rheostat” to adjust protein output. 3. Proteins down-regulated more than twofold are 10% or less of targets 3. One miRNA can repress the production of hundreds of proteins 4. Targeting is primarily through seed-matched sites with a favorable “context” score in the 3′ UTR 5. mRNA destabilization is greatest for highly repressed targets

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THE LIVER: miR-122, THE MAJOR LIVER miRNA

average miRNA effect on protein synthesis was found to be modest, in the 25–50% reduction range. Only rarely was a protein down-regulated over fourfold. In any event, the dramatic demonstration that single miRNAs can affect hundreds of proteins simultaneously is a major advance that adds an important new level to our appreciation of molecular mechanisms of gene expression.

miRNAs AND LIVER BIOLOGY Requirement of miRNAs for normal liver functions One approach to investigate the impact of miRNAs on cellular functions has been to produce mice in which Dicer is universally or conditionally knocked out. Knocking out Dicer causes the loss of all miRNAs and is lethal for early embryonic development. However, conditional Dicer knockouts in specific organs at late stages of embryonic development are generally not lethal. This latter approach has revealed that miRNAs are required for differentiation of several cell types [88]. One study has explored the effects of knocking down Dicer on neonatal functions of the liver. The study generated double transgenic mice containing a floxed Dicer gene and a Cre recombinase under control of a hybrid albumin–α-fetoprotein enhancer–promoter element. Dicer levels were reduced approximately 90% at birth and the level of key liver miRNAs, such as miR-122, was also reduced by approximately 90% [89]. Interestingly, serum levels of albumin, bilirubin, and cholesterol were essentially normal in the mice up to 100 days old. This demonstrated that many metabolic functions of the liver were maintained even with a large knockdown of all miRNAs. However, levels of miR-122, the major miRNA in the liver, have been estimated as up to 50 000 copies per hepatocyte in normal liver [18]. Therefore, the remaining 10% of miR-122 may be sufficient to allow basic metabolic functions of the liver to continue. In contrast to the minimal metabolic effects, the Dicer knockdown mice develop major pathological effects in the liver as the mice age. Newborn mice and mice up to 28 days old have normal liver histology that worsens quickly as the mice age. The liver/body weight ratio increases by 100 days, and this is the result of a very large (20–25-fold) increase in hepatocyte proliferation that is not matched by as large an increase in hepatocyte apoptosis (10–15-fold). The loss of miRNAs also causes an imbalance in cellular differentiation in the livers. There is a dramatic proliferation of CK 19-positive bile ductular cells adjacent to the portal tracts and the 60-fold increase in α-fetoprotein (a stem/progenitor marker) [89]. Furthermore, the fetal liver growth factor insulin-like growth factor 2, which promotes hepatocarcinogenesis in livers with oval cell proliferation [90], is increased 60-fold. Taken together,

these data strongly suggest that liver stem cell proliferation is actively occurring in the Dicer knockdown model. Overall, the model reveals a critical need for miRNAs to maintain normal liver functions in the mouse during its lifetime. The pathology that develops resembles auto-immune hepatitis since inflammation also occurs in the livers. As in other miRNA-directed genetic changes, the alterations in gene expression measured by microarray analysis are nearly all less than twofold. These data support an important role of miRNAs as “rheostats” that fine tune gene expression in the liver [89].

PROFILING miRNAs IN THE LIVER The original method for discovery and characterization of miRNAs was based on cloning and sequencing small RNAs that were isolated from mammalian tissues [11]. This experimental approach still remains the most specific approach to miRNA profiling [91]. It has the great advantage of being capable of distinguishing between miRNA family members and of discovering new miRNAs. Therefore, in this section we present profiles of miRNAs that were obtained using miRNA cloning/sequencing protocols of liver. Tables 63.3 and 63.4 present the 25 most frequently cloned miRNAs from human liver [91]. A complete accounting of all the miRNAs cloned from human liver can be obtained from [91] (supplementary data). It must be emphasized that experimental evidence on the functions of miRNAs in the liver is very limited at present. Therefore, most of the functional references are inferred from the effects of the specific miRNA in tissues other than liver. To some extent, this seems to be justified for miRNAs that are widely expressed in many tissues, since proteomic studies have shown common miRNA-regulated genes in different cell culture systems [20, 87]. A full accounting of all of the reported functions of the miRNAs expressed commonly between the liver and other tissues is beyond the scope of this chapter. This chapter focuses on the major liver miRNA, miR-122, and a few other miRNAs for which functions in the normal liver and in liver tumors have begun to emerge.

miR-122, THE MAJOR LIVER miRNA A principle of miRNA biology is that genes that are highly expressed in a tissue generally do not contain target sites for miRNAs that are expressed in the tissue. Experimental support for this concept is strong [28]. One example is studies that have shown that overexpression of miRNAs that are specific for muscle or brain in HeLa cells have caused a more “muscle-like” or “brain-like”

63: miRNAs AND LIVER BIOLOGY

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Table 63.3 Putative functions and targets of miR-122 in the liver miR-122 effect Over-expression in hESC delayed general differentiation Over-expression in HCC decreases cell viability, increases capase-3, targets anti-apoptosis gene Bcl-w Reduce cholesterol in blood of primates Up-regulated in HCC IFNb reduces miR-122 expression Positive regulation of HCV replication Maximum level of 50 000 copies per cell before birth miR-122 down in HCCs from choline-deficient diet-fed rats Inhibits HCC cells kept under stress conditions Reduced plasma cholesterol, increased FA oxidation, decreased steatosis in mice upon knockdown in liver of mice

Reduction in plasma cholesterol, no change in FFA, bile acids, or triglycerides upon knockdown in liver of mice

Not significantly changed in primary HCC

gene expression profile in the HeLa cells [19]. Genes that remained active in the transfected cells had fewer “seed sites” for the miRNAs that were transfected and were genes that are normally highly expressed in the specific tissue type. In contrast, genes that were repressed by the treatments were enriched for target sites of the transfected miRNAs and were generally low in each tissue type [161]. miR-122 expression is detected in the embryonic mouse liver at e12.5 [18]. This is an early stage of liver development in which hepatoblasts are present along with more differentiated hepatocytes. Bile duct formation has not occurred at this stage. According to the above hypothesis, it is predicted that the early and high-level expression of miR-122 in the fetal liver may serve to establish a “hepatocyte-like” gene expression profile in hepatoblasts. Many hepatocyte-specific genes do not contain miR-122 sites which allow them to escape miR-122 regulation (http://www.targetscan.org/). For example, both the human albumin and α-fetoprotein genes have short 3′ UTRs, and have no conserved or non-conserved target sites for miR-122, -126, -16, -22 or the Let-7 family of miRNAs which comprise over 90% of the liver miRNAs. However, experimental evidence for miR-122 establishment of the hepatocyte expression profile is currently lacking. In contrast, extensive evidence points to the role of specific growth factors and transcription factors in the establishment of hepatocyte-specific gene expression profiles [162]. The effect of miR-122 on the liver specific transcription factors is also currently unknown. A relevant

Targets (direct or indirect)

Reference

No targets Bcl-w

[92] [93]

— — — CAT-1, HCV viral RNA CAT-1 — CAT-1 GLYS1 SLC7A1 ALDOA CCNG1 P4HA1 Aldo-A Ndrg3 Ipgap1 Hfe2 Tmed3 Lass6 Slc35a4 Tmem50b Gpx7 Cs —

[94] [95] [96] [97] [18] [98] [99] [39]

[40]

[100]

observation in this area is that all liver-derived cell lines, whether they are of stem cell origin or hepatocellular carcinoma (HCC) origin, have very low levels of miR-122 compared with liver [18, 91, 97, 100] . However, these cell lines maintain various degrees of differentiated liver functions. Overexpression of miR-122 in an HCC cell line decreased cell viability, increased caspase-3, and targeted the anti-apoptosis gene Bcl-w [163]. Also, miR-122 inhibits the growth of HCC cells subjected to stress conditions that could be equivalent to those experienced by the cell culture process [99, 164]. Therefore, high levels of miR-122 appear to be detrimental to cultured liver cells, whereas low levels appear to be sufficient to maintain liver-specific gene functions in culture. The Dicer knockdown experiment demonstrates that higher levels of miR-122 in primary liver are needed for the long-term maintenance and stability of the liver. Another approach to the problem of miR-122 function in the liver has been to knock down its level in intact mouse liver using “antagomirs” to eliminate miR-122 from the liver of adult mice. To develop a pharmacological approach for silencing miRNAs in vivo, Krutzfeldt et al. [40] designed chemically modified cholesterol-conjugated single-stranded RNA analogs complementary to miRNAs that have been termed “antagomirs”. In order to test the role of miR-122 in liver, an antagomir against miR-122 was synthesized and was injected intravenously into mice. Localization studies showed that the bulk of the antagomirs were taken up by the liver and that they were maintained at high

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THE LIVER: miR-122, THE MAJOR LIVER miRNA

Table 63.4 Putative functions and targets of microRNAs in liver

MicroRNA

Percentage of liver expression

Mir-122 Mir-126

72.4 6.0

mir-16

3.3

mir-22 mir-143

1.1 1.1

Let-7a

1.4

Mir-125b

1.1

Mir-99a Let-7b Let-7c Mir-451 Mir-194 Mir-181a

0.9 0.9 0.9 0.6 0.5 0.51

Mir-30d Mir-24

0.4 0.4

Mir-29a Mir-193 Mir-15b

0.4 0.4 0.4

Mir-23b

0.3

Mir-27b

0.3

Mir-26b Mir-30a Mir-15a Mir-26a

0.3 0.3 0.2 0.2

Function

Confirmed targets

Reference

Angiogenesis, angiogenic signaling, vascular integrity, invasiveness, cell growth suppression, inhibition of PI3K signaling Control of cell survival, invasion, proliferation, tumor suppressor

VCAM-1, Hox A9, IRS-1, P85beta, Crk

[101–105, 104, 106–108]

Wnt3a, BCL, cyclin D1, COX-2, c-Myb, cyclin D3, cyclin E1, CDK6, Ngn3, Nodal, IGSF4 BMP-7, PPARA

[109–120]

Ras, HMGA2, Myc caspase 3, integrin b3, PRDM1/blimp-1, TRIM71 Smoothened, ERBB2 and ERBB3, lin-28, TNFα, 3′ UTR of HIV,

[124–129]

Inflammation, osteoarthritis Adipogenesis, growth inhibition, tumor suppressor Tumor suppressor

Differentiation arrest, suppress cell growth, oncogene Tumor suppressor

[121] [122, 123]

[130–137]

Hmga2, cyclin D1 Lin-41 GATA2

[138, 139] [140] [141–143]

Tumor suppressor, cell fate in immune system, T cell receptor signaling, muscle differentiation — Inhibits erythropoiesis, methotrexate resistance Type 2 diabetes

HoxA-11

[134, 144–146]

— P16 (ink4a), ALK4, DHFR

[147–149]

Stem cell self-renewal, apoptosis, pancreas development Maintenance of the glomerular filtration barrier Sprout formation in endothelial cells, tumor suppressor, anti-apoptosis

Bcl-2, Ngn3

Erythroid maturation, tumor suppressor

—

levels in the liver for the length of the study, which was approximately 1 month. Assays for miR-122 revealed that it was completely eliminated from the liver for prolonged periods. Mechanistic studies have since shown that antagomirs interact with their target miRNAs in a cytosolic compartment distinct from P bodies and cause degradation of the target miRNAs while the antagomirs remain for a long period [165]. Antagomir-122 treatment caused a large change in liver gene expression, including 363 transcripts that were up-regulated at least 1.4-fold. These genes had a 2.6-fold increased frequency of miR-122 target sites in their 3′ UTRs. The genes in this group included members of gene families that are normally repressed in hepatocytes,

[150] [151, 115, 117] [152] Cyp1b1, ZBTB10/RINZF, RYBP/DEDAF

[153–155]

SOD2 Bcl2, Ccnd1, Wnt3a, c-Myb Myc, enhancer of Zeste homology2, Smad1

[156] [109, 157, 120] [158–160]

such as aldolase-A (aldo-A), N-Myc regulated gene 3 (Ndrg3) and a GTPase-activating protein (Iqgap1). Interestingly, the antagomir-122 injection experiment also revealed 305 transcripts that were down-regulated. The 3′ UTRs of these genes had a 2.7-fold decrease in miR-122 target sites, showing that they are not likely to be direct miR-122 targets. Such indirect effects could be due to the suppression of a transcriptional repressor or to chromatin remodeling. Most significantly, the top-ranking functional category of the repressed genes was “cholesterol biosynthesis.” Eleven genes involved in cholesterol biosynthesis were decreased (1.4–2.3-fold), including the rate-limiting enzyme of cholesterol biosynthesis

63: miRNAs AND LIVER BIOLOGY

3-hydroxy-3-methylglutaryl-CoA-reductase (Hmgcr), which had a 45% decrease in live enzyme activity. These gene changes led to a specific 40% reduction in plasma cholesterol that lasted for at least 2 weeks. At the same time, triglycerides, bile acids, glucose, ALT, and free fatty acids remained normal in the mice. A second study treated mice for 4 weeks with an miR-122 antisense oligo (ASO) that contains 2′ -O-methoxyethylphosphorothioate-modified nucleotides for stability [39]. This study confirmed a similar level of serum cholesterol reduction as in that by Krutzfeldt et al. [40], and in addition showed that triglycerides and glucose levels were reduced during a longer term treatment protocol. The longer term treatment caused a nearly twofold increase in fatty acid oxidation and a reduction in fatty acid synthesis in hepatocytes isolated from the mice and kept in short-term cell culture. In the same report, additional mice were kept on a high-fat diet for 19 weeks and then were treated for 5.5 weeks with the same miR-122 ASO. At the end of treatment, control mice had severe steatosis and there was a substantial reduction in steatosis and reductions in lipogenic gene expression in the miR-122 ASO-treated mice. Therefore, contrary to what might be expected, inhibition of miR-122 produced beneficial effects on plasma cholesterol, hepatic fatty acid metabolism, and improved liver steatosis in a high-fat fed mouse model [39]. A third study [94] tested the effect of systemic delivery of an unconjugated PBS formulated locked-nucleic acid (LNA)-modified oligonucleotide against miR-122 in non-human primates. Intravenous injections led to the inactivation of internal miR-122 by forming stable duplexes that could not function in RISC in the liver. These treatments also caused a dose-dependent lowering of plasma cholesterol without histopathological changes in the liver. Therefore, the special LNA–antiR-122 molecules were found to be very effective and were proposed for further therapeutic testing [94]. Together, these studies point out the importance of miR-122 in the maintenance of normal hepatic functions. They leave open the question of the role of miR-122 in the initial establishment of the hepatocyte-specific transcriptome. They also leave unanswered the question of why the liver has so much miR-122 when a reduction in miR-122 can improve health by lowering cholesterol and reducing fatty liver. These questions bring out the newness of this field to liver biology.

miR-122 ANC HEPATITIS C VIRUS (HCV), A CURIOUS RELATIONSHIP Hepatitis C virus (HCV) is a single-stranded RNA virus that infects the liver and establishes persistent infections. This virus has evolved a mechanism to take advantage

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of the major miRNA in the liver, miR-122, to promote its own replication. The HCV RNA genome contains a 5′ UTR, a long polyprotein open reading frame, and a 3′ UTR. The 5′ UTR has a very unique secondary structure and a bioinformatics search identified a perfect seed match for miR-122 near the 5′ end of the viral genome (Figure 63.2) [97]. The HCC cell line Huh7 has a low but detectable level of miR-122, and this cell line has been used to study the replication of HCV by transfection of a specially constructed HCV replicon that contains the 5′ UTR region. Transfection of this cell line with an miR-122, 2′ -OMe-modified antisense inhibitor, reduced endogenous levels of miR-122, and reduced replication of HCV. The reverse experiment, transfection of wild-type miR-122 mimics into the cells, caused an increase in HCV replication. Mutations were engineered into the miR-122 seed sequence in the 5′ UTR so that its interaction with endogenous miR-122 would be disrupted. Transfection of the mutant (designated p1) reduced HCV replication. A second mutant (p3-4) was constructed with a double mutation in the seed, and replication of this mutant was also reduced in Huh 7. The addition of a wild-type miR-122 mimic did not restore replication of the p3-4 mutant, as expected (Figure 63.2). Finally, an miR-122 double-stranded mimic with mutations in the seed that were complementary to the seed mutations in the HCV replicon (designated mimic p3-4) was transfected along with the HCV p3-4 mutant. This combination restored replication above that of wild-type HCV in the Huh 7 cells (Figure 63.3). These experiments established that direct binding occurs between miR-122 and the HCV replicon, and stand as an excellent example of how miRNA target validation is carried out. Further work using the mutant viruses suggested that miR-122 did not affect translation or RNA stability of HCV. Therefore, it was concluded that miR-122 was likely to facilitate replication of the viral RNA, and that miR-122 may represent a target for antiviral intervention.

MINOR miRNAs IN THE LIVER miR-126 is the second most abundant miRNA in liver. Studies of this miRNA have linked it to roles in angiogenesis, angiogenic signaling, vascular integrity, and invasiveness. Validated miR-126 target genes that may be related to these functions include VCAM-1, Hox A9, IRS-1, P85beta, and Crk. This information suggests that miR-126 may be expressed in the endothelial cells and sinusoidal lining cells in the liver (Table 63.4). Let-7a is the third highest expressed miRNA in the liver. However, the Let-7 family of miRNAs, including members Let-7a, b, c, d, f, and g, are all expressed in the liver and together account for nearly 4% of cloned liver miRNAs. Let-7 miRNAs have been linked to

Hepa1-6

30 miR-122 20

U6 snRNA 1

2

3

4

5

6

7

8

(a) 5′

3′

UGUUUGUGGUAACAGUGUGAGGU

5′

miR-122

(b)

HCV 5′VTR

5′

3′

GC C A A U U P5 CG UG AG CG GU A UA A HCV 5′VTR G A U G A AU P6 C G P7 GA CG AU CG UU GU U GC UG CA U UG C C U A A G GC GC II AU CG U GC UA P4 C G UG GCCA GACACUCGACGAUAGACGUCG ACGCCGCUGC GC 5 C G 20 38 CG CG CG In CG

IIa

NNeoC-5B cured

NNeoC-5B replicon

Huh7

HepG2

HeLa

Human liver

THE LIVER: MINOR miRNAs IN THE LIVER

Rat liver

1038

U U GA

ACUCUCCA

p6

ACACUGCA

p3

ACACAGCA

p3-4

ACACUCCG

p1

(c) HCV miR-122

wt -

wt wt

p1 -

p6 wt

p6 p3-4 p3-4 p6 wt p3-4 HCV

3′ G AUGGG GC C ACACUCCA UGUUU UG U UGUGAGGU G G miR-122

A A

5′

G C

Actin

A

(d)

(e)

Figure 63.2 miR-122 directly interacts with HCV RNA and promotes HCV replication. (a) Northern blot analysis of miR-122 expression in total RNA extracted from (1) rat, (2) human liver, (3) human HeLa, and (4) HepG2 cells, (5) na¨ıve Huh 7 cells, (6) replicon and (7) cured human Huh7 cells, and (8) mouse Hepa1-6 cells. Expression of U6 small nuclear (sn) RNA was used as a loading control. (b) Sequence of miR-122 with the seed sequences surrounded by a box. (c) Secondary structure of the 5′ non-coding regions of the HCV genotype 1a strain H77c, with predicted miR-122 binding sites indicated by a box. The seed matches are enclosed in boxes. SL, stem-loop; UTR, untranslated region. (d) Position of the mutations introduced into the 77c full-length RNA. The locations of single or double substitution mutations in the 5′ non-coding region seed match (p1, p3, p6, and p3-4) are shown. The mutated nucleotides are enclosed in boxes. (e) RNA was synthesized by in vitro transcription and introduced into Huh cells by electroporation, and HCV RNA levels were determined by Northern blotting 5 days later. Levels of actin mRNAs were determined as loading controls. Cells were transfected with synthetic duplexes corresponding to wild-type miR-122 (wt) or miR-122 with mutations in the seed complementary to the seed match mutations, with the opposite strand of the duplex based on the miR-122 precursor hairpin. The duplexes were introduced into cells 1 day before electroporation with wild-type H77c RNAs or mutant RNAs, and again at the first and third day post-electroporation. Total RNA was harvested 5 days post-electroporation, and HCV and actin RNA levels were determined by Northern blotting. Adapted from Jopling et al. (2005) Science 309: 1577–1581, with permission from AAAs

developmental and cell differentiation controls and Let-7 has been designated a tumor suppressor [124]. Validated Let-7 targets include Ras, HMGA2, Myc, caspase 3, integrin b3, PRDM1/bli, mp-1, and TRIM71. The targeting of both Ras and Myc protooncogenes by the Let-7 family of miRNAs is very significant because

these two genes were the first two protooncogenes to be shown to function synergistically to promote cell growth and malignant transformation [166]. In normal liver, hepatocytes are in G0 and both Ras and Myc are either absent or in present at very low levels. Let-7 family miRNAs are most likely important in maintaining the low

average relative expression level

average relative expression level

63: miRNAs AND LIVER BIOLOGY

6

mir-17-92

1.2

5

1.0

4

0.8

3

0.6

2

0.4

1

0.2

0

1039

miR-23b cluster

0.0 16.5

17.5

P1

AL

18

16.5

17.5

P1

AL

1.2 hematopoetic cluster

16

let 7 family

1.0

14 12

0.8

10

0.6

8

0.4

6 4

0.2

2

0.0

0 16.5

17.5

P1

AL

days

16.5

17.5

P1

AL

days

Figure 63.3 Coordinate expression of families and polycistronic clusters of miRNAs from e16.5 through adult mouse liver. Examples of miRNA microarray expression data are from livers of murine embryos at e16.5 and 17.5, 1-day-old pups (P1) and adult (AL). Relative expression levels at e16.5, 17.5, and P1 miRNAs are normalized to the adult liver signal set at 1.0. mir-17-92: average signal of the following miRNAs: miR-17, -18a, -19, -20, -92, -93, -106. All of these miRNAs are from the miR-17-92 polycistron or its paralogs illustrated in Plate 63.1. miR-23b cluster: average signal of miR-23b, -27b, and -24. Hematopoietic cluster: miR-181, -151. Let-7 family: average signal of Let-7a, b, c, d, e, and f

levels that allow hepatocytes to stay in G0. In this context, the down-regulation of multiple Let-7 miRNAs in HCC is easily understandable for the growth requirements of the tumors. Mir-16 is the fourth most abundant miRNA in the liver. miR-16 is ubiquitously expressed at high levels in mammalian cells. Therefore, its functions are most likely not specific to liver. Early studies focused on roles of miR-16 in cancer because it is down-regulated in many cancers, but incidentally, it is not significantly altered in HCC. Roles for miR-16 so far include control of cell survival, invasion, and proliferation. It has been characterized as a tumor suppressor miRNA, and some of its validated targets that fit these possible functions include Wnt3a, BCL-2, cyclins D1, D2, and E1, CDK 6 (cyclin-dependent kinase), Cox-2, c-Myb, Nfn3, Nodal, and IGSF4. miR-22 is the fifth highest cloned miRNA in liver. The current functions assigned to this miRNA are roles in control of inflammation and in osteoarthritis and two targets include BMP-7 and PPARA. It will be necessary to determine what cells within the liver express this miRNA. If Kupfer cells express it selectively, perhaps it will also have a role in inflammation in the liver.

miR-143 is the sixth most frequently cloned miRNA in the liver. This miRNA has been linked to adipogenesis and growth inhibition and validated targets have not been reported. This miRNA may therefore control intermediate metabolism in hepatocytes and also help maintain them in the G0 state. miR-143 was cloned less frequently from HCC, but due to variability from tumor to tumor it was not significantly down. HCCs vary tremendously in growth rate and lipid metabolism, both of which may be differentially regulated due to variability of miR-143. miR-125 is the sixth most frequently cloned miRNA in the liver and it is dramatically down in HCC cell lines [91]. One of its validated targets is tumor necrosis factor alpha (TNF-α). TNF-α is central to hepatocyte growth during liver regeneration and therefore the loss of miR-125 makes sense for tumors that are dependent upon TNF-α for growth. This picture becomes complicated by other functions, such as differentiation arrest and suppression of cell growth, which are also attributed to miR-125 in other experimental systems. miRs 181a, 451, and 24 comprise 1.5% of the liver clones. These miRNAs have been linked to B cell differentiation, erythroid maturation, and erythropoiesis, respectively. Validated targets include HoxA-11 (miR-181a),

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THE LIVER: miRNA PROFILING OF THE FETAL LIVER AND HEPATOBLAST DIFFERENTIATION IN VITRO

GATA2 (miR-451), and ALK4, P16, and DHFR (miR-24). Hence the functions of these miRNAs may be primarily to repress non-hepatocyte genes and help maintain a liver-specific transcriptome. Alternatively, these miRNA could be highly expressed in the few hematopoietic cells that may reside in the adult liver. miRs 23b, 27b, and 24 are referred to as the miR-23b cluster because they are all encoded by a single polycistronic gene and are coordinately expressed. Together these miRNAs comprise 1.1% of the liver clones. Mir 24 is implicated in erythropoiesis, miR-23b in maintenance of glomerular function, and miR-27b in formation of endothelial cells. A common theme for these miRNAs is therefore control of cellular differentiation. New work has linked these three miRNAs to down-regulation of TGFβ signaling and regulation of bile duct differentiation in fetal liver stem cells. These effects are regulated through targeting Smads that are key mediators of TGFβ signaling, and are described below.

miRNA PROFILING OF THE FETAL LIVER AND HEPATOBLAST DIFFERENTIATION IN VITRO miRNA profiling of the fetal murine liver from e16.5 to birth has identified families of miRNAs and clusters of unrelated miRNAs that are coordinately regulated in different patterns (Figure 63.3). The large set of Let-7 family of miRNAs that are transcribed form different loci in the genome are strongly up-regulated from e16.5 to adult liver (Figure 63.3). The Let-7 miRNAs are often increased in developmental stages associated with cell differentiation and a reduction in cell growth [169], both of which are major features of the liver from e16.5 to birth. Let-7 miRNAs have also been characterized as tumor suppressors because they target at least one protooncogene [169]. miRs-30a, b, and c are another family of miRNAs that are strongly up-regulated from murine fetal development day e12 through e18. These miRNAs are expressed in the developing bile ducts in the fetal liver and not in hepatocytes. Knocking them out in zebrafish embryos by antisense injection caused a blockage in bile duct development throughout the liver. Hence miR-30 miRNAs appear to be necessary for normal bile duct development in the liver [170]). Another set of miRNAs that are highly up-regulated in the fetal liver from e16.5 to birth is the miR-23b cluster that includes miR-23b, -27b, and -24 (Figure 63.4). These miRNAs are not related in sequence, but originate from the same pri-miRNA. In contrast to the miR-30 family, these miRNAs are highly expressed in hepatocytes and not in bile ducts. Functional studies described below implicate them in cell fate decisions in the fetal liver.

ROLES OF miRNAs DURING FETAL LIVER DEVELOPMENT The embryonic mouse liver from day 16.5 of gestation to birth is a fluid environment in which growth, apoptosis, and differentiation of committed liver progenitor cells are occurring [167]. Specifically, cholangiocytes in ductal plates undergo remodeling and form bile ducts while the hepatocyte compartment expands. During this window of development, hepatoblasts near the portal mesenchyme form the ductal plates and remodel to form the bile ducts, while hepatoblasts that do not come in contact with portal mesenchyme form hepatocytes. TGFβ is a major cytokine that functions in this cell fate decision [168].

(a)

(b)

(c)

Figure 63.4 Illustration of the phenotypic changes of fetal liver stem cells (HBC-3 cells) undergoing hepatocytic and bile ductular differentiation. (a) undifferentiated HBC-3 cells on STO feeder layer. Reproduced from Rogler, LE (1997) Am J Pathol 150: 591–602 (b) hepatocytic differentiation induced by DMSO. Reproduced from Rogler, LE (1997) Am J Pathol 150: 591–602 (c) bile duct differentiation on Matrigel. Reproduced from Ader et al., Mech. Dev. 123: 177–194. Copyright (2006), with permission from Elsevier

63: miRNAs AND LIVER BIOLOGY

In contrast, miRNAs present in the miR-17–92 polycistron and its paralogs are highly down-regulated in the liver from e16.5 to birth. High expression of miR-17–92 miRNAs has been linked to rapid cell proliferation and malignant transformation, and down-regulation of miR-17–92 is consistent with an overall reduction of liver growth rate from e16.3 to adult liver. Another cluster of miRNAs, which are associated with hematopoiesis (miRs 181b, 136, and 154) are transiently up-regulated from e18.5 to birth (Figure 63.3) [31, 170]. This is also consistent with the liver serving transiently as a hematopoietic organ during this same period. Interestingly, miR-122, which is the major miRNA in liver [91], did not vary in its expression from e16.5 to adult. This is consistent with previous work showing that miR-122 is induced during liver specification around e12.5 and is fully expressed in hepatoblasts at e16.5 [18]. A cluster of three miRNAs that are also up-regulated during fetal liver development is the miR-23b cluster. This cluster includes miR-23b, -27b, and -24 and it is located on mouse chromosome 13. HBC-3 cells are a non-immortalized, clonal, euploid hepatoblast cell line, isolated by culturing hepatoblasts from an e9.5 mouse embryo [171]. This stem/progenitor cell line is maintained in an undifferentiated state on STO feeder layers and can be induced toward the hepatocytic lineage by treatment with dimethyl sulfoxide (DMSO) or sodium butyrate, and toward the bile ductular lineage by plating on Matrigel (Figure 63.4) [171]. HBC-3 cells can be transplanted into recipient livers and form both hepatocytes and bile duct cells [91, 172]. Furthermore, gene expression profiling has shown that liver-specific gene expression patterns are induced by DMSO and bile duct genes are expressed in the cells on Matrigel [172, 173]. Genes associated with other tissue types (e.g. muscle, brain, lung, pancreas) that are expressed in undifferentiated HBC-3 cells are repressed during hepatocytic differentiation. HBC-3 cells were used to investigate the role of miR-23b cluster miRNAs in hepatocytic and bile ductular differentiation. The TGFβ/BMP signal transduction pathway is preferentially activated during bile duct morphogenesis in HBC-3 cells (Plate 63.2). In contrast, this same pathway is markedly down-regulated during hepatocytic differentiation of HBC-3 cells [91, 172, 173]. A bioinformatics approach was used to identify candidate miRNA mir-23b cluster targets. This approach identified candidate target sites for miR-23b and 27b in the Smad 3, 3′ UTR. The significance of this was further investigated because Smad 3 is a key signaling intermediate in the TGFβ pathway that controls bile duct differentiation. A pattern match approach revealed that expression of the gene containing miR-23b miRNAs went up in HBC-3 cells during hepatocytic differentiation at the same time as Smad 3 levels went down (Figure 63.5). In contrast, the miR-23b miRNA gene expression went down during

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bile duct differentiation when Smad 3 levels went up (Figure 63.5). This suggested that Smad 3 was a target for miR-23b cluster miRNAs. To test this, HBC-3 cells were treated with ASO inhibitors of miR-23b cluster miRNAs during hepatocytic differentiation. Normally, Smad 3 levels are low during hepatocytic differentiation. As predicted, ASO treatment with the miR-23b cluster miRNAs caused a large increase in Smad 3 levels in HBC-3 cells. Since TGFβ is required for bile duct differentiation, an experiment was conducted to determine whether miR-23b miRNAs can block bile duct differentiation by their targeting of Smads. The approach was to transfect HBC-3 cells undergoing bile duct differentiation (cultured on Matrigel) with mimics of miR-23b cluster miRNAs and determine the effect on bile duct differentiation. The transfection of mimics was shown to reduce the endogenous Smad 3 level and at the same time it dramatically blocked bile duct differentiation of the cells (Figure 63.5). Together, these experimental approaches support a role for miR-23b miRNAs in the control of HBC-3 cell differentiation. According to the model, high levels of the miRNAs during hepatocytic differentiation block TGFβ signaling by reducing Smads and allow hepatocytes to form. During bile duct differentiation, miR-23b cluster miRNAs are low and allow TGFβ signaling to proceed and bile ducts to form. Generally, TGFβ suppresses tumor progression through its regulation of cytostasis, differentiation, and apoptosis [174]. A role for TGFβ in suppressing epithelial growth is revealed in the context of tissue injury and oncogenic stress [174]. Therefore, activation of the miR-23b cluster miRNAs could promote malignant progression by knocking out the tumor suppressive effects of TGFβ. In this context, the miR-23b miRNAs would be classified as oncogenes. Indeed, a recent cloning survey of miRNA in many tumor cell types has revealed the presence of miR-23b cluster miRNAs at significant levels in a broad range of tumor cells [91]. Therefore, miRNA-based regulatory mechanisms will likely help define the contextual dependence and pleotropic effects of in a wide range of TGFβ effects in biological systems.

PROFILING OF miRNAs IN HEPATOCARCINOGENESIS Some of the earliest applications of miRNA profiling were in the field of cancer. These studies were pioneered by Carlo Croce and colleagues, who constructed an miRNA microarray that contained oligonucleotides for mature miRNAs and also oligonucleotides encoding pre-miRNAs [47]. These studies and those of many other groups [34, 175] have established several important principles of miRNA biology and cancer. Briefly, these studies have shown that miRNA levels are generally lower in

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THE LIVER: PROFILING OF miRNAs IN HEPATOCARCINOGENESIS 2 Matrigel

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Figure 63.5 Identification of candidate target genes by pattern matching. (a) Bile duct differentiation in Matrigel: Smad 3 RNA (black diamonds) is increased vs miR-23b (open squares) is decreased. (b) Hepatocytic differentiation in DMSO: Smad 3 RNA (black diamonds) is decreased vs miR-23b (open squares) is increased. Treatments are for 0–7 days in each graph

cancers than matching normal tissues. Second, miRNAs can target oncogenes, in which case they are functioning in a tumor-suppressive role, and they can target tumor suppressor genes and thus act in an oncogenic role [46]. Profiling has identified small sets of miRNAs that are upor down-regulated specifically in cancers [33, 176]. Some miRNA profiles have also been correlated with clinical outcomes [177]. Work on miRNAs involved with hepatocarcinogenesis has generally lagged behind research on other tumors with a higher prevalence in America. However, several miRNA profiling studies using oligonucleotide arrays have identified sets of miRNAs that are differentially expressed in HCCs compared with peri-tumor liver and normal liver [98, 178–182]. Some commonly regulated miRNAs were identified by these studies and further work will be required to resolve differences in the profiles.

As stated earlier, the authors prefer to utilize the data from miRNA cloning studies to compare miRNA profiles. The major miRNAs that were significantly increased or decreased in clone frequency between primary HCCs and normal human liver are shown in Figure 63.6 [100]. This list reveals major qualitative differences in miRNA frequencies. In HCCs, the seven most highly cloned miRNAs were not cloned from normal liver, suggesting that they are not present. The wide range of studies on roles of miRNAs in cancer has defined sets of interrelated pathways affected by miRNAs in cancer. Although these studies have generally not focused on HCC, many of the miRNAs and pathways that they identified are common with HCC. Whereas the target genes identified for specific miRNAs in different cancers would be the same, the functional downstream outcome may or may not be the same in HCCs. Some of the pathways affected by miRNAs are briefly reviewed below.

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(a)

(b)

Figure 63.6 MiRNAs that are differentially cloned from primary HCCs versus liver. (a) The 25 most increased in HCC vs Liver. (b) The 25 most decreased HCC vs Liver. 95% confidence intervals are displayed. * = P 0.05 and ** = P 0.001 Calculated using clone counts from the pooled HCC samples and liver. Reproduced from Connolly et al . (2008) Am. J. Pathol. 173: 856–864

miR-21 miR-21 is one of the most abundant miRNAs in primary HCCs, and its up-regulation is a hallmark of many different cancer types [91, 183] STAT-3 is a major mediator of IL-6 signaling and it participates in cellular transformation through suppression of apoptotic signaling [184].

Recent studies have demonstrated that STAT-3 is able to bind the promoter of the miR-21 primary transcript, leading to its activation [185]. Mir-21 is also up-regulated by TGFβ by a unique mechanism. The processing of the mir-21 pri-RNA is enhanced by binding of Smad 4 protein directly to the pri-RNA [186]. Interaction of Smad4 with the Dicer complex causes an increased level of processing of this miRNA in cells treated with TGFβ.

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THE LIVER: ROLE FOR miRNAs IN HEPATIC CYSTOGENESIS

The tumor suppressor PTEN is a direct target of miR-21. PTEN is a phosphatase that normally dephosphorylates FAK. The activated form of FAK is phosphorylated and activated FAK in HCCs correlates with more aggressive tumor behavior [187, 188]. Thus, miR-21 knockdown of PTEN leads to an increase in FAK phosphorylation and a more invasive phenotype of the HCC [180]. miR-21 knockdown also reduced HCC growth in soft agar and increased apoptosis of the cells [100] In contrast, enhanced miR-21 expression, by transfection with precursor miR-21, increased tumor cell proliferation, migration, and invasion [180].

miR-34 A major tumor suppressor that is inactivated in a large majority of tumors is p53. Surveys of p53 mutations in HCC have revealed a hotspot for p53 mutations in many HCCs. In cases where p53 is not mutated, other mutations in p53 response pathways often occur. Recent reports have shown that members of the miR-34 family of miRNAs are directly up-regulated by p53 [150, 197, 198]. However, in both the cloning study [100] and the other miRNA profiling studies of HCC [132, 179], miR-34 family miRNAs are not significantly up-regulated. This supports the overall understanding that p53 or its downstream pathways are inactivated in HCC.

miR-17–92 The list of up-regulated miRNAs in HCCs includes five out of six of the miRNAs expressed from the miR-17–92 polycistron that has been referred to as the first human “oncomir” for its early role in lymphoma (Plate 63.1) [189]. A survey of 68 primary HCCs by Northern blot demonstrated up-regulation of miR-17–92 miRNAs and miR-21in 100% of the tumors [100]. The miR-17–92 locus was also over-expressed in liver cirrhosis, suggesting an early role in hepatocarcinogenesis [100]. Two genetic mechanisms have been linked to the increased expression of the miR-17–92 polycistron in cancers. These include gene amplification and c-Myc activation [189, 190] . One study has mapped c-Myc binding sites in the 5′ promoter of the human miR-17–92 gene and shown that c-Myc binding activates transcription of the gene [191]. cMyc also activates the cell cycle regulatory gene E2F1 [192, 193]. Interestingly, two of the miRNAs from the miR-17–92 polycistron, miR-17-5p and miR-20, have also been shown to target E2F1 through binding to its 3′ UTR [192, 194]. Therefore, these regulatory pathways interact in a positive–negative regulatory loop illustrated in Plate 63.3. According to the model, c-Myc positively regulates both miR-17–92 and E2F1. The E2F1 activation is then attenuated by the down-regulation of miR-17-5p and miR-20a. Excessively high levels of E2F1 induce apoptosis, and regulation of E2F1 by miR-21 is seen as a way to maintain the correct level of E2F1 to promote growth as opposed to apoptosis. The miR-17–92 locus is up-regulated by c-Myc and this can increase angiogenesis in tumors [195]. However, is has also been shown that c-Myc expression can cause a wide range of miRNAs to be down-regulated [196]. Interestingly, several of the miRNAs shown to be down-regulated by c-Myc are expressed in normal liver but are not detected in primary HCCs which express c-Myc (Plate 63.3, Table 63.4). These findings add to the complexity of the web of interactions and show that c-Myc can up-regulate oncogenic miRNAs (miR-17–92) while at the same time down-regulating tumor suppressive miRNAs (Let-7, for example) (Plate 63.3).

miR-15a/16 Another example is the loss of miR-15a and miR-16 in chronic lymphocytic leukemia (CLL) [47]. These miRNAs target the anti-apoptotic gene Bcl2 and their loss can promote cancer cell survival. In HCCs, miRs 15a/16 continue to be expressed at levels equivalent to normal liver, and HCCs have evidently found their mechanisms to avoid apoptosis.

Let-7 family Let-7a is down-regulated in lung cancer. Since Let-7a targets the H-Ras gene, its down-regulation leads to increases in cell proliferation [169]. The Let-7 family of miRNAs are also significantly down-regulated in HCCs. Many earlier studies of primary HCCs have noted that H-Ras mRNA levels were normal and concluded that the H-Ras pathway was not down-regulated. Clearly, re-evaluation of the miRNA status of the HCCs may reveal an increase in H-Ras protein. This could occur by elimination of translational repression while mRNA levels are unchanged from normal. Another important direct target of Let-7 is an RNA-binding protein called IMP-1 [199]. IMP-1 has growth-promoting activities through stabilization of both c-Myc and IGF-2 mRNAs [200]. Thus reduction of Let-7 during malignant transformation can lead to coordinate activation of IGF-2 and Myc proteins. Interestingly, coordinate activation of Myc and IGF-2 in precancerous nodules is one of the hallmarks of hepatocarcinogenesis in some animal models [90, 201, 202].

ROLE FOR miRNAs IN HEPATIC CYSTOGENESIS Hyper-proliferation of bile duct epithelial cells (cholangiocytes) is a key feature of cystogenesis in polycystic

63: miRNAs AND LIVER BIOLOGY

liver diseases (PCLDs). A cholangiocyte cell line (CCL) from a rat model proliferates faster than normal rat cholangiocytes (NRCs). miRNA profiling identified 12 miRNAs present in NRCs and not in CCL and 39 miRNA present only in CCL [127]. The most highly down-regulated of the miRNAs in CCL was miR-15a, which was decreased 37-fold. miR-15 was also decreased in the proliferative cholangiocytes that line the liver cysts in both human and rat livers with polycystic liver disease. A bioinformatics approach identified a conserved miR-15a site in the dual specificity phosphatase CDC25A. CDC25A is a member of a family of dual-specificity phosphatases that play an essential role in cell cycle progression by activating CDKs. CDC25A is principally responsible for G1–S and G2–M transitions [203]. CDC25A protein levels were threefold higher in CCL cells than NRCs. When miR-15a mimics were transfected into the CCLs, the CDC25A levels were reduced 50%. A luciferase reporter assay was used to test whether miR-15a directly interacted with the candidate site in the CDC25A 3′ UTR using the CCL cell line. A mutant reporter construct in which the miR-15a seed site was mutated was tested along with the wild-type. This test showed that miR-15a only down-regulated the reporter with the wild-type site and not the mutant site. Therefore, miR-15a regulates CDC25A protein expression through direct binding to the 3′ UTR. Interestingly, the repression was observed on the protein level and not on the mRNA level, in a classic miRNA pattern. Finally, the effect of miR-15a over-expression was tested by assaying the formation of cysts in cultures of CCL cells. These cells spontaneously form cyst-like structures in cell culture in less than 2 weeks. When miR-15a was transfected into CCLs, there was a dramatic reduction in cyst formation in vitro [127]. The authors concluded that although no miRNA therapies of any liver pathological conditions have been reported, their data suggest that the modulation of miRNA expression should be considered a potential therapeutic approach in benign hyper-proliferative diseases such as PCLD.

SUMMARY AND NEW HORIZONS When the Lewis and Clark expedition explored the great American West, they mapped out some of the main rivers in the frontier, but many fascinating places in the West remained to be discovered. In many ways, the field of miRNA biology is in a similar state at this period in history. Although some of the major mechanistic pathways of miRNA action have been identified, the field still remains as a broad new frontier in which exciting new discoveries are waiting at every turn of the river. The importance of gene regulation for the proper development of multicellular organisms has been understood for decades and the study of mechanisms that control the

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spatial and temporal expression of genes during development and in disease have been a central theme of research. Therefore, it was a huge surprise that a major level of gene regulation had been completely unknown until the relatively recent discovery of small non-coding RNA molecules know known as microRNAs[17, 204]. Since then, there has been a race to understand miRNA functions and amazing progress has been made in a very short time. This chapter has provided a brief overview of progress in understanding mechanisms of action of miRNAs based on large numbers of original papers and outstanding reviews. So far there are over 600 human miRNAs, most of which are highly conserved in evolution. miRNAs are expressed in all tissues and all developmental stages of plants and animal and, therefore, our understanding of their tentacles in biology is at the very early stages. With regard to the liver, miRNA expression profiles during fetal development are just beginning to be characterized. The major miRNA expressed in liver, miR-122, comprises approximately 70% of all the miRNAs in liver. Although it is expected that miR-122 may have hundreds of direct targets, only a few have been validated. Experiments that knocked down miR-122 in mouse liver have given us a glimpse of the wide range of genes that it may regulate and the metabolic consequences of its loss. However, the mechanisms of miR-122 actions in the liver are virtually unknown. Roles for miRNAs in cell fate decisions in the fetal liver are just beginning to become known, as are interactions of miRNAs with signal transduction pathways in the liver. Also, miRNA roles in liver disease are just beginning to be reported. miRNA profiling of HCCs has only begun to define clearly which miRNAs are increased or decreased during malignant transformation of hepatocytes. Early work has identified miRNAs that target tumor suppressor genes and others that target oncogenes and several of these are differentially regulated during hepatocarcinogenesis. The simple flow diagram of interactions between miRNAs and oncogenes and tumor suppressor genes presented is certainly only the very beginning of a much more complex web of miRNA regulation in cancer. Some miRNAs are globally expressed whereas others are tightly developmentally regulated. It is therefore expected that globally expressed miRNAs may have a broader range of functions that act to modulate many target genes. Tissue-specific miRNAs may have roles in turning off a few key target genes that help define a specific tissue metabolic phenotype or function. Therefore, the presence of a very high abundance miRNA in the liver, miR-122, and many other significantly expressed miRNAs, presents a great challenge to liver biologists.

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Imaging Cellular Proteins and Structures: Smaller, Brighter, and Faster Erik Snapp Department of Anatomy and Structural Biology, Albert Einstein College of Medicine, Bronx, New York, NY, USA

ADVANCES IN FLUORESCENT DYES AND PROTEINS

INTRODUCTION Hepatocytes have long been used to explore fundamental problems in cell biology, including cell polarization, vesicular transport, protein trafficking, and organelle biogenesis [1–5]. While biochemistry and fixed-cell imaging methods, including histochemistry and electron microscopy (EM) [6], have provided many important insights, live cell microscopy methods have enabled investigators to follow trafficking of a specific vesicle in real time [7], to observe how specific proteins change their distribution during cell polarization [8], and the dynamics of organelle movements [7]. Recently, several important advances have been made in the resolution of cell structures and cell processes by light microscopy imaging. It is now possible to resolve organelle structures and the proteins within them to the level of single molecules. In addition, new classes of fluorescent proteins and advances in high-speed microscopic imaging have recently provided investigators with powerful tools for the characterization of protein localization, dynamics, and fate. In this chapter, the principles and implications of these advances for liver cell biologists are discussed.

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

An exceptionally important tool for studying organelle and protein structure and dynamics is the fluorescent protein tool box [9]. Cells and proteins are generally too small to detect with the naked eye, relatively transparent when imaged by light microscopy, and are highly dynamic. To overcome these limitations, microscopists have developed a number of methods to enhance the contrast of cell structures and resolve individual organelles and proteins better. The primary contrast method used in cell biology today is fluorescence. Proteins or other molecules are labeled with a dye that can be excited with a light source. The dye absorbs the energy of the exciting light and can emit photons of a longer wavelength than the exciting light source. Popular fluorophores include fluorescein isothiocyanate (FITC), rhodamine, Alexa dyes, and green fluorescent protein (GFP). GFP and other fluorescent proteins have revolutionized cell biology [10–13]. These genetically encoded fluorophores can be attached to any protein using standard molecular biology methods and the fluorescent protein folds and forms the fluorophore in the

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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absence of any special cofactors, other than oxygen [13]. The cloning and optimization of GFP have enabled cell biologists to label cells in tissue culture and in whole animals, label proteins on the cell surface and in the lumena of organelles, and follow the dynamics of cells in animals and proteins in cells [7, 11]. Since the cloning of GFP [14], the number of new fluorescent proteins has increased exponentially. Brighter and spectrally diverse blue, green, red, and far-red proteins have been identified and been made available to investigators [11, 12]. The different colors of fluorescent proteins are only the beginning of the fluorescent protein revolution. With the first reports of photoactivatable and timer fluorescent proteins, it is now possible to optically mark and distinguish different populations of the same protein within the same individual cell or cellular compartment. The investigator can now perform a biochemistry-style optical pulse-chase experiment in a single cell with sub-second temporal resolution. Patterson and Lippincott-Schwartz reported the first practical photoactivatable fluorescent protein, when George Patterson mutated wild-type (wt) GFP into photoactivatable green fluorescent protein (PA-GFP) [15]. GFP can be excited at 488 nm and emits a fluorescent signal between 488 and 550 nm. After wt GFP is briefly pulsed with intense blue light (about 400–413 nm), the fluorescence emission intensity of the photoactivated GFP, when now excited with 488 nm light, increases about threefold over background, a modest increase. Patterson’s modified PA-GFP fluorescence emission is indistinguishable from background fluorescence transfected cells when excited at 488 nm. Photoactivation induces an up to 70-fold increase in fluorescence intensity when excited with 488 nm light following photoactivation (Figure 64.1). This dramatic improvement in fluorescent signal over background autofluorescence now permits optical marking of proteins in cells. The number of different types of PA proteins has dramatically multiplied [11, 16] and many of these proteins are appropriate for studies in hepatocytes. The photoactivation of most PA proteins is irreversible, although exceptions, such as kindling proteins [17] and DRONPA [18], can be reversibly or repeatedly activated. Photoactivation is exceptionally powerful. A whole cell can be photoactivated and the movement of that cell or daughter cells from that particular cell can be followed visually. Equally importantly, optical marking of a pool of proteins permits the observation of the changes in a pool of protein’s distribution with time or even the turnover rate of a protein in single cells. In Figure 64.1b, the rate of export of a PA-GFP-tagged viral membrane protein from the Golgi complex in a single cell can be quantitated by following the rate of depletion of the activated protein from the Golgi complex. Unfortunately, GFP is not without its issues. First, GFP is not small. The 5 nm diameter is sufficiently bulky potentially to block protein interactions sterically [19]. Second, GFP is not always sufficiently bright for imaging

Photoactivating UV light

(a) Preactivation

Photoactivated

5 min

(b)

Figure 64.1 Photoactivation. (a) Illustration of photoactivation. A cell expressing a photoactivatable protein is subjected to intense UV light from either a laser or an arc lamp, converting the photoactivatable protein from a dark state into a visible or differently colored state. The activated protein’s behavior and fate can be followed over time in the cell. (b) The viral membrane protein, vesicular stomatitis virus G protein, was fused to PA-GFP, transfected into Cos-7 cells, and imaged. The pool of protein in the Golgi complex was photoactivated with 413 nm laser light and imaged with 488 nm laser light. Photoactivation reveals the pool of protein in the Golgi complex and the vesicles that traffic toward the plasma membrane over time. Image provided courtesy of Dr. George Patterson

of low-expression proteins in cells [20, 21]. Finally, GFP does not instantaneously fold and form its fluorophore. The time delay for fluorophore formation can range from half-times of 15 minutes to hours, depending on the fluorophore [22]. Several alternatives to fluorescent proteins have been made available for live cell imaging. Quantum dots (qdots) [23] are selenium derivative dyes that are two to three orders of magnitude brighter than GFP. The exceptionally bright qdots have other useful properties, including size-dependent fluorescence emission (the larger the qdot, the longer is the emission wavelength) and all qdot colors are excited by the same wavelength of light. The chemistries of qdots have recently been mastered and qdots are relatively well tolerated by cells. In addition, qdots can be conjugated with other proteins such as antibodies to label cell proteins indirectly in live cells. Qdots are typically twice the size (or more) of GFP and are not membrane permeant. Therefore, qdots have issues with steric hindrance of labeled proteins and lumenal proteins remain generally inaccessible to qdots. Another class of fluorescent labels [24] includes FlAsH/ReAsH peptides (∼2 kDa) [25], single-chain antibody proteins (∼25 kDa), and dihydrofolate reductase [26] small proteins. The domains can be tagged to proteins of interest. The domains lack a genetically encoded fluorophore. However, the domains can bind membrane-permeant dyes with high affinity and thus fluorescently label the protein of interest. These smaller proteins are less prone to steric effects from the tag

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compared with the 5 nm GFP tag. In addition, this class of fluorescent tags allows investigators to perform another form of optical pulse labeling, similar to photoactivation. All proteins are labeled with a fluorophore at time zero and then protein stability can be followed over time or a second fluorophore can be incubated at a later time to reveal how much new protein has been synthesized since time zero or to compare the organization of old and new proteins [25]. Furthermore, ReAsH can be used for correlative EM. A fluorescent image of the tagged protein distribution is captured, the cell is fixed, a DAB chemical reaction is performed to create an electron-dense precipitate wherever the ReAsH protein is localized, and it can be visualized by transmission electron microscopy (TEM) [25]. Not all of the alternative fluorescent proteins are appropriate for all cellular compartments. For example, FlAsH/ReAsH dyes bind free cysteines and do not work in the oxidizing environment of the endoplasmic reticulum or Golgi complex lumena [27]. However, the potential for incorporating organic dyes that are much brighter than GFP into specific intracellular proteins makes this an important technology to follow.

RESOLUTION Resolution is a measure of a microscope’s ability to separate optical features (spatial) or temporal events. Increasing resolution permits two structures, points, or events in a cell to be visually and quantitatively distinguished. In practical terms, increased optical resolution permits an investigator to delineate compartments, organelles, or even proteins, while temporal resolution permits an investigator to visualize finer increments of a cellular process such as the movement of a vesicle or diffusion of a pool of proteins across a cell. For thorough discussions of the fundamentals of light microscopy, the reader is referred to excellent reviews by Davidson and Abramowitz (http://micro.magnet.fsu.edu/ primer/opticalmicroscopy.html) and Petty [28], and Murphy’s light microscopy book [29]. Optical resolution is distinct from magnification, which simply means increasing the size of an image. Magnification of a high-resolution image will reveal increasing levels of detail, whereas a magnified low-resolution image will appear blocky (Figure 64.2). Optical resolution is independent of magnification and instead depends on the numerical aperture (NA) of the objective and the wavelength of light used for imaging. Over 100 years ago, Ernst Abbe calculated the “limit” of light microscopy resolution. The equation resolution = 0.61λ/NA indicates that with the best objective available (NA ≈ 1.4) and visible light (λ = 400–600 nm), a resolving power of two fluorescent points of the same color is 200 nm in the lateral plane (x − y) is a best-case scenario. In the axial

1X

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20X

Figure 64.2 Magnification. A Cos-7 cell expressing an endoplasmic reticulum (ER)-GFP was imaged with a 63× oil NA 1.4 objective. The region indicated with the white square was magnified 20-fold and simply appears pixilated. Compare this pair of images with the images in Figure 64.8, in which the same magnification objective was used, but with two very different resolutions using confocal microscopy and PALM

(z ) plane, the resolution is much poorer, 500–800 nm. The inability to resolve particles perfectly below these dimensions is due to light diffraction.

DIFFRACTION Light microscopy resolution is largely a function of diffraction, the phenomenon in which light waves bend around barriers or objects and spread out at angles that are neither parallel nor perpendicular (Figure 64.3a). As a consequence of diffraction, a ray of light does not bounce off a point-like object as a reflection. Instead, light waves bend around point-like objects forming an observed spot much larger than the point itself. Furthermore, the diffracted light is not simply a single larger spot. The wave-like nature of light generates a series of maxima and minima that form a bull’s-eye-like pattern termed an “airy disk” in two dimensions (Figure 64.3b). Detection of the maxima and minima depends on wavelength of the light and the size of any slits or apertures in front of the detector. The use of apertures with detectors is the basis of confocal microscopy and will be discussed below. Microscopic objects are not two-dimensional and airy disk patterns are not simple spheres. Rather, diffracted spots tend to be circular in the lateral (x − y) plane and elongated (poorer resolution) in the axial (z ) direction that occurs in the optical axis of the microscope. A very small, spherical bead, when imaged with a microscope, will appear as peak with successive small ripples around it (Figure 64.3b). The three-dimensional pattern around the bead is termed a point spread function (PSF). The PSF depends on the wavelength (λ) of light and the NA of the microscope objective. The higher an objective’s NA, the smaller is the PSF and the better the resolving power of the objective. The practical consequences of diffraction are that a fluorescence image pixel does not necessarily

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High S/N incoming light waves

Moderate S/N

Low S/N

Airy Disk

q

(a)

PSF diffracted light waves

(a)

(b)

Figure 64.3 Diffraction. (a) Planar light waves are disturbed by diffraction upon encountering an object similar in size or smaller than the wavelength of light. The waves bend or scatter at an angle θ that is inversely proportional to the size of the particles. (b) Illustrations of the two-dimensional airy disk and three-dimensional point spread function that result from illumination of a fluorescently labeled particle

represent a single fluorophore or fluorescently labeled molecule within the space of the pixel. For example, one can readily visualize a microtubule in a cell by fluorescence microscopy, but microtubules are only 25 nm in diameter, well below the diffraction limit of 200 nm. While fluorescence permits visualization of cellular structures and molecules, fluorescence should not be used in place of EM to measure fine structures below 200 nm. Additional confounders include fluorophore labeling efficiency of a sample and detector exposure time, which will further affect the apparent size of a fluorescent object.

SIGNAL-TO-NOISE RATIO Image quality can be enhanced by increasing the signal-to-noise ratio (S/N) (Figure 64.4a). A bright cell against a dark background will have a higher S/N than a dimly labeled sample with a high degree of autofluorescence (Figure 64.4b). Any method that boosts the signal of the fluorescent label or decreases the background or removes other sources of noise will improve the S/N. Improving the S/N should not be confused with improved resolution, even though image quality can be greatly improved by increasing the S/N (compare high and low S/N images in Figure 64.4b). Deconvolution, confocal, and total internal reflection fluorescence (TIRF) microscopies, and also charge-coupled device (CCD) binning, represent examples of improved S/N over standard wide-field fluorescence. A wide-field fluorescence image (termed wide-field because the entire specimen is illuminated simultaneously), collected on a standard fluorescence microscope, consists of the signal from a fluorescently labeled cell(s), illuminated simultaneously over the sample field, and

(b)

Figure 64.4 Signal-to-noise ratio (S/N). (a) Illustration of consequences of high and low S/N. Note how the high S/N sample reveals intensity details for more squares and presents a greater dynamic range than the moderate and low S/N samples. (b) Images of a Cos-7 cell expressing an endoplasmic reticulum localized GFP with different S/N. The low S/N image is obviously of poor quality. In contrast, it is easier to see the entire ER structure in the moderate S/N image than in the high S/N image. However, the range of detectable details is lower in the moderate S/N image. For quantitative imaging, it is essential to be able to detect the greatest range of differences in fluorescence intensities

collected by a camera attached to a microscope. Most light microscope cameras are cooled CCDs that collect the entire image at once (Figure 64.5a). The signal from the sample is converted by the CCD into an array of pixels and the resolution of each image pixel is a function of the size of each pixel or well on the camera detector, the wavelength of light, and the NA of the objective. Camera architecture will be discussed more in Section on High-speed Imaging. As a consequence of illuminating the whole sample field and capturing the entire image at once, the fluorescent sample will have considerable levels of noise. The two main sources of noise are (i) the out-of-focus light from above and below the sample plane and (ii) the overlap of PSF signals from adjacent fluorophores within the plane (poorly resolved signal). Both sources of noise can be reduced by methods that produce images that approach the diffraction limit: deconvolution and confocal microscopy.

Deconvolution Deconvolution is a software-based solution to separate the PSF signals both within the x − y plane and from the out-of-focus light [30]. First, the user must measure the PSF for the particular microscope–camera–illumination setup. Next, the user collects a Z series [a 2D (x − y) series of images stepping through successive focal planes along the optical axis of the microscope objective] of a fluorescently labeled cell. Then, the experimentally determined PSF is iteratively applied to the Z series data

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Binning

CCD

(a) Point Scanner

(c)

(b) Line Scanner

(d)

Figure 64.5 Image acquisition modes. Comparison of how photons for each pixel of an image are acquired using different acquisition modes. (a) A CCD typically collects photons for each pixel of the image simultaneously. (b) To enhance the signal of a CCD, a block of pixels [here a 2 × 2 block of pixels from (a)] can be summed in a process called binning. The image loses resolution, but significantly increases the S/N. (c) In a point scanner system (i.e. a laser scanning confocal microscope with a PMT detector), photons are collected for one pixel at a time in a linear direction, followed by a return to the first pixel point of the next line of the image. (d) A line scanner combines aspects of a point scanner and a CCD. A CCD is also used, but the CCD consists of a single line of pixels. A galvanometer moves an illuminating laser shaped like a line up and down the sample to collect all of the photons for a line of pixels simultaneously. The line scanner increases acquisition speed and can be combined with a slit to achieve a significant degree of confocality

to remove the out-of-focus light that corresponds to the planes above and below each focal plane. The resulting deconvoluted image exhibits a dramatic reduction in noise and increased resolution approaching the Abbe diffraction limit. Although deconvolution can be more time consuming than confocal microscopy (see below), it has two important advantages. Typically, deconvolution is much less expensive than purchasing a confocal microscope and, because the user is not throwing away photons, dimmer samples can be imaged with less photobleaching. As will be seen in the section on super-resolution, other microscopies, even confocal microscopy, can potentially benefit from deconvolution.

Confocal and Multiphoton Microscopy For thick samples, such as tissue samples with several layers of cells, confocal microscopy is required to reduce the noise in focal planes well into the cell layers. In theory, confocal microscopy improves image resolution by a modest 30% in lateral and axial planes under ideal conditions. The confocal microscope acquires images differently from a wide-field system, by exploiting a pinhole. The laser light source is focussed into an hourglass-like

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volume with the narrowest point coinciding with the focal plane. Although a sample is illuminated through the entire optical axis of the microscope, the sample is only illuminated at a single point in the lateral plane. The point-like light source substantially reduces noise from the adjacent lateral regions of the sample. Most samples consist of more than a single point. To acquire the whole image, the point source is moved back and forth in a rastering motion (Figure 64.5c) over the sample. The spatial and temporal precision with which a sample can be illuminated have been exploited for other applications, such as fluorescence recovery after photobleaching (FRAP), photoactivation, and fluorescence correlation spectroscopy (FCS) [10]. The pinhole, positioned in front of the detector, excludes light from above and below the focal plane. The pinhole is adjustable and must be closed down to resolve a single airy disk. By incorporating a motorized stage with the microscope system, one can collect images for each focal plane and then reconstruct the entire sample in three dimensions, without having physically sliced the sample. Successive 3D stacks of fluorescently labeled live cells can be captured over time to perform 5D imaging (three spatial dimensions, a fluorescence color, and time). For very bright samples, it is possible to close the pinhole to less than the size of a single airy disk and achieve resolution slightly better than the Abbe diffraction limit. The small increase in resolution and large increase in S/N come at the expense of the loss of 90% or more of the photons from the sample. Practically, this means that only moderately to very bright live samples can be imaged by confocal microscopy. Also, illumination through the entire thickness of the sample results in photobleaching of regions outside the plane of focus. However, the multifunctional nature of confocal microscopes (5D imaging, colocalization, photobleaching, and excellent image quality) makes these instruments the microscopes of choice for core facilities. Although confocal microscopy is excellent for imaging samples up to 50 µm thick, there are several instances in which an investigator would like to image even thicker samples such as tissue or even in live animals. Light scatter in tissue seriously degrades the utility of confocal microscopy, which utilizes mostly visible light wavelengths. In contrast, infrared light is more transparent in tissues and can penetrate deep into tissues. The primary method for delivering infrared light into tissues, up to 400 µm deep, is multiphoton imaging [31]. The physical basis of multiphoton microscopy is beyond the scope of this chapter. The relevant feature of multiphoton imaging is that only fluorophores in the focal plane and focal volume are excited. Similarly to confocal microscopy, multiphoton imaging employs a rasterized point source of illumination to create essentially the same volume as a confocal PSF. A major advantage of multiphoton imaging is high S/N due to the lack of illumination outside the focal volume. The

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resulting images are similar to confocal images. Multiphoton imaging requires a multiphoton laser and highly sensitive detectors, which can be relatively expensive. Regardless of the cost, multiphoton imaging remains the primary method for imaging cells in live animals. In addition, multiphoton microscopy is relevant to super-resolution microscopy, as several methods utilize multiphoton lasers to achieve sufficiently high S/N without loss of fluorescent signal.

TIRF Microscopy TIRF microscopy is an exceptionally powerful imaging technique for investigators studying questions close to the cell plasma membrane [32, 33]. TIRF microscopy dramatically improves the image S/N. Unlike confocal or wide-field instruments, TIRF microscopes do not directly illuminate the sample. Instead, the light is brought in at an incident angle in which the light source reflects off the coverslip. Under certain conditions, an evanescent wave is created at the coverslip. The evanescent wave will only excite fluorophores within 100 nm of the coverslip, which will include the plasma membrane, the cytoskeleton, and vesicles close to the plasma membrane. The results are impressive (Figure 64.6) with an S/N even better than in confocal microscopy. TIRF microscopy does not throw away photons and illuminates a slice even thinner than the actual focal plane. No out-of-focus light is produced, which reduces the threshold for detection of single fluorophores, such that a TIRF microscope can detect single GFP molecules. The lateral PSF of the fluorophore is the same as in wide-field, so despite the ability to detect the signal of a single GFP, TIRF cannot resolve two GFP molecules any better than wide-field microscopy. TIRF microscopy is exceptionally popular for studying cell surface membrane trafficking and cytoskeletal questions. In addition, it can be combined with photoactivatable proteins and computational methods to achieve super-resolution, as

Widefield

TIRF

Figure 64.6 TIRF. MTLn3 rat adenocarcinoma cells transiently transfected with EB1-GFP (microtubule plus-end binding). (a) A wide-field fluorescence image and (b) the same cell imaged by TIRF. Note the absence of the cytoplasmic GFP haze and low background in the TIRF image. Image courtesy of Vera DesMarais

will be seen for photoactivated localization microscopy (PALM).

INCREASING MICROSCOPE RESOLUTION Achieving higher temporal or spatial resolution frequently occurs at the expense of each other. For example, the highest spatial resolution microscopies, EM and atomic force microscopy (AFM), are generally incompatible with live cell imaging. EM is performed under a vacuum, which is not compatible with live liver cells or tissue, and AFM is so slow that only extremely small areas can be scanned to follow a live cell process. Some groups have worked around these limitations by combining technologies. To resolve a fluorescent cellular structure in high detail, a cell can be imaged by fluorescence microscopy and then rapidly fixed and imaged by TEM in a method termed correlative microscopy [6]. The method does not provide higher resolution information concerning the fluorescently labeled molecules. Now, methods such as PALM have ushered in a new era for correlative microscopy by providing high resolution of both cellular structures and the fluorescently labeled molecules within the structure. The types of questions that can now be addressed with the new high-resolution technologies are exciting and important. Furthermore, the high-resolution fluorescence technologies have made the transition into living cells, meaning that very few questions will be off limits to imaging.

SUPER-RESOLUTION MICROSCOPY Since the late 1990s, cell structures have been resolved below the Abbe diffraction limit with such regularity that it might seem that there is no longer a diffraction limit. However, the “super-resolution” microscopies do not violate Abbe’s equation. Rather, super-resolution methods circumvent the limit through new microscope designs, new fluorescent probes, and/or computer algorithms. There are two general types of super-resolution microscopies: (i) “near-field” methods that do not image beyond the surface or close to the surface of the sample and (ii) “far-field” techniques, such as stimulated emission depletion (STED), structured illumination, and 4pi, can image as deep into a sample as standard confocal microscopy, at least the thickness of one cell layer. In the following sections, the general principles and capabilities of the different super-resolution microscopies will be described.

4pi One way to increase resolution is to redesign the microscope. With a standard light microscope, the user obtains

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only a fraction of the total possible image data. This is because the PSF is obtained from only one half of the sample. The objective does not collect the light from the side of the sample that radiates in the opposite direction of the objective. Stefan Hell and colleagues reasoned that collection of the other half of the data could improve the PSF, particularly in the axial plane. By using a modified slide that sandwiches the sample between two coverslips and placing microscope objectives on both sides of the sample, the 4pi method improves Z resolution by about fivefold to 100 nm (Figure 64.7c). Lateral resolution is unaffected. The 4pi system uses a point scanning raster mechanism. The main drawbacks of 4pi microscopy are the cost of the commercial system (Leica) (nearly US$1 million at the time of writing this chapter, mostly due to the expensive multiphoton laser and accompanying optics) and the production of lobes or ghost images in the image. Fortunately, the lobes can be removed with software algorithms. Deconvolution can further improve 4pi resolution to 140 nm in x − y and 90 nm in z . The enhanced axial resolution of 4pi can be especially useful for resolving cellular structures such as cytoskeletal components and vesicles.

Structured Illumination Mats Gustafsson and John Sedat pioneered patterned or structured illumination (SI) (also called I5S) [34], a super-resolution method that can improve wide-field images from 200 × 500 to ∼100 × 280 nm or even Widefield

Confocal, Multiphoton and Deconvolved Widefield

Axial

Lateral

800 nm

500 nm 200 nm

300 nm (b)

(a) 4pi 200 nm

Structured Illumination

90 nm

50–100 nm

(c)

(d)

STED, NSOM, PALM/STORM/fPALM 10–20 nm (e)

Figure 64.7 Point spread functions. Comparative illustrations of the relative PSFs of the same fluorescent bead using different microscopy methods, including (a) wide-field, (b) confocal and deconvoluted wide-field, (c) 4pi, (d) SI, and (e) NSOM, STED, and PALM/STORM/fPALM. Axial (z ) and lateral (x − y) dimensions are indicated. Note that deconvolution produces a result comparable to confocal microscopy

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100 × 100 nm (Figure 64.7d) [35]. There are several variations of SI [34–37], but the basic method consists in illuminating a sample with patterned light, similar to a polarized light grating filter. The pattern of the sample interferes with and is multiplied by the filter pattern to create a Moir´e pattern. A computational restoration algorithm decodes the pattern and measures the fringes in the Moir´e pattern. The SI pattern remains constrained by diffraction and cannot be focused to anything smaller than half the wavelength of the excitation light, which limits resolution enhancement to a factor of 2. Resolution can be further increased in multiple ways, including saturating illumination of the sample and a 4pi-like setup with placement of microscope objectives on both sides of the sample [35]. However, saturating illumination causes sample photobleaching and is therefore impractical for live cell imaging. SI is a wide-field technique, which makes it one of the faster super-resolution microscopies. However, multiple images frequently must be collected for a single final image. Therefore, the method remains best for fixed samples or relatively slow processes. SI is an excellent alternative to and improvement over confocal microscopy for colocalization experiments. A commercial SI system (Apotome from Carl Zeiss) that improves the S/N, but not resolution, is available and is less expensive than many laser scanning confocal systems. A commercial super-resolution SI system is offered by Applied Precision.

NSOM Of all of the super-resolution methods, the near-field scanning optical microscopy (NSOM) instrument least resembles a traditional microscope design. There is no recognizable objective on the system. To break the diffraction barrier, NSOM employs both a light source and a detector in a nanometer-sized tip, which is placed close to the structure to be detected [38]. The design circumvents issues of diffraction, which is a far-field effect. When the NSOM tip is brought to within nanometers of a molecule, the resolution is no longer limited by diffraction, but by the size of the tip aperture. Effectively only one fluorescent molecule will be exposed to the light emanating from the tip to achieve resolutions of 10 nm or less (Figure 64.7e). Images are built by raster scanning of the tip over the surface of the sample. The primary limitations of NSOM are (i) the limited number of illuminating photons emanating from the small tip, (ii) the low photon collection efficiency, and (iii) the current need to use fixed samples to maintain a steady distance of the tip from the sample. The ultrafine scanning and low efficiency of sample photon collection will substantially increase image collection time over 4pi or confocal techniques. NSOM remains restricted to the analysis of cell surface molecules and remains an experimental technology for cell biologists.

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PALM and STORM A limitation for resolution of individual fluorescent molecules in samples is that all of the fluorescent molecules in a field are illuminated and imaged simultaneously. The different super-resolution methods employ different strategies to resolve the fluorophores. Confocal, 4pi, SI, and STED techniques (see below) reduce the size of the PSF to varying degrees. NSOM avoids the PSF altogether and interrogates fluorophores one at a time. In PALM [39], fluorescence photoactivation localization microscopy (fPALM) [40], and stochastic optical reconstruction microscopy (STORM) [41, 42] (different acronyms for the same basic method that will be referred to as PALM in this chapter), the physical PSF is not mechanically modified and the image is collected in wide-field. Instead, individual fluorophores are visualized over the entire field. Visualizing single fluorophores is achieved by employing photoactivatable fluorophores that are initially turned off and are randomly turned on a few at a time. The activated fluorophores are imaged until they photobleach. Then a few more fluorophores are activated successively until all of the fluorophores in the field have been eventually photoactivated and photobleached to assemble the image. As noted above, wide-field and confocal imaging are hindered by substantial autofluorescence and out-of-focus light, which prevents detection of single fluorophores in cells. However, TIRF can provide sufficient S/N to detect single fluorophores in cells. The final component of PALM is to localize each individual fluorophore, and this can be accomplished by performing a Gaussian fitting routine to pinpoint the center of the PSF for each fluorophore (Figure 64.8a). Gaussian fitting has been used by several groups [43] to assign fluorophore positions with nanometer precision. The robustness of the technique depends on the number of photons detected for each fluorophore, which slows the image acquisition rate and requires the use of relatively bright fluorophores. Technically, Gaussian fitting methods do not qualify as optical resolution. Rather, the fitting methods permit measurement of the distance between fluorophores. By distinguishing the position of each fluorophore in PALM, it is possible to achieve localization of fluorophores with a precision of ∼10–40 nm (Figure 64.7e) and stunning images (Figure 64.9) [39]. The constraints of TIRF restrict PALM to molecules at or close to the cell surface. Physical sectioning of fixed samples permits the application of PALM methodology to any region of a fixed cell or tissue. Image acquisition time remains the main hurdle for PALM. The image (Figure 64.9) generated for the first PALM publication took 12 hours to acquire, in addition to computer processing time to pinpoint PSF centers and sample processing [39]. Hess et al. recently acquired live cell images of influenza hemagglutinin (HA) on the cell surface and could localize individual HA molecules

10 nm

actual CCD image

Ideal PSF

Gaussian-fitted point

(a) 20 nm

excited spot

STED

effective spot

(b)

Figure 64.8 Super-resolution methods. (a) PALM employs TIRF to reduce background noise and then selectively activates and illuminates a few single photoactivatable fluorescent proteins at a time. Photons are collected with a CCD as in standard TIRF imaging, summed for each intensity spot, and the center of fluorescence emission for each spot is determined. This is achieved by statistically fitting the measured photon distribution with an ideal PSF, which can have sufficient accuracy to produce 10 nm resolution. (b) In STED, a focused excitation beam is superimposed with a second donut-shaped laser beam that quenches excited fluorophores outside of the donut hole. The resulting spot can be resolved to 20 nm

to 40 nm spots. The fPALM approach yielded new insights into the organization of lipid raft proteins and challenges to the lipid raft model [44]. Imaging of a live sample, as the authors pointed out, required live cell proteins with exceptionally low diffusion coefficients (i.e. 0.09 µm2 s−1 for HA compared with 0.4 µm2 s−1 for most membrane proteins and 25 µm2 s−1 for cytoplasmic proteins) to collect sufficient numbers of photons before the molecules diffused away. Currently, PALM is most useful for imaging large molecular assemblies such as nuclear pores or chromatin and dramatically improved correlative microscopy. When coupled with TEM, the positions of individual molecules can be assigned and one can determine whether molecules are distributed in a homogeneous or heterogeneous manner over a vesicle or organelle. The correlative microscopy methods should be exceptionally useful for studying cargo sorting in vesicles and the organization of large molecular assemblies. A commercial PALM system has been developed recently by Carl Zeiss.

STED Stefan Hell and colleagues developed a second super-resolution method, STED. Two laser pulses are used to generate each pixel. The first nanosecond pulse is focused into a diffraction-limited spot and is at the absorption wavelength of the fluorophore of interest.

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1.0 mm (a)

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1.0 mm (b)

0.2 mm

0.5 mm (c)

(d)

Figure 64.9 PALM images. Comparison of summed-molecule images by (a) TIRF and (b) PALM. Images of the same region within a cryo-prepared thin section from a Cos-7 cell expressing the lysosomal transmembrane protein CD63 tagged with the PA-fluorescent protein (FP) Kaede. The same objective was used for both images. The TIRF image can resolve the gross distribution of lysosomal structures (note that individual vesicles are too small, ∼50 nm, to be well resolved by standard methods and each pixel within the TIRF image can contain multiple lysosomes). The larger boxed region in (b), when viewed at higher magnification (c), reveals smaller associated membranes that may represent interacting lysosomes or late endosomes that are not resolvable by TIRF. In a region where the section is nearly orthogonal to the lysosomal membrane, the most highly localized molecules fall on a line of width 10 nm (inset). In an obliquely cut region [(d), from the smaller boxed region in (b)], the distribution of CD63 within the membrane plane can be discerned. From Betzig et al ., Science 313: 1642–5 (2006). Reprinted with permission from AAAS

A second intense multiphoton donut-shaped laser pulse immediately follows the first pulse and is at the emission wavelength. The second pulse stimulates excited fluorophores to fall back to the ground state and not emit any photons in the depletion pulse (Figure 64.8b). Only fluorophores in the outer part of the diffraction-limited spot are depleted, while the small center molecules still fluoresce. The size of the donut directly regulates the resolution and can produce resolution of 10–20 nm in x , y, and z (Figure 64.7e). Deconvolution methods can enhance resolution further [45]. Unlike PALM and NSOM, STED is a far-field method. Thus, any part of the cell can be studied. STED is currently achieved with a multiphoton laser, which suggests that the method is effectively confocal and can be used for 3D reconstruction. However, since the initial excitation is single photon, the complications of diffraction will

probably restrict STED to the depth range of confocal microscopes (0–50 µm). Although STED images are acquired through raster scanning, Hell and colleagues have reported video rate [28 frames per second (fps) or about 35 ms per frame] imaging of vesicle movement within a neuronal synapse [46]. Due to the small size of each pixel, achieving video rate required restricting the scan area to 1.8 × 2.5 µm. However, if coupled with a Z motor, a cubic volume of this area could be a useful area for following 5D localized dynamics. That is, a vesicle that translates across a cell at 1 µm s−1 would be too fast to follow. In contrast, localized processes or assemblies such as organization of a nuclear pore or dynamics of the Golgi complex dynamics or bile canaliculi would be excellent subjects for investigation with STED. A STED system is commercially available from Leica, though the million dollar price tag, at the time of writing this chapter, means

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that investigators will want to seek out collaborations with imaging centers that can afford the instrument. Regardless of price, the power of STED as a super-resolution far-field solution makes it the most powerful general solution for achieving super-resolution images of cells.

Beyond Super-resolution Unfortunately super-resolution microscopies cannot detect interactions between proteins or the dynamics of the interactions. Currently, no light microscopy technique can resolve direct protein interactions, as most proteins are below 5–10 nm in diameter and individual amino acids are smaller than 0.5 nm. However, biophysical fluorescence methods, such as fluorescence resonance energy transfer (FRET), permit investigators to detect molecular proximities below 10 nm in living cells. FRET has been the subject of numerous reviews [10, 47, 48] and remains the method of choice for detecting the subnanometer proximities of molecules in cells. It should be possible to couple FRET probes with super-resolution imaging and this has been achieved with NSOM [49]. Hence it is likely that as more investigators gain access to super-resolution instruments, new combinatorial imaging modes will be developed to study the organization and interactions of proteins in cells.

HIGH-SPEED IMAGING A number of cellular processes occur on fast (millisecond) time scales including: calcium waves, diffusion of cytoplasmic proteins, and vesicle motility. Imaging these processes requires a combination of rapid image acquisition and a sensitive detector. Although all imaging generally benefits from higher sensitivity in a detector, fast imaging is especially demanding because shorter acquisition times translate to fewer photons collected. One solution is to use brighter fluorophores, that is, qdots [23]. A complementary approach is to increase detector sensitivity. In the past several years, detectors have become dramatically more sensitive. The two main types of microscope detectors are CCDs and photomultiplier tubes (PMTs). PMTs, while capable of detecting and counting single photons [31], are less useful for high-speed imaging because they normally detect only a single pixel at a time (Figure 64.5c). If a full 512 × 512 pixel image is not needed, reducing the number of pixels to be acquired will require less time to scan. Despite this limitation, a PMT coupled with a resonant scanner can achieve 25 fps, near video frame rates (30 fps). Resonant scanners also employ a pair of mirrors and a galvanometer to sweep the point illuminator pixel by pixel over the sample. However, the resonant scanner moves the mirrors at a single maximum speed while maintaining the spatial resolution

of confocal sample acquisition. Both Nikon and Leica have incorporated resonant scanners into their confocal systems. CCD detectors are inherently faster image collectors, because CCDs typically collect the entire image field at once (Figure 64.5a). There are two types of CCDs for light microscopy, CCDs and electron multiplying charge-coupled devices (EMCCDs). The latter are more sensitive and have higher spatial resolution due to smaller pixel size [28]. The CCD S/N can be further enhanced by collecting more photons per pixel through binning (Figure 64.5b), in which photon counts for multiple collected camera pixels are combined into a single pixel. Binning will reduce image resolution by whatever factor is chosen for the bin size, but the tradeoff is individual pixels with the combined photons of 4, 9, or X 2 pixels, which substantially boosts the S/N. Standard scanning confocal microscopes remain limited in their image acquisition rates due to imaging only one pixel at a time. However, confocal imaging can be coupled with the speed of CCDs. The first implementation of this combination was spinning disk confocal microscopy [29]. Instead of a single point scanner [also referred to as a confocal laser scanning microscope (CLSM)], the microscope illuminates the entire sample field through a disk that contains numerous microlenses. The fluorescence emission from the sample travels through a second disk that contains multiple non-adjustable pinholes and projects onto a CCD or EMCCD. The two disks spin rapidly and produce confocal images at video rates (30 fps). The sensitivity of an EMCCD is essential for higher spinning disk image acquisition speeds. More recently, a hybrid approach has been developed that couples partial confocality with a single line of pixels of a CCD. The Zeiss Live captures a whole line of pixels and scans up and down a sample (Figure 64.5d) to achieve frame rates up to 180 fps. Through the use of slits, in a manner similar to a confocal pinhole, the PSF y-axis is equivalent to confocal, the x -axis is ∼10% greater, and the z -axis is ∼20% greater [50]. These small reductions in PSF resolution do not significantly reduce image quality (Figure 64.10a). The sensitivity of the CCD and a microscope with high quantum efficiency dramatically improve image quality over previous generation confocal PMTs scanning an image at 5 fps (Figure 64.10b). By achieving 180 fps, it is possible to follow practically any cellular process. EMCCD imaging, spinning disk confocal, resonant scanning confocal, and line scanning confocal techniques now permit outstanding temporal resolution of vesicular trafficking, calcium waves, 5D imaging of many processes on the scale of the whole cell, and protein diffusion in the cytoplasm (see below).

High-speed Photomanipulation The spatio-temporal coordination of protein location and movement permits proteins to regulate virtually

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Figure 64.10 High-speed photobleaching. Comparison of photobleaching data for (a) a high-speed CLSM (Zeiss Duo) imaging at 180 fps and (b) a standard point scanning CLSM imaging at 5 fps. Cos-7 cells expressing cytoplasmic GFP were briefly photobleached and the fluorescence recovery was monitored and quantitated. The bleach region is indicated by the white box. Note the distinctly visible photobleach region in the high-speed CLSM post-bleach image that is visible only as a slight dimming in the standard CLSM post-bleach image. In addition, the high-speed system collected 140 recovery data points (filled squares) compared to the two recovery data points (empty circles) for the standard CLSM. As cytoplasmic GFP is the fastest (25 µm2 s−1 ) fluorescent protein to be used for photobleaching applications, the data reveal that the high-speed CLSM can monitor the mobility of any cellular GFP-fusion protein. In addition, the use of higher sensitivity detectors dramatically enhances the quality of the high-speed CLSM image, even at 180 fps

all dynamic processes in living cells. Protein function depends on the availability of a protein to interact with substrates or partner proteins in the cellular environment. By labeling proteins with GFP variants and performing photomanipulation (fluorophore destruction/photobleaching or highlighting/photoactivation) [51]

with a CLSM, it has become possible to quantitate protein availability in live cells. Selective photobleaching [i.e. FRAP and fluorescence loss in photobleaching (FLIP) [52]] or photoactivation of GFP-labeled proteins permits investigators to measure a protein’s mobility (diffusion coefficient), molecular size, and the percentage of mobile

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proteins (mobile fraction), and to follow a photoactivated protein’s fate. Although photomanipulation applications have provided many insights into protein dynamics [10], a serious deficit in this methodology has been the relatively slow rates of data acquisition (two to five images per second) and the substantial delay between the end of a photomanipulation event and the collection of the first post-manipulation images. These two factors are especially problematic for cytoplasmic or lumenal proteins that can move several microns in the space of milliseconds. Wide-field fluorescence imaging solutions are available, but often require exceptionally powerful lasers to photobleach or photoactivate a region of interest rapidly. In contrast, confocal microscope scanners concentrate laser light and selectively apply it to discrete regions of a cell, which results in rapid photobleaching or photoactivation. Although CLSM systems offer superior resolution relative to a wide-field microscope, they often have less sensitive detectors, and use the same scanner for photomanipulation and imaging (Figure 64.10b). Coupling photomanipulation with high-speed microscopes has broken this temporal barrier with remarkable image quality, spatial resolution, quantum efficiency, and speed (Figure 64.10a). High-speed FRAP of cytoplasmic GFP reveals that it is now possible to follow the dynamics of virtually any fluorescently labeled cellular protein with exceptional temporal resolution. Furthermore, high-speed photomanipulation experiments can be performed in 5D, which permits the monitoring of events throughout the cell and how much of the cellular volume was affected by photobleaching or photoactivation.

could be used to follow the lineage and fates of individual transplanted stem cells or developing transgenic cells in liver [54]. It is not always necessary to use multiphoton microscopy to image cellular phenomena in live animals. Thiberge et al. [55] recently applied high-speed spinning disk confocal microscopy to follow the malarial sporozite gliding on liver sinusoid, crossing the sinusoidal barrier, invasion, and development within hepatocytes in real time. Similar kinds of approaches could be used to visualize invasion steps by other liver pathogens, including viruses and bacteria. Super-resolution microscopies hold great promise for studies of vesicular trafficking in hepatocytes. Wakabayashi et al. [56] used confocal microscopy and FRAP to visualize trafficking of the bile salt export pump in primary hepatocytes. Wide-field fluorescence microscopy studies of purified hepatocyte-derived vesicles by Murray et al. revealed additional mechanistic insights into the segregation of endocytic cargo [4]. STED or PALM could provide precise spatial details of how individual cargo proteins are sorted within vesicles. Similarly, super-resolution studies of changes in the dynamics and distribution [57, 58] of cell–cell communication proteins (i.e. connexins) could define cellular microenvironments and provide insights into the differentiation and life cycle of hepatocytes [59]. The power of super-resolution optical and high temporal resolution microscopies and their recent translation to live cells are making these technologies vital to the future of liver cell imaging and our understanding of liver biology.

REFERENCES THE FUTURE OF LIGHT MICROSCOPY Light microscopes have become especially powerful tools for cell biologists. What were experimental technologies 10 years ago are now commercially available. Many of the technologies described in this chapter are already commercially available or will become so within the next few years. Resolution has become a tractable problem. The next technological breakthrough that will be needed is the creation of smaller and brighter fluorescent proteins and brighter fluorescent dyes. Such reagents will improve cell imaging with shorter image acquisition times to collect adequate numbers of photons for faster imaging and higher S/N. Super-resolution and high-speed imaging technologies will both benefit from brighter fluorophores. Many of the technologies described in this chapter have the potential to provide new insights into fundamental aspects of liver biology. For example, STED or PALM methods could be used to investigate the trafficking and high-resolution distributions of sensory proteins in cholangiocyte primary cilia [53]. Imaging live liver tissue with multiphoton microscopy coupled with photoactivatable fluorescent proteins and/or additional fluorescent proteins

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9. Giepmans, B.N. et al. (2006) The fluorescent toolbox for assessing protein location and function. Science, 312, 217–24. 10. Lippincott-Schwartz, J. et al. (2001) Studying protein dynamics in living cells. Nat Rev , 2, 444–56. 11. Shaner, N.C. et al. (2007) Advances in fluorescent protein technology. J Cell Sci , 120, 4247–60. 12. Shaner, N.C. et al. (2005) A guide to choosing fluorescent proteins. Nat Methods, 2, 905–9. 13. Tsien, R.Y. (1998) The green fluorescent protein. Annu Rev Biochem, 67, 509–44. 14. Chalfie, M. et al. (1994) Green fluorescent protein as a marker for gene expression. Science, 263, 802–5. 15. Patterson, G.H. and Lippincott-Schwartz, J. (2002) A photoactivatable GFP for selective photolabeling of proteins and cells. Science, 297, 1873–77. 16. Lukyanov, K.A. et al. (2005) Innovation: photoactivatable fluorescent proteins. Nat Rev , 6, 885–91. 17. Verkhusha, V.V. and Lukyanov, K.A. (2004) The molecular properties and applications of Anthozoa fluorescent proteins and chromoproteins. Nat Biotechnol , 22, 289–96. 18. Ando, R. et al. (2004) Regulated fast nucleocytoplasmic shuttling observed by reversible protein highlighting. Science, 306, 1370–73. 19. Andresen, M. et al. (2004) Short tetracysteine tags to beta-tubulin demonstrate the significance of small labels for live cell imaging. Mol Biol Cell , 15, 5616–22. 20. Niswender, K.D. et al. (1995) Quantitative imaging of green fluorescent protein in cultured cells: comparison of microscopic techniques, use in fusion proteins and detection limits. J Microsc, 180, 109–16. 21. Snapp, E. (2005) Design and use of fluorescent fusion proteins in cell biology. In: Current Protocols in Cell Biology (eds J.S. Bonafacino et al.), John Wiley & Sons, Inc., Hoboken, NJ, Unit 21.21. 22. Shaner, N.C. et al. (2004) Improved monomeric red, orange and yellow fluorescent proteins derived from Discosoma sp. red fluorescent protein. Nat Biotechnol , 22, 1567–72. 23. Arya, H. et al. (2005) Quantum dots in bio-imaging: revolution by the small. Biochem Biophys Res Commun, 329, 1173–77. 24. Prescher, J.A. and Bertozzi, C.R. (2005) Chemistry in living systems. Nat Chem Biol , 1, 13–21. 25. Gaietta, G. et al. (2002) Multicolor and electron microscopic imaging of connexin trafficking. Science, 296, 503–7. 26. Calloway, N.T. et al. (2007) Optimized fluorescent trimethoprim derivatives for in vivo protein labeling. ChemBioChem, 8, 767–74. 27. Gaietta, G.M. et al. (2006) Golgi twins in late mitosis revealed by genetically encoded tags for live cell imaging and correlated electron microscopy. Proc Natl Acad Sci U S A, 103, 17777–82. 28. Petty, H.R. (2007) Fluorescence microscopy: established and emerging methods, experimental strategies, and applications in immunology. Microsc Res Tech, 70, 687–709.

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29. Murphy, D.B. (2001) Fundamentals of Light Microscopy and Electronic Imaging, Wiley-Liss, New York. 30. Wallace, W. et al. (2001) A working person’s guide to deconvolution in light microscopy. BioTechniques, 31, 1076–97. 31. Zipfel, W.R. et al. (2003) Nonlinear magic: multiphoton microscopy in the biosciences. Nat Biotechnol , 21, 1369–77. 32. Axelrod, D. (2001) Total internal reflection fluorescence microscopy in cell biology. Traffic, 2, 764–74. 33. Axelrod, D. (2003) Total internal reflection fluorescence microscopy in cell biology. Methods Enzymol , 361, 1–33. 34. Gustafsson, M.G. (1999) Extended resolution fluorescence microscopy. Curr Opin Struct Biol , 9, 627–34. 35. Shao, L. et al. (2008) I5S: widefield light microscopy with 100-nm-scale resolution in three dimensions. Biophys J , 94 (12), 4971–83. 36. Gustafsson, M.G. et al. (2008) Three-dimensional resolution doubling in widefield fluorescence microscopy by structured illumination. Biophys J , 94, 4957–70. 37. Gustafsson, M.G. (2005) Nonlinear structuredillumination microscopy: wide-field fluorescence imaging with theoretically unlimited resolution. Proc Natl Acad Sci U S A, 102, 13081–86. 38. Edidin, M. (2001) Near-field scanning optical microscopy, a siren call to biology. Traffic, 2, 797–803. 39. Betzig, E. et al. (2006) Imaging intracellular fluorescent proteins at nanometer resolution. Science, 313, 1642–45. 40. Hess, S.T. et al. (2006) Ultra-high resolution imaging by fluorescence photoactivation localization microscopy. Biophys J , 91, 4258–72. 41. Huang, B. et al. (2008) Three-dimensional super-resolution imaging by stochastic optical reconstruction microscopy. Science, 319, 810–13. 42. Rust, M.J. et al. (2006) Sub-diffraction-limit imaging by stochastic optical reconstruction microscopy (STORM). Nat Methods, 3, 793–95. 43. Park, H. et al. (2007) Single-molecule fluorescence to study molecular motors. Q Rev Biophys, 40, 87–111. 44. Simons, K. and Ikonen, E. (1997) Functional rafts in cell membranes. Nature, 387, 569–72. 45. Donnert, G. et al. (2006) Macromolecular-scale resolution in biological fluorescence microscopy. Proc Natl Acad Sci U S A, 103, 11440–45. 46. Westphal, V. et al. (2008) Video-rate far-field optical nanoscopy dissects synaptic vesicle movement. Science, 320, 246–49. 47. Bunt, G. and Wouters, F.S. (2004) Visualization of molecular activities inside living cells with fluorescent labels. Int Rev Cytol , 237, 205–77. 48. Snapp, E.L. and Hegde, R.S. (2006) Rational design and evaluation of FRET experiments to measure protein proximities in cells. In: Current Protocols in Cell Biology (eds J.S. Bonafacino et al.), John Wiley & Sons, Inc., Hoboken, NJ, Unit 21.21. 49. Ha, T. et al. (1996) Probing the interaction between two single molecules: fluorescence resonance energy transfer between a single donor and a single acceptor. Proc Natl Acad Sci U S A, 93, 6264–68.

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50. Wolleschensky, R. et al. (2006) High-speed confocal fluorescence imaging with a novel line scanning microscope. J Biomed Opt , 11, 064011. 51. Lippincott-Schwartz, J. et al. (2003) Photobleaching and photoactivation: following protein dynamics in living cells. Nat Cell Biol , 5 (Suppl), S7–14. 52. Snapp, E. et al. (2003) Measuring protein mobility by photobleaching GFP-chimeras in living cells, in Current Protocols in Cell Biology (eds J.S. Bonafacino, et al.), Unit 21.21, John Wiley & Sons, Inc. 53. Masyuk, A.I. et al. (2008) Cholangiocyte primary cilia in liver health and disease. Dev Dyn, 237 (8), 2007–12. 54. Oertel, M. and Shafritz, D.A. (2008) Stem cells, cell transplantation and liver repopulation. Biochim Biophys Acta, 1782, 61–74. 55. Thiberge, S. et al. (2007) In vivo imaging of malaria parasites in the murine liver. Nat Protoc, 2, 1811–18.

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Zebrafish as a Model System for the Study of Liver Development and Disease Randolph P. Matthews Division of Gastroenterology, Hepatology and Nutrition, The Children’s Hospital of Philadelphia and Department of Pediatrics, University of Pennsylvania School of Medicine, Philadelphia, PA, USA

INTRODUCTION Although many advances have been made in the understanding of liver disease and physiology using rodent models and cell culture, recent studies using the zebrafish have also been able to contribute to a greater understanding of the liver in health and in disease. Zebrafish are an excellent model to study development, and fish in general make an excellent model to study drug effects given the ease of administration into the water. Developmental biologists have become increasingly enamored of zebrafish over the past decade, owing to several features. Zebrafish develop rapidly, outside the mother, reaching a mature enough state to swim and eat by 5 days post-fertilization (dpf), and are optically clear, allowing easy visualization of internal organs, including the liver (Figure 65.1). Furthermore, they can breed frequently and prolifically. The ability to generate large numbers of fish facilitates the use of zebrafish in forward genetic screens, in which investigators are able to screen large numbers of families for interesting phenotypes, and subsequently identify the causative gene. Furthermore, genetic manipulation by microinjection of morpholino antisense The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

oligonucleotides or messenger RNA (mRNA) allows relatively rapid determination of the effects of knocking down or overexpressing specific genes. Compared with homologous recombination techniques in mice, morpholino injections lead to less dramatic and only temporary reduction of the targeted gene product, but can be accomplished in a fraction of the time and cost required for similar experiments in mice. Interestingly, maternal gene and protein expression persists through later stages of development than in mammals; this allows for the survival of mutants past critical stages of early development that would not be possible without lingering maternal mRNA or protein. Hence zebrafish are a powerful model to study genetic influences on development. Given the optical clarity of the zebrafish, investigators can utilize transgenically labeled fish to study cell lineage and organogenesis. Transgenic lines expressing green fluorescent protein (GFP) or red fluorescent protein (RFP) in specific developing cell types or organs allow visualization of cell movement and organogenesis during development. These transgenic fish, when injected with morpholinos, treated with a drug, or crossed with mutant lines, allow the investigator to view in vivo the effect of the genetic or pharmacological manipulation on

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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Figure 65.1 Visible structures in a 5 dpf zebrafish larva. Left-sided view of a 5 dpf zebrafish larva showing the visible digestive organs and other notable features

development of the tissue or organ of interest. Not only can such powerful techniques lead to novel findings, but also the results are often esthetically captivating. As the number of zebrafish researchers continues to grow, additional techniques are being developed that will further establish the zebrafish as an outstanding animal model. Several genetic engineering techniques have been developed that allow targeting of specific genes, similar to techniques to generate null and conditionally null mice. Bioinformatics approaches such as expression microarrays and proteomics are becoming more readily available and more frequently utilized. Further techniques for labeling and following or ablating individual cells during development will enhance developmental studies. Underlying all of the technical advances is another facet that attracts investigators to zebrafish: it is becoming increasingly clear that there is a high degree of conservation between zebrafish and mammals in numerous biological processes. Despite 350 million years of evolutionary distance between humans and teleost fish, multiple pathways are shared, including those important in liver development, disease, and response to injury. Recent advances in understanding these processes using zebrafish are outlined below.

USE OF ZEBRAFISH TO STUDY LIVER DEVELOPMENT Early liver development In mammals, the initial signals to the foregut that lead to hepatic specification and liver bud formation are derived from the cardiac mesoderm and septum transverse mesenchyme, and involve members of the fibroblast growth factor (FGF) and bone morphogenic protein (BMP) families. These signals induce expression of sonic hedgehog (shh), which inhibits pancreatic development but not liver development. The interplay between the developing heart and liver does not appear to occur in zebrafish, as the developing heart and liver are not in close proximity during liver specification or morphogenesis. Signaling via FGFs

and BMPs is clearly important in zebrafish, however, as inhibition of FGF or BMP activity inhibited hepatic specification [1]. Using transgenic zebrafish expressing inhibitors of FGF or BMP activity driven by inducible promoters, Shin et al. demonstrated that the importance of BMP and FGF signaling in specification occurs during a relatively narrow developmental time, and that while inhibition of either factor after that time appears to inhibit growth of the liver, specification is normal [1]. Furthermore, this study demonstrated that increased BMP activity, by way of overexpressing bmp2b, could overcome a lack of FGF activity. These studies exemplify some of the more powerful features of doing such experiments in zebrafish, namely using transgenic lines to express genes or inhibitors at specific time points during development. Although hedgehog activity does appear to be involved in liver development in zebrafish, its role appears different than in mammals. Whereas mammalian shh expression is permissive for liver development, in zebrafish shh expression inhibits liver formation. Global inhibition of hedgehog signaling using cyclopamine or via genetic inactivation of smoothened inhibits liver formation [2, 3]. The effect of cyclopamine was evident if administered at 12 hours post-fertilization (hpf) or earlier, but had no effect if administered later. Thus, unlike in mammals, hedgehog signaling plays an important role in specification of the zebrafish liver. Furthermore, liver specification in zebrafish occurs well before gut tube formation (24 hpf), as evidenced by expression of the liver markers hhex and other genes rostral to the developing gut at 18 hpf [3]. Field et al. examined liver formation using a transgenic zebrafish with GFP expressed in the developing gut [4]. At around the same time that BMP and FGF signaling is critical (24–28 hpf), GFP-positive cells rostral to the gut tube, in the position of liver progenitors, proliferate, and extend laterally. As development proceeds, the gut tube, which eventually forms the intestine [3], loops left and ultimately becomes contiguous with the extrahepatic duct [4]. The gut-GFP line used by Field et al. was also used in a forward genetic screen to look for phenotypes in which there was abnormal gut development. The mutant prometheus was identified as having an almost absent liver, suggesting defects in liver specification; like the defects in BMP and FGF signaling that impair liver development, these defects were transient [5]. The causative gene for prometheus was identified as wnt2bb, suggesting a role for Wnt signaling in early liver development. Importantly, the wnt2bb gene is expressed in the lateral plate mesoderm, suggesting that in zebrafish, as in mammals, signals derived from the surrounding mesoderm drive early liver development. This importance of Wnt signaling in early liver development had not been appreciated in mammalian models, most likely because mouse mutants affecting Wnt signaling were lethal prior to the initiation of liver development, while the zebrafish mutants were able to survive these stages because of the presence of wild-type maternal mRNA and/or protein.

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The role of Wnt signaling in early liver development is clearly more complex, however, as studies in Xenopus demonstrated that inhibition of Wnt activity in the anterior endoderm was required for normal liver development, and that induced repression of Wnt activity in posterior endoderm resulted in ectopic liver formation [6]. An explanation for this apparent contradiction was provided by further zebrafish studies from Goessling et al., who demonstrated that very early wnt8 activation led to an inhibition in the expression of early liver markers and a lack of hepatocyte differentiation, whereas a slightly later wnt8 activation led to an increase in expression of markers and an increase in the size of the developing liver [7]. These studies in zebrafish and Xenopus suggest that Wnt signaling is critical in early development, a finding consistent with the known importance of Wnt signaling in the generation of hepatoblastoma. Interestingly, zebrafish and mammals share several liver-specific genetic markers, while some markers in mammals are not in fact present in the fish. The ceruloplasmin (cp) gene is the earliest marker for the developing zebrafish liver [8]; the albumin and α-fetoprotein genes used for early markers in mammals appear to not be present in the fish. Other markers, such as prox1 and hhex , are present in both zebrafish and mammals [3], as is transferrin (tfa) [9], although expression of tfa initiates later than cp. Injection of morpholinos directed against hhex lead to a dramatic reduction in the size of the developing liver, suggesting that hhex expression is important for the initial stages of growth of the developing liver [10]; these findings are similar to those for experiments done in mammals.

Later liver and hepatobiliary development Hepatoblast differentiation into hepatocyte and bile duct cell is a process that has been studied extensively in rodent and cell culture models, but there have also been contributions from zebrafish. There are clear distinctions between zebrafish and mammalian biliary development, however [12]. Whereas the mammalian liver is organized into lobules with portal triads containing bile ducts, portal vein, and hepatic artery, there is no such organization in the zebrafish liver. The teleost liver maintains a more acinar organization, with an interconnecting network of bile ducts. Unlike in mammalian liver, where canaliculi form from adjacent hepatocytes, zebrafish canaliculi are invaginations of the hepatocyte plasma membrane. These canaliculi drain bile directly into the interconnected intrahepatic ducts. Thus, whereas mammalian bile ducts form via the developing ductal plate around a nascent portal vein, there is no ductal plate intermediate in the developing zebrafish liver (see Plate 65.1 for a depiction of zebrafish hepatobiliary anatomy). Despite this discrepancy

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in anatomy, there is considerable conservation of biliary development at the molecular level, and the relationship to the developing vasculature is probably still important. Recent work demonstrates that the developing intrahepatic vasculature leads to the establishment of hepatocyte apical/basolateral polarity, and this polarity in turn leads to developmental placement of the intrahepatic bile ducts [11]. The endothelial vasculature enters the developing liver around 50 hpf, and polarity appears to be initiating by 55 hpf and complete by 60 hpf. Disruption of the developing hepatic vascular endothelial cells by chemical inhibition of vascular endothelial growth factor (VEGF) or genetic inhibition of heart of glass or valentine, genes encoding for likely signaling molecules between the endothelium and hepatocytes, results in abnormal hepatocyte polarization and abnormal intrahepatic bile duct development. Interestingly, however, a complete lack of vasculature results only in subtle polarity abnormalities and normal intrahepatic biliary development [11, 12]. Thus, VEGF and genes that direct vascular development may have a direct role in biliary development independent of their effects on the developing vasculature. The establishment of hepatocyte polarity occurs prior to detection of the first intrahepatic bile ducts, as one would expect. Intrahepatic bile ducts, as delineated by cytokeratin immunostaining, appear around 60 hpf. These ducts are initially short and not interconnected; connections develop over the next 24–36 hours, and by 5 dpf the intrahepatic ductal network has a relatively mature appearance, with clear anastamoses with the hepatocytes and drainage into the extrahepatic biliary tree [12, 13] (see Figure 65.2 for the timeline of zebrafish hepatobiliary development). The extrahepatic biliary tree in zebrafish is identical with that in mammals, with the cystic duct leading to the gallbladder, and the common bile duct being joined by the pancreatic duct prior to emptying into the intestine. Thus, the development of the intrahepatic bile ducts in zebrafish, although not going through a ductal plate stage, does therefore have gross similarities with the process in mammals. The similarities with mammalian intrahepatic biliary development continue at the molecular level. The Onecut transcription factors Hnf6, OC2, and OC3, and the homeodomain protein Hnf1β play a critical role in mammalian intrahepatic biliary development, being essential for early specification of bile duct cells and late remodeling of the bile ducts [14, 15]. This latter role is conserved in zebrafish, as morpholino-mediated knockdown of the zebrafish orthologs of hnf6 and oc3 led to abnormalities in the later stages of intrahepatic biliary development [13, 16]. As in mammals, hnf1b appears downstream of hnf6 , and in zebrafish, forced expression of hnf1b rescued the defects caused by hnf6 knockdown, suggesting that most of the remodeling effects of hnf6 are mediated through hnf1b in zebrafish [13]. Interestingly, whereas in mammals Hnf6 acts earlier than Oc2 or Oc3 , in zebrafish there is no Oc2 ortholog, and oc3 appears to act earlier

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THE LIVER: USE OF LARVAL ZEBRAFISH TO GENERATE MODELS OF HUMAN LIVER DISEASE

Figure 65.2 Timeline of hepatobiliary development in zebrafish. The timeline of development is depicted, with hours post-fertilization in the thick black bar. Above the bar are noted milestones in hepatobiliary development, and below the bar are milestones of zebrafish development in general

than hnf6 [16]. This suggests that although there is overall functional conservation, specific roles of these genes have diverged. This distinction also holds true for members of the Jagged and Notch signaling pathway. Mutations in JAGGED1 or NOTCH2 lead to Alagille syndrome in humans, a disease characterized by bile duct paucity and presumed defects in intrahepatic biliary development. Pack’s group established an Alagille syndrome phenocopy in zebrafish using morpholino-mediated knockdown, but the specific morpholinos required to generate the Alagille-like phenotype included combinations of jagged2 and jagged1b, jagged2 and notch2 or notch5 , or notch2 and notch5 . These results suggest the possibility that Jagged/Notch interactions with respect to biliary morphogenesis are more complex in zebrafish than in mammals, or possibly that mutations in other NOTCH or JAGGED genes may lead to bile duct paucity. Interestingly, in the morphant fish, there were defects in the initial stages of biliary development that resulted in hybrid cells resembling both bile duct cells and hepatocytes, suggesting that Jagged/Notch signaling may be important in biliary cell specification. Patients with Alagille syndrome and mouse models of Alagille syndrome demonstrate apparent defects in later stages of biliary development, so these findings represent a possible divergence of the role of Jagged/Notch in biliary development, although Jagged/Notch signaling is also important in the later stages of biliary development in zebrafish (K. Lorent and M. Pack, unpublished work). The studies regarding both the Jagged/Notch and Onecut roles in zebrafish biliary development demonstrate conservation of these pathways between zebrafish and mammalian biliary development, but have also expanded our understanding of biliary development in general. Interestingly, whereas Hnf6 −/− mice demonstrate an abnormal extrahepatic biliary network with a hypoplastic

gallbladder, the zebrafish hnf6 morphants have a normal extrahepatic biliary tract. In contrast, zebrafish fgf10 mutants demonstrate an abnormal extrahepatic duct and a smaller gallbladder relative to wild-type [17]. Interestingly, these mutants also have ectopic liver and duct staining in the pancreas and into the intestine, suggesting that fgf10 also led to suppression of liver and duct differentiation in other regions of the developing endoderm. Not surprisingly, fgf10 gene expression is highest in the surrounding mesenchyme, consistent with a role as a secreted factor similar to the role of BMP and FGFs in mammalian and zebrafish liver development mentioned above.

USE OF LARVAL ZEBRAFISH TO GENERATE MODELS OF HUMAN LIVER DISEASE As stated above, Pack’s group utilized morpholinomediated knockdown to establish a zebrafish with features similar to those of patients with Alagille syndrome. The morphant fish demonstrated an abnormal facial structure, cardiac defects, and bile duct paucity –three of the five cardinal features of Alagille syndrome. As mentioned above, this phenocopy provided further evidence of the molecular conservation of hepatobiliary development between zebrafish and mammals, and also advanced our understanding of the role of Jagged/Notch in vertebrate biliary development by demonstrating a possible role for Notch signaling in specification of biliary cells and by suggesting a possible role for other JAGGED and NOTCH genes in Alagille syndrome. Morpholino-mediated gene knockdown of vps33b resulted in a partial phenocopy of the rare human disorder arthrogryposis–renal dysfunction–cholestasis (ARC) syndrome [18]. Patients with this disorder have congenital joint contractures, kidney abnormalities, disordered bile transport, and occasionally bile duct paucity, whereas the

65: ZEBRAFISH AS A MODEL SYSTEM FOR THE STUDY OF LIVER DEVELOPMENT AND DISEASE

zebrafish morphant demonstrated only bile duct paucity. There were apparent vesicular trafficking defects in the vps33b morphant livers, as would be expected from the decreased activity of Vps33b, a protein important in intracellular trafficking. Interestingly, the vps33b morphants also demonstrated abnormal vesicles in intestinal cells, possibly consistent with fat malabsorption seen in some ARC syndrome patients. These results underscore the similarities between zebrafish and mammalian hepatobiliary disease pathogenesis and development, and demonstrate the importance of intracellular trafficking pathways in biliary development. Using mutants first uncovered in a large insertional mutagenesis screen [19], Sadler et al. found several mutant phenotypes, including a phenotype with biliary defects and hypopigmentation in which the mutation is in vps18 [20]. This screen was performed looking for mutants with hepatomegaly. As the name suggests, Vps18 is functionally similar to Vps33b, and the vps18 mutants also demonstrate intracellular trafficking defects, again showing the importance of trafficking in biliary development. In addition to vps18 , Sadler et al. also uncovered mutant phenotypes in the tumor suppressor gene nf2 , in which the fish develop choledochal cyst-like structures, and foie gras, a novel gene in which the fish develop hepatic steatosis. Hepatic steatosis is also an important feature of the recently described mutant duct-trip [21]. This mutant was originally identified in a screen for exocrine pancreas mutants [22] and was subsequently found to have hepatic steatosis, abnormal mitochondria, and progressive liver degeneration. The causative gene is ahcy, which encodes S -adenosylhomocysteine hydrolase, an important component of the methionine metabolism pathway, that when inhibited leads to an increase in S -adenosylhomocysteine, which in turn leads to a global inhibition of methylation. Patients with mutations in the AHCY gene have been described, and develop hepatic steatosis and mitochondrial abnormalities similar to duct-trip [23]. Interestingly, TNFα levels were increased in duct-trip, and inhibition of TNFα led to rescue of steatosis and degeneration. Although increased TNFα activity and inhibition of methylation have been independently linked to other models of hepatic steatosis, this is the first report demonstrating a causal link between a genetically mediated methylation defect, TNFα activation, and hepatic steatosis. Further studies on duct-trip and foie gras should yield interesting findings regarding increasingly more prevalent hepatic steatosis, as zebrafish are readily amenable to drug or alcohol treatment.

USE OF ADULT ZEBRAFISH TO MODEL DISEASE Although most recent zebrafish studies concerning the liver have examined development and disease modeling in

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the context of the developing organism, studies of adult zebrafish have also been revealing. The adult zebrafish liver is a bilobed organ, and although it is not organized similarly to mammalian liver, it performs many of the same functions. The duct-trip mutant discussed above not only exhibits hepatic steatosis and degeneration in the homozygous larvae, but the adult heterozygotes also demonstrate hepatic steatosis [21]. This suggests that even mild methylation defects alone can lead to hepatic steatosis, a finding with implications for patients at risk for hepatic steatosis. Partial hepatectomy (PH) is a method frequently used in rodent models to study liver regeneration and growth. Sadler et al. developed this technique for use in adult zebrafish, and using a forward genetic screen, identified the uhrf1 gene as an important modulator of hepatocyte growth [24]. The mutant was identified due to decreased liver growth during development, but uhrf1 is important not only in developmental liver growth, but also in the PH model. To perform PH on adult zebrafish, these investigators were able to remove up to 40% of the liver through a small ventral–lateral incision, and achieved a postoperative success rate of ∼75%. As in rodent models, the liver regrew, and interestingly this regrowth was limited in the adult uhrf1 heterozygotes, suggesting that uhrf1 is important in regeneration of the adult liver as well as in development. These studies provide a powerful tool for use in zebrafish, as studies of liver regeneration and growth can now be performed in the fish, and in conjunction with mutagenesis screens for genes affecting developmental liver growth should continue to uncover novel genes and pathways involved in liver growth. Uncontrolled growth, of course, leads to cancer, and zebrafish are also emerging as a good liver cancer model. There are numerous cancer models in zebrafish, but relatively few liver cancer models to date. Zebrafish cancer models are derived from forward genetic screens for phenotypes of overgrowth and tumor formation, and historically from chemical mutagenesis to induce non-germline mutations leading to tumors. Zebrafish carrying one mutation in apc demonstrate increased liver growth, and adult apc carriers demonstrated accelerated regrowth and increased Wnt activity after PH [7]. These findings are consistent with the known role of APC mutations in hepatoblastoma, and suggest that the importance of Wnt pathways is conserved in growth regulation and carcinogenesis. Lam et al. addressed this conservation directly by determining that gene expression profiling in four types of human liver cancer was similar to gene expression changes in zebrafish liver tumors induced by chemical mutagenesis [25]. These findings suggest that liver cancer modeling in zebrafish is a legitimate model for studying human liver cancer. Because zebrafish are amenable to both forward genetic screens and chemical treatment to induce tumors, this suggests that zebrafish should be a powerful model system to study liver cancer.

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THE LIVER: CONCLUSION

USE OF ZEBRAFISH TO STUDY DRUG AND TOXIN EFFECTS As stated above, zebrafish have been used to study toxin effects, and this experimental use of zebrafish predates the use of zebrafish as a developmental genetic model. This can be used to study liver damage, as measured by visual necrosis of the liver, histopathology of larval or adult liver, or by enzymatic assays [26]. This approach is useful for screening potentially hepatotoxic drugs or environmental toxins. Certainly one of the main advantages of screening in zebrafish is the ability to perform simple and rapid tests and keep the larvae alive; hence histopathological screens are of limited utility. The fluorescent lipid reporter PED-6 can be used to screen for hepatic and biliary anatomy and function [27]. PED-6 is a quenched phospholipid that is activated by phospholipase A2 in the zebrafish intestine, absorbed via the enterohepatic circulation, processed through the hepatocyte and excreted into the bile, where it concentrates in the gallbladder. Larval zebrafish can be easily scored by examining the intensity of gallbladder PED-6 uptake; this method was used to corroborate structural biliary defects in the vps18 mutants, and also the vps33b, hnf6 , and oc3 morphants mentioned above. PED-6 can also be used as a screen for defects in lipid metabolism and hepatobiliary development. The first mutant identified in such a screen, fat free, demonstrates a mutation in the gene cog8 , which encodes a protein important in Golgi structure and vesicular trafficking that affects lipid transport and also adversely affects intrahepatic bile ducts [28]. Additional lipid metabolism and biliary developmental mutants are likely to be uncovered using PED-6.

FUTURE APPROACHES TO STUDYING LIVER IN ZEBRAFISH While forward genetic screens offer an excellent opportunity to uncover novel genes and pathways, a similar approach can be used to find small molecules that affect such pathways, either positively or negatively. Large chemical libraries have become available, either commercially or generated within academic institutions. Application of such chemical libraries to larval zebrafish and observation for a particular phenotype allow one to screen for interesting compounds. Such approaches have been used for several years now to uncover cancer phenotypes [29]. This technology could be used to uncover phenotypes affecting the liver. Alternatively, one could screen a chemical library for compounds that reverse a mutant phenotype, such as hepatomegaly or abnormal PED-6 uptake.

Newer mutant technologies are allowing investigators to utilize reverse genetic approaches in zebrafish. Clearly, the use of embryonic stem cells and universal or conditional null alleles in mice has resulted in numerous animal models of disease and a greater understanding of various genes and pathways. Such technology is not yet available in zebrafish, but alternative methods of reverse genetics have been developed. The process of TILLING (targeting induced local lesions in genomes), in which mutagenized sperm are screened for mutations in a gene of interest, has been utilized by several groups and is becoming commercially available. Although it is not a new technology, many of the insertional mutants originally generated by the Hopkins laboratory at MIT are available through the Zebrafish International Resource Center (http://zebrafish.org/zirc/home/guide.php) [30], and mutants derived similarly may become available commercially. Finally, custom zinc finger nucleases, targeting specific genes, can be used to engineer mutants of a specific gene [31]. With these tools, zebrafish researchers will be able to order or derive mutants of specific genes and study the effects of these mutations. Technologies to track cells during development are also improving. As the most basic level, as more markers are identified, more transgenically labeled fish can be engineered to track cells during development. This, along with more powerful microscopes and software for image processing, allows for in vivo tracking of cells. A similar approach can be used to induce specific cell death during development. Curado et al. reported a method by which small populations of cells can be ablated at discrete points during development, by establishing transgenic lines with tissue-specific expression of nitroreductase [32]. Fish were treated with metronidazole, which is not harmful to the fish and can be washed away, but is toxic in the cells expressing the nitroreductase; one can then observe the effects of tissue- and stage-specific cell death. Tissue-specific expression of the zinc finger endonucleases, leading to tissue- and stage-specific mutagenesis, is also feasible.

CONCLUSION This chapter has outlined recent progress in understanding liver and biliary development and disease using the zebrafish. Zebrafish provide an outstanding model to study development, given physical and practical considerations, and the general conservation with mammalian hepatobiliary development and physiology. There are numerous existing technologies allowing these studies in zebrafish, and as more investigators enter the field and more technologies become available, there will be more discoveries using zebrafish that increase our understanding of human hepatobiliary disease.

65: ZEBRAFISH AS A MODEL SYSTEM FOR THE STUDY OF LIVER DEVELOPMENT AND DISEASE

ACKNOWLEDGMENT The author thanks Dr Michael Pack for a critical review of the manuscript.

REFERENCES 1. Shin, D., Shin, C.H., Tucker, J., Ober, E.A., Rentzsch, F., Poss, K.D., Hammerschmidt, M., Mullins, M.C. and Stainier, D.Y. (2007) Bmp and Fgf signaling are essential for liver specification in zebrafish. Development , 134, 2041–50. 2. Roy, S., Qiao, T., Wolff, C. and Ingham, P.W. (2001) Hedgehog signaling pathway is essential for pancreas specification in the zebrafish embryo. Curr Biol , 11, 1358–63. 3. Wallace, K.N. and Pack, M. (2003) Unique and conserved aspects of gut development in zebrafish. Dev Biol , 255, 12–29. 4. Field, H.A., Ober, E.A., Roeser, T. and Stainier, D.Y. (2003) Formation of the digestive system in zebrafish. I. Liver morphogenesis. Dev Biol , 253, 279–90. 5. Ober, E.A., Verkade, H., Field, H.A. and Stainier, D.Y. (2006) Mesodermal Wnt2b signalling positively regulates liver specification. Nature, 442, 688–91. 6. McLin, V.A., Rankin, S.A. and Zorn, A.M. (2007) Repression of Wnt/beta-catenin signaling in the anterior endoderm is essential for liver and pancreas development. Development , 134, 2207–17. 7. Goessling, W., North, T.E., Lord, A.M., Ceol, C., Lee, S., Weidinger, G., Bourque, C., Strijbosch, R., Haramis, A.P., Puder, M. et al. (2008) APC mutant zebrafish uncover a changing temporal requirement for wnt signaling in liver development. Dev Biol , 320, 161–74. 8. Korzh, S., Emelyanov, A. and Korzh, V. (2001) Developmental analysis of ceruloplasmin gene and liver formation in zebrafish. Mech Dev , 103, 137–39. 9. Mudumana, S.P., Wan, H., Singh, M., Korzh, V. and Gong, Z. (2004) Expression analyses of zebrafish transferrin, ifabp, and elastaseB mRNAs as differentiation markers for the three major endodermal organs: liver, intestine, and exocrine pancreas. Dev Dyn, 230, 165–73. 10. Wallace, K.N., Yusuff, S., Sonntag, J.M., Chin, A.J. and Pack, M. (2001) Zebrafish hhex regulates liver development and digestive organ chirality. Genesis, 30, 141–43. 11. Sakaguchi, T.F., Sadler, K.C., Crosnier, C. and Stainier, D.Y. (2008) Endothelial signals modulate hepatocyte apicobasal polarization in zebrafish. Curr Biol , 18, 1565–71. 12. Lorent, K., Yeo, S.Y., Oda, T., Chandrasekharappa, S., Chitnis, A., Matthews, R.P. and Pack, M. (2004) Inhibition of Jagged-mediated Notch signaling disrupts zebrafish biliary development and generates multi-organ defects compatible with an Alagille syndrome phenocopy. Development , 131, 5753–66.

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13. Matthews, R.P., Lorent, K., Russo, P. and Pack, M. (2004) The zebrafish onecut gene hnf-6 functions in an evolutionarily conserved genetic pathway that regulates vertebrate biliary development. Dev Biol , 274, 245–59. 14. Clotman, F., Lannoy, V.J., Reber, M., Cereghini, S., Cassiman, D., Jacquemin, P., Roskams, T., Rousseau, G.G. and Lemaigre, F.P. (2002) The onecut transcription factor HNF6 is required for normal development of the biliary tract. Development , 129, 1819–28. 15. Coffinier, C., Gresh, L., Fiette, L., Tronche, F., Schutz, G., Babinet, C., Pontoglio, M., Yaniv, M. and Barra, J. (2002) Bile system morphogenesis defects and liver dysfunction upon targeted deletion of HNF1beta. Development , 129, 1829–38. 16. Matthews, R.P., Lorent, K. and Pack, M. (2008) Transcription factor onecut3 regulates intrahepatic biliary development in zebrafish. Dev Dyn, 237, 124–31. 17. Dong, P.D., Munson, C.A., Norton, W., Crosnier, C., Pan, X., Gong, Z., Neumann, C.J. and Stainier, D.Y. (2007) Fgf10 regulates hepatopancreatic ductal system patterning and differentiation Nat Genet 39(3):397-402. 18. Matthews, R.P., Plumb-Rudewiez, N., Lorent, K., Gissen, P., Johnson, C.A., Lemaigre, F. and Pack, M. (2005) Zebrafish vps33b, an ortholog of the gene responsible for human arthrogryposis-renal dysfunction–cholestasis syndrome, regulates biliary development downstream of the onecut transcription factor hnf6. Development , 132, 5295–306. 19. Amsterdam, A., Burgess, S., Golling, G., Chen, W., Sun, Z., Townsend, K., Farrington, S., Haldi, M. and Hopkins, N. (1999) A large-scale insertional mutagenesis screen in zebrafish. Genes Dev , 13, 2713–24. 20. Sadler, K.C., Amsterdam, A., Soroka, C., Boyer, J. and Hopkins, N. (2005) A genetic screen in zebrafish identifies the mutants vps18, nf2 and foie gras as models of liver disease. Development , 132, 3561–72. 21. Matthews, R.P., Ma˜noral-Mobias, R., Lorent, K., Huang, Y., Gong, W., Murray, I.V.J., Blair, I.A. and Pack, M. (2009) TNFα-dependent hepatic steatosis and liver degeneration caused by mutation of zebrafish S-adenosylhomocysteine hydrolase. Development , in press Mar;136(5):865-75. 22. Yee, N.S., Lorent, K. and Pack, M. (2005) Exocrine pancreas development in zebrafish. Dev Biol , 284, 84–101. 23. Baric, I., Fumic, K., Glenn, B., Cuk, M., Schulze, A., Finkelstein, J.D., James, S.J., Mejaski-Bosnjak, V., Pazanin, L., Pogribny, I.P. et al. (2004) S Adenosylhomocysteine hydrolase deficiency in a human: a genetic disorder of methionine metabolism. Proc Natl Acad Sci U S A, 101, 4234–39. 24. Sadler, K.C., Krahn, K.N., Gaur, N.A. and Ukomadu, C. (2007) Liver growth in the embryo and during liver regeneration in zebrafish requires the cell cycle regulator, uhrf1. Proc Natl Acad Sci U S A, 104, 1570–75. 25. Lam, S.H., Wu, Y.L., Vega, V.B., Miller, L.D., Spitsbergen, J., Tong, Y., Zhan, H., Govindarajan, K.R., Lee, S.,

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Mathavan, S. et al. (2006) Conservation of gene expression signatures between zebrafish and human liver tumors and tumor progression. Nat Biotechnol , 24, 73–75. 26. McGrath, P. and Li, C.Q. (2008) Zebrafish: a predictive model for assessing drug-induced toxicity. Drug Discov Today, 13, 394–401. 27. Farber, S.A., Pack, M., Ho, S.Y., Johnson, I.D., Wagner, D.S., Dosch, R., Mullins, M.C., Hendrickson, H.S., Hendrickson, E.K. and Halpern, M.E. (2001) Genetic analysis of digestive physiology using fluorescent phospholipid reporters. Science, 292, 1385–88. 28. Ho, S.Y., Lorent, K., Pack, M. and Farber, S.A. (2006) Zebrafish fat-free is required for intestinal lipid absorption and Golgi apparatus structure. Cell Metab, 3, 289–300.

29. Stern, H.M. and Zon, L.I. (2003) Cancer genetics and drug discovery in the zebrafish. Nat Rev Cancer, 3, 533–39. 30. Amsterdam, A. (2006) Insertional mutagenesis in zebrafish: genes for development, genes for disease. Brief Funct Genomic Proteomic, 5, 19–23. 31. Meng, X., Noyes, M.B., Zhu, L.J., Lawson, N.D. and Wolfe, S.A. (2008) Targeted gene inactivation in zebrafish using engineered zinc-finger nucleases. Nat Biotechnol , 26, 695–701. 32. Curado, S., Stainier, D.Y. and Anderson, R.M. (2008) Nitroreductase-mediated cell/tissue ablation in zebrafish: a spatially and temporally controlled ablation method with applications in developmental and regeneration studies. Nat Protoc, 3, 948–54.

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The Hepatocyte and the Cancer Cell: Dr Jekyll and Mr Hyde Jean-Pierre Gillet1, Michael M. Gottesman1 and Mitsunori Okabe2 1

Laboratory of Cell Biology, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA 2 Tohoku University School of Medicine, Sendai, Japan

INTRODUCTION The liver plays a major role in metabolism, especially in the detoxification of diverse groups of substrates, both endogenous compounds and xenobiotics. Hepatic uptake of these compounds from sinusoidal blood is accomplished by transporter proteins localized in the basolateral membrane of hepatocytes, whereas hepatic efflux of bile acids, metabolites, and/or drugs is mediated by adenosine triphosphate (ATP)-binding cassette (ABC) transporters (Figure 66.1; reviewed by Alrefai and Gill [1]).

The strange case of Dr Jekyll and Mr Hyde Despite the use of numerous treatment modalities and chemotherapeutic agents, the survival rate for individuals with HCC has not improved during the past few decades. Although the reasons for this failure are multifactorial, intrinsic resistance to chemotherapy [2–4] and/or to antiviral treatment [5, 6] rank as primary. While both types of resistance deserve equal consideration and can sometimes be explained by the same mechanisms, this chapter will focus on resistance to chemotherapy. Drug resistance can be attributed to a number of mechanisms, including decreased uptake [7, 8], increased detoxThe Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

ification [9], alteration of target proteins [10–12], or increased efflux [13, 14]. Several of these pathways can lead to MDR, in which the cell becomes resistant to several drugs in addition to the initial compound administered [15]. The multidrug-resistant cancer cell often displays other properties, such as genome instability [16, 17], polymorphisms in cytochrome P450 (CYP) [18], and loss of checkpoint control [19], which complicate further therapy (Figure 66.2). Hepatocytes, in some ways, exhibit the “split personality” described in the story of Dr Jekyll and Mr Hyde. Their beneficial, healing function of filtering out harmful compounds can be transformed into the inevitable failure of chemotherapeutic treatment. The similarity in gene expression profiles of normal hepatocytes and multidrug-resistant cells is striking. Once a neoplasm develops, the particular gene expression pattern renders it well equipped to resist chemotherapeutic treatments. ABC transporters are key players in MDR and are well represented in liver cells. Since the discovery of the first ABC transporter gene (ABCB1) in 1976 [20, 21], huge amounts of data have been collected, highlighting the complexity of the mechanisms involved in MDR. However, considering the number of cancer patients who still experience treatment failure due to this phenomenon, it seems that we are still in the initial phase of understanding MDR.

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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OATP1A2 OATP1B1 OATP1B3 OATP2B1

PGT

OA−

PG

Lumen of sinusoidal capillary

NTCP

ABCC2 ATP

OC+

Na+ BS−

ATP ATP ABCG2

ABCG5/G8 PC

ATP

Cholesterol

OCT1 OCT3

GSH, X− OA−

ATP ABCB11

ABCB4

BS−

OC+ Bile canaliculus

ATP ABCC1 ABCC3 ABCC4 ABCC5 ABCC6 ATP

ABCB1

OAT2 OA−

H+ MATE1 OAT7 Butyrate

OA−

OA− BS−

OST alpha + beta

OA− BS−

Lumen of sinusoidal capillary

Figure 66.1 Transporters in hepatocytes. Bile salts, drugs, and metabolites are taken up in hepatocytes via the basolateral solute carriers OATP1A2/SLCO1A2, -1B1/SLCO1B1, -1B3/SLCO1B3, -2B1/SLCO2B1, PGT/SLCO2A1, NTCP/SLC10A1, OCT1/SLC22A1, OCT3/SLC22A3, OAT2/SLC22A7, and OAT7/SLC22A9. Hepatic efflux from the hepatocyte to the bile canaliculus is mediated through the apical membrane by the ABC transporters ABCB1, B4, B11, C2, and G2, the heterodimer G5/G8, and the solute carrier MATE1/SLC47A1. The efflux of compounds from hepatocytes to the blood is accomplished by ABCC1, C3, C4, C5, and C6, whereas the heterodimeric transporter OSTα–OSTβ is responsible for sterol reabsorption. SLC, solute carrier; SLCO, solute carrier organic anion-transporting polypeptide; OATP, organic anion-transporting polypeptide; LST, liver-specific organic anion transporter; NTCP, Na+ -taurocholate cotransporting polypeptide; OCT, organic cation transporter; OAT, organic anion transporter; PGT, prostaglandin transporter; OST, organic solute transporter; MATE1, multidrug and toxin extrusion transporter 1; GSH, X− , glutathione conjugates; OA− , anionic anions or conjugates; OC+ , organic cations; BS− , bile salts; PG, prostaglandins; PC, phosphatidylcholine

In this chapter, we will first review what we have learned in the last few decades about the genetic profiles of hepatic tumors, with emphasis on the SLC uptake transporters and ABC transporters, two of the main contributors to MDR. In the second part, we will stress recent data indicating the critical role of polymorphisms in treatment success.

Role of transporters and phase I enzymes in hepatocytes (See also Chapters 21, 23, 24 and 43) Hepatic uptake transport: the solute carriers Uptake transport in hepatocytes is mediated by transporters belonging to the solute carrier (SLC) family,

which includes approximately 360 transporters classified in 45 gene families (classification of the SLC superfamily is outlined at http://www.bioparadigms.org/slc/menu.asp). Genes of the SLC superfamily encode passive transporters, ion-coupled transporters, and exchangers (reviewed by Hediger et al. [22]). In the liver, organic anion transporting polypeptides (OATPs) encoded by the solute carrier organic anion transporting polypeptide (SLCO) gene family, organic anion transporters (OATs) encoded by the SLC22 gene family, organic cation transporters (OCTs) also encoded by the SLC22 gene family, and Na+ -taurocholate cotransporting polypeptides (NTCPs) encoded by SLC10A1 are involved in the transport of many kinds of endogenous compounds and drugs (reviewed by Shitara et al. [23]). Na+ -dependent bile acid uptake is mediated by NTCP, whereas Na+ -independent bile acid uptake is mediated by OATPs [24].

66: THE HEPATOCYTE AND THE CANCER CELL: DR JEKYLL AND MR HYDE

SLCs

ABCs

GFRs GF

Integrins /

Uptake Efflux

Wnt

SHH

FZD

PTCH Smo

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CER

AB

Cs

Phase I & II enzymes*

Signal transduction pathways

Lysosome/ endosome

Detoxification

Drug compartmentalization

MAPK Ras

PI3K

Drug resistance Cell proliferation

Glucosylceramide ax

B

Ceramide

Apoptosis

Epigenetic modifications

l2

Bc

GCS Cell cycle checkpoints DNA repair genes

p53* pRb* Transcriptional

activation Drug-induced damage

HDAC

HP1

TFs

Figure 66.2 Multidrug resistance mechanisms in cancer cells. Following drug treatment, several mechanisms result in resistance to either a small number of related drugs or to a broad range of structurally and functionally unrelated drugs, which is known as MDR. The first response to drug treatment is the activation of signal transduction pathways through integrins, growth factors (GFs), Wnt/FZD, and sonic hedgehog (SHH/PTCH), which activate genes such as Bcl2, blocking apoptosis, and glucosylceramide synthase (GCS), affecting membrane lipids. Cancer cells also harbor mutations (*) that impair cell cycle checkpoints (e.g. p53, pRB) and phase I and II enzymes, resulting in increased expression of DNA repair genes and increased drug detoxification. Epigenetic modifications generally occur after drug treatment through activation of genes including HP1 and HDAC. Lastly, transporters play a critical role in MDR. While efflux pumps (i.e. ABC transporters, ABCs) are involved in drug compartmentalization, several of these transporters are often found to be over-expressed in the cell membrane, whereas uptake transporters (i.e. solute carriers, SLCs) are down-regulated. GFRs, growth factor receptors; Wnt, wingless; FZD, frizzled receptor; SHH, sonic hedgehog; PTCH, patched; Smo, smoothed; CER, ceramide; MAPK, mitogen-activated protein kinase; PI3 K, phosphatidyl inositol 3′ -kinase; TFs, transcription factors; HDAC, histone deacetylases; HP1, heterochromatin protein 1

In the OATP family, OATP1B1 (also known as liver-specific organic anion transporter-1 (LST-1)/ OATP-2/OATP-C, which is encoded by SLCO1B1 ), OATP1B3 (also known as LST-2/OATP-8, which is encoded by SLCO1B3 ), and OATP2B1 (also known as OATP-B, which is encoded by SLCO2B1 ) are expressed in the liver (reviewed in and [25, 26]). OATP1B1 mediates the uptake of not only bile acids but also conjugated steroids, thyroid hormones, eicosanoids, and some drugs, including 3-hydroxy-3-methylglutarylcoenzyme A (HMG-CoA) reductase inhibitor, pravastatin, and the anticancer drug, methotrexate [27]. OATP1B3 has a characteristic expression pattern: less expression in normal hepatic cells (compared with that of OATP1B1) and higher expression in various human cancer tissues and in different tumor cell lines derived from the

stomach, colon, pancreas, liver, gall bladder, and lung [28]. The substrate specificity of OATP1B3 is similar to that of OATP1B1, and includes methotrexate. The expression pattern and substrate specificity of OATP2B1 are different from those of OATP1B1 and 1B3. Although OATP2B1 is mainly expressed in the liver, it is distributed in a wide range of organs such as the spleen, placenta, lung, kidney, heart, ovary, small intestine, and brain. OATP2B1 does not transport bile acid but transports sulfobromophthalein, estrone-3-sulfate, and dehydroepiandrosterone sulfate [29]. OCT1 and OAT2 belong to the SLC22 transporter family and mediate the basolateral uptake of organic cations and anions (see [30]). Substrates of OCT1 include serotonin, prostaglandin E2, F2α, and drugs such as acyclovir. OCT1 also mediates uptake of platinum drugs

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[31]. Knowledge of these transporters, such as gene localization, tissue distribution, and substrate selectivity, has rapidly increased. The transporters exhibit genetic polymorphisms, which can be an obstacle in predicting pharmacological and toxicological effects. Recent advances suggest that OATPs could play important roles in explaining inter-individual variability in pharmacokinetics.

Hepatic efflux transport: the ABC transporters (See also Chapters 23, 24 and 43) ABC transporters are responsible for active efflux transport from the hepatocyte to the bile canaliculus through the canalicular/apical membrane and to the sinusoidal blood through the basolateral membrane. Apical membrane transporters include ABCB1 /Pgp (P-glycoprotein), ABCB4 /MDR3, ABCB11 /BSEP (bile-salt export pump), ABCC2 /MRP2 (multidrug resistance protein 2), ABCG2 /BCRP (breast cancer resistance protein), and the two half transporters ABCG5 and G8 . Under physiological conditions, the bile-salt export pump [ATP binding cassette (ABC)B11] transports monovalent bile acids [32, 33], ABCB4 exports phosphatidylcholine [34], ABCC2 mediates the translocation of divalent bile acids through a co-transport mechanism with reduced glutathione (GSH), glucuronate, or sulfate [35] and the ABCG5/G8 heterodimer exports cholesterol [36, 37]. ABCB1 and ABCG2 are involved in the protection of the organism by limiting the absorption of xenobiotics (see [38, 39]). ABCC2 is also known to efflux xenobiotics, leading to resistance to chemotherapeutics [40, 41]. Five members of the ABC (MRP) transporter C subfamily are localized in the basolateral membrane; ABCC1 /MRP1 , ABCC3 /MRP3 , ABCC4 /MRP4 , ABCC5 /MRP5 , and ABCC6 /MRP6 . These transporters mediate the excretion of organic anions from hepatocytes to sinusoidal blood. ABCC1 is barely expressed in adult liver cells [42] and is found primarily in intracellular vesicles [43]. This transporter can transport a broad range of organic conjugates, including steroids and bile salt (BS) conjugates [44]. ABCC11/MRP8 shares with ABCC3 the ability to transport monovalent BSs such as cholate, taurocholate, and glycocholate, and conjugated BSs [45]. Although the function of ABCC11 in the liver has not been undisputedly demonstrated, the presence of its transcripts in the liver with the recent determination of its substrates has led to speculation concerning its potential role in bile acid homeostasis [46, 47]. While the affinity of ABCC4 for sulfated BSs and steroids was demonstrated by Zelcer et al. [48], ABCC5 and C6 mediate transport of a wide range of conjugated organic anions. These transporters also have a critical role in multidrug resistance (MDR) (reviewed by Gillet et al. [13] and Szakacs et al. [14]).

Key role of CYP450, Phase I enzymes (See also Chapter 70) Once inside the cell, compounds are converted to metabolites by CYP enzymes, key players in the mechanisms of detoxification. An up-to-date database and nomenclature for CYP450 enzymes can be found at http://drnelson.utmem.edu/CytochromeP450.html [49]. These phase I metabolism enzymes comprise a superfamily of oxidases responsible for the oxidation of numerous endobiotics and thousands of xenobiotics. Although there are 57 P450 genes in the human genome, only 10 contribute to drug metabolism, with the main contribution coming from three isoforms, CYP3A4, CYP2D6, and CYP2C9 [18]. CYP enzymes metabolize endo- and xenobiotics into reactive species, substrates for phase II enzymes, which transform them into soluble non-toxic metabolite conjugates, further excreted through the bile, and sinusoidal blood. Phase II enzymes are involved in conjugation reactions including glutathionylation [50], glucuronidation [9], and sulfation [51]. In the last decade, studies have highlighted the synergism between CYP enzymes and ABC transporters, which can be considered as phase III in the detoxification system [52]. Many conjugated metabolites are substrates of members of the ABCC/MRP subfamily of ABC transporters [44]. This synergism may also occur when metabolites produced by CYP enzymes, especially CYP3A4, are better substrates for ABCB1 than the parent compound or when ABCB1 prolongs the duration of absorption by necessitating a subsequent entry of the compound/drug into the cell [53]. This process increases exposure to CYP enzymes and could prevent kinetic saturation of these proteins [54, 55]. The co-regulation of phase I and II metabolism enzymes via ligand-activated transcription factors, such as the aryl hydrocarbon receptor (AhR), the constitutive androstane receptor (CAR), the pregnane X receptor (PXR), and nuclear factor E2-related factor 2 (Nrf2), has been demonstrated ([52, 56]; see Chapter 22). Recently, Jigorel et al. reported a complex pattern of transporter regulation by xenobiotics in human hepatocytes, where efflux and uptake transporters are synergistically up- and down-regulated following drug administration through activation of ligand-activated transcription factors [8]. The interplay between not only ABCC but also OATP transporters and CYP enzymes was recently reviewed by Nies et al. [57].

Prevalence of liver cancer and challenges associated with treatment Hepatocellular carcinoma (HCC) accounts for approximately 80–85% of primary liver cancer, whereas intrahepatic cholangiocarcinoma (∼14%) and fibrolamellar carcinoma (∼1%) are the two other types of neoplasm that

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occur [58]. Liver cancer is the sixth most common cancer and the third most deadly cancer worldwide, with an overall mortality of ∼600 000 deaths per year [58]. In most cases, HCC arises as a consequence of underlying liver disease, usually a viral hepatitis [hepatitis B virus (HBV) or hepatitis C virus (HCV)] [59, 60]. However, cirrhosis from alcohol or non-alcoholic steatohepatitis, hereditary tyrosinemia, and primary hemochromatosis are other pathologies that predispose individuals to the development of HCC [59, 60]. The variety of diseases that give rise to liver cancer render the choice of treatment difficult and challenging; for an excellent review, see Llovet and Bruix [61]. Briefly, surgery is the mainstay treatment for patients with early-stage tumors. Either resection or transplantation is advocated if the HCC is within the Milan criteria [62]. Non-surgical treatments, including percutaneous ethanol injection (PEI) [63], radiofrequency ablation (RFA) [63, 64], and transcatheter arterial chemoembolization (TACE) [65], are used as adjuvant therapy to surgery but also to treat unresectable HCC. Although a meta-analysis showed a beneficial survival effect for patients with intermediate HCC treated by chemoembolization/TACE using doxorubicin and cisplatin, the survival benefit of systemic chemotherapy for the treatment of liver cancer is marginal at best. Indeed, systemic administration of doxorubicin has been evaluated in more than 1000 patients in clinical trials. There is a partial response in only around 10% of the cases, without any evidence of survival advantage [66, 67]. Lastly, a phase III randomized placebo-controlled trial (the SHARP trial) demonstrated that sorafenib, a tyrosine kinase inhibitor, improves survival in patients with advanced HCC. This will likely be established as the first line of therapy for advanced HCC [68, 69]. Further analysis will reveal whether sorafenib contributes alone or in combination with other chemotherapy to a decrease in the morbidity of liver cancer.

IMPACT OF GENE EXPRESSION PROFILING ON MOLECULAR CHARACTERIZATION OF HEPATOCELLULAR CARCINOMA The challenge of clinical oncology has been to target specific therapies to well-defined distinct cancer types to maximize efficacy and minimize toxicity of treatment. Significant advances in treatment are usually spurred on by technological developments. In this case, high-density DNA microarrays have revolutionized clinical oncology, unraveling the heterogeneity of cancers, and therefore refining the classification of many cancers that had previously been diagnosed on the basis of staging systems that did not incorporate biological information concerning the tumor [70–72]. A striking example was reported

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by Golub et al., who demonstrated the feasibility of cancer classification based solely on gene expression [73]. They distinguished between acute myeloid leukemia (AML) and acute lymphoblastic leukemia (ALL) without previous knowledge of these classes [73]. A number of studies subsequently confirmed the valuable contribution of genome-wide analyses of various tumors, including HCC (reviewed by Dunphy [74], Thorgeirsson et al. [75], and Quackenbush [76]).

Lack of consensus on reliable clinical staging systems and gene signatures Classification of HCC is hampered by similar issues brought to light in the leukemia and lymphoma field in the early 2000s. Numerous clinical staging systems, such as Barcelona-Clinic Liver-Cancer (BCLC) [77], the French model Groupe d’Etude de Traitement du Carcinoma Hepatocellulaire (GRETCH) [78], and Japan Integrated Staging (JIS) [79], have been suggested to predict patient survival with HCC, but none of these models predicts outcome accurately. Although some studies have claimed that the BCLC staging systems are superior [80–82], the current literature is conflicting. Indeed, while some studies did not report the superiority of any of the current clinical staging models [83], others supported one staging system over the others [84–87]. These conflicting reports can be explained to some extent by the complexity of the pathology and lack of molecular information in the models used. Data have been generated in the last 8 years in an attempt to unravel the molecular pathogenesis of HCC (see Chapters 60, 61). Laurent-Puig et al. highlighted two groups of patients with good and poor prognosis based on chromosomal stability status and genetic mutations [88]. More recently, this group reported an unsupervised study of more than 100 HCC samples that identify six groups of patients according to transcription patterns, chromosomal stability, promoter methylation status, and genetic mutation analysis [89]. Ye et al. reported a supervised study that allowed classification of patients with metastatic HCCs and highlighted genes such as osteopontin as highly correlated with metastasis and poor patient survival [90]. The same researchers recently published a 20-microRNA (miRNA) signature that may assist in identifying patients with HCC who are likely to develop metastases [91]. Another miRNA-based study reported a 19-miRNA signature in HCC associated with patient survival from cirrhosis and hepatitis [92]. Lee et al. tested the prognostic value of gene signatures obtained from 91 HCC samples. Those tumors were sub-classified into two groups strongly associated with patient survival [93]. Recently, Wurmbach et al. analyzed 75 samples representing the stepwise carcinogenic process from preneoplastic lesions (cirrhosis and dysplasia) to HCC, including neoplastic stages (very early HCC to metastatic

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tumors) from patients with HCV infection [94]. The study revealed specific gene signatures that accurately reflect the pathological progression of disease at each stage [94]. For reviews on molecular profiling of HCC, see Villanueva et al. [95], Iizuka et al. [96] and Chapters 60, 61. Although genomic-based arrays have significantly impacted our understanding of the molecular pathogenesis of HCC, to date only a few studies have comprehensively integrated data from other sources [88, 89].

Integrative systems biology needed for classification and diagnosis Systems biology has the potential to refine the classification of HCC and improve diagnosis and prognosis (reviewed by Vivekanandan and Singh [97]). However, the success of such an integrative approach depends on our ability to “speak the same language.” Indeed, the variety of experimental systems developed to characterize cancers at a molecular level, such as complementary DNA (cDNA) expression profiling, comparative genomic hybridization (array CGH), promoter arrays, SNP arrays, and so on, has increased dramatically in the last 5 years. In addition, the variety of platforms proposed for each of those analytical systems is remarkable. The multitude of normalization processes proposed also renders an integrative computational and analytical approach extremely challenging [98]. A combined database would also serve as an analytical tool that could compute the degree of overlap between global signatures and specific signatures imported by a researcher, comparing his or her findings with standardized data [98]. Rhodes et al. developed Oncomine, a bioinformatics initiative aimed at collecting, standardizing, analyzing, and delivering cancer transcriptome data to the scientific community [99]. The first version of this database was released in 2003 and contained 40 microarray data sets and nearly 100 differential expression analyses, allowing users to query differential expression results for a gene of interest across collected data sets [100]. The current version compiles 18 000 cancer gene expression experiments, and automated analysis has identified gene networks activated and repressed in human cancers (http://www.oncomine.org) [99]. In addition, they further developed the concept of molecular signature maps by demonstrating their utility in generating hypotheses that link cancer types and subtypes, pathways, mechanisms, and drugs [101]. Similar resources have arisen focusing specifically on HCC. They include EHCO (Encyclopedia of Hepatocellular Carcinoma Genes Online), a repository of genes found relevant in HCC [102], and OncoDB.HCC, an integrated oncogenomic database of HCC [103]. For additional information, the reader is directed to the websites http://ehco.iis.sinica.edu.tw/ and http://oncodb.hcc.ibms.sinica.edu.tw/index.htm. A considerable amount of data has been generated that has significantly improved our understanding of HCC

pathobiology. We are now at a crossroads, where we need integrative approaches to extract biological insights efficiently from vast datasets generated on genomic, proteomic and metabolomic levels. As discussed in Chapters 60, 61 [104], this approach will further help not only to stratify patients into clinically homogeneous groups, but also to uncover the origins of tumor cells and distinct pathways involved in the molecular pathogenesis of HCC.

ROLE OF MULTIDRUG RESISTANCE IN THE INTRACTABILITY OF HCC As mentioned earlier, the survival benefit of systemic chemotherapy in the treatment of liver cancer is only marginal at best. The reason for such a high level of resistance to drug treatment has been solely explained by the over-expression of ABCB1/MDR1, an ABC transporter cloned in 1987 [105]. Since that time, a vast amount of data has been generated, bolstering our understanding of ABC transporter functions in tumor resistance and concerning the multifactorial nature of those mechanisms (see Figure 66.2; recently reviewed by Mimeault et al. [106]). With regard to the liver, an MDR-centered study, in the strict sense, has not been performed. Instead, uptake and efflux transporters have been studied for their pivotal role in hepatobiliary elimination through ADMET (absorption, distribution, metabolism, excretion, and toxicity) profiles of drug candidates involved in the drug discovery process. In this section, we stress the role of two transporter superfamilies in MDR mechanisms. We briefly review the emerging role of SLCs, drug uptake transporters, in MDR mechanisms and their role in clinical drug resistance. We also review the current knowledge on the involvement of ABC transporters in those mechanisms.

The role of solute carriers (SLCs) in multidrug resistance (see also chapter 21) Cellular entry represents the first step in the mechanism of action of anticancer agents. Transporter-related MDR could result from not only increased efflux of drugs mediated by ABC transporters but also reduced drug uptake. The net accumulation of an anticancer drug in a cell is probably influenced by the concurrent actions of uptake and efflux transporters. Relative differences in uptake and efflux might contribute to drug resistance and be a primary factor in the differential response of various tumor types to the drugs. In cancer cells, studies have shown that some uptake transporters belonging to SLC families confer sensitivity to anticancer drugs [28, 107–112]. OATP1B3, which is expressed predominantly at the basolateral (sinusoidal) surface of hepatocytes, and has

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been shown to be expressed in various human cancer tissues and also in different tumor cell lines including HCC, confers sensitivity to methotrexate in cancer cells [107]. Kullak-Ublick and co-workers reported the differential expression of OATP1B3 and OATP1B1 in HCC [113]. They showed inhibition of OATP1B3 expression in HCCs that over-express hepatocyte nuclear factor 3β (HNF3β), one of several liver-enriched transcription factors shown to be differentially expressed in HCC compared with non-tumor liver tissue. Some studies have shown that OCT1, OCT2, and OCT3 mediate cell sensitivity to platinum drugs such as cisplatin, carboplatin, and oxaliplatin [31, 109–111]. OCT1 was also shown to mediate the uptake of imatinib [108]. A clinical study conducted by Crossman et al. showed that the expression level of OCT1 messenger RNA (mRNA) prior to treatment with imatinib in non-responders was only one-eighth of that seen in responders [114]. Dose escalation of imatinib could overcome resistance to standard-dose therapy in patients with chronic myeloid leukemia (CML), as reported by Kantarjian et al., which indicates that intracellular concentration of imatinib might be crucial for the therapy of CML [115]. Knowing whether these uptake transporters are expressed and functional in specific cancers can be a powerful tool to predict response to specific therapies. Okabe et al. used a bioinformatic approach to identify SLC substrates [112]. mRNA expression of 28 members of the SLCO and SLC22 family in 60 diverse cancer cell lines (the NCI-60) used by the National Cancer Institute (NCI) to screen for anticancer activity was profiled. By correlating expression profiles with growth inhibitory profiles of 1429 compounds (including anticancer drugs and drug candidates) tested against the cells, it was confirmed that OCTN1/SLC22A4 confers sensitivity to doxorubicin in cancer cells [112]. Unfortunately, cell lines derived from HCC are not included in the NCI-60 panel. Further study is needed to elucidate the gene and/or protein expression of SLCs in the HCC cell lines and clinical samples.

ATP-binding cassette (ABC) transporters mediate multidrug resistance The ABC transporter encoding genes are widely dispersed in the genome and show a high degree of sequence identity among eukaryotes. For extensive reviews, see Gillet et al. [13] and Szakacs et al. [14]. The ABC family includes 48 members divided into seven sub-families. The nomenclature for human ABC transporter genes is provided at the website http://nutrigene.4t.com/humanabc.htm. In the late 1990s, tumor resistance to therapy was correlated with expression of three ABC transporters, namely ABCB1/MDR1 [20, 105], ABCC1/MRP1 [116], and ABCG2 [117]. For recent reviews, see Callaghan et

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al. [38], Deeley et al. [44], and Polgar et al. [39]. To date, 13 ABC transporters (ABCA2, ABCB1, ABCB4, ABCB11, ABCC1–6, ABCC11–12, and ABCG2) were associated with drug resistance. For reviews, see Gillet et al. [13] and Szakacs et al. [14]. The determination of the expression profiles of ABC transporter genes in multidrug-resistant cell lines has opened up new avenues for the diagnosis of MDR in the clinic and for monitoring expression profiles in clinical biopsies and their correlation to clinical treatment. Four microarray-based assays for the detection of ABC transporter genes have been developed [118–121]. Annereau et al. developed a high-density microarray platform containing probes specifically matching 36 ABC transporters and also 70-mer oligonucleotides, allowing the analysis of 18 000 unique human genes [118]. Huang et al. developed a similar oligonucleotide microarray [120]. However, large amounts (25 and 12.5 µg, respectively) of total RNA were required for reverse transcription to run each array. Gillet et al. developed a low-density DNA microarray for profiling the expression of 38 ABC transporter genes which required less total RNA [119]. Finally, Liu et al. designed a semi-quantitative assay to detect the expression of 47 ABC transporter genes [121]. This last approach has two main drawbacks: it is not reliable for the detection of moderate changes in expression levels, and is not applicable for quantitative detection of abundant mRNAs [121]. These technologies require relatively large amounts of sample and often have poor probe specificity for individual transporters with highly homologous gene family members, such as those involved in MDR. Quantitative real-time reverse transcriptase polymerase chain reaction (qRT-PCR) has emerged as a fast and sensitive detection method that allows reproducible quantification of very low amounts of total RNA. Two approaches have been used to quantify 47 human ABC transporters in tissue samples and tumor cell lines. The first approach, developed by Langmann et al. [122], was based on qRT-PCR using Taqman chemistry, whereas Szakacs et al. measured ABC gene expression levels by qRT-PCR using SYBR Green [123]. Although these techniques are more reliable and accurate than microarrays, they are tedious and require multiple pipetting steps, which can introduce variability. Langmann et al. recently developed an assay to quantify 47 human ABC transporters using Taqman chemistry in a high-throughput platform termed Taqman low-density array [124]. These studies suggest that more than 25 ABC transporters can be involved in chemotherapy-induced resistance [119, 123, 125–127]. The role played by ABC transporter genes in clinical treatment and tumor recurrence is a subject of debate. This is well illustrated by experimental and clinical studies related to drug-mediated resistance of leukemia patients. Leukemia has been used as a model disease in some studies (reviewed by Ross [128], van den Heuvel-Eibrink et al. [129], and Hirose et al. [130]). In contrast to adult AML, for which the role of ABCB1 gene expression in

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the drug resistance of tumors and prognosis of patients is widely accepted [131, 132], the data for adult ALL are conflicting [132–134]. Therefore, ABC transporters other than ABCB1 may have an effect on treatment response and prognosis of adult ALL. The contribution of the ABCC1, ABCC3, and ABCG2 transporters to treatment outcome of adult ALL patients has been shown by some, but not all, authors [135, 136], leaving the prognostic relevance of these ABC transporters open to discussion. In solid tumors, most attention has been directed to the role played by the ABC transporter proteins in MDR observed in breast cancers, particularly by ABCB1, ABCC1, and ABCG2. However, it is difficult to define their exact role in the clinical drug resistance observed in this disease [137, 138]. ABCG2 has been detected in at least one drug-resistant breast tumor cell line [117]. Like ABCB1 and C1, its relevance to clinical drug resistance in breast tumor is still disputed [139–141] (reviewed by Polgar et al. [39] and Robey et al. [142]). Several ABC transporters expressed in tumors that have not yet been linked to drug resistance could have prognostic relevance. This view is supported by data published by Vitale et al. exploring the expression of ABC transporters associated with antigen processing (TAP), namely TAP1 (ABCB2) and TAP2 (ABCB3) [143]. In a collection of five specimens of normal mammary tissue and 53 primary breast carcinoma lesions, TAP1 and TAP2 expression was significantly associated with tumor grading. Like normal mammary tissue, the low-grade (G1) breast carcinoma lesions showed strong staining for TAP1 and TAP2. In contrast, only a few of the high-grade (G2 and G3) breast carcinoma lesions displayed a normal expression pattern. These data demonstrate an association of human leukocyte antigen (HLA) class I antigen and TAP down-regulation with tumor progression in breast carcinoma, and suggest that loss of TAP may represent a mechanism employed by cancers to escape the host’s immune pressure or may reflect accumulation of abnormalities associated with neoplastic progression. Accumulation of defects in antigen processing and presentation may be responsible for reduced recognition of malignant cells by putative clinically relevant tumor-specific T cells. Recent data obtained on ABCB5 suggested that this transporter could be a marker of melanoma cancer-initiating cells [144]. Cancer-initiating cells, also known as cancer stem cells, might underlie the intractable nature of many human cancers, explaining why conventional cancer therapy fails in many patients. For a review, see Alkatout et al. [145]. Although the concept is exciting, knowledge of ABCB5 from the genomic to the proteomic level is rudimentary, and does not support this hypothesis. Although considerable effort has been made to understand the role of ABC transporters in clinical samples, attempts to transform those transporters into clinical targets have been unsuccessful. However, expression of many ABC transporters in HCC is at levels sufficient to confer drug resistance and may contribute to

resistance without being the limiting determinants, especially given the extraordinary variety of drug-resistance genes expressed in HCC.

IMPORTANCE OF PHARMACOGENETICS IN MULTIDRUG RESISTANCE The introduction of pharmacogenetics to the clinic is now proposed to individualize therapy [146, 147]. Indeed, inter-individual differences in drug response are major causes of adverse drug reaction and drug treatment failure. Drug bioavailability is dependent on an individual’s expression of drug transporters, such as ABC transporters or uptake transporters. Individual variations in drug plasma level are also, in part, explained by identification of single nucleotide polymorphisms (SNPs) revealing distinct phenotypes of drug-metabolizing enzymes.

Genetic variation in ABC transporters A number of recent reports have addressed genetic polymorphisms in drug transporters; for an excellent review, see Cascorbi [148]. Among the 48 ABC transporters, ABCB1 is one of the best studied and characterized, with more than 50 SNPs reported [148–150]. Hoffmeyer et al. were the first to show association of the synonymous SNP 3435C > T in exon 26 with decreased expression of ABCB1 in the duodenum of patients with the T allele (variant) compared with those with the C allele (wild-type) [151]. However, a subsequent study reported that this effect might be due to the non-synonymous SNP 2677G > T/A [152], which is frequently linked to C3435T. In contrast to these earlier reports, Gerloff et al. reported no changes in digoxin clearance between Caucasian patients carrying the variant or the wild-type allele [153]. Elevated ABCB1 expression was reported in Japanese and Caucasian patients carrying the variant allele (3435T) [154, 155]. The non-synonymous SNP 2677G > T/A is also subject to conflicting data [156]. Recently Kimchi-Sarfaty et al. analyzed the role of synonymous mutations in protein folding and function [10]. Synonymous SNPs (3435C > T, 1236C > T) and non-synonymous 2677G > T in the ABCB1 gene sequence result in a protein with altered drug and inhibitor interactions, without a change in expression levels, possibly due to altered protein folding related to a change in the rhythm of translation [10]. Based on our current knowledge, overall drug bioavailability is only moderately influenced by ABCB1 polymorphisms, as compared with variants of the drug-metabolizing enzymes (CYP family) [157, 158]. Although the findings for ABCC1, showing

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a high rate of polymorphisms, are similar to those for ABCB1, studies on ABCG2 report a significant effect of polymorphisms on ABCG2 expression and function [147, 150, 159]. The correlation of ABC transporter genetic variants with treatment outcomes is gradually being clarified, yet the overall picture is still puzzling, as much of the published data are conflicting. Nevertheless, many studies reporting correlation between SNPs and clinical outcome indicate the necessity to pursue further investigations. Initiatives such as the Pharmacogenetics Research Network could aid the development of this complex field. One of their various goals is to understand how genetic variation in membrane transporters contributes to variation in drug transport [160].

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SLC22A4 is expressed in various cancer cell lines and mediates uptake of the anticancer agent doxorubicin [112]. As described above, the OCT1 (SLC22A1 ), OCT2 (SLC22A2 ), and OCT3 (SLC22A3 ) transporters may mediate uptake of some platinum anticancer drugs [31, 109–111]. Although genetic variants of SLC22A1 and SLC22A2 showed altered transport of some of their substrates, such as metformin, which is used in therapy for type 2 diabetes mellitus [166, 167], their transport of platinum anticancer drugs has not been investigated. Unlike ABC transporters, SLCs have not been intensively focused on as candidate transporters for anticancer drugs. There is little information about the association between MDR and SNPs in the SLCs.

Genetic variation in solute carriers

CONCLUSION –DR JEKYLL OR MR HYDE?

Among the SLCs, the OCTs, OATs, and OATPs have been proposed to have critical roles in the absorption, distribution, and excretion of xenobiotics and endogenous compounds in the liver, kidney, central nervous system, intestine, and other tissues. Genetic polymorphisms of these transporters may alter drug pharmacokinetics, triggering interindividual differences in the safety and efficacy of drug therapy. OATP1B1, which is encoded by SLCO1B1 , is expressed abundantly in the liver and mediates the hepatic uptake of a broad range of organic ions, has been well studied concerning SNPs. For an excellent review, see Niemi [161]. Certain SNPs (e.g. SLCO1B1 -388A > G, 521T > C, 578T > G) have been shown to affect its surface expression and/or function in vitro and/or in vivo in humans. However, no data exist on the consequences of SLCO1B1 variants in MDR. A number of sequence variants have been reported in OATP1B3 (SLCO1B3 ), also expressed in the liver and in cancers. Letschert et al. demonstrated that SLCO1B3 -1564G > T, encoding OATP1B3-G522C, abolished the transport of bile acid but not other substrates [162]. OATP1B3 was shown to mediate uptake of paclitaxel. Investigation of the functional consequences of mutation in SLCO1B3 showed that paclitaxel pharmacokinetics were not associated with SLCO1B3 -344T > G or 699G > A [163]. OCTN1, encoded by SLC22A4 , is expressed mainly in the kidney and is thought to be a transporter responsible for renal disposition of cationic compounds. An SNP of SLC22A4 that produces the amino acid mutation L503F is associated with risk for developing Crohn’s disease [164]. The effects of the 503F variant on SLC22A4 specificity may diminish uptake of physiological compounds while increasing uptake of potential toxins. One intronic SNP in SLC22A4 also showed strong association with rheumatoid arthritis, which, like Crohn’s disease, has a pathogenesis associated with inflammation and autoimmunity [165].

In the Introduction to this chapter, we characterized hepatocytes through an analogy to the story of Dr Jekyll and Mr Hyde. Do these cells really have a “split personality?” We have learned a great deal in the last decade with the development of high-throughput genomic profiling systems and unraveling of the human genome sequence. These breakthroughs have provided new insights in cancer research, revolutionizing current classification of almost all cancers and their clinical management. HCC is a complex and heterogeneous disease. Genomic expression profiling has focused on the predictive power of gene signatures for overall survival (see Chapters 60, 61 and [88, 89, 93]). Attention has also been directed towards gene signatures associated with the carcinogenic process, from preneoplastic lesions to neoplastic stages, including very early HCC to metastatic tumors exemplified by findings reported by the Llovet group [94]. The field of ABC transporter-mediated drug resistance has also been revolutionized by technological and scientific advances. From 13 ABC transporters known to be involved in MDR, now more than 25 transporters are suggested to be involved in tumor resistance. However, a red flag has been raised regarding the study of gene families with a high degree of homology, such as ABC transporter genes. The lack of specificity and sensitivity of platforms used to profile expression of ABC transporters is most likely one reason for observed discrepancies. In work in progress in our laboratory [168], we are evaluating three unique gene-expression profiling technologies to ascertain which technology provides the best tool for drug discovery and has potential for clinical applications in personalized medicine. We determined that Taqman low-density arrays (TLDAs) are the most sensitive and selective in measuring ABC transporter gene-expression patterns in a group of intensively studied cancer cell lines. An additional major concern raised in this chapter is standardization of reported gene signatures. How can

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Figure 66.3 Systems biology model for HCC study. This model proposes four starting points: blood samples, tumor biopsies, normal liver, and in vivo–in vitro models. Technological advances allow the study of all the cellular levels from the genomic to the metabolomic through various types of microarrays, 2D electrophoresis, mass spectrometry, and so on. The challenge is now the development and expansion of integrative tools that will help to extract thoroughly all the insights of those studies. The capsules in bold type indicate the key research objectives of the studies

scientists communicate with each other without a common language? In this regard, leading research institutions have the duty to promote standardization of the data reported. A database such as Oncomine, a bioinformatics initiative aimed at collecting, standardizing, analyzing, and delivering cancer transcriptome data to the scientific community, is invaluable. Considerable data have been compiled on multiple biological levels. We need now to develop tools allowing integrative approaches to extract biological insights efficiently from vast datasets generated at the genomic, proteomic, and metabolomic levels. There is a relative paucity of research on the intractability of HCC from the perspective of MDR. Can the malignant transformation of liver cells be accompanied by multifactorial MDR gene expression? Does the genetic background of hepatocytes predispose them to transformation (i.e. in the role of Mr Hyde), or are those genes irrelevant to the pathobiology of HCC, protecting the organisms (i.e. in the role of Dr Jekyll)? Further investigation is needed to answer these questions. Figure 66.3 proposes a systems biology model for further analysis of

HCC based on what is currently in the literature and the authors’ vision of multidimensional biology now possible in the post-genomic era.

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130. Hirose, M., Hosoi, E., Hamano, S. and Jalili, A. (2003) Multidrug resistance in hematological malignancy. J Med Invest , 50, 126–35. 131. Marie, J.P. and Legrand, O. (1999) MDR1/P-GP expression as a prognostic factor in acute leukemias. Adv Exp Med Biol , 457, 1–9. 132. Wuchter, C., Leonid, K., Ruppert, V. et al. (2000) Clinical significance of P-glycoprotein expression and function for response to induction chemotherapy, relapse rate and overall survival in acute leukemia. Haematologica, 85, 711–21. 133. Tafuri, A., Gregorj, C., Petrucci, M.T. et al. (2002) MDR1 protein expression is an independent predictor of complete remission in newly diagnosed adult acute lymphoblastic leukemia. Blood , 100, 974–81. 134. Wattel, E., Lepelley, P., Merlat, A. et al. (1995) Expression of the multidrug resistance P glycoprotein in newly diagnosed adult acute lymphoblastic leukemia: absence of correlation with response to treatment. Leukemia, 9, 1870–74. 135. Suvannasankha, A., Minderman, H., O’Loughlin, K.L. et al. (2004) Breast cancer resistance protein (BCRP/MXR/ABCG2) in adult acute lymphoblastic leukaemia: frequent expression and possible correlation with shorter disease-free survival. Br J Haematol , 127, 392–98. 136. Steinbach, D., Wittig, S., Cario, G. et al. (2003) The multidrug resistance-associated protein 3 (MRP3) is associated with a poor outcome in childhood ALL and may account for the worse prognosis in male patients and T-cell immunophenotype. Blood , 102, 4493–98. 137. Larkin, A., O’Driscoll, L., Kennedy, S. et al. (2004) Investigation of MRP-1 protein and MDR-1P-glycoprotein expression in invasive breast cancer: a prognostic study. Int J Cancer, 112, 286–94. 138. Leonessa, F. and Clarke, R. (2003) ATP binding cassette transporters and drug resistance in breast cancer. Endocr Relat Cancer, 10, 43–73. 139. Burger, H., Foekens, J.A., Look, M.P. et al. (2003) RNA expression of breast cancer resistance protein, lung resistance-related protein, multidrug resistance-associated proteins 1 and 2, and multidrug resistance gene 1 in breast cancer: correlation with chemotherapeutic response. Clin Cancer Res, 9, 827–36. 140. Faneyte, I.F., Kristel, P.M., Maliepaard, M. et al. (2002) Expression of the breast cancer resistance protein in breast cancer. Clin Cancer Res, 8, 1068–74. 141. Kanzaki, A., Toi, M., Nakayama, K. et al. (2001) Expression of multidrug resistance-related transporters in human breast carcinoma. Jpn J Cancer Res, 92, 452–58. 142. Robey, R.W., Polgar, O., Deeken, J., To, K.W. and Bates, S.E. (2007) ABCG2: determining its relevance in clinical drug resistance. Cancer Metastasis Rev , 26, 39–57. 143. Vitale, M., Rezzani, R., Rodella, L. et al. (1998) HLA class I antigen and transporter associated with antigen processing (TAP1 and TAP2) down-regulation in

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159. Conseil, G., Deeley, R.G. and Cole, S.P. (2005) Polymorphisms of MRP1 (ABCC1) and related ATP-dependent drug transporters. Pharmacogenet Genomics, 15, 523–33. 160. Giacomini, K.M., Brett, C.M., Altman, R.B. et al. (2007) The pharmacogenetics research network: from SNP discovery to clinical drug response. Clin Pharmacol Ther, 81, 328–45. 161. Niemi, M. (2007) Role of OATP transporters in the disposition of drugs. Pharmacogenomics, 8, 787–802. 162. Letschert, K., Keppler, D. and Konig, J. (2004) Mutations in the SLCO1B3 gene affecting the substrate specificity of the hepatocellular uptake transporter OATP1B3 (OATP8). Pharmacogenetics, 14, 441–52. 163. Smith, N.F., Marsh, S., Scott-Horton, T.J. et al. (2007) Variants in the SLCO1B3 gene: interethnic distribution and association with paclitaxel pharmacokinetics. Clin Pharmacol Ther, 81, 76–82. 164. Peltekova, V.D., Wintle, R.F., Rubin, L.A. et al. (2004) Functional variants of OCTN cation transporter genes

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are associated with Crohn disease. Nat Genet , 36, 471–75. Tokuhiro, S., Yamada, R., Chang, X. et al. (2003) An intronic SNP in a RUNX1 binding site of SLC22A4, encoding an organic cation transporter, is associated with rheumatoid arthritis. Nat Genet , 35, 341–48. Wang, D.S., Jonker, J.W., Kato, Y., Kusuhara, H., Schinkel, A.H. and Sugiyama, Y. (2002) Involvement of organic cation transporter 1 in hepatic and intestinal distribution of metformin. J Pharmacol Exp Ther, 302, 510–15. Kimura, N., Masuda, S., Tanihara, Y. et al. (2005) Metformin is a superior substrate for renal organic cation transporter OCT2 rather than hepatic OCT1. Drug Metab Pharmacokinet , 20, 379–86. Orina, J.N., Calcagno, A.M., Wu, C.-P. et al. (2009) Evaluation of current methods used to analyze the expression profiles of ABC transporters yields an improved drug-discovery database. Mol Cancer Ther, 8, 2057–66.

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The Role of Endocannabinoids and Their Receptors in the Control of Hepatic Functions George Kunos, Douglas Osei-Hyiaman, S´andor B´atkai, P´al Pacher, Bin Gao, Won-Il Jeong, Jie Liu and Gregorz Godlewski National Institute on Alcohol Abuse and Alcoholism, National Institutes of Health, Bethesda, MD, USA

THE ENDOCANNABINOID SYSTEM The recreational and medicinal use of marijuana goes back thousands of years, yet it is only in the last few decades that we have begun to understand the physiological underpinnings of its effects, many of which have important therapeutic potential. Progress in our understanding of the biology of cannabinoids (CBs) has been marked by important milestones. The resinous exudate of hemp contains more than 60 different CB molecules, among which 9 -tetrahydrocannabinol (THC) is responsible for its psychoactive properties. The first major milestone in contemporary CB research was the resolution of the correct chemical structure of THC in 1964 by Gaoni and Mechoulam (Figure 67.1a) [1]. This has allowed the synthesis of structurally modified analogs, which could then be used to study structure–activity relationships. Such The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

studies revealed tight structural and steric selectivity in the biological actions of CBs, hallmarks of drug–receptor interactions. This was unexpected, because prevailing dogma at the time held that the hydrophobic CBs act by non-specifically perturbing the membrane lipid environment. Once the notion of specific receptors for CBs had emerged, it did not take very long before the existence of such receptors in the brain was established, first by radioligand studies [2], then by molecular cloning [3]. The brain-type CB receptor was later named CB1 to distinguish it from a second receptor, CB2 , which was first identified in lymphoid tissues [4]. Both CB receptors belong to the G protein-coupled receptor family and signal through Gi /Go proteins, although they can also activate G protein-independent signaling pathways [5]. CB1 receptors are expressed at very high levels in the brain [6], where they are localized almost exclusively presynaptically on both excitatory and inhibitory nerve terminals [7], but are also present at much lower

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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Figure 67.1 The structure of prototypical cannabinoid ligands. (a) 9 -Tetrahydrocannabinol (THC), (b) endogenous cannabinoids, and (c) CB1 receptor antagonists that have been tested or used in humans

yet functionally relevant levels in many peripheral tissues, including the liver [8]. CB2 receptors are expressed predominantly on cells of the immune and hematopoietic systems, but they too can be detected in other tissues, including the liver [9] and the brain [10, 11], where their expression is often induced in pathological conditions [9, 12]. CB1 and CB2 receptors display a low level of homology, yet their pharmacology is remarkably similar,

with most plant-derived and early synthetic ligands having similar affinities to both. It is only in the last fifteen years that agonist and antagonist ligands highly selective for CB1 or CB2 receptors have been developed [27], [28]. In recent years, functional evidence for additional CB receptors has been accumulating [13]. One such receptor may be the orphan G protein-coupled receptor (GPCR) Gpr-55, which recognizes certain CB ligands with high affinity, and utilizes signaling pathways distinct from those

67: THE ROLE OF ENDOCANNABINOIDS AND THEIR RECEPTORS IN THE CONTROL OF HEPATIC FUNCTIONS

activated by CB1 or CB2 [14–16]. The physiological function of this receptor remains to be established. The existence of specific receptors for plant-derived substances in mammalian cells raised the question of endogenous ligands. The first endogenous CB ligand or endocannabinoid to be identified, in 1992, was arachidonoylethanolamide, named anandamide from the Sanskrit term for bliss (Figure 67.1b) [17]. This was followed 3 years later by the discovery of a second endocannabinoid, 2-arachidonoylglycerol(2-AG, Figure 67.1b) [18, 19]. Although a number of additional related endogenous ligands have been identified [20], to date these two remain the most extensively characterized. Both anandamide and 2-AG are generated on demand from membrane phospholipid precursors, in response to a rise in intracellular calcium or metabotropic receptor activation [20]. Their biosynthesis may proceed along multiple, parallel pathways (Figure 67.1) [21, 22], which would make blocking their endogenous generation difficult to achieve. Unlike classical neurotransmitters, they are not stored, and the mechanism of their release from cells is not yet clear. Even when released, they remain largely membrane associated due to their lipophilicity, and they are taken back up by cells through a high-affinity transport mechanism, following which they are metabolized. Anandamide is degraded primarily by the membrane-associated fatty acid amidohydrolase (FAAH) [23], whereas 2-AG is metabolized by monoglyceride lipase (MGL) [24]. Pharmacological inhibition or genetic ablation of FAAH results in a marked increase in tissue anandamide, but not 2-AG, levels, which may unmask certain behavioral or metabolic effects resulting from tonic activation of CB1 receptors by endogenous anandamide [25]. Selective in vivo effective inhibitors of MGL are not yet available. Endogenous CBs, their receptors, and the enzymes/ proteins involved in their biosynthesis, transport, and degradation collectively make up the endocannabinoid system (ECS). The introduction of potent and selective inhibitors of CB1 [26] and CB2 receptors [27], and the generation of mouse strains deficient in these receptors [28–30] have been the key tools for uncovering the biological functions of the ECS. There is a rapidly growing list of these effects, both centrally mediated and peripheral functions, many of which can be exploited therapeutically [20]. The presence and functional importance of the ECS in the liver [8] came as a surprise, as the dominant psychoactive properties and behavioral activity of CBs, combined with the very high level of expression of CB1 receptors in the brain, have suggested that it is primarily a neuronal signaling system. Indeed, early studies of brain CB1 receptors used the liver as a negative control. However, several recent reports have now documented the presence, albeit at low levels, of functional CB1 receptors in whole liver [8, 31], and also in various subtypes of liver cells such as hepatocytes [32, 33], stellate cells [34, 35], and hepatic vascular tissue including endothelial cells [36–39]. Although CB2 receptors are absent from or

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present at very low levels in normal liver, their expression appears to be induced in pathological conditions such as non-alcoholic fatty liver disease (NAFLD) [40], embryonic state [41], and liver fibrosis [9], and both CB1 and CB2 expression have been found to be up-regulated in human hepatocellular carcinoma [31]. Endocannabinoids are also present in the liver at levels comparable to those measured in brain [8, 35]. The functional relevance of the hepatic ECS is illustrated by recent findings that implicate it in the regulation of hepatic hemodynamics, fibrogenesis, and lipid metabolism, and in the dysregulation of these functions in pathological states such as cirrhosis, NAFLD, alcoholic fatty liver, and ischemia–reperfusion (I/R) injury. This will be discussed in some detail in the following sections.

ENDOCANNABINOIDS AND THE HEMODYNAMIC CONSEQUENCES OF CIRRHOSIS THC and its synthetic analogs have long been known to have powerful cardiovascular effects, the most striking being a long-lasting and profound decrease in blood pressure, which is mediated by CB1 receptors located in the peripheral cardiovascular system [42]. This triggered investigations into the potential involvement of endocannabinoids in hypotensive states, with subsequent findings implicating the ECS in the hypotension associated with various forms of shock [43, 44], including the hypotension induced by bacterial endotoxin or lipopolysaccharide (LPS) [45]. Advanced liver cirrhosis is often associated with endotoxemia and hypotension, and the analogy with LPS-induced hypotension suggested the potential involvement of endocannabinoids. This was then documented, using different rat models of cirrhosis. Rats with cirrhosis induced by bile duct ligation or carbon tetrachloride (CCl4 ) treatment develop progressive hypotension that can be acutely reversed by a bolus dose of the CB1 receptor antagonist rimonabant [36]. Rimonabant also reduced the pathologically elevated portal venous pressure and mesenteric blood flow (Figure 67.2), which implicated endocannabinoids in these changes. Circulating macrophages can generate endocannabinoids through a CD14-dependent, LPS-inducible mechanism [45, 46]. Indeed, macrophages isolated from cirrhotic rats or hypotensive cirrhotic patients with increased plasma endotoxin levels contained elevated amounts of anandamide compared with macrophages from healthy animals or volunteers, and elicited CB1 -mediated hypotension when injected into normal recipient rats, which could be prevented by pretreatment of the donor rats with the CB1 antagonist rimonabant [36, 47]. Target organ sensitivity was also increased, as indicated by a threefold increase in the expression of CB1 receptor messenger RNA (mRNA)

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Figure 67.2 The effect of the CB1 antagonist rimonabant (SR141716A, 3 mg kg−1 iv) on arterial blood pressure, portal venous pressure, and mesenteric blood flow in a urethane-anesthetized rat. Reproduced from [36] with permission

and protein in vascular endothelial cells isolated from cirrhotic versus non-cirrhotic human liver tissue [36], or in mesenteric arteries of cirrhotic rats [38, 48], and by the increased vasodilator potency of anandamide in mesenteric arteries from cirrhotic versus control rats [38, 48]. These findings implicate anandamide and vascular CB1 receptors in the vasodilated state that accompanies advanced liver cirrhosis. Because the vasodilated state and consequent increase in mesenteric blood flow increase the risk of a rupture of varicosities and also contribute to ascites formation, its reversal through CB1 blockade may have therapeutic value by delaying such potentially fatal complications, thus keeping patients alive until a transplant becomes available. Anandamide has been shown to decrease mesenteric vascular resistance through a rimonabant-sensitive, CB1 receptor-mediated, NO-independent mechanism [49]. However, at higher doses, the mesenteric vasodilator effect was resistant to CB1 blockade [49], which could suggest the existence of additional, CB1 -independent mechanisms. Indeed, the persistence of anandamide-induced vasorelaxation in mesenteric artery segments from CB1 knockout or CB1 /CB2 double knockout mice suggested the existence of an “anandamide receptor” distinct from CB1 and CB2 [13], and such a mechanism may also contribute to anandamide-mediated mesenteric vasodilation in cirrhosis.

The peripheral vasodilation in cirrhosis triggers reflex activation of the sympathetic nervous system resulting in a “hyperdynamic” circulation with tachycardia and increased cardiac output. However, this hyperdynamic circulation masks an underlying latent heart failure with reduced cardiac contractility and reduced responsiveness to β-adrenergic stimulation, a condition termed “cirrhotic cardiomyopathy” [50]. The results of a recent study have implicated endocannabinoids and CB1 receptors in this condition. Papillary muscle isolated from the heart of rats with bile-duct ligation-induced cirrhosis displayed reduced contractile responsiveness to isoproterenol, which could be corrected by CB1 blockade [51]. Exogenous anandamide also reduced the maximum response to isoproterenol, and this effect was similarly prevented by a CB1 receptor antagonist [51]. A subsequent study in rats with CCl4 -induced cirrhosis provided in vivo hemodynamic evidence for the role of cardiac anandamide and CB1 receptors in the reduced cardiac contractile function [52]. Detailed hemodynamic analyses through the use of an intraventricular pressure–volume microcatheter system revealed decreased baseline cardiac contractility in the cirrhotic rats, and its acute reversal by a single bolus dose of the CB1 antagonist AM251. Through the use of load-independent contractile parameters, a direct, CB1 -mediated decrease in contractile function of the cirrhotic heart could be

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Figure 67.3 Effects of the CB1 antagonist AM251 on load-independent contractile parameters. Representative pressure–volume (PV) loops obtained with a P–V conductance catheter system at different preloads generated by inferior vena cava occlusion. Note that the slope of the end-systolic P–V relation (ESPVR, dotted lines) is less steep in cirrhotic than control animals, indicating decreased contractile function, and is acutely increased following treatment of the cirrhotic rats with 3 mg kg−1 AM251. Reproduced from [52] with permission

unequivocally documented and separated from secondary effects due to the increased preload and decreased afterload present in the circulation, which account for the paradoxical increase in cardiac output. The key findings are illustrated in Figure 67.3. The mechanism underlying this endocannabinoid-mediated decrease in cardiac contractility have not yet been definitively identified. CB1 receptor activation inhibits L-type calcium channels [53] and reduces cAMP levels [5], which could both contribute to decreased contractility. The mediator involved is most likely anandamide, the tissue levels of which were increased about threefold in the heart of cirrhotic versus control rats, whereas the cardiac levels of 2-AG and expression of CB1 receptors were not affected by cirrhosis [52]. As discussed in the next section, endocannabinoids acting via hepatic CB1 receptors have recently been shown to contribute to the progression of liver fibrosis [34]. This would further strengthen the case for a therapeutic trial with CB1 antagonists in advanced cirrhosis. Such a treatment not only could correct the complex hemodynamic abnormalities and the associated risk for fatal complications, but also could delay the progression of the cirrhotic process itself.

ENDOCANNABINOIDS AND LIVER FIBROSIS A rather unique feature of the cannabinoid CB2 receptor is that although it is absent from most tissues under normal conditions, it is inducible by a variety of physiological or pathological stimuli. For example, CB2 receptors could not be detected in normal human liver, but they were strongly expressed in fibrotic tissue and non-parenchymal cells from specimens of cirrhotic liver, using immunohistochemistry [9]. These receptors were functional, as

THC was able to stimulate GTPγS labeling. THC was also antiproliferative and induced apoptosis in human hepatic myofibroblasts and stellate cells, an effect mimicked by selective CB2 but not CB1 agonists and inhibited by a CB2 antagonist, and were mediated via a COX-2 induced increase in oxidative stress [9]. As activated myofibroblasts and stellate cells are required for the development and progression of fibrosis, these findings suggested that CB2 receptor activation may be antifibrotic and hepatoprotective during the fibrotic process [54]. Indeed, mice lacking CB2 receptors were found to have an exaggerated response to fibrogenic stimuli [9]. CB2 receptor activation by anandamide was also found to inhibit the hyperplastic proliferation of cholangiocytes, which commonly occurs in extrahepatic biliary obstruction, cholestatic liver diseases, and toxic liver injury. This inhibitory effect was mediated through the induction of the AP-1 complex and thioredoxin-1, resulting in increased reactive oxygen species and cell death [55]. In a subsequent study in rats with CCl4 -induced cirrhosis and ascites, daily treatment for 9 days with the CB2 -selective agonist JWH-133 resulted in a general improvement of the fibrotic condition. This was reflected in a reduction in the number of activated stellate cells, increased apoptosis of non-parenchymal cells, and reduced fibrosis. The last effect was reflected by reduced α-smooth muscle actin and collagen I and increased matrix metalloproteinase-2 expression [56]. Together, these findings present a strong case for the therapeutic use of non-psychoactive CB2 agonists in liver fibrosis. Unexpectedly, an epidemiological study of 270 patients with hepatitis C virus infection demonstrated that daily cannabis use was a risk factor for, rather than protective against, fibrosis progression in these patients [57]. In view of the hepatoprotective effect of CB2 activation described above, these findings suggested that CBs may also exert

THE LIVER: ENDOCANNABINOIDS AND HEPATIC STEATOSIS

ENDOCANNABINOIDS AND HEPATIC STEATOSIS Non-alcoholic fatty liver and related hormonal/metabolic changes Endocannabinoids acting at CB1 receptors in the hypothalamus, limbic forebrain, and brainstem stimulate appetite and are part of the leptin-regulated hypothalamic appetitive circuitry [20, 58, 59]. Although this may contribute to the recently documented effectiveness of CB1 antagonists in the treatment of obesity [60], mice deficient in CB1 receptors are resistant to diet-induced obesity, hepatic steatosis, and the associated hormonal/metabolic changes, even though their caloric intake is similar to that in wild-type mice [8, 61]. This suggested that the ECS must have effects on energy balance independent of its effect on appetite. Recently emerging evidence indicates that obesity induced by high fat diets is associated with an increase in de novo lipogenesis, despite the increased availability of dietary fats [8, 62–64]. As the liver is a major source of de novo lipogenesis, the possible role of the ECS in the regulation of hepatic lipogenesis was explored [8]. Briefly, activation of CB1 receptors

SR141716 + HU210

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in mice was found to increase the hepatic gene expression of the lipogenic transcription factor sterol regulatory element-binding protein-1c (SREBP-1c) and its targets acetyl-CoA carboxylase-1 (ACC1) and fatty acid synthase (FAS) (Figure 67.4). Treatment with a CB1 agonist also increased de novo fatty acid synthesis in the liver

SREBP1c ACC1 FAS 18S rRNA

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an opposite, profibrotic effect, which would be dominant in the case of THC and would most likely be mediated by CB1 receptors. Accordingly, in a subsequent study the same group was able to demonstrate the pro-fibrotic function of CB1 receptors in the liver [34]. CB1 receptors were found highly induced in stellate cells and hepatic myofibroblasts in human cirrhotic liver specimens, and in three different models of experimentally induced hepatic fibrogenesis in mice. Furthermore, daily treatment with the CB1 antagonist rimonabant protected mice against CCl4 -, thioacetamide-, or bile duct ligation-induced liver injury, as indicated by the reduced expression of smooth muscle actin and transforming growth factor-β, and genetic ablation of CB1 receptors had a similar protective effect [34]. Although cellular endocannabinoid levels were not measured in this study, the likely fibrogenic mediator is 2-AG, the hepatic levels of which have been shown to be preferentially increased by CCl4 treatment of mice [35] and rats [52]. In another study, 2-AG was reported to cause stellate cell apoptosis in vitro through a CB-receptor independent mechanism [35], which could suggest that 2-AG is antifibrotic [35]. However, this effect is unlikely to occur in vivo, as it required high micromolar concentrations well above those measured in cirrhotic livers. The profibrotic effects of CB1 receptor activation in the liver, along with CB1 receptor-mediated deleterious hemodynamic effects in cirrhosis, would provide a strong rationale for clinical trials with CB1 antagonists for the medical management of advanced liver cirrhosis.

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Figure 67.4 Activation of CB1 receptors increases the gene expression of SREBP-1c, acetyl-CoA carboxylase-1 (ACC1), and fatty acid synthase (FAS) in mouse liver. Mice were injected i.p. with vehicle, 20 ng g−1 of the CB1 agonist HU210, 3 µg g−1 rimonabant (SR141716), or HU210 + SR141716 1 hour prior to sacrifice and removal of the liver for isolation and quantification of mRNA by Northern hybridization. An original blot (a) and means ± standard error from five replicate experiments in each group (b) are shown. Relative mRNA levels were quantified by densitometry, corrected for 18S rRNA levels used as loading control, and expressed as a percentage of the value in vehicle-treated control. Reproduced from [8] with permission

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Figure 67.5 Presence of CB1 receptors in mouse liver. (a) CB1 mRNA is present in the liver of CB1 but not CB1 mice, as tested by reverse transcriptase polymerase chain reaction (RT-PCR). β-Actin mRNA was amplified as internal control. (b) Localization of CB1 message in normal mouse liver by in situ hybridization. (c) Immunoreactive CB1 are present in −/− +/+ (left panel) but not a CB1 mouse (right panel). The middle panel illustrates the effect hepatocytes in the liver of a CB1 of preincubation of the N-terminal antibody with its blocking peptide in a section from the same liver as in the left panel. Tissue structure is visualized in all three panels by nuclear fast red counterstaining. (d) Immunoreactive CB1 in purified liver plasma membranes was visualized in Western blots using an antibody against the C-terminus of the rat CB1 . The specificity −/− mouse. The expression of CB1 is up-regulated in of the reaction is indicated by its absence in a preparation from a CB1 mice on a high-fat (HF) diet compared to regular (R) diet. Reproduced from [8] with permission

or in isolated hepatocytes, and hepatocytes were found to express CB1 receptors (Figure 67.5). A high fat diet increased the hepatic levels of anandamide, CB1 receptor density, and basal rates of fatty acid synthesis, and the latter was reduced by CB1 blockade. These findings led to the conclusion that high-fat diets activate the ECS, and anandamide acting at hepatic CB1 receptors contributes to diet-induced obesity and fatty liver through increasing de novo lipogenesis in the liver [8]. Despite the elevated tissue levels of anandamide, its biosynthetic rate remained unchanged in mice on a high-fat diet. Instead, the rate of degradation of anandamide was reduced due to a diet-induced decrease in the activity, but not the gene expression, of FAAH [8]. This explains why the increase was selective for anandamide and not 2-AG, the latter not being an in vivo substrate of FAAH. This finding also suggests that high-fat diets induce the generation of an endogenous FAAH inhibitor, which would then contribute to the activation of ECS by reducing the degradation of endogenous anandamide. Mice lacking CB1 receptors are resistant to high-fat diet-induced obesity and hepatic steatosis, despite a caloric intake similar to that of wild-type mice, which do become obese and steatotic on the same diet [8, 61]. Furthermore, CB1 antagonist treatment of mice with diet-induced obesity results in a sustained decrease in body weight [65] and also the reversal of hepatic steatosis [66], whereas the same treatment causes only a transient reduction in caloric intake. This means that

genetic or pharmacological ablation of CB1 receptors must result in increased energy expenditure to compensate for the increase in caloric intake. Indeed, both the gene expression and the enzymatic activity of carnitine palmitoyltransferase-1 (CPT-1), the rate-limiting enzyme in fatty acid β-oxidation, were elevated in the liver +/+ −/− of CB1 mice, and were also compared with CB1 increased following rimonabant treatment of wild-type mice. Furthermore, the diet-induced suppression of CPT-1 activity observed in wild-type mice was absent in CB1 knockout mice, suggesting that the effect of the diet is mediated by endocannabinoids acting via CB1 receptors. Indeed, treatment of normal mice with a potent CB1 agonist resulted in a marked decrease in hepatic CPT-1 activity, which could be prevented by rimonabant pretreatment [67]. The ability of CB1 antagonists to increase energy expenditure in the post-prandial state has also been documented using indirect calorimetry in intact rats [68] and mice [67]. In the latter study, the rimonabant-induced marked decrease in respiratory quotient, indicating increased fat burning, was completely absent in CB1 knockout mice, confirming that even at the relatively high dose used (10 mg kg−1 i.p.), the effect of rimonabant is mediated exclusively by CB1 receptors. Although the above findings strongly suggest the role of hepatic CB1 receptors in the development of diet-induced obesity, hepatic steatosis and related metabolic changes, they do not exclude the alternative possibility that ECs may act on CB1 receptors in the central nervous system (CNS) to influence peripheral energy metabolism

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indirectly through neuronal or hormonal pathways, or at CB1 receptors at extrahepatic sites, such as the adipose tissue. The role of hepatic CB1 receptors in metabolic regulation has been more directly tested through the use of a mouse strain with a hepatocyte-selective deletion −/− −/− mice, genmice). LCB1 of CB1 receptors (LCB1 erated by crossing CB1 floxed mice with albumin Cre mice, have no CB1 receptors expressed in hepatocytes, although they have normal levels of CB1 receptors elsewhere in the body, including the brain, adipose tissue, and non-hepatocyte cells of the liver. When placed on a −/− mice developed the same degree high-fat diet, LCB1 of obesity as wild-type mice but, similarly to mice with −/− mice), they global knockout of CB1 receptors (CB1 were largely resistant to hepatic steatosis. Total caloric intake during the 14 week diet period was similar in the three groups of mice. These findings implicate hepatic CB1 receptors in diet-induced hepatic steatosis, but not in the associated increase in adiposity. Interestingly, this conclusion is supported by the results of a recent epidemiological study of over 300 patients with chronic hepatitis C infection, who are predisposed to hepatic steatosis. In these patients, daily cannabis smoking was found to be an independent risk factor for steatosis severity, but not for obesity [69]. The high-fat diet-induced hypertrygliceridemia seen in wild-type mice was only modestly attenuated in the two knockout strains, but the accompanying increase in plasma LDL cholesterol and decrease in HDL cholesterol −/− −/− mice on the and LCB1 were absent in both CB1 high-fat diet [67]. This indicates that hepatic CB1 receptors mediate diet-induced changes in hepatic lipoprotein metabolism and/or secretion. High-fat diets also result in elevated plasma insulin and leptin levels [8, 67, 70]. This is associated with hyperglycemia, indicating insulin resistance [67, 70], and there is also evidence that the elevated leptin levels reflect diet-induced leptin resistance [71, 72]. Interestingly, the diet-induced glucose intolerance and insulin resistance −/− −/− were found to be absent in both CB1 and LCB1 mice, and the diet-induced hyperleptinemia was also absent in both knockout strains [67]. There is also evidence that THC induces glucose intolerance in humans [73] and in rodents, via activation of CB1 receptors [74]. Together, these findings indicate that the diet-induced insulin and leptin resistance are mediated by endocannabinoids acting at hepatic CB1 receptors. Diet-induced insulin resistance involves adipose tissue, skeletal muscle and liver, and interactions among the three tissues through neurogenic [75] and/or humoral factors [76]. In mice, a high-fat diet induces the expression of CB1 receptors in skeletal muscle [77], and CB1 blockade increases insulin-induced glucose uptake and phosphorylation in skeletal muscle of genetically obese mice [78]. It remains to be determined how endocannabinoid action at hepatic CB1 receptors may influence insulin sensitivity in distant tissues such as skeletal muscle. Regardless of

the exact cellular mechanisms, the ability of CB1 receptor antagonists to improve insulin sensitivity and glucose tolerance may be therapeutically exploited in diabetes and prediabetes/metabolic syndrome. More recent data suggest that CB2 receptors may also be involved in diet-induced hormonal and metabolic changes. Treatment of normal Wistar rats with low doses of the selective CB2 agonist JWH-133 improved glucose tolerance following i.p. injection of glucose, whereas the CB2 -selective antagonist AM630 had the opposite effect and it also prevented the effect of subsequently injected JWH-133 [79]. These effects mediated via CB2 receptors are opposite to the glucose intolerance induced by CB1 receptor activation (see above), and could minimize the effects of mixed CB1 /CB2 agonists on glucose homeostasis. The well-documented insulin-sensitizing action of CB1 blockade [80, 81] may reflect the fact that the endocannabinoid anandamide acts primarily via CB1 receptors, having very low efficacy at CB2 receptors [82]. This is also consistent with findings that high-fat diet-induced glucose intolerance and insulin resistance are associated with a selective increase in hepatic anandamide, but not 2-AG, levels [8] (2-AG being a full agonist at both CB1 and CB2 receptors). Another study reported that CB2 receptor mRNA was undetectable in normal human liver tissue, but was strongly induced in both steatosis and non-alcoholic steatohepatitis [40], which could suggest CB2 involvement in hepatic fat metabolism. Indeed, results in a recent meeting abstract indicate that CB2 receptor knockout mice are resistant to high-fat diet-induced steatohepatitis, and also show smaller increases in plasma insulin and less insulin resistance than wild-type littermates on the same diet [83]. The CB2 -mediated impairment in insulin sensitivity suggested by these findings in mice is opposite to the insulin-sensitizing effect of CB2 agonists in rats referred to above [79], and may reflect species differences and/or multiple mechanisms, such as modulation of insulin secretion in pancreatic β-cells, altered insulin signaling, or glucose transport in different target tissues. Further studies are obviously needed to resolve these questions.

Alcoholic fatty liver Chronic alcohol use can lead to the development of fatty liver that can further progress into steatohepatitis and liver cirrhosis. The steatogenic action of ethanol has been attributed to enhanced hepatic lipogenesis [84, 85] and decreased fatty acid oxidation in the liver [86]. Obesity is also frequently associated with fatty liver and the subsequent development of cirrhosis, and high-fat diets in rodents induce obesity, hepatic lipogenesis, and steatosis. As reviewed in the previous section, diet-induced obesity and steatosis are mediated, at least in part, through

67: THE ROLE OF ENDOCANNABINOIDS AND THEIR RECEPTORS IN THE CONTROL OF HEPATIC FUNCTIONS

activation of the EC/CB1 receptor system. These similarities between the steatogenic effects of high-fat diet and of alcohol, together with evidence that chronic ethanol exposure can increase endocannabinoid levels, at least in the brain [87], suggest that the ECS may also be involved in ethanol-induced fatty liver. This possibility has gained recent experimental support. It has been reported that exposure of male mice to a low-fat, liquid ethanol diet for 3 weeks increased CB1 receptor gene expression in the liver and the hepatic levels of 2-AG but not anandamide. The increase in 2-AG occurred selectively in hepatic stellate cells, in which the expression of the 2-AG biosynthetic enzyme diacylglycerol lipase-β (DAGLβ) was also increased, whereas DAGLα or MGL (2-AG degrading enzyme) expression remained unchanged. These findings suggest that the ethanol-induced increase in 2-AG is related to its increased biosynthesis. Chronic ethanol exposure also resulted in the development of fatty liver, as verified by post mortem histological and biochemical analyses. Simultaneous chronic treatment of the ethanol-fed mice with rimonabant significantly attenuated the steatosis without affecting daily alcohol intake and blood ethanol levels, suggesting CB1 receptor involvement in ethanol-induced steatosis. This was strongly supported by the finding that mice with global or hepatocyte-specific knockout of CB1 receptors were both resistant to the steatogenic action of alcohol. In agreement with published findings [85], ethanol feeding increased the hepatic nuclear expression of SREBP1c and its target FAS, and decreased both the hepatic expression and enzyme activity of CPT-1. In both −/− −/− CB1 mice, ethanol induction of SREB1c and LCB1 and FAS expression and the parallel inhibition of CPT-1 expression were blunted or absent. Furthermore, the enzymatic activity of CPT-1 was significantly higher in both CB1 knockout strains than in their respective controls and, unlike in controls, chronic ethanol intake failed to reduce CPT-1 activity. These findings support the notion that ethanol-induced fatty liver involves increased lipogenesis and decreased elimination of fat by fatty acid oxidation from the liver, and the absence of −/− −/− these effects in CB1 mice may explain and LCB1 their resistance of to ethanol-induced steatosis. −/− The resistance of LCB1 mice to the steatogenic action of ethanol implicated CB1 receptors located on hepatocytes, whereas the ethanol-induced increase in 2-AG synthesis occurred exclusively in hepatic stellate cells. This suggests a paracrine mechanism whereby stellate cell-derived 2-AG activates CB1 receptors on adjacent hepatocytes to stimulate lipogenesis and inhibit fatty acid oxidation in the latter. Such a mechanism was supported by the finding that co-culturing stellate cells isolated from alcohol-fed mice with hepatocytes from control mice resulted in increased lipogenic gene expression in the latter. This paracrine effect triggered by the ethanol-primed

1099

Hep/HSC cocultures Hep

WT

WT

LCB1−/−

HSC

Pair-fed

EtOH

EtOH

CB1R SREBP1c FAS β-actin (a) Hepatocytes cocultures with HSC

Pair-fed

EtOH

SREBP1c FAS β-actin (b)

Figure 67.6 Paracrine regulation of hepatocyte lipogenesis by hepatic stellate cell (HSC)-derived endocannabinoids. (a) Up-regulation of CB1 receptor, SREBP1c, and FAS expression in hepatocytes from pair-fed mice co-cultured with HSC from ethanol-fed vs pair-fed mice (middle vs −/− hepatocytes co-cultured with HSC left lanes). In LCB1 from ethanol-fed mice CB1 mRNA is absent, and the induction of SREBP1c and FAS expression is blunted compared with wild-type hepatocytes (right lanes). mRNA levels were determined by RT-PCR and (b) increased SREBP1c and FAS protein levels, determined by Western blotting, in wild-type control hepatocytes co-cultured with HSC from ethanol-fed (right lanes) vs pair-fed mice (left lanes). Reproduced from [33], Copyright (2008), with permission from Elsevier

stellate cells was blunted when the hepatocytes in the −/− mice, confirming the role co-culture were from LCB1 of CB1 receptors (Figure 67.6). The findings discussed in this section suggest that CB1 antagonists may be effective in the treatment of both alcoholic and non-alcoholic fatty liver disease. Clinical trials with rimonabant for obesity/metabolic syndrome revealed a modest, but worrisome, increase in the incidence of anxiety and depression, which has been the main reason why the compound has not received approval from the US Food and Drug Administration (FDA) for use in −/− the United States. The findings in LCB1 mice discussed above suggest that a peripherally restricted antagonist may retain its efficacy in the treatment of hepatic steatosis, whereas such compounds would be less prone to produce centrally mediated side effects.

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ENDOCANNABINOIDS AND HEPATIC ISCHEMIA– REPERFUSION INJURY I/R injury may develop in conditions where blood and oxygen supply to a tissue is transiently disrupted, such as myocardial infarction or stroke. Hepatic I/R injury is a potentially fatal complication of liver surgery, including liver transplantation. Endogenous or exogenous substances that improve hypoxia tolerance have the potential to protect against I/R injury. The biological effects of CBs prominently include hypomotility and hypothermia, both of which would tend to reduce oxygen demand. Their metabolic effects, promoting energy storage and reducing energy expenditure (see above), could also contribute to such a protective effect. Recent studies in mice have provided strong evidence that the ECS, primarily through CB2 receptor-mediated mechanisms, protects against hepatic I/R injury [88, 89]. In a mouse model, segmental ischemia followed by reperfusion (but not ischemia alone) resulted in marked increases in the hepatic levels of anandamide and 2-AG. This rise in endocannabinoid levels correlated with the degree of tissue damage as indicated by rises in serum AST/ALT levels, and also increases in both serum and tissue levels of TNFα, MIP1α, and MIP2α [88]. The I/R-induced tissue damage was significantly reduced by pretreatment of mice with the CB2 -selective agonist −/− JWH-133, whereas it was significantly greater in CB2 mice than in their wild-type littermates. Also, the protective effect of JWH-133 could be prevented by a selective −/− CB2 antagonist and it was absent in CB2 mice. Neutrophil infiltration and lipid peroxidation are important factors in I/R injury, an indicator of the former being tissue levels of myeloperoxidase, whereas the latter is quantified by measuring tissue malondialdehyde levels. The I/R-induced rise in both of these markers was attenu−/− ated by JWH-133 and increased in CB2 compared with wild-type mice [88]. The role of CB2 receptors was more extensively explored in a subsequent study by the same group [89], using the highly potent and selective CB2 agonist HU-308 [90]. It was found that, in addition to the effects reported earlier, CB2 receptor activation attenuates the I/R-induced hepatocyte apoptosis and also mitigates the TNFα-induced expression of the cell adhesion molecules ICAM-1 and VCAM-1 in hepatic sinusoidal endothelial cells [89]. Since adhesion molecules mediate the initial attachment of neutrophils to the vascular endothelium, inhibition of their expression by CB2 receptor activation must play a key role in the protection against I/R injury. By additionally suppressing the production of TNFα, a strong stimulator of neutrophil infiltration and generator of oxidative stress, CB2 activation may afford protection at multiple levels against I/R injury. In contrast to psychoactive CB1 receptor agonists, selective agonists

of CB2 receptors do not exert significant behavioral effects, making them therapeutically more acceptable.

HEPATIC ENCEPHALOPATHY, AUTOIMMUNE HEPATITIS Hepatic encephalopathy is a neuropsychiatric syndrome associated with acute liver failure. Although several pathogenic factors have been identified, including the accumulation of neurotoxic metabolites, such as ammonia, alterations in various central neurotransmitter systems, and altered cerebrovascular function, the mechanisms underlying the altered neuro-cognitive state are not completely understood. Recent findings have implicated endocannabinoids and both CB1 and CB2 receptors in this condition. In a mouse model of thioacetamide-induced fulminant liver failure, brain levels of 2-AG were found to be markedly increased [91]. Furthermore, administration of exogenous 2-AG or the CB2 -selective agonist HU-308 improved the neurological score, activity, and cognitive function, and these effects could be prevented by the CB2 antagonist SR144528A. Similarly to CB2 agonists, the CB1 antagonist rimonabant also improved the neurological score [91]. In a subsequent study by the same group, it was found that thioacetamide treatment or bile duct ligation induced the expression of CB2 receptors in the brain, and also resulted in activation (by phosphorylation) of the AMP-activated protein kinase (AMPK). The neuroprotective effect of THC in these mice could be attributed to a CB2 -mediated increase in AMPK activity, as both the neuroprotective effect and AMPK activation were absent in CB2 knockout mice [92]. The well-known immunomodulatory effect of THC led to its recent testing in a murine model of concanavalin A (ConA)-induced autoimmune hepatitis. THC administered following a ConA challenge was found to inhibit hepatitis, as judged by its ability to decrease liver enzymes and inflammatory cytokines and reduce tissue injury [93]. Non-selective CB1 /CB2 agonists, but not selective agonists, mimicked the hepatoprotective effect of THC, which could be attenuated by either CB1 or CB2 antagonists, implicating both CB1 and CB2 receptors. Interestingly, mice deficient in the anandamide-degrading enzyme FAAH showed reduced hepatic damage in response to ConA treatment, suggesting hepatoprotection by endogenous anandamide [93].

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Telomeres and Aging, Cancer, and Hepatic Fibrosis Hans L. Tillmann1, Ruben R. Plentz2, Yvonne Begus-Nahrmann3, Andr´e Lechel3 and Lenhard K. Rudolph3 1

Duke Clinical Research Institute, GI/Hepatology Research Program, Division of Gastroenterology, Durham, NC, USA 2 Department of Gastroenterology, Hepatology and Endocrinology, Medical School Hannover, Hannover, Germany 3 Institute of Molecular Medicine and Max Planck Research Group on Stem Cell Aging, Ulm University, Ulm, Germany

INTRODUCTION Aging is associated with a decrease in organ maintenance and function, impaired regenerative capacity, and an increased cancer risk. One of the underlying mechanisms is cellular aging. The proliferative capacity of human cells is limited to a finite number of cell divisions. On a molecular level, cellular aging is characterized by an accumulation of DNA damage, activation of checkpoints, and alterations in gene expression, leading to cellular dysfunction, cell cycle arrest, or apoptosis. Telomere shortening represents the main mechanism of DNA damage accumulation, limiting the proliferative lifespan of human cells in culture. Telomeres shorten in almost all tissues during human aging. In addition, chronic diseases, such as hepatitis, accelerate the rate of telomere shortening and cellular aging in the affected organs by increasing the rate of cell turnover. In this chapter, we summarize the role of telomere shortening in cellular aging and its influence on aging, disease progression, and carcinogenesis in the liver. The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

TELOMERE SHORTENING LIMITS THE PROLIFERATIVE CAPACITY OF HUMAN CELLS It was first recognized by Leonard Hayflick that the proliferative capacity of human cells is limited to a finite number of cell divisions [1]. When human fibroblasts are grown in cell culture, they lose proliferative potential after 50–70 cell divisions at a stage named replicative senescence. Senescent cells are permanently arrested in the cell cycle and fail to initiate DNA-synthesis in response to growth stimuli. Fibroblasts show typical morphological alterations at the senescence stage, including an enlarged cytoplasm and an increased activity of β-galactosidase at pH 6 (senescence-associated β-galactosidase = SA-βGal). Using nuclear transplantation experiments, Hayflick recognized that the cell nucleus carries the memory that limits the proliferative capacity of human cells. However, it took a further 30 years for

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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Telomerase can synthesize telomere sequences de novo [15]. The enzyme consists of two essential components: (i) the telomerase reverse transcriptase (TERT) is the catalytic subunit of the enzyme [16–18], and (ii) the telomerase ribonucleic acid component (TERC) serves as a template for the synthesis of telomere sequence [19–21]. In addition, there is evidence that processing of TERC and TERT is required for generation of a functional enzyme complex [22, 23]. In human cells and tissues, TERC is expressed ubiquitously. In contrast, telomerase activity is only detectable in embryonic tissues and is repressed in most somatic tissues after birth [24]. Postnatal suppression of telomerase activity correlates with a repression of TERT expression, which is only detectable in germ cells and some stem cells in adult humans [25, 26]. It has been reported that somatic cells can express very low levels of TERT during the S-phase of the cell cycle [27]. However, the low expression of TERT cannot prevent telomere shortening in cultured fibroblast [5, 12]. When TERT is over-expressed in adult human fibroblast or fetal hepatocytes, telomerase is activated, leading to telomere stabilization and immortal proliferation [2, 3]. Today, it is well accepted that the lack of TERT expression is responsible for telomere shortening and the limited lifespan of human cells. Of note, the re-expression of TERT does not lead to transformation of human cells into cancer cells [28]. However, there are reports that extensive culture of TERT-transduced cells can lead to chromosomal instability, gene mutations, and transformation of human cells [29, 30]. Together, these studies indicate that a transient reactivation of telomerase over short periods of time could be a valuable tool to expand genetically stable cells that

scientists to disclose the molecular mechanism responsible for this phenomenon. Today, it is known that telomere shortening is the underlying cause limiting the proliferative capacity of primary human cells, including hepatocytes [2, 3]. Telomeres form the ends of human chromosomes [4]. They consist of small tandem DNA repeats (TTAGGGn in human cells). Telomeres do not encode for a protein product. The main function of telomeres is to cap the chromosomal ends. Telomere capping is essential to distinguish the chromosome ends from DNA breaks within the chromosome. Human telomeres are 5–15 kb long [5]. It has been shown that a minimum telomere length of 84 bp (14 TTAGGG-repeats) is required for telomere capping [6]. Moreover, telomeres need to form a tertiary structure (telomere loops, G-quadruplexes) to cap the chromosome ends [7]. The tertiary structure of telomeres is stabilized by telomere binding proteins that specifically bind to telomeric DNA [8]. There is emerging evidence that alterations in the expression of these proteins can be the cause of diminished organ maintenance and increased cancer formation [9]. Telomeres shorten during each round of cell division because of (i) the end replication problem of DNA polymerase and (ii) the processing of telomeres during the cell cycle [10, 11]. Human telomeres shorten at a rate of 50–100 bp per cell division [12]. It has been shown that telomere shortening correlates with the limited proliferative capacity of primary human cells [13]. Experiments on re-expression of the enzyme telomerase have proven that telomere shortening is the underlying cause limiting proliferation of human cells [2] (Figure 68.1).

ATM

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Cell Division Figure 68.1 Two checkpoints (mortality stages) limit the lifespan of cells in response to telomere shortening [14]. Telomeres shorten as a consequence of cell division. When a sentinel of telomeres lose capping function, the senescence checkpoint is activated. The figure shows a simplified cartoon on some of the major signaling components inducing senescence. Abrogation in this pathway can elongate the lifespan of cells. However, telomeres continue to shorten and a second checkpoint (crisis, mortality stage 2) is induced in response to an accumulation of telomere dysfunction and genomic instability. The crisis checkpoint leads to cell death but the signaling components inducing this checkpoint are not yet disclosed

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could be used for cell therapies. Most studies on activation of telomerase have used viral gene transfer, resulting in permanent integration of a TERT expressing DNA construct. This technique harbors the risk of insertional mutagenesis and the permanent expression of telomerase may further increase the cancer risk in this context. Pharmacological or molecular approaches for non-toxic, transient activation of telomerase still remain to be developed.

SENESCENCE AND CRISIS: CHECKPOINTS INDUCED BY TELOMERE DYSFUNCTION When telomeres reach a critically short length, they lose capping function and DNA damage checkpoints are activated, including the p53-signaling pathway [31] (Figure 68.1). It has been shown that a subset of dysfunctional telomeres is sufficient to induce replicative senescence [32]. The main characteristic of senescence is the irreversible and permanent loss of cell proliferation. The induction of senescence is p53 and Rb dependent and an inhibition of these pathways abrogates the senescence checkpoint [31]. A second checkpoint can limit survival of cells with dysfunctional telomeres. When cells bypass the senescence checkpoint, further telomere shortening leads to an accumulation of telomere dysfunction, inducing chromosomal instability, activation of p53-independent DNA damage responses, and cell death [31]. This checkpoint is named crisis. The molecular signaling pathways that induce crisis are less understood than senescence. It is possible that crisis is a more heterogeneous checkpoint induced by a variety of different signaling pathways, depending on the combination of genetic lesions induced by telomere dysfunction. These lesions likely show great cell-to-cell variability. Crisis is a tight checkpoint and in human fibroblast cultures only 1 in 107 cells can bypass this checkpoint by spontaneous immortalization [14]. Cells that bypass crisis activate a mechanism of telomere stabilization. Studies on human fibroblast have shown that two-thirds of surviving clones show a spontaneous reactivation of telomerase – the enzyme that can synthesize telomeres de novo [14, 31]. The remaining one-third of the clones activates an alternative mechanism of telomere elongation [alternative lengthening of telomeres (ALT)]. The molecular basis of ALT is not yet completely understood, but it involves the activation of DNA recombination, leading to telomere exchange between different chromosomes and a highly heterogeneous telomere length [33]. Senescence and crisis checkpoints are thought to act as tumor suppressor checkpoints, limiting the proliferative capacity of genetically unstable cells with dysfunctional telomeres. As a downside, the same checkpoints may limit the regenerative reserve of tissues and organs harboring

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short telomeres as a consequence of aging or chronic disease.

THE INFLUENCE OF TELOMERE SHORTENING ON ORGAN REGENERATION AND AGING: LESSONS FROM MOUSE MODELS Studies in telomerase knockout [mouse telomerase RNA component (mTERC−/− )] mice have provided experimental evidence that telomere dysfunction can induce premature aging, especially affecting organ systems with high rates of cell turnover [34, 35]. In addition, telomere shortening limited the regenerative capacity of injured tissues and organs including the liver [36]. In response to partial hepatectomy, telomere shortening reduced the number of regenerating liver cells in mTERC−/− mice compared with wild-type mice [37]. The liver cells that did not participate in liver regeneration contained shorter telomeres and stained positive for senescence-associated β-galactosidase activity [37]. These data provided a proof-of-principle that telomere shortening can limit organ regeneration by reducing the number of regenerating cells in an organ. In response to chronic liver injury, telomere dysfunction accelerated the activation of stellate cells, fibrotic scarring, and the induction of steatosis [36]. In addition, telomere shortening had a dominant negative effect on survival in mice with chronic liver damage, although the rate of hepatocellular carcinoma (HCC) formation was reduced [38]. These data suggest that an improvement in regenerative reserve could have positive effects on overall survival in the context of telomere dysfunction and chronic liver damage – a situation relevant to human cirrhosis (see below). Impaired organ maintenance in telomere dysfunctional mice is mediated by DNA damage pathways. It has been shown that p53 deletion rescued germ cell apoptosis in TERC−/− mice [39]. However, p53 deletion cooperated with telomere dysfunction to induce chromosomal instability and the mice died prematurely due to the formation of malignancies. In contrast, deletion of p21 – a downstream target of p53 inducing cell cycle arrest – improved organ maintenance and survival of telomere dysfunctional mice without accelerating cancer formation [35]. This rescue was associated with improved proliferation and a rescue in maintenance and function of adult stem cells in aging telomere dysfunctional mice. Deletion of p21 left apoptosis checkpoints in response to telomere dysfunction intact, providing a possible explanation of why tumor rates were not increased. Similarly, studies on human fibroblasts have shown that deletion of p21 elongated the lifespan of human fibroblasts, but apoptosis checkpoints remained intact and the genome of the p21−/− cells with an extended lifespan remained stable [40].

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Another gene that was shown to mediate the adverse effects of telomere dysfunction on organ maintenance was exonuclease-1 (Exo1) – a 5–3 prime exonuclease. Studies in yeast have shown that Exo1 is necessary to induce senescence arrest in response to telomere dysfunction [41]. In mice, Exo1 deletion rescued the induction of DNA damage signals in response to telomere dysfunction. This rescue was upstream of p53 and rescued both cell proliferation and apoptosis in the intestinal epithelium of TERC−/− mice [42]. Interestingly, Exo1 deletion extended the lifespan of telomere dysfunctional mice and did not accelerate cancer formation, although both cell cycle arrest and apoptosis checkpoints were abrogated [42]. A possible explanation is that Exo1-dependent processing of dysfunctional telomeres is necessary to initiate DNA damage response. This processing may also be necessary to activate DNA repair responses leading to the formation of chromosomal fusions and the evolution of chromosomal instability in response to telomere dysfunction. In line with this interpretation, Exo1 deletion reduced the formation of anaphase bridges – a morphological correlate of chromosomal fusion – in the intestine of telomere dysfunctional mice [42]. Together, these studies indicated that DNA damage signals limit cellular lifespan and organ maintenance in response to telomere dysfunction. The studies on Exo1 and p21 have provided a proof-of-principle that an inhibition Increased Hepatocyte Turnover

of checkpoints can improve stem cell function, organ maintenance and lifespan of telomere dysfunctional mice without accelerating the cancer risk. Given the regenerative defects and the prevalence of critically short telomeres in human cirrhosis (see below), these findings could point to novel treatment approaches for this disease stage.

TELOMERE SHORTENING IN HUMAN AGING, LIVER DISEASE AND CIRRHOSIS There is growing evidence that telomere shortening and cellular aging can limit the regenerative reserve of organs and tissues during chronic diseases and aging. Telomere shortening occurs in almost all tissues and organs during human aging [5, 43]. In addition, there is evidence that chronic diseases that accelerate the rate of cell turnover also increase the rate of telomere shortening. A classical example is the human liver. There is little cell division in healthy liver correlating with limited telomere shortening during aging [44, 45]. However, chronic liver diseases accelerate the rate of hepatocyte turnover and are associated with accelerated telomere shortening in liver of affected patients compared with age-matched control patients without liver disease (Figure 68.2) [46–48].

Chronic Liver Disease

Telomere Shortening Telomere Dysfunction

Activation of DNA DamageCheckpoints

Senescence

Impaired Regeneration

Increased Cell Death

Cirrhosis Stellate Cell Activation Fibrosis Steatosis

Figure 68.2 Telomere shortening induces cirrhosis formation. Chronic liver disease increases the rate of hepatocyte turnover by inducing hepatocyte death followed by regeneration of remaining hepatocytes. This increase in cell turnover leads to telomere shortening and finally to an accumulation of dysfunctional telomeres in hepatocytes. Dysfunctional telomeres induce cell-intrinsic DNA damage checkpoints that impair liver regeneration by inducing cell cycle arrest (senescence). In addition, the activation of DNA damage checkpoints can increase the sensitivity of hepatocytes to chronic insults and the rate of hepatocyte death. Together, these mechanisms disturb the balance between liver regeneration and hepatocyte death, thus leading to an activation of stellate cells, fibrogenesis, and the evolution of cirrhosis. According to this model, eradication of chronic liver disease, stabilization of hepatocyte telomeres, or inactivation of checkpoint responses could improve the balance between liver regeneration and hepatocyte death. According to this model, the potential to reverse cirrhosis after eradication of chronic liver disease could be telomere length dependent

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Moreover, cirrhosis is characterized by critical telomere shortening [44–47], a decline in cell proliferation [48, 49], and an up-regulation of senescence markers (Sa-βGal and p21) [44, 50–55]. Together, these data suggest that telomere shortening and senescence inhibit hepatocyte proliferation, thus promoting the evolution of cirrhosis at the end stage of chronic liver disease. In agreement with this hypothesis, there is genetic evidence that telomere shortening influences human aging and organ maintenance. It was shown that telomerase mutations in humans lead to premature telomere shortening, impaired organ maintenance, and reduced survival. The first clinical example was dyskeratosis congenita (DKC). The autosomal dominant form of this disease is caused by mutation in the RNA component of telomerase [56]. Another form of the disease is caused by mutation in the dyskerin gene, which is necessary for the processing of small nuclear RNAs. Dyskerin also processes the RNA component of telomerase and DKC patients with dyskerin mutations show reduced levels of TERC expression [22]. DKC patients have abnormally short telomeres and die from bone marrow and intestinal failure [57]. Interestingly, DKC patients also exhibit an increased risk of developing cirrhosis and cancer. Telomerase mutations have also been linked to other diseases, such as aplastic anemia [58], myelodysplastic syndrome (MDS) [59], and idiopathic lung fibrosis [60, 61]. Together, these studies show that human telomere reserves are limited and that a heterozygous mutation of telomerase can have deleterious effects on organ homeostasis and survival early in life. It is conceivable that telomere shortening can also limit organ maintenance at old age during “natural” aging or in response to accelerated cell turnover during chronic diseases. Along these lines, telomere length in human leukocytes has been associated with survival and risk of cardiovascular events [62, 63]. In addition to age, leukocyte telomere length decreases with smoking and obesity and both factors are associated with premature aging [64]. In agreement with a role of telomere dysfunction in human aging, it has been shown that marker proteins of telomere dysfunction and DNA damage associate with human aging, age-associated disease, and diseases that are associated with telomere shortening such as cirrhosis and myelodysplastic syndromes [65]. Chronic liver disease is associated with progressive fibrosis deposition, most likely due to a misbalance where fibrosis-inducing factors outrange fibrosis-removing factors such as TIMPs (tissue inhibitor of metalloproteinases). The end stage of this process is liver cirrhosis, characterized by fibrosis and impaired organ function. Increased fibrosis leads to portal hypertension with its clinical complications, such as ascites and esophageal varices. In addition, reduced liver function results in clotting disorders, icterus, and hepatic encephalopathy. Hence if fibrosis can be removed but the potential of liver cells to regenerate is exhausted due to telomere shortening, the course of disease might not be substantially improved. It appears that two

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obviously different, although not independent factors, are present in end-stage liver disease, increasing fibrosis and impaired regeneration. This observation could be highly relevant for clinical therapies, as only targeting both mechanisms may be effective for treatment of cirrhosis. Telomere shortening and, even more important, telomere dysfunction [65] may not only represent therapeutic targets in chronic liver disease, but could also serve as biomarkers predicting disease outcome. It is conceivable that, at similar levels of inflammation, livers with increased levels of telomere dysfunction show lower regenerative activity, and may thus be more prone to chronic or sudden hepatic insults. In agreement with this assumption, it was shown that hepatitis C virus (HCV)-positive patients with elevated transaminases have shorter telomeres compared with those with normal alamin-aminotransferase levels [66]. The latency of chronic hepatitis leading to cirrhosis formation is shortened in older patients [67]. Hepatic telomere shortening in older humans [44, 45] could provide a molecular explanation for this association. Accelerated telomere shortening in diabetes patients [68] could also explain the increased risk of cirrhosis and HCC in diabetes patients with chronic liver disease compared with non-diabetic patients with chronic liver disease. Another interesting, yet not well-explained, finding is the higher frequency of men compared with women in patients with progressive liver disease [67, 69, 70]. Both male gender and age are significantly associated with shorter survival in patients with chronic liver disease [71]. There is some clinical evidence for a higher regenerative potential in women compared with men [72, 73]. This may partially be explained by longer telomeres in aging women compared with men [74]. One reason for this gender difference in telomere length might be estradiol, which was shown to reduce telomere shortening in human hepatic cells in vitro and in rat livers in vivo exposed to carbon tetrachloride [75]. The classical hypothesis of cirrhosis development indicates that cirrhosis emerges as a consequence of continuous tissue damage followed by matrix deposition [76]. The telomere hypothesis of cirrhosis formation indicates that cirrhosis is the consequence of telomere shortening, hepatocyte senescence, and impaired regenerative reserve in response to chronic injury [77] (Figure 68.2). While cirrhosis as a consequence of chronic alcohol abuse usually does not reverse even if alcohol is completely avoided [78], effective treatment for viral hepatitis can result in the reversal of cirrhosis in some patients. Especially in hepatitis B, many patients listed for transplantation have shown remarkable improvement, allowing removal from waiting lists for liver transplantation [79]. It remains to be tested whether hepatic telomere reserves can indicate patients’ outcome and the chances of cirrhosis reversal (Figure 68.2).

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TELOMERES, TELOMERASE, AND CANCER: LESSONS FROM MOUSE MODELS Telomere shortening limits the lifespan of human cells by induction of senescence or crisis. Both senescence and crisis represent powerful tumor suppressor checkpoints. In agreement with this model, most human malignancies (including hepatocellular and cholangiocellular carcinoma) show an activation of telomerase indicating that stabilization of telomeres is a critical step in cancer formation [25]. Although telomerase itself is not an oncogene [28], in vitro experiments on human fibroblast have provided experimental evidence that the activation of telomerase (induced by TERT re-expression) is a critical step for the immortalization and the tumor-forming capacity of transformed human cells [80]. Specifically, TERT expression was necessary for the immortalization of oncogene-transformed cells and only TERT-expressing clones efficiently form tumors after xenotransplantation in nude mice [80]. Experiments on telomerase knockout mice have provided the first in vivo evidence for a critical role of telomerase during tumor formation (Figure 68.3). TERC knockout mice (TERC−/− ) lack telomerase activity and show critical telomere shortening in late generations (G3–G6 TERC−/− ) [81]. Late-generation TERC−/− mice showed reduced formation of macroscopic tumors in different organ compartments in genetic and chemical models of tumor formation. Telomere shortening also suppressed

the formation of macroscopic HCCs in carcinogen-treated mice [82, 83] and in models of chronic liver injury [38, 84]. The suppression of tumor formation in telomere dysfunctional TERC−/− mice was accompanied by an activation of p53-dependent DNA damage pathways, increased tumor cell apoptosis, and suppression of tumor cell proliferation [82–85]. However, suppression of macroscopic liver tumors also occurred in mice carrying a homozygous deletion of p53 in the liver. Tumor suppression in p53−/− TERC−/− liver was associated with very high levels of chromosomal instability and DNA damage accumulation in tumor cells [84]. This study showed that p53-independent checkpoints can limit HCC formation in vivo in response to telomere dysfunction and accumulation of chromosomal instability. These checkpoints might be similar to the crisis checkpoint observed in virus-transformed human fibroblast cultures that bypass the senescence checkpoint [31]. In contrast to the suppression of macroscopic tumor formation, studies in TERC−/− mice have also revealed evidence that telomere shortening can increase the rate of tumor initiation (Figure 68.3). Aging TERC−/− mice on a mixed genetic background showed increased rates of spontaneous cancer formation during aging [34]. Moreover, genetic and carcinogen-induced formation of microscopic tumors was increased in the intestine [85] and in the liver [83] of telomere dysfunctional TERC−/− mice compared with TERC+/+ mice. However, the increased initiation of microscopic tumors did not translate to increased formation of macroscopic tumors. The

Hepatocellular Carcinoma

Chromosomal Instability Tumor Initiation

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clonal selection of cells with mutant checkpoints

Impaired Liver Regeneration Cirrhosis

Figure 68.3 Telomeres shorten as a consequence of aging and chronic diseases. Telomere dysfunction leads to the induction of chromosomal instability and cancer initiation. Intact checkpoints prevent proliferation of genetically instable cells but the loss of DNA damage checkpoints can cooperate with telomere dysfunction to induce chromosomal instability and cancer initiation. In contrast to its tumor suppressor function, the induction of DNA damage checkpoints can limit liver regeneration and promote cirrhosis, resulting in a loss of proliferative competition and environmental alteration promoting the selection of abnormal proliferating precancerous cells. Genetically unstable cancer cells need to activate a mechanism of telomere stabilization to allow tumor progression, since ongoing telomere dysfunction would ultimately result in genetic chaos and tumor cell death

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formation of macroscopic adenomas in the intestine [85] and macroscopic HCC in the liver [83, 84] was suppressed in telomere dysfunctional TERC−/− mice compared with TERC+/+ mice. The increase in tumor initiation in TERC−/− mice was associated with an increase in chromosomal instability in tissues and precancerous lesions [34, 85] (Figure 68.3). When critically short telomeres lose capping function at the chromosome end, DNA damage pathways are activated, including DNA repair pathways. The most common pathway for DNA break repair in mammalian cells is non-homologous end joining. In contrast to DNA breaks within the chromosome, dysfunctional telomeres cannot be repaired unless telomere-free chromosome ends are fused to one another. This process leads to the formation of chromosomal fusions in telomere dysfunctional cells. However, these fusions are unstable when cells with fused chromosomes enter the cell cycle. The fused chromosomes break during anaphase as chromosome separation proceeds – a morphological correlate of this process is the formation of anaphase bridges between the segregating chromosome plates [34, 36]. This process results in chromosomal breakage and generation of telomere-free chromosome ends in the daughter cells representing the substrate for the formation of new chromosomal fusions. It has been shown that a single dysfunctional telomere can induce a cascade of fusion–bridge–breakage cycles resulting in chromosomal instability and an accumulation of chromosomal gains and losses [86]. The formation of chromosomal instability is one of the hallmarks underlying cancer formation in humans, including the formation of HCCs [87]. In agreement with the role of p53-dependent senescence in restricting the lifespan of cells with short telomeres [31], studies in TERC−/− mice have revealed evidence that deletion of p53 can cooperate with telomere dysfunction to induce chromosomal instability and cancer formation [88] (Figure 68.3). Specifically, the heterozygous deletion of p53 resulted in an increased formation of epithelial cancers in telomere dysfunctional mice [88]. Epithelial tumors rarely occur in aging laboratory mice but are common in aging humans [89]. Interestingly, human epithelial cancers and HCC are characterized by short telomeres and often show dysfunction of the p53 signaling pathway. These data indicate that telomere dysfunction and p53 mutation may especially cooperate in epithelial tissues to induce cancer formation. It is conceivable that loss of p53 checkpoint function allows proliferation and survival of genetically unstable cells with short telomeres, thus increasing the accumulation of chromosomal gains and losses and the rate of cellular transformation. The relative contribution of p53-dependent cell cycle arrest and apoptosis for tumor suppression in the context of telomere dysfunction remains to be investigated. Studies on BCL2 (B-cell lymphoma 2, an anti-apoptotic protein) overexpression in a mouse model of lymphoma formation have shown that p53-dependent senescence can

1111

suppress tumor formation in vivo [90]. However, deletion of p21 did not increase tumor formation in telomere dysfunctional mice but improved organ maintenance and lifespan of the mice [35]. Together, these findings indicate that loss of p53-dependent cell cycle arrest is not a major tumor-promoting factor in telomere dysfunctional mice. It is possible that functionality of either p53-dependent senescence or apoptosis is sufficient to suppress tumor formation in the context of telomere dysfunction. However, the mechanisms of tumor suppression/promotion in the context of telomere dysfunction are likely more complex and could also involve environmental alterations. It is conceivable that loss of checkpoint function in single cells may result in a different outcome compared with loss of checkpoint function on tissue or organ level. The loss of checkpoint function in single cells could confer a growth advantage of mutant cells in organ system harboring telomere dysfunction. Clonal selection may not only involve cell intrinsic mechanisms but could also be driven by alterations in the environment [91, 92] and the loss of replicative competition of surrounding cells [93, 94]. According to this model, the inhibition of checkpoints on organ level could release selective pressure, thus inhibiting the clonal expansion of mutant cells [95] (Figure 68.3). In addition to the functional role of cell cycle checkpoints and apoptosis, it will be important to identify other genetic alterations that can cooperate with telomere dysfunction to induce cancer initiation and progression. Among possible candidates are the telomere-binding proteins. These proteins specifically bind to telomeric DNA and are essential to maintain the three-dimensional structure of telomeres and telomere capping function. Studies in telomere dysfunctional mice suggest that alterations in the expression level of telomere-binding proteins can accelerate the evolution of chromosomal instability and the initiation of cancer as a consequence of telomere shortening [96]. Moreover, there is emerging evidence that the expression of telomere binding proteins is altered in human cancer, including HCC [97]. Together, the studies in telomere dysfunctional mice indicate that telomere dysfunction, DNA damage checkpoints, and telomerase have a complex role in cancer formation, influencing both cancer initiation and progression (Figure 68.3).

TELOMERE SHORTENING AND TELOMERASE ACTIVATION IN HUMAN HEPATOCARCINOGENESIS Studies in telomerase knockout mice have shown that telomere dysfunction can increase chromosomal instability and cancer initiation, but telomerase activation is necessary for tumor progression. Studies on telomere length

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and telomerase activity in human liver samples indicate that the same mechanisms operate during human hepatocarcinogenesis.

Telomere shortening characterizes human hepatocarcinogenesis The risk of liver cancer development is very low in normal human liver not affected by chronic diseases. The most unifying risk factor for liver cancer formation is cirrhosis and the yearly risk of liver cancer formation in cirrhosis patients is 3–6% [98]. It is conceivable that telomere shortening at the cirrhosis stage is associated with an increased liver cancer risk, since telomere shortening induces chromosomal instability and cancer initiation in mice. In agreement with this hypothesis, it has been shown that telomeres are shorter in non-cancerous liver tissue of HCC patients compared with biopsies from cirrhosis patients without HCC [99]. In other cancer-prone diseases (e.g. colitis ulcerosa and myelodysplastic syndromes), an association between telomere shortening and the evolution of chromosomal instability [100] and malignant transformation [101] has been demonstrated. In addition, there is evidence that individuals with short telomeres in white blood cells have an increased risk for various types of cancer [102]. A prospective analysis on the association between telomere shortening and HCC risk remains to be evaluated. Such an association could help decision-making in cirrhosis patients waiting for liver transplantation. Support for a causative role of telomere shortening for the induction of HCC comes from telomere measurements in human HCC samples. A variety of studies have detected shorter telomeres in HCC compared with normal liver or compared with surrounding non-cancerous liver tissue from HCC patients [47, 51, 103–112]. It has been demonstrated that telomere shortening in HCC correlates with the evolution of chromosomal instability at the single-cell level [112], indicating a causal link between telomere shortening and the development of chromosomal instability during hepatocarcinogenesis. Furthermore, telomere shortening occurs in precancerous lesions (dysplastic nodules, small cell changes) in cirrhotic liver and during early malignant stages of hepatocarcinogenesis [51, 107], indicating that telomere shortening could act as a tumor-initiating event in human hepatocarcinogenesis. This hypothesis stands in line with the observation that telomere shortening increases liver tumor initiation in carcinogen-treated mice. Studies in mouse models have shown that loss of checkpoint genes, especially loss of p53 checkpoint function, can cooperate with telomere dysfunction to induce chromosomal instability and tumor initiation. Investigations on human carcinogenesis have revealed evidence that the p53 checkpoint is defective in 70–100% of human HCC [95,

113, 114]. In addition, loss of DNA damage checkpoint function has been associated with telomere shortening during early stages of human hepatocarcinogenesis (small cell changes) [51]. Together, these results indicate that telomere dysfunction and loss of DNA checkpoints could cooperate during cancer initiation in human hepatocarcinogenesis.

Telomerase activation characterizes human hepatocarcinogenesis Studies in mouse models and on primary human cells have shown that activation of telomerase is an important step for tumor progression, allowing the immortalization of transformed cells. In agreement with these experimental data, the majority of human cancers show a reactivation of telomerase (for a review, see [77, 114]). A small percentage (80% of human HCC. Telomerase activation is required to stabilize critically short telomeres in HCC, thus allowing tumor progression. In contrast, telomerase activity is merely detectable in normal liver. These results predict that an inhibition of telomerase could selectively impair tumor cell viability without affecting regenerative

Chromosomal Instability Tumor Initiation

Telomere Shortening Telomere Dysfunction Telomerase Activators

Activation of DNA Damage Checkpoints

Telomerase Inhibitors

Telomerase Activation Tumor Progression

Checkpoint Inhibitors

Cirrhosis

Clonal Selection

Tumor Progression

Figure 68.4 Telomeres and telomerase represent therapeutic targets in chronic liver disease and hepatocarcinogenesis. (i) Telomerase inhibition could impair progression of HCC but would have no side effects on the regenerative capacity of telomerase-negative liver tissue. (ii) Telomerase activators could improve liver regenerative reserve by preventing critical telomere shortening and telomere dysfunction in hepatocytes. Telomerase activators could prevent the induction of chromosomal instability and tumor initiation but could promote the progression of early (telomerase-negative) tumor lesions. (iii) Checkpoint inhibitors could improve liver regeneration and the progression of cirrhosis in liver with short telomeres. Checkpoint inhibitors could induce chromosomal instability and tumor initiation but could ameliorate the selective pressure for an outgrow of malignant clones at the cirrhosis stage

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reserve of non-transformed liver cells. Xenotransplantation experiments on human HCC cell lines have provided a proof-of-principle that telomerase inhibition can impair HCC progression [145]. In addition, it is possible that telomere shortening in response to telomerase inhibition could increase the chemosensitivity of HCC, as was shown in transformed murine cells [146]. Therefore, the combination between chemotherapy and telomerase inhibitors could be a promising approach for cancer treatment. Telomerase inhibitors are currently being tested in phase I/II clinical trials in lymphoma and breast cancer [147]. Study results and initiation of trials in HCC are awaited. 2. Telomerase activation/telomere stabilization for treatment of cirrhosis: Experimental data from mouse models and human liver biopsies indicate that telomere shortening limits the regenerative capacity of hepatocytes, thus promoting the development of cirrhosis in response to chronic liver disease. The activation of telomerase could represent a therapeutic approach stabilizing telomeres in chronically damaged liver in order to improve liver regeneration and prevent cirrhosis formation (Figure 68.4). Experiments on telomerase knockout mice have provided a proof for this concept, showing that adenoviral-mediated, transient activation of telomerase can improve liver regeneration and impair the activation of stellate cells and fibrogenesis in telomere dysfunction mouse liver exposed to chronic damage [36]. The clinical use of telomerase activation strategies needs to be carefully evaluated, given the association of telomerase activation and human cancer. It is possible that telomerase activation would allow progression of early telomerase-negative cancer lesions. It is conceivable that a transient activation of telomerase would be safer than long-term activation of the enzyme. A transient activation of telomerase, increasing the regenerative capacity of cells by only a few cell divisions, could have a significant effect on liver regenerative reserve, but should have little effect on progression of microscopic early cancer lesions. A technical drawback is that pharmaceutical compounds for efficient activation of telomerase are lacking. Adenoviral-mediated gene transfer does not represent an option for clinical use because of toxic side effects of adenoviral vectors, including hepatotoxicity. 3. Inhibition of DNA damage checkpoints for treatment of cirrhosis (Figure 68.4): Studies on telomerase knockout mice have shown that DNA damage checkpoints mediate the adverse effects of telomere dysfunction on organ maintenance and lifespan. These studies have shown that deletion of some checkpoint components can improve stem cell function, organ maintenance, and lifespan of telomere dysfunctional mice without increasing the rate of cancer formation [35, 42]. Given the evidence for telomere shortening and checkpoint activation in human cirrhosis [44,51–55],

an inhibition of checkpoint components could represent a promising approach for treatment of cirrhosis patients. Pharmacological compounds for checkpoint inhibition have yet to be identified and tested in preclinical studies. In addition to these therapeutic approaches, telomeres and telomerase could serve as biomarkers indicating the risk of disease progression and cancer development. A variety of studies indicate that telomerase activity may serve as a diagnostic and prognostic marker for HCC [115–138]. Given the role of telomere dysfunction in limiting cell proliferation and inducing chromosomal instability, it is conceivable that telomere shortening may serve as a biomarker predicting the progression of chronic liver disease and the risk of cirrhosis or HCC development. Prospective trials have yet to be conducted. Biomarkers of telomere dysfunction and DNA damage have been identified and serum levels of these biomarkers show an increased expression in cirrhosis patients compared with non-cirrhotic patients with chronic hepatitis [65]. Studies in mouse models have shown that telomere dysfunction correlates better with regenerative defects compared with mean telomere length [148], indicating that biomarkers of telomere dysfunction should be included in future studies. Together, there is increasing evidence for a functional role of telomere shortening and telomerase in chronic liver disease, cirrhosis, and hepatocarcinogenesis. A chapter on telomeres and telomerase was first included in the “Horizon” section of the last edition of The Liver: Biology and Pathobiology in 2001 (Chapter 67). The field has made substantial progress since then and we anticipate seeing translational approaches and clinical trials investigating potential use of telomerase/telomere therapeutics for the treatment of cirrhosis and HCC over the coming years.

ACKNOWLEDGEMENT This work was supported by a grant of the Deutsche Krebshilfe (tumor stem cell consortium).

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87. Wilkens, L., Flemming, P., Gebel, M. et al. (2004) Induction of aneuploidy by increasing chromosomal instability during dedifferentiation of hepatocellular carcinoma. Proc Natl Acad Sci U S A, 101, 1309–14. 88. Artandi, S.E., Chang, S., Lee, S.L. et al. (2000) Telomere dysfunction promotes non-reciprocal translocations and epithelial cancers in mice. Nature, 406, 641–45. 89. DePinho, R.A. (2000) The age of cancer. Nature, 408, 248–54. 90. Feldser, D.M. and Greider, C.W. (2007) Greider short telomeres limit tumor progression in vivo by inducing senescence. Cancer Cell , 11, 461–69. 91. Ju, Z., Jiang, H., Jaworski, M. et al. (2007) Telomere dysfunction induces environmental alterations limiting hematopoietic stem cell function and engraftment. Nat Med , 13, 742–47. 92. Krtolica, A. and Campisi, J. (2003) Integrating epithelial cancer, aging stroma and cellular senescence. Adv Gerontol , 11, 109–16. 93. Bilousova, G., Marusyk, A., Porter, C.C. et al. (2005) Impaired DNA replication within progenitor cell pools promotes leukemogenesis. PLoS Biol , 3, e401. 94. Marusyk, A. and DeGregori, J. (2008) Declining cellular fitness with age promotes cancer initiation by selecting for adaptive oncogenic mutations. Biochim Biophys Acta, 1785, 1–11. 95. El-Serag, H.B. and Rudolph, K.L. (2007) Hepatocellular carcinoma: epidemiology and molecular carcinogenesis. Gastroenterology, 132, 2557–76. 96. Blanco, R., Mu˜noz, P., Flores, J.M. et al. (2007) Telomerase abrogation dramatically accelerates TRF2-induced epithelial carcinogenesis. Genes Dev , 21, 206–20. 97. Oh, B.K., Kim, Y.J., Park, C. et al. (2005) Up-regulation of telomere-binding proteins, TRF1, TRF2, and TIN2 is related to telomere shortening during human multistep hepatocarcinogenesis. Am J Pathol , 166, 73–80. 98. Sangiovanni, A., Del Ninno, E., Fasani, P. et al. (2004) Increased survival of cirrhotic patients with a hepatocellular carcinoma detected during surveillance. Gastroenterology, 126, 1005–14. 99. Isokawa, O., Suda, T., Aoyagi, Y. et al. (1999) Reduction of telomeric repeats as a possible predictor for development of hepatocellular carcinoma: convenient evaluation by slot-blot analysis. Hepatology, 30, 408–12. 100. O’Sullivan, J.N., Bronner, M.P., Brentnall, T.A. et al. (2002) Chromosomal instability in ulcerative colitis is related to telomere shortening. Nat Genet , 32, 280–84. 101. Ohyashiki, J.H., Ohyashiki, K., Fujimura, T. et al. (1994) Telomere shortening associated with disease evolution patterns in myelodysplastic syndromes. Cancer Res, 54, 3557–60. 102. Wu, X., Amos, C.I., Zhu, Y. et al. (2003) Telomere dysfunction: a potential cancer predisposition factor. J Natl Cancer Inst , 95, 1211–18.

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119. Takahashi, H., Kitamoto, M., Takahashi, S. et al. (2000) Precancerous hepatic nodules had significant levels of telomerase activity determined by sensitive quantitation using a hybridization protection assay. Cancer, 88, 312–17. 120. Koutoula, V., Hytiroglou, P., Pyrpasopoulou, A. et al. (2002) Expression of human telomerase reverse transcriptase in regenerative and precancerous lesions of cirrhotic livers. Liver, 22, 57–69. 121. Komine, F., Shimojima, M., Moriyama, M. et al. (2000) Telomerase activity of needle-biopsied liver samples: its usefulness for diagnosis and judgement of efficacy of treatment of small hepatocellular carcinoma. J Hepatol , 32, 235–41. 122. Kojima, H., Yokosuka, O., Kato, N. et al. (1999) Quantitative evaluation of telomerase activity in small liver tumors: analysis of ultrasonography-guided liver biopsy specimens. J Hepatol , 31, 514–20. 123. Harada, K., Yasoshima, M., Ozaki, S. et al. (2001) PCR and in situ hybridization studies of telomerase subunits in human non-neoplastic livers. J Pathol , 193, 210–17. 124. Kawakami, Y., Kitamoto, M., Nakanishi, T. et al. (2000) Immuno-histochemical detection of human telomerase reverse transcriptase in human liver tissues. Oncogene, 19, 3888–93. 125. Takahashi, S., Kitamoto, M., Takaishi, H. et al. (2000) Expression of telomerase component genes in hepatocellular carcinomas. Eur J Cancer, 36, 496–502. 126. Toshikuni, N., Nouso, K., Higashi, T. et al. (2000) Expression of telomerase-associated protein 1 and telomerase reverse transcriptase in hepatocellular carcinoma. Br J Cancer, 82, 833–37. 127. Nakashio, R., Kitamoto, M., Tahara, H. et al. (1997) Significance of telomerase activity in the diagnosis of small differentiated hepatocellular carcinoma. Int J Cancer, 74, 141–47. 128. Hytiroglou, P., Kotoula, V., Thung, S.N. et al. (1998) Telomerase activity in precancerous hepatic nodules. Cancer, 82, 1831–38. 129. Nakayama, J., Tahara, H., Tahara, E. et al. (1998) Telomerase activation by hTRT in human normal fibroblasts and hepatocellular carcinoma. Nat Genet , 18, 65–68. 130. Ide, T., Tahara, H., Nakshio, R. et al. (1996) Telomerase in hepatocellular carcinogenesis. Hum Cell , 9, 283–86. 131. Kishimoto, K., Fujimoto, J., Takeuchi, M. et al. (1998) Telomerase activity in hepatocellular carcinoma and adjacent liver tissues. J Surg Oncol , 69, 119–24. 132. Iizuka, N., Mori, N., Tamesa, T. et al. (2003) Telomerase activity and Nm23-H2 protein expression in hepatocellular carcinoma. Anticancer Res, 23, 43–47. 133. Nouso, K., Urabe, Y., Higashi, T. et al. (1996) Telomerase as a tool for the differential diagnosis of human hepatocellular carcinoma. Cancer, 78, 232–36. 134. Ohta, K., Kanamaru, T., Morita, Y. et al. (1997) Telomerase activity in hepatocellular carcinoma as a predictor of postoperative recurrence. J Gastroenterol , 32, 791–96.

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135. Satra, M., Gatselis, N., Iliopoulos, D. et al. (2007) Real-time quantification of human telomerase reverse transcriptase mRNA in liver tissues from patients with hepatocellular cancer and chronic viral hepatitis. J Viral Hepat , 14, 41–47. 136. Hiyama, E., Yamaoka, H., Matsunaga, T. et al. (2004) High expression of telomerase is an independent prognostic indicator of poor outcome in hepatoblastoma. Br J Cancer, 91, 972–79. 137. Kobayashi, T., Kubota, K., Takayama, T. et al. (2001) Telomerase activity as a predictive marker for recurrence of hepatocellular carcinoma after surgery. Am J Surg, 181, 284–88. 138. Llovet, J.M., Chen, Y., Wurmbach, E. et al. (2006) A molecular signature to discriminate dysplastic nodules from early hepatocellular carcinoma in HCV cirrhosis. Gastroenterology, 131, 1758–67. 139. Suda, T., Isokawa, O., Aoyagi, Y. et al. (1998) Quantitation of telomerase activity in hepatocellular carcinoma: a possible aid for a prediction of recurrent diseases in the remnant liver. Hepatology, 27, 402–6. 140. Wang, J., Xie, L.Y., Allan, S. et al. (1998) Myc activates telomerase. Genes Dev , 12, 1769–74. 141. Oh, B.K., Kim, Y.J., Park, Y.N. et al. (2006) Quantitative assessment of hTERT mRNA expression in dysplastic nodules of HBV-related hepatocarcinogenesis. Am J Gastroenterol , 101, 831–38.

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142. Liu, Y.C., Chen, C.J., Wu, H.S. et al. (2004) Telomerase and c-myc expression in hepatocellular carcinomas. Eur J Surg Oncol , 30, 384–90. 143. Ferber, M.J., Montoya, D.P., Yu, C. et al. (2003) Integrations of the hepatitis B virus (HBV) and human papillomavirus (HPV) into the human telomerase reverse transcriptase (hTERT) gene in liver and cervical cancers. Oncogene, 22, 3813–20. 144. Paterlini-Br´echot, P., Saigo, K., Murakami, Y. et al. (2003) Hepatitis B virus-related insertional mutagenesis occurs frequently in human liver cancers and recurrently targets human telomerase gene. Oncogene, 22, 3911–16. 145. Djojosubroto, M.W., Chin, A.C., Go, N. et al. (2005) Telomerase antagonists GRN163 and GRN163L inhibit tumor growth and increase chemosensitivity of human hepatoma. Hepatology, 42, 1127–36. 146. Lee, K.H., Rudolph, K.L., Ju, Y.J. et al. (2001) Telomere dysfunction alters the chemotherapeutic profile of transformed cells. Proc Natl Acad Sci U S A, 98, 3381–86. 147. Harley, C.B. (2008) Telomerase and cancer therapeutics. Nat Rev Cancer, 8, 167–79. 148. Hemann, M.T., Strong, M.A. et al. (2001) The shortest telomere, not average telomere length, is critical for cell viability and chromosome stability. Cell , 107, 67–77.

69

Treatment of Cirrhosis with Vitamin A-coupled Liposomes Carrying siRNA against Heat Shock Protein 47 Yoshiro Niitsu1, Yasushi Sato2, Kazuyuki Murase2 and Junji Kato2 1 Department

of Molecular Target Exploration, Sapporo Medical University, Sapporo, Japan 2 Fourth Department of Internal Medicine, Sapporo Medical University, Sapporo, Japan

INTRODUCTION Cirrhosis is the ultimate pathological feature of all forms of chronic hepatic injury, including viral infection, immune-driven process, metal overload, non-alcoholic steatohepatitis, alcoholic hepatitis, or biliary obstruction. Irrespective of cause, destruction of hepatic architecture and vascular structures with deposition of fibrotic tissue leads to functional decompensation [1]. Most therapies for chronic liver disease target the causative factors; however, once fibrosis advances to cirrhosis, they are rarely effective and there is currently no

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

approved antifibrotic therapy for liver cirrhosis in humans, mainly due to a lack of specificity to target molecules and/or cells relevant to fibrogenesis. The principal cell type responsible for liver fibrosis is the hepatic stellate cell (HSC), a resident perisinusoidal cell that stores vitamin A (VA) and, after stimulation by reactive oxygen intermediates and/or inflammatory cytokines, actively secretes procollagen protein with the aid of a collagen-specific chaperone, heat shock protein 47 (HSP47) [2]. Based on these considerations, we recently developed a novel strategy to treat cirrhosis, employing

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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small interfering RNA (siRNA)–HSP47 encapsulated in VA-coupled liposomes which were specifically delivered to HSC after intravenous injection.

is also protease hypersensitive has been reported with HSP47−/− embryonic stem cells, suggesting that HSP47 is required for maturation of both type I and IV collagen in ER [5]. Upon activation, HSCs produce not only collagen, but also metalloproteinase [mainly metalloproteinase 2 (MMP-2) and membrane-bound metalloproteinase, MT1-MMP (MMP-14), and their inhibitor (tissue inhibitor of metalloproteinase, TIMP] [6–9]. When the liver undergoes injury, the net balance of over-produced collagen plus TIMP to the increased MMP activity becomes positive, resulting in accumulation of collagen in the extracellular matrix. A characteristic feature of quiescent HSCs is cytoplasmic storage of VA which is gradually lost during the activation process [10]. Although the biological implication of VA storage, particularly in fibrogenesis, is not fully understood, it serves as a good marker for HSCs in normal liver [3]. The mode of uptake of VA by HSC has long been debated. In circulation, VA exists as a complex with retinol-binding protein (RBP) and in cytoplasm with cellular retinol-binding protein (CRBP). The transmembrane transfer of VA from RBP to CRBP has been postulated to be receptor-driven uptake or passive diffusion [11]. Recent studies, however, are mostly indicative of the former mechanism based on the receptor-binding assays of GRX cells (HSC cell line) [12] and most recently, the receptor gene for RBP–VA complex was cloned [13], although its role in HSCs has not been directly demonstrated. It has also not been elucidated whether HSCs lose expression of the receptor as they lose cytoplasmic VA droplets during the activation process.

MECHANISM OF HEPATIC FIBROSIS WITH SPECIAL REFERENCE TO THE ROLE OF HSC AND HSP47 HSCs comprise approximately 5% of resident cells in normal liver and, following injury, undergo proliferation and activation in response to the stimulation by reactive oxygen species (ROS) and cytokines from inflammatory cells, Kupffer cells, and HSCs themselves [3]. Activated HSCs in turn synthesize and secrete procollagen of which both the N-terminus and C-terminus are extracellularly processed by procollagen endopeptidases, resulting in the formation of insoluble collagen fibrils. During maturation of procollagen molecules in the endoplasmic reticulum (ER) of HSCs, a collagen-specific molecular chaperone, HSP47, is involved in the correct folding of procollagen (Figure 69.1). HSP47 is essential for the maturation of procollagen molecules in ER because HSP47-knockout mice (HSP47−/− ) are embryonically lethal and display disrupted basement membrane, and fibroblasts of these mice poorly secrete type I collagen, which is more susceptible to protease digestion than that from normal fibroblasts [4]. Similar retarded secretion of type IV collagen which

fibrosis Hepatic stellate cell(HSC) libosome Procollagen mRNA

Collagen Heat shock protein47(HSP47) mRNA

Type I~V, procollagen α chain

HSP47 Secretion control HSP47

ER

procollagen

Figure 69.1 Role of HSP47 in collagen production by HSCs. HSP47 is a specific chaperone for collagen and facilitates proper folding of the procollagen molecule in ER.

69: TREATMENT OF CIRRHOSIS WITH VITAMIN A-COUPLED LIPOSOMES

PREVIOUS ANTIFIBROTIC APPROACHES The fact that liver fibrosis in animals [14] and humans [15] can regress when the cause of chronic hepatic damage is eliminated suggests that fibrosis can be reversed, most likely by activity of MMP. Based on this, a number of therapeutic approaches have been explored; several are currently undergoing clinical trials. They may be categorized by mechanisms such as attenuating HSC activation [16, 17], inducing HSC apoptosis [18–21], inhibiting collagen synthesis [22], and accelerating ECM degradation [14]. However, there are no therapeutic drugs licensed for primary treatment of liver fibrosis. The most critical obstacle in developing primary anti-fibrotic modality is the difficulty of assuring specific targeting of therapeutic means(s) or drug(s) to molecules and/or cells which are responsible for fibrosis. Because patients with cirrhosis are supposed to undergo continuous or repeated therapy, modalities without specificity are always at risk of causing sever adverse effects.

TREATMENT OF RAT LIVER FIBROSIS WITH VA-LIPOSOME-SIRNA HSP47 Suppression of collagen by siRNA–HSP47 Since excess deposition of collagen in cirrhotic liver results from an imbalance between collagen synthesis and degradation by MMP, we therefore proposed to treat cirrhosis by amending the imbalance of collagen metabolism. We initially targeted messenger RNA (mRNA) of HSP47, a specific chaperone of a diverse range of collagen types, with ribozyme in which the hammerhead sequence was ligated to a tRNAval promoter [23]. Treatment of human primary fibroblasts (HPFs) with the ribozyme brought about not only cleavage of HSP47 mRNA but also substantial suppression of type I procollagen secretion. However, since the target sequence for ribozyme of this type on HSP47 mRNA of rodents, which may be used for pre-clinical studies, was not identifiable, we chose a strategy to knock down HSP47 mRNA by siRNA, which is easier to construct than ribozyme and of which target sequences were identified on both human and rodent HSP47 mRNAs. To examine the silencing effect of siRNA on HSP47 mRNA expression, we transduced normal rat kidney (NRK) cells, which have bean used to study collagen production, with three siRNA (types A, B, and C) against mRNA encoding rat gp46, a homolog of

1123

HSP47 (Figure 69.2) [24]. The results obtained suggested that type A is the most potent siRNA (Figure 69.2a). Using type A siRNA, we confirmed the dose dependence of the silencing effect, durative silencing effect (Figure 69.2b), and suppressive effect on collagen production (Figure 69.2c). Of note, our attempt to shut off collagen synthesis by applying siRNA to procollagen mRNA failed, for unknown reasons.

Targeting HSC with VA–liposome To deliver siRNA gp46 specifically to HSC, we took advantage of the fact that HSCs accumulate VA; and targeted HSCs with VA-coupled liposome in which siRNA gp46 was encapsulated (VA–lip–siRNA gp46) (Figure 69.3). Uncertainty with this strategy was whether activated HSCs in fibrous tissue would take up VA–lip–siRNA gp46 since HSCs lose VA droplets in cytoplasm as they undergo activation. We therefore examined uptake of VA–lip–siRNA gp46 coupled to carboxyfluorescein (FAM) by primary rat HSCs (pHSCs) which were activated during incubation for 10 days. In cells treated with VA–lip–siRNA gp46–FAM, fluorescence appeared as a faint granular pattern in the cytoplasm at 30 minutes, and as a denser granular pattern in the perinuclear region at 2 hours. In contrast, no green fluorescence was seen in the cytoplasm of cell treated with lip-siRNA gp46–FAM after 30 minutes. Moreover, perinuclear fluorescence at 2 hours was very faint (Plate 69.1). FACS analyses also demonstrated a significantly higher fluorescence intensity of pHSCs treated with VA–lip–siRNA gp46–FAM in the presence of RBP than that in the absence of RBP, indicating that activated pHSCs take up VA–lip through the RBP receptor [24]. To assure specificity of VA–lip–siRNA gp46–FAM uptake by HSCs, we conducted a control experiment. Primary fibroblasts isolated from rat skin, which lack RBP receptor on the plasma membrane, did not show any cytoplasmic fluorescence after treatment with the complex [24]. Transduced siRNA gp46 also effectively suppressed collagen secretion from pHSCs [24]. On the basis of these in vitro data, we extended exploration to confirm specific delivery of intravenously (i.v.) injected VA–lip–siRNA gp46–FAM to HSCs in cirrhotic liver of rats treated with dimethylnitrosamine (DMN). Fluorescence of siRNA gp46–FAM was identified in liver predominantly in the region that stained positive for α-smooth muscle actin (α-SMA; an indicator of activated HS cells) (Plate 69.2) [24]. In other organs, such as lung and spleen, few cells, which were identified as macrophages, were positive for fluorescence [24]. The retina, which requires VA for photoreceptor function, showed almost no fluorescence emission [24]. Specific delivering of VA–lip–siRNA gp46 to HSCs in cirrhotic

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THE LIVER: TREATMENT OF RAT LIVER FIBROSIS WITH VA-LIPOSOME-SIRNA HSP47

(a)

(b)

(c)

Figure 69.2 Suppressive effect of siRNA gp46 on gp46 expression and collagen synthesis (a) Western blotting was used to analyze the expression of gp46 or β-actin (normalization control) in NRK cells transfected with three types of siRNA gp46 (A, B, and C) or siRNA random using Lipotrust. (b) Dose-dependent inhibition of gp46 expression with siRNA gp46A. (c) Collagen synthesis in NRK cells treated with siRNA gp46 or with siRNA random was assayed by the [3 H]proline incorporation method. [3 H]Proline-incorporated collagen levels in supernatants were assayed 2 days after transfection. (d) Collagen deposition on tissue culture plates in NRK cells was assayed 1 day after transfection by the dye-binding method. Data were expressed as mean ± SD calculated from five times transfections and as a percentage of untreated control. ∗p < 0.01 vs siRNA gp46. NS, not significant [24]. Reproduced from Sato et al ., Nat Biotechnol . 26 (4) (2008)

Figure 69.3 Scheme of RBP receptor-mediated uptake of VA–lip–siRNA gp46 on HSCs. VA–lip–siRNA gp46 is taken up by HSCs via RBP receptors on the cell membrane.

liver was further supported by FACS analysis of cellular components (hepatocyte, Kupffer cell HSCs) isolated from the liver of DMN rats infected with VA–lip–siRNA gp46–Cy5 (cyanine 5) [24], and by tissue-distribution study using radiolabeled liposome carrying siRNA gp46 with [3 H]VA [24]. DMN-treated cirrhotic (day 24 rat) or normal rats (n = 3 per group) received a single i.v. injection of 200 µCi [3 H]VA–lip–siRNA gp46 via the

tail vein. Tissue biodistribution was analyzed 24 hours later.

Fibrosis-resolving effect Treatment of fibrosis was then performed in three rat models of cirrhosis, involving induction by DMN, CCl4 or bile

69: TREATMENT OF CIRRHOSIS WITH VITAMIN A-COUPLED LIPOSOMES

1125

35 × 106

25 × 106 20 × 106 15 × 106 10 × 106

Cirrhotic rat

In te s

e Ey

ar t

le e

He

n

s Sp

ng

ey

Lu

Ki dn

Li ve

r

0

tin e

Normal rat

5 × 106

Br ai n

Radioactivity (cpm/tissue)

30 × 106

Figure 69.4 Effect of iv injected VA–lip–siRNA gp46 on liver histology of cirrhotic rat. Representative photomicrographs of Azan–Mallory-stained liver section (a) obtained from DMN cirrhotic rats treated with five injections of VA–lip–siRNA gp46 (siRNA doses of 0.75 mg kg−1 , three times per week) and from control rats (n = 12 per group). Arrowheads indicate collagen fibrils. Bars represent 100 µm. (b) Hydroxyproline content in the livers. Mean ± SD of 12 rats per group. ∗p < 0.001 vs VA–lip–siRNA gp46. Liver tissues were obtained from rats treated as described in (a) [24]. Reproduced from Sato et al ., Nat Biotechnol . 26 (4) (2008)

Figure 69.5 Survival of DMN-treated rats that were injected i.v. with VA–lip–siRNA gp46 at siRNA doses of 0.75 mg kg−1 three times per week (n = 12 per group). In the control groups, DMN-treated rats were injected with Lip-siRNA gp46, VA–lip–siRNA random, VA–liposomes, VA, PBS or siRNA gp46A (n = 12 for group of 0.75 mg kg−1 siRNA, three times per week). Life-table analyses are presented as a Kaplan-Meyer plot. (∗ ∗ p < 0.0001 compared with control rats) [24]. Reproduced from Sato et al ., Nat Biotechnol . 26 (4) (2008)

duct ligation (BDL). In the DMN model, gp46 expression in liver was significantly suppressed and fibrosis was significantly resolved histologically after five treatments (Figure 69.4) [24]. Treatment also resulted in an improved survival rate (Figure 69.5) [24], suggesting that hepatic insufficiency resulting from an impaired blood supply and regurgitation of toxic bile, due to a massive stricture of the portal area by fibrosis, was reversed by treatment. Nearly complete restoration of normal hepatic architecture was observed at day 70 (44 and 46 days after administration of DMN and VA–lip–siRNA gp46, respectively,

ceased) [24], suggesting that activation of hepatic stem cells occurred during this process. Thus, in addition to the partial hepatectomy model [25], this treatment model may also provide a means by which to study hepatic regeneration. Virtually the same effect with respect to shrinkage of fibrosis was seen in cirrhosis induced by CCl4 [24]. It was previously demonstrated that at longer periods of time (12 weeks) than in our study (8 weeks) the liver remains cirrhotic [26]. It has also been observed that areas of fibrosis that do not undergo remolding are extensively

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THE LIVER: EFFICACY

cross-linked, suggesting that the cross-linked ECM is resistant to MMP [26]. Effect of our approach on 12 weeks fibrosis of CCl4 model is presently under investigation. Effectiveness of treatment was also verified in cirrhosis induced by BDL [24]. An important issue clarified by the BDL experiment was that fibroblasts, which are responsible for fibrosis of the portal area, can also take up VA in a similar manner to HSCs in centrilobular areas, although the two cell types are phenotypically distinct [27].

Molecular mechanism for fibrosis-resolving effect To prove the involvement of MMP, we first measured MMP activity in the cirrhotic liver specimen from DMN rats which had been treated with the complex three times and found that MMP activity in the specimen remained as high as in normal rat liver [24]. We then treated DMN rats with a TIMP-1 expression vector that was encapsulated in VA–lip to suppress MMP activity in the tissue and subsequently with VA–lip–siRNA gp46. Animals all died on the same experimental days as non-treated DMN rats (unpublished observation). Hence suppression of collagen secretion and resolution of predeposited collagen in combination are considered to be a mechanism for fibrosis-resolving effect of our modality (a primary mechanism). However, when we carefully considered the fact that the resolution of fibrosis occurred unexpectedly rapidly and efficiently with only five times treatment [24], we proposed an additional mechanism that might be related to the fate of HSCs themselves. Therefore, we next examined the expression of α-SMA, a marker for activated HSCs, by immunohistochemical staining and found a significantly impaired expression in the treated DMN rats, compared with that in non-treated DMN rats [24], indicating the disappearance of activated HSCs in the treated rats. The disappearance of the activated HSCs was due to apoptosis since tunnel-positive HSCs were increased in the liver of DMN rats treated with the complex [24]. Further in vitro experiments suggested that activated HSCs required a critical signal from collagen for their survival and therefore underwent apoptosis under the situation in which MMP degraded collagen. Such apoptosis induced by anchorage loss may be referred to as “anoikis.” The survival signal from collagen was found to involve the PI3 kinase/AKT pathway (unpublished observation). With respect to the sensitivity of quiescent HSCs to anoikis, it should be very low, if it even exists, since near-normal architecture with the presence of very few quiescent HSCs in the centrilobular area was retained after treatment in all three types of rat models [24] and no apparent histological damage of the liver was observed in

normal rats treated with repeated injections (12 times) of VA–lip–siRNA gp46 (unpublished observation). The fact the quiescent HSCs are insensitive to anoikis not only raises an important question as to the molecular basis for the difference between survival signals of quiescent HSCs and activated HSCs, but also indicates minimal adverse effects of this approach in future clinical use.

Off-target effect and immune response There are multiple barriers to be overcome in siRNA therapeutics. Major considerations are off-targeting due to silencing of genes sharing partial homology with the siRNA, and immune stimulation due to components of the innate immune system by the siRNA duplex [28]. We used three independent siRNAs (types A, B, and C) against the same target (gp46) mRNA and found comparable gene silencing efficacy in vitro (Figure 69.2), and antifibrotic effects in vivo [24], suggesting that the phenotype observed with down-regulation of gp46 was related to gp46 knockdown and not to a bystander effect (off-target effect) of the siRNA sequence. To circumvent immune responses such as induction of interferon-α (IFN-α), we used siRNA with a 2-nucleotide 3′ overhang, shown to impair activation of the transcription factor IRF3 [29]. In addition, our siRNA did not contain the 5-triphosphate of the T7-transcript (manufacture’s information) or distinct sequences of the so-called CpG motif, which reportedly plays a role in IFN induction [30]. In fact, there was no elevation of either IFN-α mRNA in the liver or tumor necrosis factor-α (TNF-α) or IL-12 in the circulation of rats treated with VA–lip–siRNA gp46 [24]. However, as these modifications of siRNA do not prevent triggering of all immune responses, it is possible that the low immune activation in the present investigation is relevant to RBP receptor-mediated uptake of siRNA, in addition to modifications of siRNA structure.

EFFICACY Although surmounting the obstacles described is critical to the use of siRNAs as therapeutic agents, designing siRNA constructs which efficiently suppress the target molecule is also important. Because synthetic RNA duplexes 25–30 nucleotides in length are up to 100-fold more potent than are corresponding conventional 21-mer siRNA, we used a 27-nucleotide RNA duplex with 2-nucleotide 3′ overhangs [31]. The VA-coupled liposome system for delivery of siRNA appeared to be efficient compared with previously reported methods using conventional liposomes. VA-coupled liposomes produced biological effects at 0.75 mg kg−1 liposome, whereas in previous reports [32–35] ∼8–160 mg kg−1 were necessary. Furthermore,

69: TREATMENT OF CIRRHOSIS WITH VITAMIN A-COUPLED LIPOSOMES

most other studies [36] used approximately a 1:10 ratio of solution to total blood volume (i.e. 200 µl/2 ml blood volume in mice; 2000 µl/20 ml blood volume in rats) to dissolve liposomes containing siRNA for a single bolus injection. However, we used only 200 µl per injection for 200 g rats (1:100 blood volume) [24]. This suggests that hydrodynamic pressure did not contribute to in vivo transduction of siRNA, whereas, in previous studies, siRNAs may have been forcedly transduced by hydrodynamic pressure. This may explain why background fluorescence for FAM was weak and showed a low non-specific distribution of radioactive VA-coupled liposomes in tissues other than liver. Furthermore, the siRNA dose used was much less than doses previously shown to have in vivo therapeutic effects [32, 33, 37]. This may also be related to the use of VA-coupled liposomes to deliver siRNA gp46 preferentially to HSCs.

POSSIBLE CLINICAL APPLICATION Although the approach being proposed will provide novel and potent therapy for cirrhosis, human trials must be reserved until several tasks have been completed. The design of siRNA most suitable for human HSP47 and the construction of a stable and relatively uniform carrier for siRNA are mandatory. Substitution of liposome with a better carrier particle, such as biodegradable polymer, may be one option. Other important tasks include confirmation of the silencing effect of siRNA HSP47 on HSP47mRNA in the human HSC cell line and the adverse effect of siRNA HSP47 in primates. As discussed above, the effect of on old scarring fibrosis which contains cross-linked collagen is crucial for clinical application, since fibrotic alterations in rodent models may be minor compared with established cirrhosis in humans.

POTENTIAL USE OF THIS COMPLEX TO DELIVER OTHER AGENTS TO HSCS Our complex may be used to deliver other agents specifically to HSCs. These agents include other siRNAs relevant to collagen metabolism or HSC survival, or antifibrotic small molecules. Some cytokines or growth factors which have antifibrotic activity, such as hepatic growth factors, may also be delivered specifically to HSCs by VA–lip. More importantly, since HSCs appear to play an essential role in hepatic regeneration [38], VA–lip may be useful for delivering factors which facilitate repair of injured liver. In addition, merging of siRNA gp46FAM (green) with a smooth muscle expression in HSCs (red)

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in Plate 69.1 suggests the potential use of this targeting method for imaging hepatic fibrosis.

APPLICATION OF OUR STRATEGY TO OTHER ORGAN FIBROSIS Recent immunohistochemical and transcriptome studies revealed cells similar to HSCs in other organs, such as the pancreas, lung, kidney, and intestine [39–42], in which pathways of fibrogenesis are thought to be the same. These cells store VA droplets in the same manner as do HSC, and undergo activation when tissues are injured. Hence it is highly plausible that our modality is also effective to against fibrosis in these organs. In a preliminary experiment, we demonstrated that in the dibutyltin dichloride (DBTC) model (pancreas), after fibrosis was established, the injection of VA–lip–siRNA gp46 for 20 days dramatically improved fibrosis with a significant recovery of whole pancreas weight (unpublished observation).

CONCLUSION Employing VA–lip–siRNA gp46, we demonstrated successful resolution of fibrosis and induction of apoptosis of HSCs with regeneration of damaged livers in three rat cirrhosis models. In this approach, we suppressed de novo production of collagen, thereby increased the relative activity of MMP-resolved predeposited collagen in the tissue, and induced anoikis of HSCs. Alternatives, in this context, could be suppression of the MMP inhibitor (TIMP) by siRNA, or forced over-expression of MMP by its transgene; both siRNA and transgene carried in VA–liposome. Hence the feasibility of these alternatives in combination with or without our modality should be tested. Further tests are crucial to ascertain whether such approaches are effective for the resolution of cross-linked collagen before clinical application can be conducted. The proposed technology provides the possibility to improve our understanding, diagnosis and treatment of fibrosis.

REFERENCES 1. Schuppan, D. and Afdhal, N.H. (2008) Liver cirrhosis. Lancet , 371, 838–51. 2. Friedman, S.L. (2000) Molecular regulation of hepatic fibrosis, an integrated cellular response to tissue injury. J Biol Chem, 275, 2247–50. 3. Friedman, S.L. (2008) Hepatic stellate cells: protean, multifunctional, and enigmatic cells of the liver. Physiol Rev , 88, 125–72.

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4. Nagai, N., Hosokawa, M., Itohara, S. et al. (2000) Embryonic lethality of molecular chaperone hsp47 knockout mice is associated with defects in collagen biosynthesis. J Cell Biol , 150, 1499–506. 5. Matsuoka, Y., Kubota, H., Adachi, E. et al. (2004) Insufficient folding of type IV collagen and formation of abnormal basement membrane-like structure in embryoid bodies derived from Hsp47-null embryonic stem cells. Mol Biol Cell , 15, 4467–75. 6. Theret, N., Lehti, K., Musso, O. et al. (1999) MMP2 activation by collagen I and concanavalin A in cultured human hepatic stellate cells. Hepatology, 30, 462–68. 7. Benyon, R.C., Hovell, C.J., Da Gaca, M. et al. (1999) Progelatinase A is produced and activated by rat hepatic stellate cells and promotes their proliferation. Hepatology, 30, 977–86. 8. Arthur, M.J., Friedman, S.L., Roll, F.J. et al. (1989) Lipocytes from normal rat liver release a neutral metalloproteinase that degrades basement membrane (type IV) collagen. J Clin Invest , 84, 1076–85. 9. Iredale, J.P., Benyon, R.C., Arthur, M.J. et al. (1996) Tissue inhibitor of metalloproteinase-1 messenger RNA expression is enhanced relative to interstitial collagenase messenger RNA in experimental liver injury and fibrosis. Hepatology, 24, 176–84. 10. Friedman, S.L., Wei, S. and Blaner, W.S. (1993) Retinol release by activated rat hepatic lipocytes: regulation by Kupffer cell-conditioned medium and PDGF. Am J Physiol , 264, G947–52. 11. Flower, D.R. (2000) Beyond the superfamily: the lipocalin receptors. Biochim Biophys Acta, 1482, 327–36. 12. Fortuna, V.A., Martucci, R.B., Trugo, L.C. et al. (2003) Hepatic stellate cells uptake of retinol associated with retinol-binding protein or with bovine serum albumin. J Cell Biochem, 90, 792–805. 13. Kawaguchi, R., Yu, J., Honda, J. et al. (2007) A membrane receptor for retinol binding protein mediates cellular uptake of vitamin A. Science, 315, 820–25. 14. Iimuro, Y., Nishio, T., Morimoto, T. et al. (2003) Delivery of matrix metalloproteinase-1 attenuates established liver fibrosis in the rat. Gastroenterology, 124, 445–58. 15. Poynard, T., McHutchison, J., Manns, M. et al. (2002) Impact of pegylated interferon alfa-2b and ribavirin on liver fibrosis in patients with chronic hepatitis C. Gastroenterology, 122, 1303–13. 16. Jonsson, J.R., Clouston, A.D., Ando, Y. et al. (2001) Angiotensin-converting enzyme inhibition attenuates the progression of rat hepatic fibrosis. Gastroenterology, 121, 148–55. 17. Benedetti, A., Di Sario, A., Casini, A. et al. (2001) Inhibition of the Na+ /H+ exchanger reduces rat hepatic stellate cell activity and liver fibrosis: an in vitro and in vivo study. Gastroenterology, 120, 545–56. 18. Wright, M.C., Issa, R., Smart, D.E. et al. (2001) Gliotoxin stimulates the apoptosis of human and rat hepatic stellate cells and enhances the resolution of liver fibrosis in rats. Gastroenterology, 121, 685–98.

19. Taimr, P., Higuchi, H., Kocova, E. et al. (2003) Activated stellate cells express the TRAIL receptor-2/death receptor-5 and undergo TRAIL-mediated apoptosis. Hepatology, 37, 87–95. 20. Anan, A., Baskin-Bey, E.S., Bronk, S.F. et al. (2006) Proteasome inhibition induces hepatic stellate cell apoptosis. Hepatology, 43, 335–44. 21. Oakley, F., Meso, M., Iredale, J.P. et al. (2005) Inhibition of inhibitor of kappaB kinases stimulates hepatic stellate cell apoptosis and accelerated recovery from rat liver fibrosis. Gastroenterology, 128, 108–20. 22. Galli, A., Crabb, D.W., Ceni, E. et al. (2002) Antidiabetic thiazolidinediones inhibit collagen synthesis and hepatic stellate cell activation in vivo and in vitro. Gastroenterology, 122, 1924–40. 23. Hagiwara, S., Nakamura, K., Hamada, H. et al. (2003) Inhibition of type I procollagen production by tRNAVal CTE-HSP47 ribozyme. J Gene Med , 5, 784–94. 24. Sato, Y., Murase, K., Kato, J. et al. (2008) Resolution of liver cirrhosis using vitamin A-coupled liposomes to deliver siRNA against a collagen-specific chaperone. Nat Biotechnol , 26 (4), 431–42. 25. Fausto, N., Campbell, J.S. and Riehle, K.J. (2006) Liver regeneration. Hepatology, 43, S45–53. 26. Issa, R., Zhou, X., Constandinou, C.M. et al. (2004) Spontaneous recovery from micronodular cirrhosis: evidence for incomplete resolution associated with matrix cross-linking. Gastroenterology, 126, 1795–808. 27. Tuchweber, B., Desmouliere, A., Bochaton-Piallat, M.L. et al. (1996) Proliferation and phenotypic modulation of portal fibroblasts in the early stages of cholestatic fibrosis in the rat. Lab Invest , 74, 265–78. 28. Jackson, A.L., Bartz, S.R., Schelter, J. et al. (2003) Expression profiling reveals off-target gene regulation by RNAi. Nat Biotechnol , 21, 635–37. 29. Marques, J.T., Devosse, T., Wang, D. et al. (2006) A structural basis for discriminating between self and nonself double-stranded RNAs in mammalian cells. Nat Biotechnol , 24, 559–65. 30. Marques, J.T. and Williams, B.R. (2005) Activation of the mammalian immune system by siRNAs. Nat Biotechnol , 23, 1399–405. 31. Kim, D.H., Behlke, M.A., Rose, S.D. et al. (2005) Synthetic dsRNA Dicer substrates enhance RNAi potency and efficacy. Nat Biotechnol , 23, 222–26. 32. Zimmermann, T.S., Lee, A.C., Akinc, A. et al. (2006) RNAi-mediated gene silencing in non-human primates. Nature, 441, 111–14. 33. Yano, J., Hirabayashi, K., Nakagawa, S. et al. (2004) Antitumor activity of small interfering RNA/cationic liposome complex in mouse models of cancer. Clin Cancer Res, 10, 7721–26. 34. Walsh, T.J., Finberg, R.W., Arndt, C. et al. National Institute of Allergy and Infectious Diseases Mycoses Study Group (1999) Liposomal amphotericin B for empirical therapy in patients with persistent fever and neutropenia. N Engl J Med , 340, 764–71. 35. Takahashi, N., Tamagawa, K., Shimizu, K. et al. (2003) Effects on M5076-hepatic metastasis of retinoic acid and

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36. 37.

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N -(4-hydroxyphenyl)retinamide, fenretinide entrapped in SG-liposomes. Biol Pharm Bull , 26, 1060–63. Behlke, M.A. (2006) Progress towards in vivo use of siRNAs. Mol Ther, 13, 644–70. Song, E., Lee, S.K., Wang, J. et al. (2003) RNA interference targeting Fas protects mice from fulminant hepatitis. Nat Med , 9, 347–51. Yang, L., Jung, Y., Omenetti, A. et al. (2008) Fate-mapping evidence that hepatic stellate cells are epithelial progenitors in adult mouse livers. Stem Cells, 26, 2104–113. Bachem, M.G., Schneider, E., Gross, H. et al. (1998) Identification, culture, and characterization of pancreatic

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stellate cells in rats and humans. Gastroenterology, 115, 421–32. 40. Zhang, K., Rekhter, M.D., Gordon, D. et al. (1994) Myofibroblasts and their role in lung collagen gene expression during pulmonary fibrosis. A combined immunohistochemical and in situ hybridization study. Am J Pathol , 145, 114–25. 41. Razzaque, M.S., Ahsan, N. and Taguchi, T. (2000) Heat shock protein 47 in renal scarring. Nephron, 86, 339–41. 42. Nagy, N., Holven, K., Roos, N. et al. (1997) Storage of vitamin A in extrahepatic stellate cells in normal rats. J Lipid Res, 38, 645–58.

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The “Green Liver” and Transcriptional Regulation of Phase II Detoxification Genes Christopher Johnson and Jonathan Arias Department of Biological Sciences, University of Maryland, Baltimore County, Baltimore, MD, USA

INTRODUCTION Plants and mammal cells confront similar needs for transporting chemical substrates across the plasma membrane for subsequent detoxification and elimination. In vertebrates, hepatocytes are primarily responsible for mediating these physiological functions, whereas in plants, cells of the apical and root meristems evidence a similar role. Some importers, exporters, transporters, channels, and pumps of plants and mammals have similar functions and structures, although others appear to be unique or differ in their regulation. This chapter focuses on a major class of detoxification enzymes, the glutathione transferases, which illustrate the shared and specific features of plants and mammals in detoxification reactions against potentially damaging xenobiotic (foreign) chemicals. Chemical detoxification in eukaryotes typically involves the sequential activities of enzymes that first transform (phase I) and then conjugate (phase II) most chemicals to more water-soluble derivatives. In the liver, a major site of detoxification in vertebrates, these conjugates are mobilized by phase III enzymes from the intracellular to extracellular environment for excretion through the bile duct and digestive tract. By contrast, phase III enzymes in plants mobilize xenobiotic conjugates to The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

latent forms in cell vacuoles or the cell wall for long-term storage. Given that plants represent an important source of human nutrition, the uptake and concentration in latent form of toxic environmental xenobiotics (i.e. pollutants) by edible plants may adversely impact public health. In 1977, Sandermann [1] coined the term “green liver” to emphasize the primary role of plants as catalysts for xenobiotic detoxification in the environment, analogous to the part played by the liver among the organs of the body. While plants and animals have many conserved structural and functional properties in xenobiotic detoxification, the transcriptional regulatory mechanisms which govern the expression of detoxification genes vary considerably between plants and animals. One such example is the activation of expression of phase II genes, including those encoding for members of a large and conserved superfamily of glutathione S -transferase (GST) isoenzymes, by xenobiotic-induced oxidative stress signaling through a promoter DNA sequence termed the antioxidant response element (ARE). This cis-regulatory element serves as the binding site for protein complexes which modulate transcription in response to oxidative stress signal pathways. While general features of this detoxification response are conserved between plants and animals, downstream

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

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transcriptional processes which govern these gene responses are surprisingly divergent.

GLUTATHIONE S-TRANSFERASES IN THE PHASE II DETOXIFICATION RESPONSE The coordinate expression of phase II genes contributes to protection against cellular damage caused by a wide array of xenobiotic chemicals, abiotic agents, and environmental conditions that generate free radical species [2–4]. Key among these protective genes are those that encode for proteins of a conserved super-family of GST isoenzymes, which detoxify xenobiotic lipophilic substrates containing electrophile centers by catalytic conjugation with the tripeptide glutathione [5]. Although biological roles for most GSTs are commonly unknown or not yet fully assigned, the general ability of these conserved enzymes to protect eukaryotic organisms from cellular damage caused by xenobiotic compounds is well established [5, 6]. As generally noted for other phase II enzymes, GSTs are important components of antioxidant defense against endogenous reactive metabolites formed during oxidative stress [5, 7–10]. As such, GST-mediated catalysis results in hydrophilic conjugates that are mobilized by adenosine triphosphate-binding cassette (ABC) transporters during the phase III response to the extracellular environment for excretion or, in the case of plants, to the vacuole or cell wall compartments for long-term storage [11, 12]. Moreover, GST peroxidase reactions can scavenge toxic organic hydroperoxides and protect cells against oxidative damage [8]. It is noteworthy that GSTs evolved long before organisms were exposed to xenobiotic chemicals of anthropogenic origin (e.g. herbicides and pollutants), and thus are presumed to play an ad hoc role in the cellular detoxification of these compounds. Mammalian GSTs represent seven cytosolic families (alpha, mu, pi, sigma, theta, omega, and zeta), the mitochondrial family kappa, and four microsomal families involved in the biosynthesis of eicosanoids [13, 14]. Various GST families can form heterodimers, thus extending the potential functional diversity of this group of enzymes. As in other metazoans, the expression of individual GST isoforms in humans can be restricted to specific organs. For example, in liver, kidneys, and ovaries, GST class alpha dominates, whereas GST class pi is the major form in lungs, muscles, brain, pancreas, erythrocytes, and skin [15]. Plants also encode for a large and robust superfamily of GST enzymes, including members of several classes not found in animals. For example, the genome of the higher plant Arabidopsis thaliana encodes at least 47 members of the GST family which can be divided into four distinct classes, two of which are present in mammals (theta and zeta) and two that are unique to plants (tau

and phi) [14, 16, 17]. Of these plant GST classes, theta and tau members are preferentially expressed in response to xenobiotic safeners, which are agricultural chemicals that enhance the tolerance of monocotyledonous plants to herbicides by inducing the expression of detoxifying enzymes such as GSTs [18]. In addition, several members of the phi class of GSTs are induced by diverse types of exogenous chemical cues including hydrogen peroxide, ozone, and herbicides, thus suggesting their protective role in oxidative stress [7]. As with humans and other vertebrates, plants show highly restricted patterns of expression of specific GST genes, in line with the notion that these enzymes have highly specialized biochemical roles in specific tissues or organs where they occur [18].

THE ARE CIS-ELEMENT AND GST EXPRESSION The ability of certain xenobiotics to activate rapidly and transiently the expression of GST genes has prompted efforts to identify common or conserved regulatory DNA sequence(s) and cognate trans-acting factor(s) that mediate this stimulus-inducible response. Such efforts have led to discovery of the ARE, first identified in the promoter of a rat GST gene, GSTA2 [19]. It subsequently became evident that the ARE is widely distributed in the promoters of other GST genes (Figure 70.1). Sequence comparisons of these AREs suggest a basic core consensus sequence, 5′ -TGAC/T A/T A/C AGC-3′ , which is essential for mediating basal and/or inducible transcriptional activities of the corresponding GST promoters. This sequence motif occurs either singly, as in the case of mouse Gsta1 , or as a tandem repeat separated by six nucleotides in the promoters of several human and rat GST genes [2, 3, 19, 20]. Deeper phylogenetic comparisons of 57 functional ARE sequences revealed that some nucleotides were partially conserved across all AREs and all three species (human, mouse, and rat), whereas others showed near 100% conservation; extended sequences outside these positions were thought to contain species-specific information [20].

PLANT ARE’S ARE-like elements have also been identified in the promoter regulatory regions of diverse plant genes [24–27]. These elements, variously termed as-1 , ocs, or nos, were initially discovered in the promoters of genes of certain viruses and agrobacteria that invade plants, where they presumably play a role in host colonization and pathogenesis. Plant as-1 -type elements have a core TGACG sequence, that is conserved with animal AREs (Figure 70.1), although flanking sequences of these motifs differ considerably. In the case of as-1 elements, this core sequence occurs as tandem copies with a restricted spatial

70: THE “GREEN LIVER” AND TRANSCRIPTIONAL REGULATION OF PHASE II DETOXIFICATION GENES

Animal promoters H.s. GSTP1 M.m. GST-Ya M.m. Gsta1 R.n. GSTA2 R.n. GSTP

GCGCCGTGACTCAGCACTG TAATGGTGACAAAGCAACT TAATGGTGACAAAGCAACT TAATGGTGACAAAGCAACT TCACTATGATTCAGCAACA 2

ARE consensus:

1 0

Plant promoters N.t. GNT1

ATAGCTAAGTGC---TTACATCT

N.t. GNT35

TTAGCTAAGTGC---TTACATCT

A.t. GST6

TTAGCTCATTGA---TGACGACC

A.t. GSTF8

TTATGTCATTATTGATGACGACC

T.a. GST-A1

ATCCGTACCAAC---GCACGTGT

S.c. GST

CAACGTCAGAGT---ATACGTAT 2

as-1-type consensus:

1 0

Plant pathogen promoters CaMV 35S as-1

TGACGTAAGGGA---TGACG-CA

A.tum. nos

TGAGCTAAGCAC---ATACGTCA

A.tum. ocs

TGACGTAAGCGC---TTACGTAC

A.tum. mas

TGACGTAAGTAT---CCTAGTCA 2

as-1 consensus:

1 0

Figure 70.1 Sequence alignments of functional ARE cis-elements of animals and as-1 -type cis-elements in promoters of plant nuclear (GST) and pathogen genes. Abbreviations: H.s., Homo sapiens (human); M.m., Mus musculus (mouse); R.n., Rattus norvegicus (rat); N.t., Nicotianum tabacum (tobacco); A.t., Arabidopsis thaliana; T.a., Triticum aestivum (wheat); S.c., Silene cucubalus; A.tum., Agrobacterium tumefaciens; and CaMV 35S, cauliflower mosaic virus 35S promoter. Sequences used here were obtained from the following sources: [19–21]. All sequences shown are from 5′ to 3′ orientation, with bold type indicating core ARE and as-1 -type (tandem repeats) sequences. Consensus logo sequences were generated using WebLogo v. 2.8.2 software [22, 23]

separation, that is, required for the complete element to function [28]. As-1 -type elements were later identified in the promoters of plant GST genes (Figure 70.1), whose expression is activated by xenobiotic stress agents (herbicides, safeners, and plant hormone analogs) which result in cellular production of reactive oxygen species [7, 16, 18, 29]. The evident sequence and functional similarities between as-1 -type and ARE elements of plants and animals, respectively, supports the notion that these cis-regulatory elements and their cognate transcription factors have diverged from an ancient protective gene regulatory system to oxidative stress.

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TRANSCRIPTION FACTORS AND ARES As noted above, ARE-type cis-elements are primary determinants of xenobiotic and oxidative stress-induced activation of GST genes. Moreover, DNA-binding proteins which preferentially bind to these elements play a key role in mediating this transcriptional stimulus-response. The prototypical ARE-binding factor is nuclear factor-erythroid 2 p45-related factor 2(Nrf2), which has a CNC (cap’n’collar) motif with basic region/leucine-zipper (bZIP) DNA-binding and dimerization domains [30, 31]. Nrf2 binds the ARE in vivo as a heterodimer with a bZIP factor of the small Maf-protein family, which includes MafF, MafG, and MafK [30]. According to current dogma (Figure 70.2a), oxidative stress induced by xenobiotic compounds causes Nrf2 to translocate from the cytoplasm to nucleus, where it encounters and forms a dimer with a small Maf protein. This complex binds the ARE of corresponding GST and other phase II genes to recruit subsequently the transcriptional co-activators creb binding protein (CBP)/p300 to the promoter region [3]. Unequivocal evidence that Nrf2 and Maf play these roles comes from knockout experiments in mice, where elimination of these transcription factors has been shown to inhibit xenobiotic induction of phase II detoxification and GST expression [32]. Nrf2 also regulates genes involved in the recognition and repair/removal of damaged proteins in the liver [33], thus involving this factor in the role of protecting and maintaining hepatocyte function.

REDOX REGULATION OF KEAP1 AND NRF2 In response to certain xenobiotics, the redox status of the cell is typically altered to produce a pro-oxidant state resulting in oxidative stress. This state is crucial for inducing the activity of signal transduction pathways and transcription factors that regulate ARE-dependent transcription. It was therefore hypothesized that activation of the ARE involves an intracellular redox sensor that possesses reactive cysteine residues that are sensitive to modification. One candidate for this redox sensor comes in the form of the Keap1 protein [36, 37]. Keap1 functions as a cytoplasmic repressor of Nrf2, by binding this trans-activator through a hydrophilic region in the Neh2 domain of Nrf2. The Keap1 protein contains a number of potential reactive cysteine residues which are modified in response to oxidative stress. This redox change results in the release of Nrf2 and translocation of this factor to the nucleus, where it activates transcription through the ARE (Figure 70.2a). In this regard, electrophilic compounds that are thiol reactive promote dissociation of Nrf2

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THE LIVER: UBIQUITINATION AND THE RATE OF KEAP1 AND NRF2 TURNOVER

Animal pathway Oxidative Stress (ROS, Electrophiles)

Plant pathway Oxidative Stress (ROS, Electrophiles)

ULC Cul3 Rbx1 Keap1

Ubiquitination and proteolysis

Nrf2

Cytoplasm Nucleus

Keap1

p120

Cytoplasm Nucleus

TGA

p120

Maf CBP/P300 Maf Nrf2 TGACnnnGC

GST

ARE (a)

TTACGTA [N4-7] TTACGTC

GST

As-1 (b)

Figure 70.2 Summary of major xenobiotic detoxification pathways in animals and plants. (a) In animals, the steady-state levels of Nrf2 in the cytoplasm are controlled by Keap1-dependent ubiquitination of Nrf2. Indeed, the cytoskeletal protein Keap1 functions as an adaptor for Cul3, which, in combination with Rbx1, assembles into a functional E3 ubiquitin ligase complex (ULC) that targets the N-terminal Neh2 domain of Nrf2 for ubiquitin conjugation and eventual protein degradation. This quenching effect is mitigated following cellular exposure to oxidative stress signals in the form of ROS and electrophiles, whereby Nrf2 is released from Keap1 and thus escapes proteasomal degradation. Following accumulation and subsequent nuclear import of Nrf2, chromatin remodeling by CBP/p300 facilitates the recruitment of the Nrf2-Maf complex to cognate ARE cis-elements, resulting in transcription of cytoprotective genes, including GSTs. (b) In plants, the TGA transcription factors are localized in the nucleus, existing most likely as inactive dimers bound to a repressor protein, termed p120. Following exposure to oxidative stress, p120 is likely targeted for modification and dissociates from TGA factors. Stable binding of DNA at cognate as-1 -type cis-elements by active dimers of TGA factors then results in transcription of cytoprotective genes, including GSTs [26, 34, 35]

from Keap1. In the absence of Keap1, mice develop normally but die shortly after birth due to hyperkeratotic constrictions of the esophagus and fore-stomach, and show constitutive activation of Nrf2-dependent genes [38]. This postnatal lethality is reversed in mice that lack both Nrf2 and Keap1, thus indicating that constitutive up-regulation of Nrf2 target genes under such conditions is responsible for the aforementioned pathology. Keap1 contains several regulatory regions, including the N-terminal Bric-a-brac, Tramtrack, Broad-complex (BTB) domain, a conserved linker region and the C-terminal Kelch domain. The last domain binds to the Neh2 region of Nrf2 and, together with sequences located in the linker domain, enables Keap1 to sequester Nrf2 in the cytoplasm [36]. The Kelch domain of Keap1 is also able to bind actin, and an intact actin-based cytoskeleton is required for cytoplasmic sequestration of Nrf2 by Keap1. In addition, as described below, Keap1 associates with Cul3 and Rbx1 to form a functional E3 ubiquitin ligase complex that targets Nrf2 for ubiquitination both in vivo and in vitro. Because it functions as a substrate adaptor for a Cul3-dependent E3 ubiquitin ligase complex, Keap1 serves to regulate turnover of itself and

Nrf2 through a mechanism involving ubiquitin-mediated proteolysis ([37]; Figure 70.2a).

UBIQUITINATION AND THE RATE OF KEAP1 AND NRF2 TURNOVER The ability of Keap1 to function as a substrate adaptor protein for Cul3 and thereby target Nrf2 for ubiquitin conjugation provides an efficient mechanism for repression of Nrf2 steady-state levels and of Nrf2-dependent transcription. In such cases, exposure of cells to a wide variety of chemicals that induce oxidative stress results in elevated steady-state levels of Nrf2 and enhanced expression of ARE-regulated genes [2, 4]. When the ability of Keap1 to target Nrf2 proteins efficiently for degradation is inhibited by redox changes of key cysteine residues in the BTB domain, newly synthesized Nrf2 proteins will no longer be bound by Keap1 and instead accumulate in the nucleus. Mutant Keap1 proteins containing serine substitutions in these residues are impaired in their

70: THE “GREEN LIVER” AND TRANSCRIPTIONAL REGULATION OF PHASE II DETOXIFICATION GENES

ability to target Nrf2 for ubiquitination and to repress Nrf2-dependent gene expression in transfected cells [37]. Therefore, multiple cysteine residues in Keap1 are capable of undergoing redox-dependent alterations that affect its ability to affect the ubiquitination and subsequent proteolysis of Nrf2.

PLANT TGA FACTORS AND THEIR MEDIATOR PROTEINS TGA factors belong to a multi-gene family of transcriptional regulators which play key roles in protective responses of plants [39–42]. Like other bZIP-type transcription factors, TGA factors are modular and share a nearly identical basic domain that confers DNA-binding specificity for as-1 -type cis-elements, which are structurally and functionally homologous to AREs of animals. Given this relationship and their proposed ancient common origin, it is not surprising that the basic regions of the DNA-binding domains of Nrf2 and TGA factors share only a modest overall sequence similarity (Figure 70.3). While the majority of bZIP transcription factors form homo- or heterodimers [43, 44] through the action of complementary leucine-zipper motifs, TGA factors also require a conserved carboxy-terminal (CT) domain for stable dimerization to occur [39, 45]. The CT domain further serves as a recognition site for mediator proteins, such as p120 [39] and non-expresser of PR genes (NPR1) [42, 46–49], whose stimulus-reversible associations affect in vivo recruitment of specific TGA transcription factors to target cis-elements [41, 50, 51]. Although both these types of mediator proteins affect the ability of TGA factors to bind DNA, they apparently do so by mechanisms different from those previously described for Keap1. For example, basic region

while the subcellular distribution of Nrf2 and MafK is modulated by xenobiotic cues, TGA factors are constitutively localized to the nuclear compartment [40, 41]. However, as in the case of the animal mediator protein Keap1 and its regulatory bZIP target Nrf2, changes in the cells redox state affect the interaction between NPR1 and TGA factors. NPR1, also known as NIM1 and SAI1 [52–55], is a positive mediator of the host defense mechanism known as systemic acquired resistance (SAR). Under basal conditions, NPR1 is present as an oligomer in the cytoplasm [56]. As the cell returns to a more reduced state following an initial oxidative burst due to chemical stimulus, NPR1 is reduced to its monomer form, accumulates in the nucleus, and binds TGA factors. Two conserved cysteines that are located in the CT domain of TGA1 (C260 and C266) control the interaction with NPR1, and mutation of these residues permits constitutive interaction with NPR1 in vivo [50]. Moreover, the redox status of cysteines in TGA1 and/or the closely related TGA4 became predominantly reduced upon stimulus induction. Thus, strong interaction of TGA1 and/or TGA4 with NPR1 is correlated with the reduced state of these cysteines. Redox regulation of NPR1 affects transcription through an as yet unidentified mechanism, which may involve changes in the dimerization stability of target TGA factors (C. Johnson, unpublished data). In contrast to the situation described above, transcriptional regulation of GST genes appears to involve the activity of TGA factors with other mediator proteins, as npr1 mutant plants show normal GST gene induction in response to agents that induce xenobiotic stress [26, 57]. One such mediator may be a nuclear protein termed p120 (Figure 70.2b), which reversibly interacts with a tobacco TGA factor (TGA1a) that binds to the promoter regulatory regions of GST genes (e.g. GNT1 and GNT35 ) in response to xenobiotic stress cues [29, 40]. leucine-zipper region

Human Nrf2 Mouse Nrf2 Rat Nrf2

RDIRRR-GKNKVAAQNCRKRK------LE----NIVELEQDLDHLKDEKEKLLKEKGENDKSLHLLKKQL RDIRRR-GKNKVAAQNCRKRK------LE----NIVELEQDLGHLKDEREKLLREKGENDRNLHLLKRKL RDIRRR-GKNKVAAQNCRKRK------LE----NIVELEQDLGHLKDEREKLLREKGENDRNLHLLKRRL

Tobacco TGA1a Arabidopsis TGA1 Arabidopsis TGA4 Bean TGA1.1

--VLRRLAQNREAARKSRLRKKAYVQQLENSKLKLIQLEQEL --IQRRLAQNREAARKSRLRKKAYVQQLETSRLKLIQLEQEL --IQRRLAQNREAARKSRLRKKAYVQQLETSRLKLIHLEQEL --IQRRLAQNREAARKSRLRKKAYVQQLESSRLKLMQLEQEL

Rat Nrf2: Plant TGA consensus:

1135

RDIRRR-GKNKVAAQNCRKRK------LE----NIVELEQDLDHLKDEKE KLLKEKGENDKSLHLLKKQL IQRRLAQNREAARKSRLRKKAYVQQLETSRLKLIQLEQEL

Figure 70.3 Alignment of the basic and leucine-zipper regions of animal and plant bZIP transcription factors thought to be involved in xenobiotic detoxification. Alignments of the Nrf2 and plant TGA transcription factors were done using EMBOSS pairwise alignment of the plant consensus sequence against rat Nrf2 sequence. There is very high conservation of the basic/leucine-zipper regions within kingdoms: ∼90% identical and 98% similar for Nrf2 orthologs, and ∼90% identical and 95% similar for plant TGA transcription factors. However, the apparent conservation of bZIP regions across kingdoms is relatively low (i.e. 21% identity and 33% similarity

1136

THE LIVER: REFERENCES

Specifically, p120 binds the CT domain of TGA1a, which is necessary and sufficient for the interaction. Previous studies have identified a “dimer stabilization domain” (amino acids 178–373) within this CT domain, which enhances the ability of TGA1a to dimerize and bind as-1 [41, 45]. Interestingly, p120 appears to be directly modified in response to xenobiotic stress, consistent with the notion that it serves as a redox sensor and therefore as a stimulus-responsive regulator of TGA1a DNA-binding activity. The molecular mechanism by which p120 mediate this activity, however, is currently unknown.

CONCLUSION Plants and animals avoid cytotoxic effects of xenobiotic stress by activating the transcription of genes for phase II detoxification enzymes. These genes are evolutionarily conserved, consistent with the notion that they arose from a distant common eukaryotic ancestor. However, despite these similarities, protective responses across this phylogeny incorporate different regulatory strategies for governing the expression of phase II genes. In particular, plant and animal cis-elements and transcription factors which govern these genes evidence little similarity in their respective structures or mechanisms. In addition, mediator proteins (e.g. Keap1, NPR1, and p120) that modulate the DNA-binding and trans-activation potentials of these transcription factors have subcellular locations and functional roles that are also quite distinct. Hence although the “green liver” concept serves as an ecological analogy of xenobiotic detoxification between plants and the liver, it lacks broader application to phylogenetically diverse mechanisms that regulate this evolutionarily conserved adaptive response.

ACKNOWLEDGMENTS Research cited here from the authors’ laboratory was supported by grants from the National Science Foundation (USA). The authors contributed to this article in their personal capacity. The views expressed are their own and do not necessarily represent those of the National Institutes of Health or the United States Government

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Index Note: All entries pertain to the liver unless otherwise specified. index entries under ‘liver’ have been kept to a minimum and readers are advised to seek more specific topics. Page numbers in italics refer to figures; page numbers suffixed by ‘t’ refer to tables. This index is in letter-by-letter order, whereby spaces and hyphens between words are excluded from alphabetization (e.g. ‘B cells’ comes before Bcl-2 family). Abbreviations used in index subentries AFLD—alcoholic fatty liver disease AP—apical membrane BL—basolateral CLD—chronic liver disease ECM—extracellular matrix ER—endoplasmic reticulum GST—glutathione S-transferase HBV—hepatitis B virus HCC—hepatocellular carcinoma HCV—hepatitis C virus HGF—hepatocyte growth factor ITR—inverted terminal repeat NAFLD—non-alcoholic fatty liver disease NPC—nuclear pore complex PEPCK—phosphoenolpyruvate carboxykinase PH—partial hepatectomy ROS—reactive oxygen species RT—reverse transcription/transcriptase TGN—trans-Golgi Network UPR—unfolded protein response

A A1/Bfl, 786 AAA type proteins, 33, 45, 48 2-AAF/PH model, 581, 583 Aagenaes syndrome (lymphedema– cholestasis syndrome), 668t, 671 AAV-2 see adeno-associated virus (AAV-2), recombinant Abbe’s equation, 1055, 1058 ABCB1 (MDR1; Pgp), 340, 349, 683t, 691, 1075, 1078 acute myeloid leukemia, 1081–1082 apical recycling from intracellular pool, 349, 352, 352 , 354 bile transport protein defect and, 682 breast cancer, 1082 cholesterol efflux, 278 CYP enzyme synergy, 1078

direct trafficking to canaliculi, 350, 350 drug resistance in HCC, 1080, 1081 hepatic efflux of bile acids, cholestasis, 690, 691 non-muscle myosin 2 role, 36 overexpression, drug resistance in HCC, 1080 single nucleotide polymorphisms, 1082–1083 tubulovesicular movement, 351 ABCB1 gene, 340 ABCB4 (MDR3), 254, 342 , 349, 683t, 691, 691 activation by bile salts in bile, 280 apical recycling from intracellular pool, 352, 352 , 354 defects/mutations, 341, 666 hepatic efflux, 280, 341, 1078 of bile acids in cholestasis, 690, 691 phospholipid (biliary) excretion, 341, 342 progressive familial intrahepatic cholestasis, 254 trafficking to canaliculi, 350, 351, 352 Abcb4 (mice), 341 ABCB4 disease (PFIC-3), 341, 664t, 666 ABCB5, 280 ABCB8, 280 ABCB11 (bile salt export pump (BSEP)), 325, 339, 349, 355, 683t, 692–693 apical recycling, 36 cAMP and taurocholate affecting, 351, 352 clathrin-mediated endocytosis, 354–355 constitutive cycling, 352 , 352–353 HAX-1 role, 354 myosin Vb role, 354 rab11a role, 352, 352 , 353, 355 ATP stimulation, 296 encoded by ABCB11 see ABCB11 gene gene, bile acid synthesis regulation, 297

The Liver: Biology and Pathobiology (Fifth Edition)  2009 John Wiley & Sons, Ltd

hepatic efflux of bile acids, 280, 296, 339, 341, 1078 in cholestasis, 690, 692–693 defective, ABCB11 defects, 298 intracellular pool, 351, 352, 352 progressive familial intrahepatic cholestasis, 254 regulation by Fxr/FXR, 325, 692 trafficking, post-Golgi compartments, 350 , 350–351 ABCB11 disease (PFIC-2), 663, 664t, 665 ABCB11 gene, 339, 663, 665 bile acid synthesis regulation, 297 mutations, 296, 663, 665 carriers, 665 ABCC1 (MRP1), 1078 breast cancer, 1082 drug resistance in HCC, 1081 ABCC2 (MRP2), 340, 349, 352, 665, 691–692, 1078 bilirubin glucuronide transport, 253, 253 , 254 deficiency (Dubin-Johnson syndrome), 254, 664t, 665–666, 691 hepatic efflux of bile acids, cholestasis, 690, 691–692 intracellular pool and recycling, 352, 354 ABCC2 disease (Dubin–Johnson syndrome), 254, 664t, 665–666, 691 ABCC2 gene, 339, 665 ABCC3 (MRP3), 682, 684, 690, 1078 ABCC4 (MRP4), 683t, 1078 ABCC5 (MRP5), 1078 ABCC6 (MRP6), 684t, 1078 ABCG1, 278, 1081 ABCG2 (BCRP), 340, 349, 684t, 693, 1078 breast cancer, 1082 hepatic efflux of bile acids in cholestasis, 690, 693 ABCG2 gene, 340

Arias, Alter, Boyer, Cohen, Fausto, Shafritz and Wolkoff, Eds

1139

1140 ABCG5, 342–343, 690, 1078 as “liftase”, 342–343 Abcg5 (mice), 340, 342–343 mutants/mutations, 340–341 ABCG8, 273, 342–343, 690, 1078 as “liftase”, 342–343 Abcg8 (mice), 340, 342–343 mutants/mutations, 340–341 ABC transporters, 131, 354, 662, 1078 apical recycling between pools and canaliculi, 349–350, 351–352, 352 cAMP and taurocholate affecting traffic, 351, 352 canalicular biogenesis link, 355, 355 clathrin-dependent endocytosis, 354–355 constitutive cycling, 352 , 352–353 HAX-1 role, 354 mechanisms, 353–355 myosin Vb role, 353–354 PI3 kinase role, 353 rab11a role, 352, 352 , 353, 355 canalicular (canalicular membrane), 339, 340, 349–358 trafficking/targeting to, 350 , 350–351 cytochrome P450 enzymes synergy, 1078 expression profiles in HCC, 1081–1082, 1083 family and sub-families, 1081 gene distribution, 1081 genetic variation/polymorphisms, 1082–1083 GST mobilization and, 1132 intrahepatic pools, 351–355, 352 MRP transporter C subfamily, 1078 multidrug resistance, 1078, 1081–1082 mutations, 1082–1083 newly synthesized, trafficking, 350 , 350–351 proteins associated with targeting of, 354 ABT-737, 796 acarbose, 610 acetaldehyde, 744, 747–748 actions, 747–748 alcoholic liver disease, 747 , 747–748 COLIA2 gene and, 743, 744 , 745, 747 signal transduction pathways, 747–748 α-acetaminofluorene (2-AAF), 581, 583 acetaminophen calcein quenching, 513, 514 non-toxic dose, lesions, 775 overdose, 601, 601 toxicity, pathogenesis, 379–380, 774 acetoacyl-CoA, 258 acetyl-aminofluorene-partial hepatectomy (AAF/PH) model, 460 acetylcholine (AC), calcium signals induced by, effects, 360–361 acetyl-CoA, 193, 257 acetyl-CoA:α-glucosaminide N-acetyltransferase, 177 acetyl-CoA carboxylase (ACC), 257 ACC1, 263, 538 CB1 receptor activation, 1096 , 1096–1097 decreased transcription by AMPK, 541 decreased levels, 260 isoforms (ACC1 and ACC2), expression, 263, 538, 539 knockout mice, 263–264 regulation by AMPK, 538, 539 , 541

INDEX therapeutic target in NAFLD, 732 N -acetylcysteine, 611 acidification, gap junction closure, 211 acid sphingomyelinase (A-SMAase), 790, 794 acini (acinus), 4, 8 actin agents disrupting, 396 bile canaliculi, 354 in clathrin-mediated endocytosis, 110–111 depolymerization, membrane fusion, 396 filamentous (F-actin), 30 depolymerization, sinusoidal obstruction syndrome, 379 polymerization, and energy for, 30, 31 structure and asymmetry (polar ends), 31, 46 function, 34 globular (G-actin), 30, 31 myosin binding, 47 non-muscle, 31, 34 see also actin filaments “actin barrier”, 396 actin-binding proteins, 31 actin filaments, 45 disruption, barrier function of tight junctions reduced, 212–213, 215 fenestrae regulation, 395–396, 396 gap junction regulation, 212 historical aspects, 29–30 localization in hepatocytes, 32 networks, 32 organization in epithelial cells (hepatocytes), 32, 46 parallel bundles, 32 role in tight junctions, 206, 212–213, 213 , 215 tight junction protein trafficking, 215 see also actin activated partial thromboplastin time (APTT), 646, 647, 980 activating transcription factor-1 see ATF-1 (activating transcription factor-1) activating transcription factor 6 (ATF6), 167, 265, 728, 788 activator of CREM in testis (ACT), 525 activin A, 559 acute injury of liver see liver injury acute kidney injury (AKI), 619 acute liver failure (ALF) brain dysfunction, 597, 598 brain edema, intracranial hypertension, 603–604 brain glucose metabolism changes, 604–605 cerebral blood flow changes, 608, 610 fibrinolysis, and abnormalities, 650 hepatic encephalopathy, 597–600, 600t, 601 treatment, 610, 611 infection and inflammation, 609 acute phase reactant, fibrinogen as, 647 acute phase response, ECM in liver inflammation, 455, 456 acute renal failure, 619 acyl-CoA, esterification, 258 acyl-CoA: cholesterol acyltransferase (ACAT), 273, 277 acyl-CoA:1-acylglycerol-sn-3-phosphate acyltransferase (AGPAT), 258 acyl-CoA:glycerol-sn-3-phosphate acyltransferase (GPAT), 258

acyl-CoA synthetases, 258 acylthiourea compounds, NS4A-NS3 inhibitors (HCV therapy), 912 ADAM(s), 559 ADAM 17 (TACE, TNFα converting enzyme), 559 ADAMTS2, 461–462 ADAMTS13, 642, 643, 644 changes in alcoholic hepatitis, 646 deficiency, 642 structure and function, 643, 644 adaptins (APs), 109 clathrin-coated pits, 108–109 adaptor protein-1A (AP-1A) see AP-1A adefovir dipivoxil (HepSera), 866, 901, 902t, 903, 903t combination therapy in hepatitis B, 904 post-transplant in hepatitis B, 927 adenine nucleotide translocator (ANT), 513, 785 adeno-associated virus (AAV-2), recombinant, 968t, 970–971, 971 applications, 975 generation, 970, 971 genome, and ITRs, 970, 971, 971 persistence, 971 Rep protein, 970–971 serotypes and receptors, 970 adenomatous polyposis coli protein see APC (adenomatous polyposis coli) gene/protein adenosine, and A(2A) receptor, 412 adenosine diphosphate (ADP), biliary, 362–363 adenosine monophosphate-activated protein kinase see AMPK (AMP-activated protein kinase) adenosine triphosphate see ATP S-adenosyl L-methionine, 756, 759 S-adenosyl L-methionine depletion (SAMe), 759, 790 adenoviruses gene therapy vector, 966, 968t, 972–973, 986 telomerase activation mediated by, 1114 adenylyl cyclase (AC; adenylate cyclase), 362, 521–522 AC1-AC9 types, 362, 521–522 cholangiocytes expressing, 362 adherens junctions (AJ) hepatocytes, polarity development, 83, 84, 85t–86t, 91 proteins associated, 84, 85t–86t, 91 see also tight junctions adhesion kinase, 796 adhesion molecules, 748, 1100 adipocytes activation, 723–724 size/number increase, weight gain, 723 see also adipose tissue adipocytokines increased in NAFLD, 626–627 proinflammatory, prothrombotic, profibrogenic, 626 , 626–627 released by adipose cells, 624t see also adipokines; specific adipocytokines (see pg. 624t) adipogenesis, inhibitor, 263 adipokines, 262, 746–747 alcoholic liver disease, 746–747 AMPK activation, 537 hepatic fibrosis, 439

INDEX see also adipocytokines; adiponectin; leptin; resistin adiponectin, 263, 624–626 actions/effects, 624t, 729–730, 746 on podocytes, 625, 625 adipose tissue secreting, 624t, 725–726, 745, 746 alcoholic liver disease, 746–747 AMPK activation, 537, 625, 625 , 725 decreased production in NAFLD, 624–626, 723–724, 729–730 deficiency, 625, 625 fatty acid oxidation, 725–726 hepatic fibrosis pathogenesis, 439 hepatoprotective/antifibrogenic actions, 746–747 insulin sensitization, 625 in non-alcoholic steatohepatitis, 729–730 receptors and interactions with, 625 , 625–626 signaling and PPAR-α, 626 stellate cells expressing, 744 stellate cells transdifferentiation, 412, 746–747 up-regulation by pioglitazone, 745 adiponectin receptor 1 (AdipoR1), 625, 745, 746 adiponectin receptor 2 (AdipoR2), 625–626, 745 adipose tissue, 719, 745 adipocytokines released, 624t, 745 adiponectin secretion, 624t, 725–726, 745, 746 brown, 296 factors secreted, 745 macrophage infiltration, inflammatory response, 722 , 723, 746 metabolic functions, 261–263, 262 , 744 response to energy excess, 722–724, 723 triglyceride storage, 261–263, 262 adipose tissue triglyceride lipase/desnutrin (ATGL; PNPLA2, iPLA2), 258 adipsin, released by adipose cells, and effects, 624t ADP, biliary, 362–363 ADP ribosylation factor (ARF), 126 α-adrenergic agonists, hepatic vascular resistance increase, 708 ADRP, 258 deficiency, 258–259 adverse drug reactions (ADRs) dose-related hepatotoxicity, 775–776 idiosyncratic, 775, 776 animal models, 776–777 inflammatory stress mechanisms, 777–778, 778 AE2 (chloride/bicarbonate exchanger), 339, 366, 693 AF-20, monoclonal antibody to, 966 afadin, adherens junction association, polarity development, 84, 86t, 91 afferent blood vessels, 3, 5–6, 7 hemodynamics, 6, 8 aflatoxin B1, toxicity, 775 age chronic hepatitis C outcome, 885 muscle mitochondrial impairment, lipid accumulation, 473 aging free-radical theory, 143 lysosomal system changes, 184–185 mitochondria and, 143 protein degradation, 183 , 183–184

pseudocapillarization (fenestrae reduction), 398 telomeres and, 1105, 1108 , 1108–1109 in yeasts, Sir2 and see sirtuin 2 (Sir2); yeast agrin, 459 AICAR (5-aminoimidazole 4-carboxamide-1β-4-ribofuranoside), 538, 545 AICAR response-element binding protein (AREBP), 541 airy disk, two-dimensional, 1055, 1056, 1056 A-kinase anchoring proteins (AKAPs), cholangiocyte cilia signaling, 362 Akt and Akt signaling, 215 Akt2, therapeutic target in NAFLD, 731 hepatic fibrosis, 440 insulin resistance, 544, 726 , 727, 731 phosphorylation, 260 see also PI3K/AKT pathway ALADIN gene, 152 Alagille syndrome (ALGS), 20, 254–255, 662, 666–668, 667t JAGGED1 and NOTCH2 mutations, 20, 1070 mouse and zebrafish models, 1070 types 1 and, 2 666, 667t alanine aminotransferase (ALT) HAV infection, increase in, 809 HBV infection, 836, 899 HCV infection, 820, 886, 924 chronic, 883, 886 hepatocellular carcinoma, 871 alb1 transcriptional enhancer, 18, 18 albumin bile acids binding, 296, 305 bilirubin binding, 253 dialysis, 611 hepatorenal syndrome therapy, 621 secretion by rat hepatocytes in culture, 938 synthesis rate in hepatocytes, 75 tagged, liver space available to, 7 albumin (Alb)–IFN-α2b, 909 albumin–organic anion complexes, 305 albuminuria, regulation by adiponectin via AMPK, 625, 625 alcohol, 747 chronic hepatitis C outcome, 887 effect on fenestrae of sinusoidal endothelial cells, 398, 398t epigenetic changes in liver injury, 759 liver injury mechanisms, 744 metabolism pathways, 743–744, 747 see also CYP2E1 synergistic action with iron, 754 alcohol dehydrogenase, 743 alcoholic cirrhosis, chemokine expression, 749 alcoholic fatty liver, 744–745 endocannabinoid role, 1098–1099 see also alcoholic liver disease (ALD) alcoholic hepatitis apoptosis in, 794 chemokine expression, 749 coagulation changes, 645–646 alcoholic liver disease (ALD), 743–772 antibody formation and, 755 antioxidants, effects, 755–756 dual-hit hypothesis, 748 endocannabinoid role, 1098–1099 epigenetic changes, 759

1141 future research directions, 759–760 hepatic fibrosis, 434–435, 746, 748 management, 434–435 histology, 744 pathogenesis and risk factors, 744–750 acetaldehyde, 747 , 747–748 adipokines, 746–747 bacteria, endotoxin and TNFα, 749–750 cytokines, chemokines, growth factors, 748–749 insulin resistance, 745–746 iron overload, 245, 750–751, 754 lipid alterations, 744–745 obesity, 745 oxidative stress, 139, 750–751, 754 SREBPs role, 630, 744 proteasome function changes, 757–758 reactive nitrogen species, 756–757 ROS generation, 750–755 sources, 750–753 species/types, 753–755 unfolded protein response, 758–759 alcoholic steatohepatitis (ASH), 745, 746 aldehydes, reactive, alcohol-induced, 744 aldo-keto reductases, 290 aldolase, as glucose sensor and V-type ATPase reassembly, 66 alginate scaffolds, 941 allyl alcohol, toxicity, 775 alpha-1-antitrypsin deficiency autophagy dysfunction, 183 , 184 ER stress, 788 hepatic fibrosis due to, 435 hepatocyte transplantation, 589 alpha-adrenergic agonists, hepatic vascular resistance increase, 708 alpha-fetoprotein (AFP), hepatocellular carcinoma, 871, 999 alpha oxidation, in peroxisomes, 191, 192–193 alternative lengthening of telomeres (ALT), 1107, 1112 Alzheimer’s disease, neuroprotection by sirtuin 1 (SIRT1), 958 Alzheimer Type II astrocytosis, 601, 601 α-amanitin, 788 amidates, 291, 295 amiloride, 444 amino acids, from protein degradation, role in starvation, 173, 182–183 amino acid transporter, at lysosomal membrane, 177 ammonia, 602–603 astrocyte glutamate release, 606, 607 brain, accumulation, 602 effect on brain glucose metabolism, 604, 605 effects on brain function, 603 hepatic encephalopathy pathogenesis, 602–603, 603 , 604, 605 management approaches, 610 inter-organ trafficking, 602, 603 production, agents decreasing, 610 removal, 602, 610–611 AMOT (angiomotin), 88t AMP-activated protein kinase kinase (AMPKK), 537 amphiphysin 1, clathrin-coated vesicle uncoating, 111 amphiregulin, liver regeneration, 559 amphisomes, 180

1142 AMPK (AMP-activated protein kinase), 141, 535–548 activation, 537, 541 by adiponectin, 537, 625, 625 , 725 CB2 receptors, in hepatic encephalopathy, 1100 energy stress causing, 535, 536, 537 by metformin, 479, 537, 541, 544, 545 by stress, hormones or drugs, 537 by thiazolidinediones, 537 catalytic (α) subunit, 536, 536 , 538 AMPKα2 knockout, 540–541 discovery, 535–536 downstream targets gluconeogenesis enzymes, 539–541, 540 lipogenesis enzymes, 538–539, 539 , 541, 732 mTOR pathway, 542–543, 543 phosphorylation of, 539–540, 540 , 541 functions, 535 acute metabolic response regulation, 538–539 cell growth regulation, 542–543 gluconeogenesis enzyme regulation, 539–541, 540 insulin signaling, 543 , 543–544 lipid metabolism regulation, 539, 543–544, 744–745 lipogenesis inhibition, 541 metabolic adaptation, 539–542, 540 mTOR regulation, 542–544, 543 transcriptional control, 539–542, 540 glycogen-binding domains (GBDs), 536–537 hepatic fibrosis, 442–443 metformin mechanism of action, 479, 537, 544 mouse models, 541t polarity mediator, 88t, 92 PPAR interactions, 541–542 reduced activity after feeding, 539 regulatory (β and γ) subunits, 536 , 536–537 structure and genes coding, 536, 536 as therapeutic target, 544–545 upstream regulators (LKB1 and CAMKK), 537–538 AMPKK (AMP-activated protein kinase kinase), 537 AMP-PNP, kinesin inhibition, 50, 51 amyloid β-peptide, 958 anandamide, 444, 1093 degradation, reduced, high-fat diet, 1097 hepatic ischemic-reperfusion injury, 1100 receptor, 1094 vasodilatation, in cirrhosis, 1094 anatomy of liver, 5–6 anemia of chronic disease, elevated hepcidin, 244–245 iron refractory, 245 angiogenesis by activated stellate cells, 437 inhibition, tumor therapy, 966 therapeutic implantable liver devices, 944 tumors, miR-17–92 up-regulated by c-Myc, 1044 up-regulation by HCV, 1002 angiomotin (AMOT), 88t

INDEX angiopoietin II, 410 angiopoietin-like protein, 4 721, 721 angiotensin, calcium signaling, 492 angiotensin II activated stellate cells stimulated, 437 hepatic fibrosis pathogenesis, 437, 439, 440 hepatic vascular resistance increase, cirrhosis, 707 NADPH oxidase role in stellate cells, 752 released by adipose cells, and effects, 624t type 1 receptor, blockage, 730 angiotensin receptors type 1 (AT1), hepatic fibrosis, 439, 440 inhibition, 439, 445 animal models dose-related hepatotoxicity, 775–776 hepatocellular carcinoma, 993–994 idiosyncratic adverse drug reactions, 776–777 implantable therapeutic construct assessment, 944 liver repopulation, 579 , 579–581, 580 telomeres and telomerase see telomerase; telomere(s) see also specific models and conditions “anoikis”, 796, 1126 antagomirs, 1035, 1036 anti-apoptotic proteins, 490, 789 antibiotics, poorly absorbed, 610 antibodies alcohol-induced oxidative injury and, 755 to HBV, 839–840 defective synthesis, HBV persistence, 840 to HCV, 844–845, 881, 882 persistent infections, 881–882 anticoagulant, protein C action, 646 anticoagulation factors, 648–649 antifibrogenic drugs/agents, 407, 413 antifibrotic therapies, 433, 443–445, 444t CB2 agonist, 1095 see also hepatic fibrosis, therapies antigen(s), foreign, removal mechanism, 10 antigen presentation, 10, 755, 967 sinusoidal endothelial cells, 10, 376 see also dendritic cells anti-inflammatory drugs, 444, 611, 777 antioxidant response element (ARE), 1131 activation, redox sensor (Keap1), 1133–1134, 1134 discovery and action, 1132, 1133 plants, 1132–1133, 1133 transcription factors controlling, 1133, 1134 antioxidants alcoholic liver disease, 755–756 bilirubin action, 252 GST role, 1132 hepatic fibrosis management, 444 mitochondrial permeability transition prevention, 605–606 NO availability, in cirrhosis, 706 reduced capacity in Wilson disease, 227 stellate cell activation prevention, 412 α2-antiplasmin (α2-AP), 641, 648 antisense oligonucleotides (ASOs), 264, 967 Ntcp translation inhibition, 306

protein kinase Cε, hepatic insulin resistance, 476–477 targeted gene modification by, 976–977 antithrombin (AT), 640, 648 antithrombin III (AT III), 648 α1-antitrypsin deficiency see alpha-1-antitrypsin deficiency AP-1, 109, 126 activation, HCC development, 994 c-Fos dimerization with c-Jun, 529 clathrin-coated trans-Golgi network-derived vesicles, 129, 130 function, 109 induction by CB2 receptor, antifibrotic effects, 1095 stellate cell activation, 413t subunits, 129 AP-1A (adaptor protein-1A) basolateral protein recognition, 81 basolateral protein sorting signal, 81, 82–83 isoforms, 82–83 AP-2, 109, 110 AP-3, 109 AP-4, 109 Apaf-1 (apoptotic protease activating factor, 1) 784, 784 APC (adenomatous polyposis coli) gene/protein apc mutations in zebrafish, 1071 inactivation, HCC, 1005 mutations in hepatoblastoma, 1071 polarity mediator, 89t, 92 apical domain, hepatocytes, 74, 74 apical plasma membrane (APM), 73 ABC transporter targeting to, 350 , 350–351, 1078 actin filament organization in hepatocytes, 32 canalicular ABC transporter recycling see ABC transporters cholangiocyte, purinergic receptors, 361 dynamic nature, 75 exocytosis, 35 hepatocyte, 74, 74 , 75 protein biogenesis/trafficking “indirect” route, 75, 77 MDCK cells, 77, 78 recognition mechanisms, 81–82 sorting on “rafts”, 82 sorting signals, 81, 82 proteins, abnormal localization (ARC syndrome), 669 transporters, 1078 regulation, 349 see also ABCB1 (MDR1; Pgp); ABCB4 (MDR3); ABCB11 (bile salt export pump (BSEP)) apical sodium-dependent bile salt transporter see ASBT apoB see apolipoprotein B apoB editing complex-1 (apobec-1), 273 apo-ceruloplasmin, 226–227 apolipoprotein(s), 271 apolipoprotein A (ApoA) apoA-I, 278 apoA-II, 278 apolipoprotein B (ApoB) apoB48, 273, 275 apoB100, 273, 275 in LDL, 277, 277 gene, 273 intravascular metabolism, 275 , 275–276

INDEX lipoprotein assembly, 273–275, 274 lipoproteins containing see chylomicrons; lipoprotein(s); low-density lipoprotein (LDL); very low-density lipoproteins (VLDL) mRNA editing, 273, 273 , 965 regulation, role of protein quality control/degradation, 167 apolipoprotein B mRNA editing enzyme (APOBEC-1), 965 apolipoprotein C (ApoC) apoC-I, 275 apoC-II, 275 apoC-III, 275, 276 apolipoprotein E (ApoE), 275, 276–277 apoproteins see specific apolipoproteins apoptosis, 140, 783–802 of activated stellate cells, 1126 induction by drugs, 444 resistance to apoptosis, 436 spontaneous recovery from fibrosis, 436 in alcoholic hepatitis, 794 anti-apoptotic drugs/agents, 796 anti-apoptotic mechanisms, 795–796 anti-apoptotic proteins, 786 , 786–788 bile acid-induced, 794, 795 calcium signaling role, 490 in cholestasis, 793–794 copper-induced, 227 definition, 783 detachment-induced (anoikis), 796, 1126 drug-induced, 143 ER stress and, 788, 796 inhibition by gelsolin, 456, 458 inhibitor of (IAP), 750, 790 initiation, TNF-α signaling and, 750, 790–792, 791–792 intrinsic mitochondrial pathway, 783, 784 , 784–786 activated by bile acids, 793 activation, 788–789 activation in steatohepatitis, 728 Bcl-2 proteins, 786 , 786–788 mechanism, 784 , 784–786 see also Bcl-2 family in liver diseases, 793–794 lysosomal pathway, 728 activation in steatohepatitis, 728 TNFR1 signaling initiating, 783, 790, 791 TRAIL role, 783, 793 mitochondrial permeability transition, 513, 785 NAFLD/NASH, 727, 728, 794 as therapeutic target, 733 pathways/mechanisms, 442, 783–793, 784 anti-apoptotic proteins, 786 , 786–788 Bcl-2 proteins, 728, 784 , 786 , 786–788 death receptors, 783, 790–793 extrinsic death receptor, 783–784, 784 intrinsic mitochondrial see above pro-apoptotic proteins, 786, 786 regulation, mitochondrial role, 140, 785 signaling pathways, hepatic fibrosis, 442 TNF-mediated, 750, 791–792 , 791–793 protection by NFκB, 555, 791 , 791–793 Type I and Type II cells, 783 apoptosis inducing factor (AIF), 140 apoptosome, 784, 784

AQP1, 366 arachidonic acid, 758 2-arachidonoylglycerol (2-AG), 1093, 1096, 1099 architecture of liver see liver organization ARC syndrome, 668–669, 668t zebrafish model, 1070–1071 ARE see antioxidant response element (ARE) AREBP (AICAR response-element binding protein), 541 ARF (ADP ribosylation factor), 126 ARF (ADP ribosylation factor)-binding proteins (GGAs), 129 ArfGAP1 lipid packing sensor (ALPS), 149 L-arginine, 756, 757 reperfusion injury association, 380 arginine-glycine-aspartate (RGD) motif, 457 Argonaute proteins, 1030 , 1032 Ago2 protein, 977 Arp1 filament, 48 Arp2 and Arp3 proteins, 31 array comparative genomic hybridization (aCGH), 992, 992 , 994, 1001 HCC metastases, 1004–1005 arrays see microarrays arthrogryposis, renal dysfunction and cholestasis see ARC syndrome aryl hydrocarbon receptor (AhR), 1078 ASBT (apical sodium-dependent bile salt transporter), 341 bile acid transport regulation, 297, 300, 329, 341, 682 expression in rabbits vs rats after cholesterol feeding, 324 functions, 682 increased expression after SC-435, 331 inhibition by SC-435, 331 regulation by FXR and bile acid flux, 329 ascites fluid, fibrinolysis associated, 650 asialoglycoprotein receptor (ASGPR), gene therapy, 974, 980 asialo-orosomucoid (ASOR), 51, 973 L-aspartate, 610 aspartate–glutamate carrier, mitochondrial, 670–671 astrocytes Alzheimer Type II, 601, 601 glutamate release, ammonia effect, 606, 607 glutamate uptake, 606, 606 hepatic encephalopathy, 601, 603, 605 swelling, brain edema, 603, 606, 606 AT-61, 861–862 ATF-1 (activating transcription factor-1), 523–524 structure, 524, 524 ATF-6 (activating transcription factor, 6) 167, 265, 728, 788 ATG (autophagy-related genes), 179 Atg5, Atg12 and Atg8, 179 Atg6/beclin-I, 180 ATG7 gene, 181 Atg11, 198 Atg30, 198 atherosclerosis, 271, 377 ATM protein kinase, cell cycle control, 1016 atomic force microscopy (AFM), 1058 Atox1, copper transport, 224, 226

1143 ATP AMPK as sensor of levels, 535, 539 see also AMPK (AMP-activated protein kinase) Ca2+ waves induced, 494 calcium signaling response to, 361 cell volume changes, 498 consumption/utilization, AMPK suppressing, 535 F-actin polymerization, 31 hydrolysis motor protein action, 45 V-type ATPase mechanism, 65 synthesis AMPK stimulating, 535 ATP synthase and, 60, 62 by mitochondria, 138–139 ATP7A and ATP7B, 57, 60 ATP8B1 (PFIC-1), 254, 663–665, 664t cholestasis, role in, 343, 664, 693 defects/gene mutations, 341, 343, 663–665 flippase function, 343, 345, 693 see also progressive familial intrahepatic cholestasis (PFIC) ATPases, 32 chaperones as, 162 copper-transporting see copper-transporting ATPases F-type see ATP synthase myosins and kinesins, 45 P-type see P-type ATPases soft-metal-ion-transporting, 57, 58t, 60 V-type see V-type ATPases ATP-binding cassette (ABC) transporters see ABC transporters ATP synthase, 60–63 assembly, 63 bacterial, 62, 63 F0 motor, 61, 63 Asp-61, 62 subunits a and c, 62 F1 motor, 61–62, 63 α- and β- subunits, 62 mechanism of action, 61 , 62 mitochondrial, 61, 63 regulation, 63 rotary catalytic mechanism, 61 , 62–63 rotary motors (F0 and F1 ), 61–62 stalks (central and peripheral/‘stator’), 62 structure, 61 , 61–62 ATR protein kinase, cell cycle control, 1016 Australia antigen, 803–804, 864 see also hepatitis B virus (HBV), HBsAg autoimmune hepatitis (AIH) endocannabinoid action, 1100 hepatic fibrosis due to, 435 murine model, 394 autoimmune liver diseases, hepatic fibrosis due to, 435 autophagolysosomes, 180, 181 autophagosome, 180, 181 formation inhibition, 184 lysosome fusion, 180 PEX14 recruitment of, perophagy, 198 autophagy/autophagic pathways, 179–183, 180 , 567 in aging, 183 , 184–185 carcinogenesis and, 183 , 184 chaperone-mediated, 180 , 182–183, 185

1144 autophagy/autophagic pathways (continued) definition, 179 in liver disease, 183 , 183–184 macroautophagy see macroautophagy microautophagy, 180 non-selective and selective, 181, 182 ultrastructure of components, 181 autophagy-related genes (ATG), 179 autosomal dominant polycystic kidney disease (ADPKD), 364, 364t autosomal recessive polycystic kidney disease (ARPKD), 364, 364t, 365, 365 , 366 auxilin, clathrin-coated vesicle uncoating, 111 avihepadnaviruses, 810 5-aza-dC, hepatocellular carcinoma, 1004

B BAAT mutations, 669–670 bacteria bile acid modification, 292, 295 gram negative alcoholic liver disease, 749–750 inflammation, endotoxin translocation, 749, 773 bacterial artificial chromosome (BAC), 1001 bacterial overgrowth, bile acid changes, 299 baculovirus, recombinant, 973 Bad protein, 750, 787, 788 bafilomycin, 515 bafilomycin-induced calcein quenching, 513, 514 , 515 Bak/BAK protein, 140, 143, 783, 786 activation by p53, 788 BAMPI (bone morphogenic protein and activin membrane-bound inhibitor), 439 Bardet–Biedl syndrome, 364, 364t bariatrics, NAFLD management, 730 barmotin (7H6), 87t basal domain, hepatocytes, 74, 74 basement membrane, 453, 455 see also extracellular matrix (ECM) basic leucine zipper domain see bZIP transcription factors basolateral plasma membrane (BLM), 73 bile acid transporters, adaptive regulation, 682, 687–688 dynamics of endocytosis/secretion, 75 exocytosis and transcytosis, 35 hepatic efflux of bile acids in cholestasis, 690–693 hepatocytes, 74, 74 , 75 Ntcp retention/targeting, 308, 687 see also Ntcp/NTCP oatp1a1 (oatp1) expression, 313, 314, 688 organic anion transporters see organic anion transporter(s) protein delivery route, 77 G823D mutation, 83 MDCK cells, 77, 78 , 81, 82 recognition mechanisms, 81, 82–83 sorting signals, 79, 81, 82 Bax/BAX protein, 728, 750, 783, 786 activation by p53, 788 apoptosis in NASH, 728, 794 mitochondrial pore formation, 140

INDEX mitofusin interaction, 143 phosphorylation, 786 BAY41–, 4109 863 B-cell lymphoma 2-associated X protein see Bax/BAX protein B-cell lymphoma 2-interacting mediator of cell death see Bim B-cell lymphoma 2 protein family see Bcl-2 family B cells, 10 HBV infection, 837 HCV infection, 844–845 viral infections, 835–836 Bcl-2 family, 140, 786 , 786–788 anti-apoptotic, 140, 786 Mcl-1, 490, 786, 787 overexpression, p53-dependent senescence and, 1111 apoptosis role, 783 Bcl-2 protein (anti-apoptotic protein), 140, 786 BH3 see BH3 proteins calcium signaling and apoptosis, 490 pro-apoptotic, 786, 786 see also Bak/BAK protein; Bax/BAX protein steatosis progression to steatohepatitis, 728 Bcl-2 homology domains (BH1–, 4) 140 Bcl-xL , 786–787, 795 BCRP see ABCG2 (BCRP) benign recurrent intrahepatic cholestasis type I (BRIC-I), 254, 663–665, 664t benign recurrent intrahepatic cholestasis type II (BRIC-II), 254, 663, 664t, 665 beta-cells, 19, 22 beta oxidation, 258 disorders, 193 fatty acids, 725 , 726 in peroxisomes, 191, 193 BH3 proteins, 786 , 787 mitochondrial apoptosis pathway, 140, 784 , 785 proteins binding to, 787, 787t bicarbonate, in bile, 339 Bid and t-Bid, 750, 783, 787, 790 biguanides, 478–479, 537 see also metformin “bi-hormonal hypothesis”, 478 bile bicarbonate, 339 bile salts see bile acid(s) canaliculi see bile canaliculi cholesterol secretion into, 277–278, 280, 342–343, 345 composition, 363 conjugated bile acids in, 295 drug/toxin and waste product transport into, 339–341 ductal, 363 flow, 202, 375 Ca2+ signaling role, 497 inborn errors, 661–679 obstruction, 661 pulsatile, 202, 363 flow rate, cholangiocyte cilia response, 361, 363 formation, 280, 339, 359 ductal, and cholangiocyte cilia, 363 steps/phases, 681–682, 687–690 transport proteins involved, 339, 340, 340 , 349, 681, 683t–684t functions, 339, 355

glutathione, 339 osmolarity, 339, 363 primary, 363 secretion, 273, 339–348, 497, 663, 681 cAMP effect, 351 in cholestasis, 682 hepatocyte plasma membrane role, 75 inborn errors, 661–679 regulation, 355 wormannin effect, 353 see also canalicular membrane solutes in, and concentrations, 339 bile acid(s), 289–304 abnormal, production of, 662 activation of FXR for regulation, 325 N-acylamidation (amidation), 291, 293 , 295 albumin-bound, 296, 305 amidation and amidates, 291, 295 in cholestasis, 690 antimicrobial properties, 292 bacterial modification, 292, 295 biliary, 295 biosynthesis, 289–291, 293, 293 , 662, 688–689 “acidic/alternative” pathway, 291, 688–689 from cholesterol, 193–194, 280, 289–290, 290 , 323 defects, 291 enzymes, 685t failure, disorders, 662, 663t 7α-hydroxylation, 280, 281, 290 increased by bile salt sequestrants, 281, 300 increased in ileal dysfunction, 297 inhibited in cholestasis, 682, 688–689 negative feedback, 297, 323, 324, 325, 327, 332, 689 “neutral” pathway, 291, 688–689 nuclear receptors regulating see FXR; LXR; nuclear receptors peroxisomes role, 193–194 rates and pool sizes, 290 reduced in cholestasis, 682, 689 regulation, 297, 323–337, 688–689 regulation, future challenges, 332 site (pericentral hepatocytes), 290, 297 see also cholesterol 7α-hydroxylase (CYP7A1) C24 , 291, 292t, 293 , 296 C27 , 291, 293 Ca2+ signaling induced by, functions, 498–499 chemistry, 289–292 in cirrhosis, 299, 412–413 conjugated, 295 hepatic uptake, 296 intestinal conservation/absorption, 294–295 uptake mediated by Ntcp, 307 conjugation, 291, 293 , 294, 681, 689 in cholestasis, 689–690 defects, 298 cytokines in macrophage induced, 333 damage and repair during recycling, 295 deconjugation, 292, 295 deficiency, intestinal, 299 dehydroxylation, 292 7-deoxy bile acids, 292 enterohepatic circulation see enterohepatic circulation epimerization, 295

INDEX evolution and phylogeny, 291 excretion/loss, 294 , 295, 341, 682 transporters involved, 682 upregulation in cholestasis, 682 fecal, 295 functions, 289, 297–298, 298t, 323, 341 lipid secretion, 280, 298 glucuronidation, 291, 293 , 295, 689–690 glycogen phosphorylase activated, 496 hepatic efflux in cholestasis, 681, 690–693 hepatic uptake, 295–296, 305–312, 681 defects, 299 impairment, familial hypercholanemia, 669–670 reduced in cholestasis, 310, 682, 687 transporters involved, 682 hepatic uptake, basolateral membrane, 305–312 in cholestasis, 687–688 microsomal epoxide hydrolase, 311–312 sodium-dependence, 305–306, 311–312, 687–688 see also Ntcp/NTCP hepatocyte affinity, 305–306 hydrophobic, apoptosis due to, 793, 794 3α-hydroxy group, 295, 306 hydroxylation, in cholestasis, 689 ileal transport, 294–295 in cholestasis, 684 defects, 298 feedback control on CYP7A1, 324 inhibitors, 300 regulation, 297 inborn errors of metabolism, 291, 299 increased intrahepatocyte levels, 661 intermediates, accumulation, 662 intestinal conservation, 280, 281, 294–295 regulation, 329–330 intrinsic apoptotic pathway activation, 793 iso (3β-hydroxy), 295 malabsorption, 299 effects on ASBT and pool size, 331 SC-435 role, 331 membrane asymmetry as resistance against, 343–345, 344 metabolic disturbances/disorders, 298–299 mixed micelle formation, phospholipid excretion, 341–342 modified β-oxidation of side chain, 290 NAFLD and renal disease, 630–631 , 632–633 negative feedback control, 297, 323, 324, 325, 327, 332 nuclear receptor activation see nuclear receptors 7-oxo bile acids, 295 p38 MAPK phosphorylation, 794 plasma, 296 elevated levels, 299, 310 pruritus association, 299 pool, 293, 294 in cholestasis, 684 cholesterol feeding effect, 330 measurement methods (animals), 330 , 330–331, 331 negative feedback by, 297, 323, 324, 325, 327, 332

in rabbits vs rats after cholesterol feeding, 324 size, effect of ASBT inhibitor, 331 primary, 292 reabsorption, intestinal, 280, 281, 294–295, 341 dietary cholesterol effect, 329–330 inhibitors, 281, 300 regulation, 329–330 receptor, G-protein-coupled see TGR5 recycling, 280, 293–296, 294 damage and repair during, 295 frequency, 294 see also enterohepatic circulation, bile salts retention in hepatocytes, 298 root, CDCA see chenodeoxycholic acid (CDCA) secondary, 292 secretion (flux) into intestine, 280, 294, 296 sensors, FXR as, 325, 332 structure, 290, 290 side chains, 290, 291, 293 therapeutic application, 299–300 toxicity, protection against, 341 transport, disorders, 662–666, 664 transporters see bile acid transporters trihydroxy, 291 unconjugated, 295, 689 absorption in intestine, 295 complexity/variety, 295 hepatic uptake, 295–296 bile acid agonists, 299–300 bile acid-CoA:amino acid N -acyltransferase, 685t, 689 deficiency, 663t, 669 bile acid-CoA ligase, 296 deficiency, 663t bile acid-CoA synthetase, 685t, 689 bile acid congeners, 299–300 bile acid glucuronides, 295 bile acid response element (BARE), 325–326, 327, 687 bile acid sequestrants, 281, 300 bile acid transporters, 329, 341, 681, 683t–684t abnormal, disorders, 662–666, 664 , 682–693 adaptative regulation in cholestasis, 682 , 682–693 basolateral membrane, 682, 687–688, 690 canalicular membrane, 690, 691–693 gene regulation, 687 hepatic efflux, 690–693 nuclear receptors regulating, 684–687, 686t transcription factors regulating, 684–686 see also specific transporters (see pg. 683–684) ASBT see ASBT intestinal, in cholestasis, 684 regulated by FXR, 329 renal, in cholestasis, 684 sodium-dependent, 306 see also Ntcp/NTCP types, location and functions, 683t–684t uptake/excretion of bile acids, 682 bile alcohols, 291, 293 C27 , 291 bile canaliculi, 9, 74, 108 , 375

1145 bile secretion into, 663 bilirubin glucuronide transport, 254 contraction, Ca2+ signaling role, 497 disruption in cholestasis, 355 formation, canalicular recycling system and, 355, 355 membrane see canalicular membrane peristalsis, 497 tight junctions around, 201–202 zebrafish, development, 1069 bile duct(s) development, 4, 1069 epithelium, 454 zebrafish, 1069 intrahepatic formation, 584 hypoplasia see Alagille syndrome (ALGS) proliferation, in cholestasis, 682 secretion from, Ca2+ signaling role, 499–500 terminal, 9 see also entries beginning biliary bile duct cells Ca2+ signaling, 496 epithelial development, 454 hepatocyte differentiation into, 580 hyper-proliferation, miRNA role, 1044–1045 production from “oval cells”, 581 see also cholangiocyte(s) bile fistula, 330, 330 , 331 bile salt export pump (BSEP) see ABCB11 (bile salt export pump (BSEP)) bile salts see bile acid(s) bile salt sequestrants, 281, 300 biliary atresia, 661 extrahepatic, 661–662 biliary fibrosis, secondary, 435 biliary nucleotides, 363 biliary obstruction bile acid metabolic changes, 298, 310 prolonged, hepatic fibrosis due to, 435 bilirubin, 251 albumin binding, dissociation/transport, 253 antioxidant action, 252 blood–brain barrier passage, 252 conjugated, accumulation, 253, 254 conjugation, 252, 253 degradation, 254 “direct”-reacting, 252 formation, 251 benefits/reasons, 252 glucuronidation, 252, 253 measurement, 252, 253 metabolism, 251–256 disorders, 252–255 enzymes, 685t excretion, 254 regulation, 254 placental extraction, 252 as reactive oxygen species scavenger, 756 serum levels elevated in neonates, 251, 252 inverse relationship to disease, 252 sodium-independent transport/uptake, 312 structure, 252 “total”, 252 toxicity, 252

1146 bilirubin (continued) transport protein, 253 unconjugated, 252, 253, 254 elevated levels, 252 bilirubin glucuronides, 252, 253, 254 transport, 253, 254 bilirubin IXα, 252 bilitranslocase, 312 biliverdin, 251–252 biliverdin IXα, 252 biliverdin reductase, 252 Bim (B-cell lymphoma 2-interacting mediator of cell death), 728, 787–788 ER stress apoptosis mediated by, 788 binning, signal-to-noise ratio improvement, 1056, 1057 bioartificial liver devices (BAL), extracorporeal, 935, 936, 937 Biological Expression Network Discovery (BLEND), 1008 biomatrix, hepatocyte culture, 938 bioreactor(s), cell-based, designs, 936, 937 bioreactor cultures, 939–940 flat plate, 939–940 BiP (IgG Binding Protein), 164, 167 chaperone role, 162 bipotential mouse embryonic liver (BMEL) cell line, 937 bitter melon, AMPK activation, 537 blastema, formation, 549, 550 bleeding diatheses, 647 blood–biliary barrier, 217 blood–brain barrier, 340 alterations, hepatic encephalopathy, 602 bilirubin passage, 252 permeability change, brain ammonia increase, 602–603 blood flow, 6 regulation, 6–7 by stellate cells, 409 blood supply, 5–6, 375 arterial, 3, 375 therapeutic implantable liver devices, 944 blood vessels anatomy, 3, 5–6 development, 4–5, 21 hemodynamics, 3–4, 6–7 B lymphocytes see B cells Bmf protein, 787 BMP4, hepatic endoderm development, 19 BMP–SMAD pathway, hepcidin signaling and regulation, 240–241, 241 boceprevir, 911, 912 bone marrow cells expressing MMPs, 461 liver cell regeneration and, 11 stem cells, liver repopulation by, 585–586 transplantation, 11 bone morphogenetic protein (BMP), 244 BMP4, hepatic endoderm development, 19 liver development, 584, 584 activation, miR-23b cluster action, 1041 zebrafish, 1068 bone morphogenetic protein receptor (BMPR), 240 bone morphogenic protein and activin membrane-bound inhibitor (BAMBI), 439 bosentan, 707

INDEX Boyer’s theory, 62 bradykinin, InsP3 R regulation, 495 brain edema, 603–604 hepatic encephalopathy, 597, 601, 607–608 gene expression, hepatic encephalopathy, 609 branched-chain amino acids, supplementation, 610 BrdU, stem cell identification, 583 breast cancer, 994 ABC transporters, 1082 breast cancer resistance protein (BCRP) see ABCG2 (BCRP) brefeldin A, 211 bromodeoxyuridine, 210 brown adipose tissue, 296 BSC1L mutations (GRACILE syndrome), 667t, 670 BSEP, bile acid uptake see ABCB11 (bile salt export pump (BSEP)) BSP (sulfobromophthalein), sodium-dependent/-independent transport of, 307, 312, 315 BSP/bilirubin binding protein, 312 Budd–Chiari syndrome, 435, 649 bulk critical concentration, microtubules, 30 bush teas, 377–378 tert-butylhydroperoxide (TBH), 515, 515 Byler disease (PFIC-1), 663–665, 664t, 693 bystander effect, 966 bZIP transcription factors, 523 Maf-protein and Nrf2 binding, 1133, 1134 TGA factors in plant xenobiotic detoxification, 1135, 1135

C C1q receptor, 847 C3, activation after PH, regeneration, 554 C3a, priming phase of liver regeneration, 553–554 C5a, priming phase of liver regeneration, 553–554 Ca2+ see calcium ions Ca2+ -ATPase, 57, 495 plasma membrane (PMCAs), 58 Ca2+ -induced Ca2+ release (CICR), 489, 490 Caco-2 cell line, plasma membrane protein/traffic research, 76, 77t, 79 CAD-based rapid prototyping, scaffolds, 943 cadherins, adherens junctions association, polarity development, 84, 85t cadmium, hypercoagulation due to, 649 Caenorhabditis elegans RNA interference, 1029 Sir2 orthologs, 955–956, 957 calcein quenching, 513, 513t, 514 bafilomycin-induced, 513, 514 , 515 calcineurin, 488, 499 calcium channel blockers, fenestrae regulation, 397 calcium ion binding protein, 499 calcium ions calcium release induced by (CICR), 489, 490 channels, 486–489, 493 novel, 493

see also inositol 1,4,5-triphosphate (IP3 ) receptor; ryanodine receptor chelators, 499 cytosolic (Cai 2+), 485 levels, 141 dyes, measurement, 491 ethanol increasing entry, 752 fenestration regulation, 395, 396–397 free, in mitochondria, 490 functions, 485 gap junction channel closure, 211 gap junction-mediated signaling, 208, 209 , 494 plasma membrane channels, 493 release, 486, 487 , 489, 493 from endoplasmic reticulum, 141 in mitochondria, 490 as second messenger, 141, 485 signaling see calcium signaling spikes (oscillations), 492–493, 499 storage pumps, accumulation in nuclear envelope, 495 tight junction regulation, 215 uptake by mitochondria, 491 waves, 361, 491, 493–494 propagation via gap junctions, 208, 209 , 494 calcium pumps, 57 calcium-sensing molecule (CaM), 211 calcium signaling, 485–510 acetylcholine induced by, 360–361 in cholangiocyte cilia, 360–362, 500 activation, 361, 363 ductal bile formation, 363 functions/effects, 360–361, 485, 496–500 bile flow and paracellular permeability, 497 canalicular contraction, 497 cell proliferation, 499 cell volume regulation, 498–499 ductular secretion, 499–500 exocytosis, 497–498 glucose metabolism, 496–497 signals induced by bile acids, 498 gap junction-mediated, 208, 209 , 494 hyperproliferative cholangiocytes, 365–366 intercellular signalling, 208, 494–495 mechanisms, 485–491 growth factors and initiation, 486 hormone receptors and initiation, 485–486 inositol 1,4,5-trisphosphate receptor, 361, 486–489 in mitochondria, 489–491 ryanodine receptor, 489 nuclear, 495–496 organization/regulation, 491–496 in bile duct cells, 496 nuclear signaling, 495–496 patterns in hepatocytes, 492 , 492–493 spikes (oscillation), 492–493, 499 spread from cell to cell, 208, 209 , 493, 494–495 subcellular signals and waves, 491, 493–494 see also calcium ions, waves response to ATP, 361, 495 signal detection in hepatocytes, 491–492, 494 calmodulin, 395, 488

INDEX calmodulin-dependent protein kinases (CaMKK) AMPK regulation, 537–538, 539 isoforms, 537–538 calnexin (CNX), 162, 163 , 166 caloric restriction, 185, 398 AMPK activity increase, 537, 538 chronological aging, Sir2 in yeast, 956, 957, 957 lifespan extension via Sir2 in yeast, 955, 956 , 956t calreticulin (CRT), 162, 163 , 166 cAMP see cyclic AMP (cAMP) cAMP/CFTR pathway, ductal bile formation, 363 cAMP-guanine exchange factors (cAMP-GEFs), 795 Can-10 cell line, 76 canalicular cell adhesion molecule (cCAM105; HA4), 350 canalicular fluid, composition, 202 canalicular membrane, 9, 32 ABC transporters in, 339, 340 cycling between pools and, 351–352, 352 trafficking, 350 , 350–351 asymmetry, resistance against bile salts, 343–345, 344 bile formation/secretion, 339–348 constituents, 342 , 344, 345 lipid translocator types, 341–342, 342 phospholipid excretion, 341 , 341–342 transporters, 339, 340, 340 , 349 ABC see ABC transporters (above) apical recycling, 349–358 in cholestasis, 690, 691, 693 hepatic efflux of bile acids, 690 ion transporters, 693 organic solute transporters, 691–692 regulation, 349 solute and lipid transporters, 693 canalicular multi-organionic transporter (cMOAT), 227, 683t, 691 canaliculi see bile canaliculi canals of Hering, 9, 582–583, 583 cancer abnormal cell cycle regulation, 1019, 1021 therapeutic target, 1021, 1022 hepatocellular see hepatocellular carcinoma (HCC) mitochondria and, 143–144 multidrug resistance see multidrug resistance stem cells, 1082 telomere shortening, 1110 , 1110–1111 increased chromosomal instability, 1111 uncontrolled cell proliferation, 1016, 1019 see also tumor(s) cannabinoid(s) (CB), 1091–1092 endogenous, 1092 , 1093 see also endocannabinoids hepatic fibrosis, 439–440, 1095–1096 plant-derived, 1091, 1092 cannabinoid (CB) receptors, 1091–1092 CB1 , 439–440, 1091–1092 activation pathway, 1096 , 1096–1097 alcoholic fatty liver, 1098–1099, 1099 autoimmune hepatitis, 1100 cAMP reduced by, 1095

cirrhosis, hemodynamic changes, 1093–1095, 1094 ethanol-induced 2-AG increase, 1099 expression in brain, 1093, 1097–1098 expression in liver, 1093, 1097 fibrogenesis activation in NASH, 730 hepatic encephalopathy, 1100 hepatic steatosis, 1096–1098 high-fat diet-induced obesity and, 1096–1097 in hypotension and endotoxemia, 1093 insulin and leptin resistance, 1098 NAFLD, 1096 , 1096–1098 NAFLD/NASH therapeutic target, 730, 1097–1098, 1099 pro-fibrotic action, 1096 role in metabolic regulation, 1096–1097, 1098 CB1 antagonists, 730, 732 AM251, 1095 hepatic steatosis treatment, 730, 1099 obesity treatment, 1096 see also rimonabant CB2 , 439–440, 1092 agonists, antifibrotic actions, 1095 autoimmune hepatitis, 1100 diet-induced hormonal/metabolic changes, 1098 hepatic encephalopathy, 1100 hepatic ischemic–reperfusion injury, 1100 HU-308 agonist, 1100 induction by pathological stimuli, 1093, 1095 insulin sensitivity impairment, 1098 JWH-133 agonist, 1100 low level normal expression, 1093 NAFLD/NASH therapeutic target, 730, 1099 protective role in hepatic fibrosis, 1095–1096 SR144528A agonist, 1100 ligands, 1092 , 1092–1093 cannabis chronic hepatitis C outcome, 887 hepatic fibrosis and, 1095–1096 steatosis risk factor, 1098 capillarization, 377, 398, 458, 704 activated stellate cells and, 377, 438 matrilin-2 synthesis, 460 capsule of liver, 6 cells, gap junctions, 206, 208 embryology, 21 CAR (constitutive androstane receptor), 685, 686t, 1020, 1078 CAR (coxsackie and adenovirus receptor), 86t, 204, 972 carbohydrate metabolism, regional differences/hepatocyte zonation, 10–11 carbohydrate response element binding protein (ChREBP), 260, 629 activation, excess energy, 723, 723 inhibition by FXR, 633, 633 lipid/glucose homeostasis, role, 724 , 725 lipogenic enzyme regulation, 541 NAFLD hepatic fat accumulation and gut microbiota, 721, 721 renal disease pathogenesis, 629, 631 as therapeutic target, 732

1147 carbon monoxide (CO) actions, 252 hepatic vascular resistance in cirrhosis, 706 production, in bilirubin formation, 251 vasodilatation, in portal hypertension, 711–712 carbon tetrachloride (CCl4 ) injury CB1 receptors and hemodynamic changes due to, 1093, 1094 cirrhosis induced by, 398, 417 c-met knockout mice and liver regeneration, 558 fibrosis shrinkage, siRNA–gp46 (HSP47), 1125–1126 oncostatin M protecting from, 557 γ-carboxylation, vitamin K-dependent coagulation, 649, 650 carcinogenesis autophagy involvement, 183 , 184 telomeres/telomerase role see telomerase; telomere(s) cardiac fibrosis, 435 cardiolipin, mitochondrial membranes, 137–138 cardioprotection, Sirt1 role, 958 L-carnitine, 610 carnitine palmitoyltransferase, 732 carotenoids, 756 caspase(s), 140, 783 effector, 783, 784 inhibitor, 796 initiator, 783, 788 activation, 783, 784 “induced proximity model”, 783 caspase, 2 788 caspase, 8 783, 789–790 caspase, 9 784 caspase, 10 783, 789–790 caspase, 12 759, 788 caspase recruitment domain (CARD), 783, 879 catalase, 756 overexpression, 143 β-catenin adherens junctions association, polarity development, 84, 86t phosphorylation, HGF role, 558 catenin(s), isoforms, adherens junctions association, 86t cathepsin B, 728, 790, 794 cathepsins (lysosomal proteases), 176 Cav-1-Tyr14, 114 caveolae, 112–114 biogenesis, 113–114 endocytosis of, 113–114 functions, 113 internalization/liberation from plasma membrane, 113–114 lipid composition, 113 structure and distribution, 112–113 caveolin (Cav), 112–113 liver regeneration, 559 types, 113 caveolin-1, in fenestrae, 392–393, 397 reduced, pseudocapillarization, 398 CB1 /CB2 see cannabinoid (CB) receptors CCAAT/enhancer binding protein, stellate cell activation, 413t CCL2, 749 CCL5 (RANTES), 438 CCL19, 749 CCR7, acute HCV infection, 842–843

1148 CD4 T cells defective, chronic HBV, 839 HBV infection acute, 837 chronic, 839 chronically evolving, 838 duration of response, 837–838 HCV infection acute phase, 843–844 chronic, 844 impairment, 843–844 viral hepatitis, 804 CD8 T cells, 748 defective, chronic HBV infection, 839 HBV infection, 835 acute, 837 acute, duration of response, 837–838 chronic, 839 chronically evolving infection, 838 clearance by, 837 multispecific, 838, 839 recruitment into liver, 837 HCV infection acute, 837, 842–843 chronic, 844, 845, 880 escape mutants, and persistence, 845–846, 880–881 impairment, 843, 844 multispecific, 845 HDV infection, 823 immunopathology due to, 835 recruitment into liver, 837 viral hepatitis, 804, 835 CD14, 750 CD31, sinusoidal endothelial cells, 381, 382 CD40, apoptosis and activated stellate cells, 442 CD43, 747 CD44, 456 CD63, 1061 CD81, HCV receptor/uptake, 116, 217, 909 E2 interaction, B cell response, 845 E2 interaction, NK cell activation inhibition, 842 CD94/NKG2A, 842 CD95 see Fas (CD95) CD105, sinusoidal endothelial cell isolation, 381 CD122, 846 CD127, 846 T memory cells, 837, 846 CD146 (melanoma cell adhesion molecule MCAM), 381 Cdc25, 364 CDC25A, 1045 cdc25 family, phosphatases, 1017 Cdc42, 92 KO animals, 92 polarity mediator, 89t, 92 Cdc48, 166 Cdc50 proteins, 343 cdk1, 152 cdk2, 1017 liver regeneration, 1019 cdk4 activation, by cyclin D1 in cancer, 1019 cyclin D1 binding, 1017, 1017 , 1018 see also cyclin D1–cdk4 complex liver regeneration, 1019 cdk6, liver regeneration, 1019 cdk family, 1017 cDNA microarrays, 992, 1000–1001

INDEX C/EBP, 788 celgosivir (MX-3253), 864, 913 cell(s) death see cell death growth, 1015 AMPK regulation of, 542–543 ribosomal biogenesis role, 567, 571–573, 572 see also cell cycle polarity see polarity (epithelial cells) proliferation, 12, 1015 Ca2+ signaling role, 499 loss, in senescence, 1107 telomerase overexpression and, 1106 telomere shortening limiting, 1105–1107, 1106 see also hepatocyte(s), proliferation shape, microtubule role, 30 swelling, hypo-osmotic, 498 transplantation see hepatocyte transplantation types see liver cells volume, regulation Ca2+ signaling, 498–499 hepatic encephalopathy, 603–604, 610 cell cycle, 1015–1027 applications for clinical hepatology, 1021–1022 calcium signals associated, 499 CDC25A regulating, 1045 checkpoints, 1016 loss, in cancer, 1016 G0 phase, 1016 G1 phase, 1016, 1016 , 1017 cyclin D1 role, 1017, 1019, 1020 G1 restriction point, 1016, 1016 abnormal regulation in cancer, 1019 regulation, 1017–1018, 1018 G2 phase, 1016 liver regeneration phases, 552 mitogenic stimuli (growth factors), 1016 M phase, 1016 regulation by cyclins–cdk complexes, 557, 1016, 1017 , 1017–1019, 1018 abnormal in cancers, 1019 abnormal in HCC, 1021 in liver, and liver regeneration, 1019–1021 transgenic mice, 1018 signal transduction pathways, 1016, 1017 single-stranded oligonucleotide, repair by, 981 S phase, 552, 1016 stages/principles, 1015–1016, 1016 stimulation, EGF and TNFα roles, 559 telomere shortening, 1106, 1106 uncontrolled in cancer cells, 1016, 1019 cell death chelatable iron and oxidative stress, 511–512, 512 mitochondrial permeability transition, 513, 516 necrotic, 785 see also necrosis programmed see apoptosis cell lines, hepatic, 937–938 hepatocyte development, 589 plasma membrane protein research, 76 see also individual cell lines cellular FLICE-like inhibitory protein (cFLIP), 789, 790

cellular retinol-binding protein (CRBP), 1122 “central” veins, 3, 7, 8 ceramide, 790 C-erbB-2 oncoprotein (ERBB2), 994 cerebral blood flow (CBF), 607–608 agents/therapy reducing, 611 autoregulation, 608, 611 five-phase classification, 608 hepatic encephalopathy pathogenesis, 607–608, 608 , 609, 610 management approaches, 611 increased, brain edema and hypertension, 607–608, 608 cerebral edema see brain, edema cerebral vascular resistance, 608 cerebrotendinous xanthomatosis, 663t C/ERPα, 754 ceruloplasmin, 226 copper loading, 224 iron release from hepatic cells, 239 Menkes disease, 227 synthesis in liver, 239 Wilson disease, 226 ceruloplasmin (cp) gene, zebrafish, 1069 cFLIP (cellular FLICE-like inhibitory protein), 789, 790 c-fos, activation in liver generation, 529 CFTR see cystic fibrosis transmembrane conductance regulator (CFTR) cGMP see cyclic GMP (cGMP) chaperone(s) binding/release cycles, 162, 166 collagen-specific see heat shock protein 47 (HSP47, gp46) in endoplasmic reticulum, 162–163 lectin-based, 162, 163 misfolded/damaged protein recognition, 166, 173 protein maturation/folding, 161–162 selectivity and mechanism, 161 types and examples in ER lumen, 162, 163 for yeast V-type ATPases, 66 chaperone-mediated autophagy, 180 , 182–183 age-related decrease, 185 inhibition, effects, 183 Charcot–Marie–Tooth disease (CMTX), 210 charge-coupled device (CCD), 1056, 1057 , 1062 chemical-induced injury models implantable therapeutic construct assessment, 944 see also carbon tetrachloride (CCl4 ) injury chemicals, hypercoagulation due to, 649 chemokines alcoholic liver disease, 748–749 cysteine-cysteine (CC), 438 alcoholic hepatitis, 749 hepatic fibrogenesis, 438 cysteine-X-cysteine (CXC), 438 alcoholic hepatitis, 749 secreted by activated stellate cells, 438 upregulation by IFNα, 905 see also specific chemokines beginning CCL, or CXCL chemotherapy gene therapy with, 966–967 sinusoidal obstruction syndrome due to, 378, 398–399

INDEX chenodeoxycholic acid (CDCA), 290, 294, 662 bacterial modification, 292 biosynthesis, 290, 290 , 291 cholestasis effect, 298 deconjugation, 295 FXR activation, 325 therapeutic use, 299 chimeraplast, 977, 978, 978 delivery to hepatocytes, 980 chimeraplasty, 977–982 cell culture studies, 979–980 deletion mutation correction, 980 efficiency, UGT1A1 gene replacement, 980–981 frameshift mutation correction, 980 gene repair mechanism, 978, 979 , 981 insertion mutations, repair, 981 in vivo nucleotide conversion, 980–981 mammalian studies, 979 mismatch with target genomic DNA, repair, 978, 981 missense mutation correction, 980 RecA-dependent pairing, 978, 979 site-specific nucleotide conversion, 979, 980–981 see also gene therapy, targeted gene modification chimeric RNA–DNA oligonucleotides, 977–978, 978 , 979 Chk2 and Chk2, 1016 chloride/bicarbonate exchanger (AE2), 339, 366, 693 chlorpromazine, 776–777 cholangiocarcinoma, 1078 cholangiocellular carcinoma agrin role, 459 extracellular matrix, 459, 463–464 Mcl-1 overexpression, 787 cholangiociliopathies, 363–366 pathogenesis, 364–366 types, genes/proteins affected, 364t cholangiocyte(s), 359 absorption by, hepatic cyst formation, 366 apical membrane, 360 P2Y and P2X purinergic receptors, 361 ATP release, P2Y/P2X receptor activation, 361, 363 CREB role, 528 cytoskeleton, 45 diseases affecting see cholangiociliopathies embryology, 4, 5 , 20 function, 359 growth factors/proteins released by, 359 hyperproliferation calcium signaling, 365–366 cAMP signaling, 365 cholangiociliopathies, 364 Hedgehog signaling (Hh), 366 miRNAs role, 364–365 InsP3 R isoforms, 493–494 polarized (apical/BL membranes), 360 primary cilium, 45, 52–53, 53 , 359–363, 500 calcium signaling, 360–362, 500 cAMP signaling, 362–363 cholangiociliopathy pathogenesis, 364–366 ductal bile formation, 363 formation, 52

functions, 362 genes, mutations, 364–366, 364t length, 52, 360 molecular motors, 52–53, 53 response to bile flow rate, 361, 363 sensory functions, 52, 360, 361–362 structure, 360, 360 see also calcium signaling secretion Ca2+ signaling role, 499–500 hepatic cyst formation, 366 structure, 9, 359–360, 360 cholangiocyte cell line (CCL), 1045 cholangiopathies, 363–366 see also cholangiociliopathies cholate, as FXR ligand, 327 cholelithiasis, low-phospholipid associated (LPAC), 666 cholera toxin, internalization mechanism, 113 cholestasis/cholestatic liver disease adaptive regulation of hepatocyte transporters, 681–702 apoptosis in, 793–794 ATP8B1 role, 343 bile acid conjugation, 689–690 bile acid hepatic efflux mechanisms, 690–693 bile acid hydroxylation, 689 bile acid metabolic changes, 298, 310, 793 bile acid synthesis regulation, 688–689 bile acid transporter changes, 682–693 nuclear receptors regulating, 684–687, 686t see also bile acid transporters bilirubin accumulation, 254 causes, 661 drug-induced, 665, 666 ER stress, 788 extra-/intrahepatic, tight junction disturbances, 215 inherited, syndromic forms, 666–671, 667t–668t inherited conditions associated, 671 intrahepatic, inherited forms, 662–666 hepatocyte transport disorders, 662–666 primary bile acid synthesis disorders, 662 intrahepatic, of pregnancy, 663 intrahepatic familial (FIC1), 664 Ntcp regulation, 310 Oatp1a1 expression modulation, 315 plasma bile acids, 296 pruritus and bile acid relationship, 299 recurrent hepatitis B after transplantation, 926 recurrent hepatitis C after transplantation, 922 therapy, UDCA, 299 cholestatic disorders, inheritable, 661–679 cholesterol, 271–285, 290 absorption, 273, 274 , 281 inhibitors, 281–282 balance, 280–281 biliary, reabsorption, 280, 281 biliary excretion, 342–343 biosynthesis, 273 control by SREBPs, 627 convergent feedback inhibition model, 627, 628 , 629 drugs inhibiting, 281

1149 miR-122 role, 1036–1037 peroxisome deficiency and ER stress relationship, 627, 630 in caveolae, 113 conversion to bile acids, 193–194, 280, 289–290, 290 , 323 dietary, 280–281 effect on bile acid reabsorption, 329–330 effect on FXR and LXRα, 327 response of CYP7A1 in rabbits vs rats, 323–324, 325, 327–328 loss in fecal bile salts, 280, 298 metabolism, 271–281 plasma membrane, disruption/disturbance, 113 “rafts” for apical protein sorting, 82, 113 removal, HDL role, 278, 279 reverse cholesterol transport, 278, 279 secretion into bile, 277–278, 280, 342–343 structure, 290, 290 total, 281 trafficking/transport caveolae role, 113 HDL role, 272 uptake, convergent feedback inhibition model, 627, 628 , 629 cholesterol 7α-hydroxylase (CYP7A1), 280, 290, 323 activation, 333 insulin role, 333 rabbits vs rats, 324 in cholestasis, 689 deficiency, 291, 663t dietary cholesterol response in rabbits vs rats, 323–324, 325, 327–328 downregulation, 297, 325, 326, 689 effect of bile acid pool size on, 324, 325 plasma cholesterol effect on, 324 regulation bile acid pool effect, 297, 323, 324, 325, 327, 332, 689 dietary cholesterol effect, 297, 323–324, 325, 327–328 FGF15 role, 297, 332, 689 FXR role, 297, 325–326, 328–329, 332 FXR–SHP–FTF cascade, 297, 328 , 328–329 HNF4 role, 297, 326–327, 333 insulin role, 333 LXR role, 325 mechanism via SHP and LRH-1, 325–327 by nuclear receptors, 327–329 signaling, role, 297, 333 regulators (negative) FGF15, 297, 332 FTF (LRH-1), 326–327 FXR, 297, 325, 332 see also FXR; liver receptor homolog-1 (LRH-1; FTF) regulators (positive transcriptional factors), 326, 327 see also hepatocyte nuclear factor 4 (HNF4); LXR (liver X receptor) transcription factors affecting, 297, 326, 327, 689 upregulation, 281, 326, 327, 689 by dietary cholesterol, 323, 324, 327 LXR role, 325, 328, 328

1150 cholesterol 7α-hydroxylase (CYP7A1) promoter binding factor (CPF), 325–326 cholesterol 12α-hydroxylase (CYP8B1), 325, 332, 689 cholesterol 27-hydroxylase (CYP27A), 280 cholesterol ester transfer protein (CETP), 280 cholesteryl esters, 273, 274 , 278 cholestyramine, 300 cholic acid, 290, 294, 662 bacterial modification, 292 choline, 721–722 cholylsarcosine, 300 CHOP, 759, 788, 793 ChREBP see carbohydrate response element binding protein (ChREBP) chromatin, remodeling, CREB-binding protein role, 525 chromosomal instability hepatocarcinogenesis, 1112 increased telomerase knockout mice, 1111 telomere shortening, 1111, 1112 chronic liver disease (CLD), 1002 end-stage hepatic encephalopathy, 601 neuropathology, 601 , 601–602 hepatic encephalopathy associated, 599, 600–601, 600t brain organic osmolytes and cell volume, 603 hepatocellular carcinoma development, 1001–1002 infection and inflammation, 609 men vs women, telomere length and, 1109 telomere shortening and, 1108 , 1108–1109 chylomicrons, 271, 376 activation/intravascular metabolism, 275 , 275–276 apoB48, 273 assembly and secretion, 272, 273–275, 274 characteristics, 272t receptor-mediated clearance, 276 , 276–277 remnants, 276–277, 282, 376 formation, 276 , 276–277 impaired clearance, sieve function loss, 377 sizes, fenestrae size reduction effect, 393 uptake/clearance by liver, 277, 376, 393 size, 393 cilia, primary of cholangiocytes see cholangiocyte(s) roles, 662 ciliary dyskinesia, primary, 662 cingulin, 87t cip/kip family, proteins, 1018 CIRH1A disease, 668t, 671 cirrhosis, 1121 acute renal failure, 619 advanced, hepatic regeneration and stem cell role, 462 alcoholic, chemokine expression, 749 ammonia levels, 602 bile acid secretion decrease, 299, 412–413 biomarkers

INDEX HCC vs, 1003–1004 telomere shortening and, 1109 capillarization (fenestrae loss), 398 carbon tetrachloride-induced, 398, 417 cerebral blood flow (CBF) changes, 608, 608 coagulation abnormalities, 645, 646 collagen deposits excess, 1123 compensated HCV-related, 885 cryptogenic, 720 detection methods (biopsy-free), 871–872 dimethylnitrosamine-induced, 398 fibrinolysis abnormalities, 650 hemodynamic effects, endocannabinoids and, 1093–1095 hepatic encephalopathy associated, 599, 600–601, 600t hepatic vascular resistance, 704, 705 , 705–708 hyporesponse to vasodilators, 705 , 705–707 hepatitis B see hepatitis B hepatitis C see hepatitis C hepatocellular carcinoma risk, 871, 1112 hyperdynamic circulation, 708 , 708–712 endocannabinoid role, 1094 nitric oxide deficiency, 705–706 antioxidants improving, 706 in non-alcoholic steatohepatitis, 720 North American Indian childhood (NAIC), 668t, 671 outcome, in HCV infection, 884–885 pathogenesis, telomere shortening, 1108 , 1108–1109 platelet aggregation abnormalities, 643, 645 portal hypertension see portal hypertension reversible in chronic HCV infection, 443 hepatic telomere reserves and, 1109 sepsis, coagulation abnormalities in, 645 telomere shortening, 1108 , 1108–1109 thioacetamide-induced, 398 treatment DNA damage checkpoint inhibition, 1114 telomerase activation/telomere stabilization, 1114 vitamin A-coupled liposomes with siRNA–HSP47 see siRNA–HSP47 (gp46) vasodilatation, 1094 cirrhotic cardiomyopathy, 1094 cis-Golgi network (CGN), 125 citrin, 670–671 deficiency, 667t, 670–671 L-citrulline, 756, 757 citrullinemia, type II, 667t, 670–671 c-Jun, 529 clathrin, 108, 129 basolateral protein targeting, 81, 82 heavy/light chains, 109 triskelions, coated pits, 108, 109, 129 clathrin box, 109 clathrin-coated pits, 108–109 assembly and budding, 108 cell surface area occupied by, 108 formation, 109, 110 ligand transfer rate, 108 size and number, 108 structure and coat proteins, 108–109

vesicle formation, 109 clathrin-coated vesicles, 108–111, 126 formation, 109, 110, 129 accessory proteins, 109 , 110, 130 late stages and dynamin role, 110 regulation, phosphorylation role, 110, 111 at trans-Golgi network, 129, 130 half-life, 111 role/function, 108 scission, 109 , 110 regulators, 110–111 in secretory pathway, 129–130 structure, 108, 109 trans-Golgi network-derived, 129–130 uncoating/fusion, regulators, 110–111 virus uptake, 115, 116 see also endocytosis, clathrin-mediated clathrin-mediated endocytosis (CME) see endocytosis claudin(s), 204–205 HCV uptake, 217 mutations, 216, 669 in tight junctions, 204–205, 669 claudin, 1 86t, 91 connexin32 (Cx32) interactions, 206, 207 deficiency (NISCH syndrome), 216–217, 662, 667t, 669 HCV infections, 818 mutations, 216–217, 669 claudin, 2 86t, 91 claudin, 3 216 CLDN1 mutations (claudin-1), 216–217, 669 clevudine, 901 CLMP (coxsackie-adenovirus receptor-like membrane protein), 87t clusterin (CLU), hepatocellular carcinoma, 1005 c-Met, 795 as HGF receptor, 486, 558, 795 knockout mice, effects, 558 role in liver regeneration, 558 cMOAT (canalicular multispecific organic anion transporter), 665 c-myb mutant, 21 c-Myc/c-MYC miR-17–92 binding/up-regulation, cancer, 1044 role in ribosome biogenesis, 571 telomerase reverse transcriptase (TERT) activation, 1113 coagulation, 639–658, 641 cascade model, 639, 640 excessive, 649 extrinsic pathway (tissue factor activity), 640, 640 initiation, 641, 649 intrinsic pathway (contact activation), 639–640 bridge with extrinsic pathway, 640 lipopolysaccharide increasing, 777 in liver diseases, 646 liver infection and sepsis effect, 645–646 negative feedback, thrombin role, 647 platelet activation effect, 641, 642–643 regulators and regulation, 648–649 vitamin K role, 649–650 coagulation factors, 639–640 congenital disorders, 644–645 deficiencies, 644–645

INDEX acquired, 645 in acute liver injury, 646 in liver disease, 646 release, platelet activation effect, 641, 642–643 site of synthesis and inhibitors, 642t thrombin action, 647 see also individual factors beginning factor coated pits, 108–109 formation, 109, 110 see also clathrin-coated pits cocaine, 775 coenzyme A, formation, unconjugated bile acids, 296 Coffin–Lowry syndrome, 525 COL1A1 gene, 753 promoter, 745 COL1A2 gene, 743, 744 , 745, 747, 753 promoter, 744 colchicine, 30 colesevelam, 300 colestipol, 300 collagen cells-producing, 407 see also hepatic stellate cells (HSCs) hepatic half-life, 417 synthesis inhibition, fibrosis therapy, 445 platelet binding, 641, 643–644 post-translational modification, ECM stiffness, 461 scaffolds, 941 synthesis suppression by siRNA-HSP47, 1123, 1124 , 1126 type I bile duct epithelium development, 454 degradation, hepatic fibrosis resolution, 443 degradation, MMP types, 460–461 ECM synthesis in liver development, 453–454, 454t stellate cells expressing, 411, 413, 417, 747, 1122 type III degradation, hepatic fibrosis resolution, 443 ECM synthesis in liver development, 453–454, 454t stellate cells expressing, 413, 417 type IV ECM synthesis in liver development, 453–454, 454t in non-alcoholic steatohepatitis, 460 stellate cells expressing, 1122 collagenase(s) activated stellate cells, 443 increased, liver regeneration, 462 see also matrix metalloproteinases (MMPs) collagenase 3 see matrix metalloproteinases (MMPs), MMP-13 collagen-coated dextran microcarriers, 941 collagenolytic activity, hepatic fibrosis reversal, 443 colony-stimulating factor-1 (CSF1), hepatocellular metastases and, 1005–1006 coma, 597 comparative genomic hybridization (CGH), 992, 994, 1001

array see array comparative genomic hybridization (aCGH) cross-species comparisons, 993–994 Complex Disassembling Model, peroxisome protein translocation, 197 concanavalin A, 1100 confocal laser scanning microscopy (CLSM), 1062, 1063 , 1064 confocal microscopy, 1056, 1057–1058, 1062 congenital hypotransferrinemia, 238 connective tissue growth factor (CTGF), produced by stellate cells, 409 connexin26 (Cx26), 10 decrease after PH, 208, 210 gap junctions, 203, 203 calcium signal spread, 494 hepatocytes, 206, 208 as tumor suppressor, 204 connexin32 (Cx32), 10 claudin-1 interaction, 206, 207 decrease after PH, 208, 210 Dlgh1 interaction, 206 gap junctions, 203, 203 , 208 calcium signal spread, 494 hepatocytes, 206, 208 gene (Gjb1 ) structure, 204 homomeric connexons in gap junction, 203 , 204 mutations, rodent models, 210 occludin interaction, 206 phosphorylation, 211 synthesis and connexon formation, 211 connexin43 (Cx43) gap junction, 203, 203 cell types, 206 stellate cells expressing, 408 synthesis and oligomerization, 211 ZO1 interaction, 206 connexin family, 203 structures and genes, 203–204 connexons, 202, 203 , 211 homomeric and heteromeric, 203 , 204 constitutive androstane receptor (CAR), 685, 686t, 1020, 1078 COP (coat promoter), 125, 126 COPI-coated vesicles, 126–127, 127 COPI coat, 126 uncoating, 127 COPII-coated vesicles, 127–128 COPII coat components, 127–128 formation and components, 128, 129 copper, 223–233 acquisition/uptake, 223, 226 blocked by zinc, 228 apoptosis induced by, 227 Atox1 protein and, 224, 226 biliary excretion, 227 defect, 225 chelators, 228 deficiency, 223, 227, 229–230 dietary intake, 225 reduction in Wilson disease, 229 digestive system and, 225–227, 226 idiopathic toxicosis, 229 intracellular transport, 224–225, 226, 226 metabolic disorders, 228–229 see also Menkes disease; Wilson disease metabolism, 225 enterocytes, 225–227, 226 hepatocytes, 224, 226, 226

1151 renal, 227 overload/excess, 225 , 229 renal excretion, 225, 227 supplementation, 229, 230 toxicity, 227 protection, metallothionein role, 225 transport across membranes, 223–224, 225–226, 226 transporter/transport proteins, 223–225, 226 see also Ctr1 copper chaperone for SOD (CCS), 224 copper-transporting ATPases, 57, 58t, 60, 224 ATP7A, 57, 224, 226 expression site and role, 60, 224 functions, 224, 226 in kidney cells, 227 loss, Menkes disease, 225 , 226, 229 mutations, Menkes disease, 225 , 229 in non-hepatic cells, function, 227 structure, 224 ATP7B, 57, 224, 226 expression site and role, 60, 224 functions, 224, 226 gene therapy, 229 in kidney cells, 227 localization in TGN, 226 mutations in Wilson disease, 75, 225 , 226, 228 structure, 224 phosphorylation, 224 coproporphyrin, excretion, 254 cortactin, 109 , 352 , 355 canalicular cycling of ABC transporters, 354, 355 receptor-mediated endocytosis, 109 , 111 corticosteroids hepatic fibrosis therapy, 444 liver transplantation in HCV infection, 925–926 COX-2 inhibitors, 711 cholangiocellular/hepatocellular carcinomas, 787 hepatic fibrosis management, 445 coxsackie-adenovirus receptor-like membrane protein (CLMP), 87t coxsackie and adenovirus receptor (CAR), 86t, 204, 972 CpG motif, 1126 CPT1 and CPT2, 258 CPY (cholesterol 7α-hydroxylase (CYP7A1) promoter binding factor), 325–326 CRBP (cellular retinol-binding protein), 1122 CRE (cAMP response element), 522 liver genes with CREs in promoters, 522, 522t, 527, 528 repressors binding, 526 sites/location, 522 CREB-binding protein (CBP), 478, 523 , 525 isoforms, 523–524 CRE-binding protein (CREB), 522, 523 , 523–524 angiogenesis control, 529 bZip class member, 523 functions, 524, 525 ICER induction, 528 P-box see P-box PEPCK expression regulation, 527–528

1152 CRE-binding protein (CREB) (continued) phosphorylation, 522, 524 , 524–525, 526 , 528 AMPK role, 540, 540 role in hepatocytes, 527–528 role in stellate cells/cholangiocytes, 528 structure and Q1/Q2 domains, 524, 524 , 525 transcriptional activation, 524 , 524–525 CRTC2 (TORC2) binding to, 540, 540 transducers of regulated activity (TORCs), 525 CRE-binding protein (CREB) kinase, 524 CRE-binding repressors, 526 CREB-regulated transcription coactivator 2 (CRTC2), 540, 540 , 541 CREM (cAMP-responsive element modulator), 523–524 isoforms, 524, 526 in liver regeneration, 528–529, 530 phosphorylation, 525, 526 structure, 524, 524 transcriptional activation by, 525 Crigler-Najjar syndrome type I, 253, 254 hepatocyte transplantation, 577, 587 Crigler-Najjar syndrome type II, 253, 254 crisis checkpoint, 1107, 1110 cristae, mitochondrial, 138 Crumbs protein, 85t cryptogenic fibrosis, hepatic fibrosis due to, 435 crystathione-β-synthase (CBS), 536 CTGF (connective tissue growth factor), produced by stellate cells, 409 Ctr1 (copper transporter), 223, 225, 226, 226 expression in kidney, 227 structure and function, 223 trafficking, localization in plasma membrane, 223, 224 Cul3, 1020 ubiquitin ligase, Nrf2 ubiquitination, 1134, 1134 culture, hepatic, 937–941 bioreactor, 939–940 ECM effect, 463, 938, 939 Fas susceptibility, 796 hepatocyte co-cultures, 938, 939, 940, 942 hepatocyte/non-parenchymal cell interactions, 938 limitations of models, 937–938 medium and supplements, 938 micro-bioreactor, 939, 940 microtechnology tools for optimization, 940–941 sandwich (2D), 938 three-dimensional, 938–939, 942 two-dimensional, 938 type I collagen-coated dishes, 463, 941 see also spheroids curcurmin, 413 CXCL8 (IL-8), 749 CXCL9, HBV infection, 837 CXCL10, 749, 837 CXCL11, 749 CXCR3, 749 CYCD1/CDK4 kinase complexes, 571–572 cyclic ADP-ribose (cADPR), 489, 495 cyclic AMP (cAMP), 521–534 bile secretion increase, 351 cytoprotection mediated by, 795

INDEX discovery/nomenclature, 521 forskolin-induced increase, cholangiocytes, 362 glucagon-induced increase, 528 level peaks in hepatocytes, 528–529, 530 liver gene transcription control, 522, 526–528, 527t activation mechanisms, 524 , 524–525 antagonists, 526 repression mechanisms, 525–526 permeability, homomeric connexons in gap junction, 203 , 204 proliferation of cells and liver regeneration, 528–529, 529 , 530 reduced by CB1 cannabinoid receptor, 1095 regulation, 522 as second messenger, 521, 795 signaling pathway, 362, 522, 523 , 526 cascade, 362 cholangiocyte cilia, 362–363 ductal bile formation, 363 hyperproliferative cholangiocytes, 365 in liver, 526 , 526–528 see also CRE-binding protein (CREB) synthesis, 521–522 transduction pathway, 521–522, 523 cyclic AMP (cAMP)/CFTR pathway, ductal bile formation, 363 cyclic AMP (cAMP) responsive element (CRE) see CRE (cAMP response element) cyclic AMP (cAMP)-responsive element modulator see CREM (cAMP-responsive element modulator) cyclic GMP (cGMP), 486 nitric oxide signaling, in cirrhosis, 706 permeability, homomeric connexons in gap junction, 203 , 204 cyclin(s), 557 cdk complexes, 1017 , 1017–1019, 1018 up-regulation in liver regeneration, 1019 cyclin A–cdk2 complex, 1017 cyclin B–cdk1 complex, 1017 cyclin D, 1016, 1017 roles, 557, 561 cyclin D1 (CYCD1), 571, 1017 absence, hepatocyte cell cycle progression, 1020 actions/functions, 1017, 1019, 1020 cdk4 binding, 1017, 1017 , 1018 disruption, HCC prevention, 1022 expression in HCC, 1021 expression persistence during cell proliferation, 1017 induction, liver regeneration, 1019 mRNA expression, control, 1020 overexpression in cancer, 1019, 1021 reduced, impaired liver regeneration, 1020 cyclin D1–cdk4 complex, 1017, 1018 activation, hepatocyte proliferation, 1020 in cancer, 1019 p27 binding, 1018 cyclin D1–Rb-E2F pathway, 1017, 1018 abnormal control in HCC, 1021 deregulation in cancer, 1019 cyclin D2, 1020 cyclin D3, 1019, 1020 cyclin-dependent kinase(s) see entries beginning cdk

cyclin-dependent kinase-activating kinase (CAK), 1017 cyclin-dependent kinase inhibitor (CKI), 1017–1018, 1019 cyclin E, 1017 mRNA expression, liver regeneration, 1020 mutants, 1018–1019 ribosome biogenesis role, 1020 cyclopentanes, 901 cyclophilin D (CypD), 513, 785 inhibitors, 912–913 cyclophosphamide, sinusoidal obstruction syndrome, 378 cyclosporin A, 606, 785 CYP2E1, 728, 743 alcohol metabolism, 743, 744, 752–753 ROS formation, 743, 744, 752–753, 794 cells expressing, 743 hydroxyethyl radical formation, 753–754 increased expression, proteasome role, 758 CYP7A1 see cholesterol 7α-hydroxylase (CYP7A1) CYP8B1 (cholesterol 12α-hydroxylase), 325, 332 CYP27 (mitochondrial sterol 27-hydroxylase), 324 CYP450 see cytochrome P450 enzymes cystatin B (CSTB), 1002–1003 cysteine transporter, at lysosomal membrane, 177 cysteinyl-leukotrienes, hepatic vascular resistance increase, cirrhosis, 707–708 cystic fibrosis transmembrane conductance regulator (CFTR), 363, 366, 499, 528 activation in cholangiocytes, 499 CRE in promoter, 528 targeted gene replacement, homologous recombination, 975 cystic kidney disease, 662 cystinosin (cysteine transporter), at lysosomal membrane, 177 cytochalasin D, 35, 354 fenestrae regulation, 395t, 396 cytochrome c, apoptosis, 140, 784, 784 , 785 cytochrome c oxidase, 139, 224 cytochrome P450 enzymes, 685t, 750–751, 962, 1078 ABC transporter synergy, 1078 CYP2E1 see CYP2E1 down-regulation by inflammatory mediators, 775 functions and isoforms, 962 hepatocyte cultures, 938, 939, 940 phase I metabolism, 1078 cytokeratins, 4 cytokines, 407 activation, priming phase of liver regeneration, 553 alcoholic liver disease, 748–749 anti-inflammatory, 774 antiviral, 835, 836 ECM synthesis in liver development, 453, 454t factors/proteins included, 438 HBV cccDNA destruction, 814 hepatic fibrosis, 438–440 induced by bile acids, 333 inflammatory hepatic fibrosis, 438–439

INDEX macrophage infiltration of adipose tissue, 723–724 NAFLD and renal disease, 626 , 626–627 liver development, 453, 454t, 584 in liver infection, effect on coagulation, 645 produced by stellate cells, 410, 437, 437 transdifferentiation, 412, 436, 436 regulation of, priming phase of liver regeneration, 552–553, 553 , 553–557 tight junction regulation, 214, 214 see also individual cytokines cytoplasmic dynein see dynein cytoprotection from apoptosis, 795–796 from ROS see reactive oxygen species (ROS) cytosine deaminase, 966 cytoskeleton, 29 cholangiocytes, 45 epithelial cells, 35 hepatocytes, 32, 45 see also actin; microtubule(s); myosin cytosol, as excitable medium, concept, 494 cytotoxic T lymphocytes (CTLs) viral infections, 835, 837 see also CD8 T cells

D DAG see diacylglycerol (DAG) Danon disease, 177–178 D-bifunctional protein deficiency, 193 Dcytb, 242 D-dimers, 650 death effector domain (DED), 783, 784 death interacting signaling complex (DISC), 783, 784 death receptors (DR), 783, 790–793 Fas see Fas (CD95) see also apoptosis DEBIO-025, 912–913 deconvolution, 1056–1057 deep vein thrombosis (DVT), 649 degradation pathways autophagy see autophagy/autophagic pathways ubiquitin-proteasome see ubiquitin-proteasome system (UPS) delta-bilirubin, 253 delta virus see hepatitis D virus (HDV) dendritic cells, 375 activation, by IFNα, 905 functions, 840 HBV persistence, 840 HCV persistence, 847–848 T cell priming, role, 847–848 dense fibrillar components (DFCs), nucleolus, 569 density gradient centrifugation, sinusoidal endothelial cells, 381 7-deoxy bile acids, 292 deoxycholic acid (DCA), 292, 294 biliary obstruction effect, 298 CYP7A1 regulation, 327 FXR activation, 325 rehydroxylation, 295 deoxyguanosine analogs, HBV infection, 901 dephosphorylation, transcriptional repression by, 525–526

desferal, 512, 515, 516 desmin, 10 detergent-insoluble glycolipid (DIG)-enriched domains, 113 detergent resistance, canalicular membranes, 343, 345 detoxification, 1131 animals, Nrf2, Keap1 and GSTs, 1133–1134, 1134 pathways in animals vs plants, 1134 phase I and II imbalance, in cultures, 938, 939, 940 phase I enzymes, 1076, 1077 , 1078, 1080, 1131 see also cytochrome P450 enzymes phase II enzymes, 1078, 1131 activation, 1131, 1132, 1133 plants, TGA factors and, 1134 , 1135–1136 see also glutathione S-transferase (GST); Nrf2 phase III (CYP enzyme and ABC transporter synergy), 1078, 1131 dexamethasone, 611 diabetes mellitus hepatic fibrosis due to, 434, 435 SREBPs and ChREBP increased, 629 telomere shortening, 1109 diabetes mellitus, type 2 (T2DM), 471 fasting hyperglycemia, 477–478 gluconeogenesis, 477–478 insulin resistance see insulin resistance insulin sensitizing agents, 478–480 metformin non-responders, 544 muscle mitochondrial impairment and lipid accumulation, 472–473 NAFLD prevalence, 624 diacylglycerol (DAG), 262, 726 accumulation, protein kinase C-θ, 473 biosynthesis, 258 formation, calcium signaling, 486, 487 vesicle biogenesis regulation, 131 diacylglycerol O-acyltransferase (DGAT), 258, 259, 539 DGAT1 and DGAT2 expression, 262 triglyceride synthesis, 258, 259, 273 therapeutic target in NAFLD, 732 dibutyltin dichloride (DBTC) model, 1127 Dicer, 1030 , 1032, 1034 knockouts/knockdown, 1034 diclofenac, 777 dielectrophoresis-mediated cell patterning, 943–944 diet alcoholic liver disease and, 744 cholesterol intake/levels see cholesterol NAFLD management, 730 protein restriction, 610 triglyceride source, 273 diffraction, 1055–1056, 1056 digoxin, 58 dilute mouse, 34, 36 dimercaptopropanol (British anti-lewisite), 228 dimethylnitrosamine (DMN), siRNA–gp46 (HSP47), 1123–1124, 1125, 1125 dimethyl sulfoxide (DMSO), 210, 213 diphtheria toxin, 64 dipyridyl, 513 disaccharides, non-absorbable, 610 DISC (death interacting signaling complex), 783, 784

1153 discoidin domain receptors (DDRs), 437 Discs Large protein, 84, 85t Cx32 interaction, 206 Disse, space of, 7, 9, 375, 375 , 408, 408 blood flow in, sinusoidal obstruction syndrome, 379 chylomicron remnants sequestration, 277, 376, 393 drugs entering and clearance, 376 endothelial passage of proteins/particles, 393 extracellular membrane, 454t, 455 retrograde plasma flow, 394 sinusoidal lumen interactions via fenestrae, 394 stellate cells in see hepatic stellate cells (HSCs) uptake of chylomicron/VLDL remnants, 277 disseminated intravascular coagulation (DIC), 642 disulfide bonds, protein folding, 162 divalent metal-ion transporter 1 (DMT1), 236, 239, 242, 512 DKTGTLT sequence, 60 DMT1, 236, 239, 242, 512 DNA copy number changes in cancer, 992, 994 damage checkpoint, 1016 activation in telomere shortening, 1107, 1112 inhibition, cirrhosis treatment, 1114 loss, tumor formation and, 1111, 1112 damage pathway, 1016 activation, telomerase knockout mice, 1111 impaired organ maintenance in telomere dysfunction, 1107–1108 methylation, 759 hepatocellular carcinoma, 1004 mitochondrial see mitochondrial DNA (mtDNA) repair, chimeric RNA–DNA oligonucleotides, 977, 978, 978 , 979 mechanisms, 978, 979 , 981 repair pathways, 975, 978, 979, 1016 replication, liver regeneration, 550, 551–552, 558, 559 site-specific modification, 975–976 see also gene therapy triplex, 975–976 DNA–DNA duplex, chimeraplasty, 978 DNA ribonucleases, 977 DNA vaccination, 967 docosahexaenoic acid (DHA), 282–283 L-DOPA, 607 dopamine-β-hydroxylase, deficiency, Menkes disease, 229 doxorubicin, 1079 DPPIV mutants (F344 rats), 584, 586 Drosha, 1030 , 1032 Drosophila melanogaster miRNA translational repression, 1032 Sir2 orthologs, 957 DRP1/Drp1, 785–786 drug-induced inhibition, 143 mitochondria fission induced by, 142, 785–786 overexpression, mitochondrial fission, 143

1154 drug(s) clearance by liver, sinusoidal endothelial cells, 376 dose-related hepatotoxicity (type A response), 775–776 idiosyncratic hepatotoxicity (type B response), 775, 776 animal models, 776–777 mechanisms, 777–778, 778 see also inflammatory stress inflammation interaction, 774–776, 775 peliosis hepatis due to, 380 sinusoidal obstruction syndrome due to, 378 transport into bile, 339–341 see also therapies for individual conditions drug abuse/illicit use, HBV/HCV infections, 805 Dubin-Johnson syndrome (DJS), 254, 664t, 665–666, 691 duck hepatitis B virus (DHBV), 810, 813, 815, 863 ductal plate, 4 duct-trip mutant, 1071 ductular reaction, 587 dynactin complex, 33, 48 , 48–49, 50 dynamin, 51 clathrin-coated vesicle scission, 110 structure, 110 trans-Golgi network (TGN)-derived vesicle scission, 130–131 dynamin 2 (Dyn2) caveolae liberation from cell surface, 113 clathrin-coated vesicle fission, 109 , 110, 111 , 113 clathrin-mediated endocytosis, 110 trans-Golgi network (TGN)-derived vesicle scission, 131 dynamin-like protein 1 (DLP1), 194 dynamitin, 48 , 49 dynein, 45 axonemal, 32 cytoplasmic see dynein, cytoplasmic discovery, 32 flagella and ciliary, 32 mitochondrial distribution, role, 141 nuclear envelope breakdown, 152 dynein, cytoplasmic, 32, 37t, 45, 48–49 dynein 1 (DYNC1), 32, 45, 48–49 properties, 47t dynein 2 (DYNC2), 32, 45, 48, 49 cholangiocyte primary cilium, 52 as “–” end motor protein, 33, 48, 49 in epithelial cells, 37–38 function, 46 mechanism of action, 33, 48, 48 structure, 33, 48 dysfibrinogenemia, 643, 647–648 dyskeratosis congenita (DKC), 1109 dyskerin, 1109 dyslipidemia, drugs treating, 281–283

E E2F1, cMyc activating, miR-17–92 effect, 1044 E2F transcription family, Rb binding, cell cycle control, 1017, 1017 see also cyclin D1–Rb-E2F pathway E3Karp, 354 EBP50 protein, 89t, 354

INDEX 4EBPI (eIF4E-binding protein, 1) 542 4EBPs (e-IF4E binding proteins), 569 E-cadherin, 84, 85t ECM see extracellular matrix (ECM) EDEM, 166 efferent blood vessels, 3, 7 EH domains (Eps homology domains), 110 eicosapentaenoic acid (EPA), 282–283 electron microscopy (EM), 1056, 1058 transmission EM, 1055, 1058 electron multiplying charge-coupled devices (EMCCDs), 1062 E ligases, 175, 180 elutriation, sinusoidal endothelial cell isolation, 381 embryoid bodies, 937 embryology of liver, 4–5, 5 , 17–25, 583–584, 584 cadherins, TGF-β and integrins cross-talk, 92 capsule development, 21 cell–ECM adhesion role, 92 cholangiocyte differentiation, 4, 20 control of liver regeneration, 21–22 ECM synthesis/remodeling, 453–455, 454t by embryonic days (ED8–, 17) 584, 584 endoderm, acquisition of hepatic competence, 4, 17–18, 18 in fish, birds vs mammals, 5 future developments, 22–23 growth defects, causes, 21 hepatic endoderm development, 18–19, 584 liver bud formation from, 4, 19–20, 584 signals, 19, 92 zebrafish use, 1068, 1069 hepatocyte differentiation see hepatocyte(s), differentiation miR-122 role, 1035 profiling in hepatoblast differentiation, 1039 , 1040 , 1040–1041, 1045 miRNA role, 1035, 1040, 1040 , 1045 molecular basis, 18, 18 , 19, 20, 21 , 22 mouse, 19, 20, 20 stellate cell origin, 411 transcription factors involved, 18, 18 vascular development, 4–5, 21 zebrafish model, 1069 zebrafish use see zebrafish see also stem cells embryonic stem cells (ES), 582, 582 , 586, 589 culture, hepatocyte-like cell generation, 586 liver repopulation by, 585–586, 937 EMK (MARK, Par1), 89t, 93 emtricitabine, 901, 902t, 903t hepatitis B, 901, 903 encephalopathy, hepatic see hepatic encephalopathy (HE) endocannabinoids, 444, 1091–1100, 1092 , 1093 activation, in fibrogenesis, 730, 1096 autoimmune hepatitis, 1100 biosynthesis, 1093 hemodynamic effects of cirrhosis and, 1093–1095 hepatic encephalopathy, 1100 hepatic fibrosis, 439–440, 730, 1095–1096

hepatic ischemic-reperfusion injury, 1100 hepatic steatosis, 1096–1099 alcoholic, 1098–1099 non-alcoholic, 1096 , 1096–1098 macrophages generating, 1093–1094 receptors see cannabinoid (CB) receptors vasodilatation in portal hypertension, 709 endocannabinoid system (ECS), 1093 endocytic vesicles see endosomes endocytosis, 36, 107–123 absorptive, 178 , 178–179 ASGPR-mediated, gene therapy, 974 caveolae-mediated, 112, 113–114 clathrin-independent, 112, 113–114 see also caveolae clathrin-mediated, 108–111 accessory proteins, 109 , 110 canalicular cycling of ABC transporters, 354–355 HBV uptake, 115–116 HCV uptake, 116, 818, 909–910 regulators, 110–111 see also clathrin-coated vesicles fluid-phase (pinocytosis), 178, 178 hepatocyte basolateral surface, 75 in hepatocytes, 35, 38, 50 , 50–52, 107 basolateral surface, fluid-phase endocytosis, 75 transferrin receptor trafficking, 114–115 rates/capability, 178, 179 receptor-mediated (RME), 107–108, 178 , 179 gene therapy delivery, 973, 974 iron uptake, 512–513 LDL particles, 277 sinusoidal endothelial cells, 376 regulation of clathrin-mediated endocytosis, 110–111 by lipids and phosphoinositides, 114 by Rab proteins, 114, 115 selectivity and efficiency of types, 178, 179 types, 178 , 178–179 vesicle formation, 107–108, 108 see also clathrin-coated vesicles virus uptake, 107, 108 HBV and HCV see above V-type ATPase role, 63–64, 64 endoderm, 17 anterior-ventral definitive, 4, 17–18, 18 acquisition of hepatic competence, 17–18, 18 hepatic development, 4, 18–19 liver bud formation, 4, 19–20 see also embryology of liver endophilin B, clathrin-coated vesicle uncoating, 111 endoplasmic reticulum (ER), 125, 157 abundance in hepatocytes, 167 apoB-containing lipoprotein assembly, 273 -associated degradation (ERAD) 165 , 166, 728–729 calcium release, 141 GPI anchor addition to proteins, 164 imaging techniques, S/N ratio, 1056 lumen chaperone systems, 162–163

INDEX N-glycosylation of proteins, 162 oxidizing environment, 162 protein assembly into complexes, 164–165 protein folding/maturation, 161, 162, 163 , 164–165 protein modifications, 163–164 see also chaperone(s); protein(s) membrane, signal recognition particle receptor, 159 mitochondria-associated (MAM), 141 mitochondria association in hepatocytes, 141, 490 , 491 mRNA, degradation during ER stress, 167–168 peroxisome association/biogenesis, 194–195, 195 physiological regulation of, 167–168 procollagen maturation and HSP47 role, 1122, 1122 protein degradation, marking for, 166, 167, 758–759 protein maturation, 161, 162, 163 , 164–165 protein processing capacity, 167 protein quality control, 165 , 165–167, 174 protein transport from Golgi apparatus, 126 protein transport to Golgi apparatus, 125 COPII-coated vesicles, 127–128 reactive oxygen species formation, 750–751 retrotranslocation of misfolded proteins, 165 , 166 “rough”, 157 secretory/membrane protein segregation, 157–161, 158 co-translational targeting to ER, 158–159 post-translational translocation/insertion, 161 recognition by signal sequences, 158, 159 translocation into ER by translocons, 159–161, 160 SREBPs and SCAP–SREBP complex, 627, 628 stress, 630 , 748 apoptosis of cells, 759, 788, 789 , 793 cell injury mechanisms, 789 cholestasis, 793 insulin sensitivity reduced, 727, 727 insults triggering, 788 mechanisms, 759 mRNA degradation during, 167–168 prevention, by ursodeoxycholic acid, 796 SREBP activation, 627, 630 steatosis progression to steatohepatitis, 728–729, 729 unfolded protein response see unfolded protein response (UPR) transitional (tER), 127 endoplasmic reticulum (ER)-associated degradation (ERAD), 165 , 166, 728–729 endoplasmic reticulum degradation-enhancing α-mannosidase-like protein (EDEM), 729 endoreduplication, hepatocytes, 552 endosomes, 179

acidic pH and pathogen interaction, 64 chelatable iron release, 513–515 early (EE) acidification, 63–64 acidification regulation, 66 cargo protein translocation to, 107, 179 formation, 107–108 ligand release, 64 vs late, motor protein function, 51 late (LE), 51, 107 maturation to lysosomes, 179 V-type ATPase action, 63–64, 64 endosymbiotic theory, 137, 139 endothelial cells, 6–7, 373 adhesion molecules, 748 antigen presentation role, 10 beta-cell development/function and, 19 bone marrow, 374 brain capillaries, in hepatic encephalopathy, 602 development, zebrafish model, 1069 fenestrated, 374, 374t diaphragmed, 374, 374t open, 374, 374t see also sinusoidal endothelial cell (SEC) liver embryogenesis, 4, 5, 5 , 21 liver bud development, 4, 19 proliferation, 12 sinusoidal see sinusoidal endothelial cell (SEC) splenic, 374 structure, 9–10 endothelin, 645 hepatic vascular resistance, 707 increase in cirrhosis, 645 receptors, 645 ET-A and ET-B, 439 ET-A blockers, 394, 439, 445, 707 ET-B blockers, 439, 707 endothelin-1 (ET-1), 439, 645 fenestrae regulation, 397 hepatic fibrosis pathogenesis, 439 receptor blockade, in therapy, 445 endothelium, 373, 374 continuous and discontinuous types, 374, 374t see also endothelial cells endotoxemia, 645 endotoxin see lipopolysaccharide (LPS) energy AMPK as checkpoint/sensor, 535, 539 balance, endocannabinoids role, 1096–1098 excess, adipose tissue response, 722–724, 723 harvest/storage, gut microbiota role, 721 , 721–722 homeostasis in liver, 724 , 724–726, 725 metabolism, hepatic encephalopathy pathogenesis, 604–605, 605 production by mitochondria, 138–139 for protein degradation, 173 Engelbreth–Holm–Swarm (EHS) gel, 463 enhancesome, 570 entecavir (Barraclude), 866, 902t, 903t, 904 enterocytes apoB transcription, 273, 273 , 274 canalicular transporters, 339–341, 340 cholesterol absorption, 273, 274 copper metabolism, 225–227, 226 in Menkes disease, 225 , 226

1155 enterohepatic circulation, bile salts, 280, 289, 293–296, 294 , 329, 329 , 341 bile acid damage and repair, 295 disorders/disturbances affecting, 298, 299 frequency of recycling, 294 hepatic uptake see bile acid(s), hepatic uptake intestinal conservation of bile acids, 280, 281, 294–295, 297 regulation, 329–330 regulation, 297, 329–330 enzymes synthesis as prozymogens, 174 see also specific enzymes/metabolic pathways EPACs (exchange proteins activated by cAMP), 362 EpCAM, stem-cell marker, 1006 epidermal growth factor (EGF) incorporation into scaffolds, 942–943 progression phase of liver regeneration, 559 epidermal growth factor receptor (EGFR), 790 hepatocellular carcinoma, 993 ligands, role in liver regeneration, 559 ubiquitinylation and degradation, 112 epigenetic signatures/changes alcoholic liver disease (ALD), 759 hepatocellular carcinoma (HCC), 1004 episomal DNA, gene therapy, 974 epithelial cells biliary see cholangiocyte(s) cytoskeleton, 35 intra-cellular trafficking, 35 kinesins and cytoplasmic dynein, 37–38 microtubule and actin filament organization, 32 myosin functions, 35–37 polarity see polarity see also hepatocyte(s) epithelial cell systems, plasma membrane protein research, 76 epithelial mesenchymal transition (EMT), 92 in hepatic fibrosis, 438 Snail role, and polarity loss, 214 Eps15 (EGFR protein substrate 15)/EPS15, 109 , 110 clathrin-coated pits, 110, 352 , 355 epsin, 110 epsinR, 131 ERGIC (endoplasmic reticulum/Golgi intermediate compartment), 125 ERK pathway c-met role, liver regeneration, 558 hepatic fibrosis, 441, 441 ERM proteins, 89t Escherichia coli , DNA repair pathways, 975, 978, 979 ESCRT (endosomal sorting complex required for transport), 112 Ese1 (endocytosis regulator), 110 E-selectin, 748 ether lipids (ether phospholipids), biogenesis in peroxisomes, 194 6-ethylCDCA (FXR agonist), 299 euchromatin, protection from repressing factors, NPC role, 151 evolution of liver, 4 excitatory amino acid transporter (EAAT)-2, 606–607

1156 exercise AMPK activation, 537 effect on insulin resistance, 479–480 NAFLD management, 730 exocytic secretion, 128–129 exocytosis, 35, 497 Ca2+ signaling role, 497–498 hepatocytes, 35, 128–129 multivesicular, 36 exonuclease-1 (Exo1), 1107–1108 Extended Shuttle hypothesis, protein targeting into peroxisomes, 196 extracellular matrix (ECM), 407, 453–467, 454t activated stellate cell interaction, 437, 437 , 458 in cholangiocellular carcinoma, 459, 463–464 components, 455 newly described, 458–460 degradation, 417–418, 460 see also matrix metalloproteinases (MMPs) deposition, 458 excessive in fibrosis, 433, 458, 462 healing role, 458 see also hepatic fibrosis elasticity of liver, 455, 455 functions, 407, 453, 455 as growth factor reservoir, stellate cell activation, 437 in hepatocellular carcinoma, 459, 463–464 hepatocyte adhesion, liver differentiation, 92 impact on hepatic cultures, 463, 938, 939 liver development, 92, 453–455, 454t liver fibrosis, 454t, 458, 462 excessive deposition, 433, 458, 462 stiffness preceding, 461 liver inflammation, 454t, 455–458, 457 low-density (space of Disse), 455 maturation modification, 461–462 normal liver, 454t, 455, 455 , 456 perisinusoidal, 455 portal area, 454t, 455 “provisional matrix”, 455–456 remodeling, 460–461 cells involved, 462 enzymes, and sources, 454, 461, 464 liver development, 453–455, 454t liver fibrosis, 458 liver regeneration, 462–463 response to short-term liver damage, 455–456, 460 see also matrix metalloproteinases (MMPs); tissue inhibitor of metalloproteinase I (TIMP-I) signaling, in liver fibrosis, 458 softness and resistance of liver, 455, 455 space of Disse, 454t, 455 stellate cells transdifferentiation, 414, 436, 437, 458 stiffness, 461 ECM maturation and, 461–462 in fibrosis, 458 regulation, 461–462 synthesis during liver development, 453–455, 454t in liver fibrosis, 458

INDEX extracellular-regulated kinase (ERK) see ERK pathway extracorporeal bioartificial liver devices (BAL), 935, 936, 937 extracorporeal blood depuration methods, 611 extrahepatic biliary atresia (EHBA), 661–662 ezetimibe, 281, 282

F F2-isoprostanes, 755 F344 rats, 584, 586 F-actin see under actin factor III (TF) see tissue factor (TF) factor V (FV), 640 , 647 factor Va (FVa), 640 , 643 factor V Leiden, 649 factor VII (FVII), 640, 640 deficiency, 644 factor VIII (FVIII), 640 , 647 deficiency, 644 elevated in liver disease, 646 gene therapy delivery, 974, 975 factor VIIIa (FVIIIa), 640, 640 , 643 factor IX (FIX), 639, 640, 640 , 643 in acute liver injury, 646 deficiency, 644 site-directed nucleotide conversion, 980 targeted gene therapy, chimeric oligonucleotides, 980 factor X (FX), 640, 640 decreased activity, 645 factor Xa (FXa), 640, 640 , 641, 643, 647 in acute liver injury, 646 inhibition, TFPI role, 648 factor XI (FXI), 639, 640, 640 , 647 in acute liver injury, 646 deficiency, 644 factor XIa (FXIa), 640 , 643 factor XII (plasma protease factor), 639 factor XIII (FXIII), 640 activation by thrombin, 647 deficiency, 645 Fah null mouse, 579–580, 580 , 582 FAK PTEN action, and miR-21 effect, 1044 see also focal adhesion kinase (FAK) pathway falciform ligament, 6 familial hypercholanemia, 667t, 669–670 familial intrahepatic cholestasis, 664 farnesoid X receptor see FXR S-farnesylthiosalicylic acid, 444 Fas (CD95), 783, 789–790 apoptosis mediated by, 442, 789–790 cholestasis, 793 c-Met binding, anti-apoptosis, 795 Fas-activated death domain protein (FADD), 749–750 Fas-associated protein with dead domain (FADD), 442, 789–790 FAS KO mice (FASKOL), 264 fasting lipid metabolism, 258 AMPK regulatory role, 537, 542 ribosome biogenesis reduced, 567 fasting-induced adipose factor (angiopoietin-like protein, 4) 721, 721 fat, visceral, 723, 723 fat-inducting transcripts (FITs), 259 fatless mouse, 476

FATP5, 296 fats see lipid(s) “fat-storing cells” see hepatic stellate cells (HSCs) fatty acid(s) beta-oxidation see beta oxidation biosynthesis control by SREBPs, 627, 725 increased rate in alcoholic liver disease, 745 oxidation balance, 263–264, 725, 725 branched-chain, oxidation in peroxisomes, 191, 192 circulating, hepatic insulin resistance, 476–477 de novo synthesis, 264 free, sources, for liver, 723, 723 hepatic levels in alcoholic liver disease, 744 hepatic metabolism, 257–270 adipose fat storage, 261–263, 262 fate of “old” (stored) vs “new” (dietary-derived) fat, 264 , 264–265 insulin resistance and, 261–263, 262 lipogenesis and oxidation balance, 263–264, 725, 725 PPArγ role/functions, 263 triglycerides see triglyceride(s) insulin resistance mechanism, 262, 262 , 726, 726 long-chain, 273 oxidation, 472, 725 , 725–726 adiponectin role, 725–726 biosynthesis balance, 263–264, 725, 725 increased, steatohepatitis development, 728 in peroxisomes, 191, 725, 725 see also beta oxidation peroxidation see lipid(s), peroxidation storage derived, 264 very-long-chain, beta oxidation, 193 see also lipid(s); specific fatty acids fatty acid aminohydrolase (FAAH), 1093, 1097 fatty acid binding protein, 5 732 fatty acid ethyl ester (FAEE), 743 fatty acid synthase (FAS), 257, 627 AMPK decreasing transcription of, 541 CB1 receptor activation, 1096 , 1096–1097 fatty acid transporter, FATP5, 296, 725, 725 fatty liver, fibrous scar replacement, 748, 749 fatty liver disease alcoholic see alcoholic fatty liver non-alcoholic see non-alcoholic fatty liver disease (NAFLD) pathogenesis, 748 fat accumulation, molecular events, 721–727 see also hepatic steatosis fenestrae/fenestrations, endothelial cells, 374, 374t cell culture and stiffness of ECM, 462 definition, 390–391 historical background, 389–390 fenestrae/fenestrations, sinusoidal endothelial cells, 6, 9, 374 aging effect, 398 caveolin-1 in, 392–393, 397, 398

INDEX classification by size (pits, pores, gaps), 391–392 damage affecting, 392, 395, 397–399 defenestration/loss, 398 capillarization, 377, 398 cirrhosis, 398 effect, 393 sinusoidal obstruction syndrome, 379, 398–399 detection methods, 392 diameter, 9, 374, 391–392, 392t, 393 affecting chylomicron remnant clearance, 376, 393 agents reducing, 396–397 reduced, effect on sieving coefficient, 393–394, 394 reduced, vascular resistance increase, 394 dimensions, 9, 374, 391–392, 393 dumbbell-shaped, 392 formation by membrane/pore fusion, 396 gaps and gap formation, 392, 397, 398 increased, 396 steatosis due to, 393 morphology/ultrastructure, 390 , 390–393, 392 number, 391 regulation, 395t, 396 pathophysiology, 397–399 reduction (pseudocapillarization), 377, 398 regulation, 394–397, 395t, 396 actomyosin, 395–396 calcium–calmodulin, 395–397 VEGF, 395 roles, 393–394 chylomicron remnant passage, 376, 393 hepatic lymph formation, 394 immune role of liver, 376–377, 394 sieve plates, 9, 374, 374 , 375 , 392 species with, 392t Fenton reaction, 511, 512 , 754 ferric iron conversion to ferrous iron, 512, 512 , 513, 754 release from lysosomes, 513, 515, 517 ferritin, 236, 513 functions, 236 structure, L and H chains, 236, 236 ferroportin (FPN), 239 degradation, control, 241, 242 hepcidin binding, 239–240, 241 as iron exporter, 239, 241–242 mutations, ferroportin disease, 244 SLC40A1 mutations, 244 ferroportin disease, 244 ferrous iron conversion from ferric iron, 512, 512 , 513, 754 hydroxyl radical formation, 515 fetal liver, 4–5, 5 development see embryology of liver epithelial cells, culture, 587 growth defects, 21 hepatic stem cells, 583–584, 587 liver repopulation by, 584–585 see also embryology of liver α-fetoprotein (AFP), hepatocellular carcinoma, 871, 999 fetoprotein transcription factor (FTF) see liver receptor homolog-1 (LRH-1; FTF)

FGFs see fibroblast growth factors (FGFs) fibrates, effect on cholesterol, 282 fibrillar centers (FCs), nucleolus, 569 fibrillin-1, 459 fibrin, 641 formation, 640, 647 role in drug–inflammatory stress, 777 fibrin degradation products (FDP), 641, 643 fibrinogen (FBG), 640, 647 abnormalities in liver disease, 647–648 actions, 647 deficiency, 645 ECM in liver inflammation, 455 fibrin formation, 640, 647 platelet linking, 641, 643 fibrinogenesis, accelerated, chemicals causing, 649 fibrinogen-like protein (FGL), ECM during liver inflammation, 455–456 fibrinolysis, 641, 650 abnormalities in liver disease, 650 inhibition, 641 liver role in, 650–651 oxidative stress effect, 646 fibrinopeptides (FpA and FpB), 640, 647 fibroblast growth factors (FGFs), 410 FGF3 expression, HCC, 1004–1005 FGF15 bile acid synthesis control, 297, 332, 689 CYP71A regulation, 297, 332 FGF19, bile acid synthesis control, 332, 689 FGF21, ketogenesis, 260 hepatic fibrosis, 439 liver development, 19, 584, 584 endoderm development, 19 zebrafish, 1068 produced by stellate cells, 409 receptors, 19 fibrocytes, 411 fibrogenesis, 417–418, 437, 437 , 1122, 1122 activation, non-alcoholic steatohepatitis, 730 inhibition, NAFLD therapy, 730 see also hepatic fibrosis fibrolamellar carcinoma, 1078 fibrolysis, 417–418 fibronectin (FN) bile duct epithelium development, 454 ECM in liver inflammation, 455 fetal isoform (EIIIAFn), 437 normal liver ECM, 455 fibrosis cardiac, 435 liver see hepatic fibrosis non-liver, vitamin A-coupled liposomes for therapy, 1127 fibulins (fibulin-1 and fibulin-2), 459 Fick’s law of diffusion, 375 filamin, 114 Fis1, 142 Fisher rat thyroid cell line see FRT (Fisher rat thyroid) cell line fish oil, 282–283 Fission1 (Fis1), 194 FlAsH/ReAsH peptides, 1054–1055 flavonoids, 756 flippase, 342 , 345, 693 ATP8B1, 343, 345 floppase, 341, 342 , 345

1157 Abcb4, 341 Abcg5/g8, 342 fluorescence correlation spectroscopy (FCS), 1057 fluorescence detection systems, 1053–1055 Ca2+ signal detection, 491 fluorescence loss in photobleaching (FLIP), 1063 fluorescence microscopy diffraction, 1056, 1056 wide-field, 1056 fluorescence photoactivation localization microscopy (fPALM), 1060, 1060 fluorescence recovery after photobleaching (FRAP), 1057, 1064 fluorescence resonance energy transfer (FRET), 1062 fluorescent dyes, advances, 1053–1055 alternatives to, 1054 fluorescent lipid reporter, PED-6, 1072 fluorophores, 1053–1054 foam cells, formation, 377 foamy viruses (FVs), gene therapy vectors, 969 focal adhesion kinase (FAK) pathway hepatic fibrosis, 440 integrins activating, cytoprotection, 796 PTEN action, and miR-21 effect, 1044 follistatin, 559 foregut, ventral, endoderm domain, 17–18 hepatic endoderm development, 4, 18–19 foreign antigens, removal mechanism, 10 forkhead transcription factors, 560, 727 see also entries beginning Fox forskolin, 362 forward phase protein arrays (FPPAs), 993 four-phosphate-adaptor protein (FAPP), 131 FoxA2 transcription factor, 18, 18 FOXD3, 560 FOXI1, 560 FoxM1B transcription factor, 1022 FoxO, 727 Sirt1 deacetylation of, 957, 958 FoxO1/FOXO1, 260, 261, 731 deacetylation, Sir2-mediated, 957, 958 FoxO3A, 728, 788 free radicals formation, alcohol associated, 743, 747 , 751 see also reactive oxygen species (ROS) free-radical theory, of aging, 143 Frizzled (Fz), hepatic fibrosis, 414, 415 , 416t, 442 FRT (Fisher rat thyroid) cell line, plasma membrane protein/traffic research, 76, 77t, 79 basolateral protein sorting signal, 81 FTF (fetoprotein transcription factor) see liver receptor homolog-1 (LRH-1; FTF) F-type ATPases see ATP synthase fumarylacetoacetate hydrolase (Fah), 579 deficiency, gene therapy, 974, 975 functions of liver, 3, 4, 11, 17, 935–936 FV to FXIII see specific coagulation factors (under Factor) FXR (farnesoid X receptor), 323, 326, 559–560, 684 activation, 325, 632 dietary cholesterol effect, 325, 327, 330 regulation by bile acid flux, 332, 632

1158 FXR (farnesoid X receptor) (continued) agonists, 299 antifibrogenic effects, 413 ASBT downregulation by, 329 bile acid biosynthesis regulation, 297, 299, 325, 633 as bile acid sensor, 325, 332 bile acid transporter regulation, 325, 329, 684 in cholestasis, 684–685 BSEP regulation, 692 cells expressing, 633, 633 CYP7A1 regulation, 297, 325, 326, 327 mechanism, FXR–SHP–FTF cascade, 297, 328 , 328–329 dietary cholesterol effect on, 325, 327, 329, 330 expression and actions, 325, 633 FGF15/19 interactions, 332 FXR–PPAR–ASBT cascade, 330 ligands, 325, 327 functions in cholestasis and, 684–685, 686t therapeutic effects, 633 liver regeneration and, 559–560 NFκB inhibition by, in NAFLD, 633, 633 Ntcp repression, 687 overriding effect of LXRα activation, 328 pathogenic role in NAFLD and renal disease, 632–633, 633 reduced in cholestasis, 685 SHP as target, 297, 325–326 SREBP-1 and ChREBP inhibition by, 633, 633 target genes, 325 FXYD family of proteins, 58, 60

G GABA, 607 hepatic encephalopathy pathogenesis, 607, 607 GABA-A receptor, 607 GABA-A receptor complex, 607 benzodiazepine site (BZ) site, 607, 607 , 609 partial inverse agonists, 611 neurosteroid modulatory site (NS site), 607, 607 , 609 G-actin, 30, 31 GAK, clathrin-coated vesicle uncoating, 111 galectin, 3 81 gall bladder bile acid storage, 293 embryology, 20 gallstone, dissolution, chenodeoxycholic acid, 299 gamma-aminobutyric acid see GABA Gammagard, 881 gangliosides, 75 GD3, 790 gap junctions, 10, 201–220 arrays and plaques, 202 Ca2+ and IP3 diffusion, 208, 209 , 211, 494 changes in/after hepatocyte injury, 210 channels, 203 , 204 opening and closures, 209 regulation of opening/closing, 210–212

INDEX subunits, 202, 203 components and genes, 203 , 203–204 connexins, 203 , 203–204, 206 connexin synthesis/oligomerization, 211 see also specific connexins formation, 211–212 functions in liver, 201, 202, 206–210, 208 electrical coupling of hepatocytes, 206–207, 211 growth control, 208, 210 signal transfer, 206–208, 209 hemi-channels (connexons), 202, 203 , 211 homomeric and heteromeric, 203 , 204 in liver regeneration, 208, 210 protein phosphorylation, 211 protein synthesis and folding, 211 reduced in hepatomas, 208 regulation, 210–212 between stellate cells, 408 structure, 202, 202 tight junctions interactions, 206, 207 ultrastructure, 202, 203 gaps (large fenestrations), sinusoidal endothelial cells, 392, 395, 397, 398 gastrointestinal flora see gut microbiota GATA4 transcription factor, 18, 18 GBF1, 126 gelatinase A see matrix metalloproteinases (MMPs), MMP-2 gelatinase B, 460 see also matrix metalloproteinases (MMPs), MMP-9 gelsolin apoptosis inhibition, 456, 458 as “defence protein”, 456 ECM during liver inflammation, 456 gender chronic hepatitis C outcome, 885 chronic liver disease and telomere length, 1109 gene expression profiling chronic liver disease, 1002 endoderm development, 18, 18 HCC see hepatocellular carcinoma (HCC) hepatitis B, 1002–1003 hepatitis C, 1002–1003 multidrug resistance (MDR) (cancers), 1081–1082 techniques, 992–993, 1000–1001 Gene Gating hypothesis, 151 gene modification, targeted see gene therapy gene redundancy, 22 gene repair see chimeraplasty genes, liver, cAMP action/regulation, 522t, 526–528, 527t gene therapy, 965–982 for acquired disorders, 966t chemo-/radiotherapy with, 966–967 ex vivo, 967 fenestrae role, 394 gene transfer mechanisms, 967–973 vectors see below goals, 965 indications, 965–967, 966t for inherited disorders, 966t, 967 in vivo, 967 for neoplastic disease, 966–967, 966t non-viral vectors, 968t, 973–974

lipid-based delivery systems, 973, 974 naked plasmid vectors, 974 polyplex and lipopolyplex delivery systems, 973 receptor-/liposome-mediated delivery, 973–974 Sleeping Beauty transposition system, 974 preparative irradiation, 967 targeted gene modification, 975–982 antisense oligonucleotides, 976–977 DNA ribonucleases, 977 homologous recombination, 975, 978 ribozymes, 976 RNAi and miRNA, 977 single nucleotide modification, 977–982 see also chimeraplasty single-stranded oligonucleotides, 981–982 triplex DNA, 975–976 targeting to liver, 967 targeting to tumor cells, 966 for viral infections, 967 viral vectors (recombinant viruses), 967–973, 968t adeno-associated virus (AAV-2), 968t, 970–971, 971 adenoviruses, 968t, 972–973, 986 baculovirus, 973 HSV-1, 973 hybrid viruses, 968t retrovirus-based, 967–970, 968t, 969 simian virus 40 (SV40)-based, 968t, 971–972, 972 Wilson disease, 229 genetic factors, chronic HCV infection outcome, 886 genetic screening, zebrafish use, 1072 gene transfer to liver, 967–973 pharmacological products produced, 965 genomic data, proteomic data integration, HCC, 995 GGA proteins, 129 Gilbert syndrome, 253, 254 Gleevec, 994 gliotoxin, 444 Glisson’s capsule, 6 Global Alliance for Vaccines and Immunization (GAVI), 870 glomerulonephritis hepatitis B, 622 hepatitis C, 622–623 glomerulopathies, in HIV-1 infection, 623–624, 623t glucagon cAMP level increase, 528 fasting hyperglycemia and, 478 gluconeogenesis regulation, 478 tyrosine-aminotransferase and PEPCK regulation via cAMP, 527 vasodilatation in portal hypertension, 709 glucocorticoid receptor (GR), 686t gluconeogenesis, 477 decreased by metformin, 479, 544–545 enzymes AMPK regulation, 540, 540 , 541 increase, type 2 diabetes, 478 location and activation, 496 fed vs fasted states, 477 increased in type 2 diabetes, 477, 478

INDEX insulin and glucagon controlling, 477–478 suppression, LKB1-dependent AMPK-related kinases, 545 glucose brain, metabolism, hepatic encephalopathy, 604–605, 605 enhanced production, 477 in insulin resistance, 261, 477–478 metabolism, 724–725 adipose tissue role, 262 ammonia effects on, 604, 605 brain, hepatic encephalopathy, 604–605, 605 Ca2+ signaling role, 496–497 hepatic, regulation, 477 mobilisation, ATP role, 496 non-oxidative disposal, impaired in insulin resistance, 473, 474 , 475, 475 phosphorylation, 472 regulation, AMPK role see AMPK (AMP-activated protein kinase) sensor, aldolase as, 66 transport defect in insulin resistance, 472 insulin-mediated, mechanism, 471, 474 uptake, insulin role, 720 glucose 6-phosphatase (G6Pc), 478 regulation, AMPK role, 540, 540 , 541 β-glucuronidase, gene therapy, 974, 975 glucuronidation bile acids, 291, 293 , 295, 689–690 bilirubin, 252, 253 GLUT4 glucose transporter, 262, 471, 474 glutamate hepatic encephalopathy pathogenesis, 606 , 606–607 neuron–astrocyte trafficking, 606, 606 release by astrocytes, 606, 607 glutamate–glutamine cycle, hepatic encephalopathy pathogenesis, 606 , 606–607 glutaminase, 602 glutamine cerebral efflux, in acute liver failure, 604 increased in liver failure, 602, 603 inter-organ trafficking, 602, 603 metabolism, 602 neuron–astrocyte trafficking, 606, 606 as organic osmolyte, 603 glutamine synthetase, 602, 603 γ-glutamyl transpeptidase (γGT), increased levels, 661, 669, 670 glutathione (GSH), 691, 755–756 alcoholic liver disease, 751, 755–756 in bile, 339 in copper toxicity, 227 depletion, 139 excretion, Mrp2 pathway, 691–692 mitochondrial (mGSH), 755–756 precursors, 756 glutathione peroxidase, 515, 756 glutathione reductase, 756 glutathione S-transferase (GST), 1131 cis-elements in promoters, animals/plants, 1132, 1133 expression and signaling, 1131, 1132 activation via cis-element AREs, 1132, 1133, 1133 Keap1-dependent ubiquitination of Nrf2, 1133–1134, 1134

plants and TGA factors, 1135 , 1135–1136 redox regulation of KEAP1 and, 1133–1134, 1134 transcription factors (Nrf2-Maf complex), 1133, 1134 , 1135 xenobiotics activating, 1132, 1133 see also Nrf2 isoenzymes and classes, 1132 phase II detoxification role, 1132 plant enzymes, 1132 glycan modifications apical sorting signal, 81, 82 N-linked, protein folding, 162, 163 glycerol-3 phosphate acetyltransferase (GPAT), 538–539 glycerolipids, biosynthesis, 258 glycerophospholipids, canalicular membrane, 344, 345 glycine, bile acid conjugation, 291, 294 glycogen, 257 in fed and fasted states, 477 synthesis, 496 glycogen phosphorylase, 496 glycogen synthase, 496 glycogen synthase kinase-3, 727 glycolysis, transcriptional control, 629, 631 glycoproteins, misfolded, recognition by chaperones, 166 glycosaminoglycans (GAGs), HCV receptor/uptake, 116 glycosylation, 961 asparagine-linked, 162 N-glycosylation oatp1a1 (oatp1), 313 proteins, 162, 164 glycosyl-phosphatidylinositol (GPI) anchor, 81, 164 glycyrrhizin, 445 Golgi adaptor protein-1A (AP-1A), basolateral protein sorting signal, 81 Golgi apparatus, 125 ABC transporter trafficking, 350, 350 , 351 clathrin-coated vesicles, role, 108 in hepatocytes, 74 , 74–75 myosin 6 role, 36 protein sorting role, 125–126, 176 protein transport from ER see endoplasmic reticulum (ER) cis-Golgi network (CGN), 125 trans-Golgi network (TGN) see trans-Golgi network (TGN) (under ‘trans’) golgins, 130 GPIa–IIa, 641 GPIb–IX, 643, 644 GPIIb–IIIa, 642, 643, 644 GPVI, 641 gp46 (rat homolog of HSP47), 1123–1124, 1124 see also heat shock protein 47 (HSP47, gp46) gp80, 795 gp130 receptor, 556, 795 gpat KO mice, 262 Gpbar-1 see TGR5 GPI anchor, 81, 164 GPI–transamidase complex, 164 Gpr-55, 1092 G-protein(s), 45, 46, 485–486, 522 cAMP regulation, 522 cAMP signal transduction, 522, 523

1159 chains, 522 Rab4, 51 subunits, 486, 522 G-protein-coupled receptor (GPCR), 485, 486 Gpr-55, 1092 hepatocyte cytoprotection, 795 G-protein-coupled receptor kinase-2 (GRK2), 706 GRACILE syndrome, 667t, 670 granular component (GC), nucleolus, 569 granulomatous disease, hepatic fibrosis due to, 435 green fluorescent protein (GFP), 1053–1054 limitations, 1054 Ntcp targeting to plasma membrane, 308, 309 photoactivatable (PA-GFP), 1054 zebrafish studies, 1067, 1068 “green liver”, 1131–1138 see also detoxification growth factors, 407 alcoholic liver disease, 748–749 ECM synthesis (liver development), 453, 454t hepatic fibrosis, 439 drugs inhibiting, 444 hepatocyte proliferation, 528 induction by bone marrow cells, 586 liver development, 584 mitogenic stimuli, G1 phase, 1016, 1016 receptors, calcium signaling initiation, 486 stellate cells releasing, 409, 410, 437 transdifferentiation, 412, 436, 436 tight junction regulation, 214, 214 GRP75, 141 GRP78, 759 GRP94, 162 GRX cells (HSC cell line), 1122 GS-9190, 912 GSK-3β, 795 GSTA2 gene, 1132, 1133 GTP α/β tubulin assembly/polymerization, 30, 31 COPII-coated vesicle formation, 128 hydrolysis, 486 clathrin-coated vesicle scission, 110 nuclear transport cycle, 150, 151 GTPases clathrin-coated endocytosis, 110 COPI-coated vesicle formation, 126 COPII-coated vesicle formation, 128, 129 signal recognition particle, 159 GTP-binding proteins, vesicle targeting, 82 GTP-GDP exchange factors (GEFs), 126 GTPγS, 51 GTPψS, clathrin-coated vesicle scission, 110 guanine nucleotide exchange factor (GEF), 150, 151 guanosine analog, 906 guanosine triphosphate (GTP) see GTP guanosine triphosphate (GTP)-binding regulatory proteins see G-protein(s) guanylate cyclase, 486 “guided evolution”, AAV-vectors, 971 Gunn rat model, 967, 980–981

1160 gut microbiota, 721, 721 hepatic fat accumulation, role, 721 , 721–722 insulin resistance development, 721–722, 722 NAFLD pathogenesis, 721 , 721–722 therapeutic target, 731 gut tube, formation, 17 GW182, 1032

H 7H6 protein, 87t HA4 (canalicular cell adhesion molecule (cCAM105)), 350 Hagen–Poiseuille formula, 393 halofuginone, 445 hamartin, 542 hamartomas, 542 haptoglobin, 245 H+ -ATPase, vacuolar see V-type ATPases H+ -ATPase 2 (AHA2), plant, 58–59 regulation, 60 structure, 59 H+ /K+ -ATPase, 58 HAX-1, 354 HBC-3 cells, 1041 heat shock protein 47 (HSP47, gp46), 163, 1121, 1123 rat homolog (gp46), 1123–1124, 1124 role in procollagen maturation in ER, 1122, 1122 siRNA against see siRNA–HSP47 (gp46) heat shock protein 70 (Hsp70), 162, 182 heat shock protein 90 (Hsp90), 711, 862 inhibitors, 862 Hedgehog signaling (Hh), 366 cholangiocyte hyperproliferation, 366 Hip repressor, 414, 416 ligands produced by stellate cells, 410 liver development in zebrafish, 1068 stellate cells transdifferentiation, 414, 416 HeLa cells, 36 hematopoiesis, liver role, 4 hematopoietic stem cells (HSC), 21 heme, 251 reactive, clearance by liver, 245 heme oxygenase, 251, 252, 513, 605 hemochromatosis, hereditary hepatic fibrosis due to, 435 HFE -associated, 239, 240, 243 C282Y mutation, 243 juvenile, causes, 243–244 severe early onset, mutations, 244 TfR2-associated (mutations), 243 hemodynamics, liver, 3–4, 6–7 microsegmentation of parenchyma, 7–9, 8 hemoglobin, breakdown, bilirubin formation, 251 hemoglobin–haptoglobin complexes, 245 hemojuvelin, 240 mutations, severe early onset hemochromatosis, 244 hemopexin, 245 hemophilia, 644 hemophilia A, gene therapy, 969 hemostasis, 640–641 abnormal, bleeding in chronic liver disease, 650–651 activation, in drug–inflammatory stress, 774, 777–778

INDEX coagulation cascade, 639–640, 640 in vivo model, 639, 640–641 in liver disease, 641–651 primary, 641, 643 see also platelet(s); von Willebrand factor (VWF) secondary see coagulation hepaciviruses, 816 hepadnaviruses, 810, 838 heparan sulfate proteoglycans (HSPGs), 455, 459 heparin-binding EGF-like factor, 559 heparin cofactor II, 648 heparinoids, 645 heparin sulfate proteoglycans (HSPGs), 277 hepassocin, 458 hepatectomy, partial see partial hepatectomy (PH) hepatic adenoma, iron refractory anemia, 245 hepatic artery, 5–6, 375 hyperdynamic circulation, 708 , 708–709 hepatic cysts, formation, 366 hepatic efflux transporters see ABC transporters hepatic encephalopathy (HE), 597–601, 597–617, 600t in cirrhosis associated with HCV, 885, 922 classification, 597–601, 600t endocannabinoid action, 1100 episodic, 600–601 minimal, 600 neuropathology, 601 , 601–602 pathogenesis, 602–610 altered gene expression (brain), 609 ammonia, 602–603, 603 brain glucose/energy metabolism, 604–605, 605 brain organic osmolytes and cell volume, 603–604, 610 cerebral blood flow, 607–608, 608 , 610 GABA, 607, 607 glutamate and glutamate–glutamine cycle, 606 , 606–607 hyponatremia, 609–610 infection and inflammation, 609 manganese, 609 mitochondrial dysfunction, 605–606 monoamines, 607 neurotransmitter systems, 606 , 606–607 oxidative/nitrosative stress, 605–606 persistent, 601 therapeutic approaches, 610–611 type A (acute liver failure associated), 597–600, 600t type B (portal–systemic bypass), 599, 600, 600t type C (cirrhosis/portal hypertension associated), 599, 600–601, 600t hepatic engorgement, sinusoidal obstruction syndrome, 378 hepatic failure see liver failure hepatic fibrosis, 407, 433–452 activated macrophages role, 626 , 626–627 alcoholic liver disease, 434–435, 746, 748 management, 434–435

alcohol metabolism associated, 744 animal models, 417, 1124–1125 siRNA–gp46 (HSP47) antifibrotic effect, 1124–1126, 1125 cannabis use and, 1095–1096 causes, 433–435, 434t CB1 receptor fibrotic action, 1096 CB2 receptor activation, protective role, 1095 cellular basis, 435–438, 458, 1122 see also hepatic stellate cells (HSCs), activation cytokines involved, 438–440 adipokines, 439, 745, 746 cannabinoids/endocannabinoids, 439–440, 1095–1096 growth factors, 439 vasoactive substances, 439 end-stage liver disease, telomere shortening and, 1109 excess vitamin A, 409 extracellular matrix (ECM), 417–418, 454t, 458, 462 liver stiffness and, 461 fibrogenic process, 417–418, 437, 437 , 1122, 1122 HCV infection, 884, 886, 922 CB2 effect, 1095–1096 HDV infection, 823 imaging, new method, 1127 leptin associated, 745, 746 lymphocyte role, 748 macrophage and Kupffer cell role, 749 NAFLD/NASH, 730, 733 natural history, 433, 434 neutrophil role, 749 portal hypertension pathogenesis, 704 remodeling and resolution, 443 by siRNA–HSP47(gp46), mechanisms, 1124–1126, 1125 reversibility, 417, 443 serological markers for monitoring, 460, 1122 signaling pathways, 440–443, 441 drugs inhibiting, 444 sinusoidal endothelial cell pathology, 377 stellate cell role see hepatic stellate cells (HSCs) T cell role, 748 therapies, 433, 434–435, 443–445, 444t, 461 CB2 agonist, 1095 cell-specific delivery, 445 current approaches, 1123 in NAFLD/NASH, fibrosis as target, 733 vitamin A-liposome-siRNA HSP47, 1123–1126, 1125 as wound-healing response to chronic liver injury, 433 see also fibrogenesis; hepatic stellate cells (HSCs), activation hepatic inflammation see inflammation hepatic ischemic-reperfusion injury, endocannabinoids, 1100 hepatic lipase, 277, 280 hepatic microcirculation, endotoxin effect, 750 hepatic microcirculatory subunit, 8 hepatic necrosis, 379 hypercoagulation leading to, 649 submassive, ductular reaction, 587

INDEX hepatic plates, 7 functional/structural heterogeneity along, 10–11 zones, 10 hepatic regeneration see liver regeneration hepatic steatosis, 263 chronic hepatitis C outcome, 886 development, energy homeostasis and, 724 , 724–726, 725 endocannabinoids and see endocannabinoids HCV-associated, SREBPs role, 631 increased fenestrations leading to, 393 insulin resistance and, 476–477 deterioration, 726–727 peripheral, causing steatosis, 473–475, 475 see also insulin resistance pathogenesis, SREBPs and ChREBP, 260, 629 zebrafish model, 1071 see also fatty liver disease; non-alcoholic fatty liver disease (NAFLD) hepatic stellate cells (HSCs), 375, 375 , 407–432 acetaldehyde-responsive elements, 747 activated, 436–437, 437 , 528, 704, 753 angiogenesis promotion, 437 apoptosis and spontaneous recovery, 436, 442, 443, 1126 apoptosis promotion by drugs, 444 biological properties, 436–437, 437 cell-specific drug delivery, 445 chemokines secreted, 438 cytokines produced by, 410, 437, 437 disappearance, siRNA–HSP47 and, 1126 ECM degradation and, 443 ECM interaction, 437, 437 gene expression changes, 412, 413 growth factors produced by, 409, 410, 437, 437 hepatic inflammation, role in, 437, 729 integrins expressed, 437 migration, stimuli for, 436–437 portal hypertension, 704 procollagen secretion, 1122, 1122 proliferation, CREB phosphorylation controlling, 528 removal/fibrosis resolution, 443 resistance to apoptosis, 436 activation/transdifferentiation, 377, 408, 408 , 409, 411–413, 433 acetaldehyde role, 747–748 adiponectin role, 412, 746–747 bile acid role, 298 by capillarization, 377, 438 defenestration leading to, 398 ECM density increase leading to, 458 factors initiating, 411–412, 436, 436 , 437, 439, 441, 753 fibrogenesis in NASH and, 730 Hedgehog pathway, 414, 416 hydrogen peroxide formation, 411, 412, 438 hypoxia role, 416 identification/immunohistochemistry, 434 initiation phase, 411–412, 435–436, 436 lymphocytes role, 414–415, 438

oxidative stress causing, 436 perpetuation by growth factors/cytokines, 409, 411, 412, 436, 436 , 437, 439 physical nature of ECM role, 414 prevention, 412, 413, 415, 444–445 as reversible process?, 416–417, 443 signaling pathways, 414, 415 , 416 , 416t, 440–443, 441 SMA expression, 412, 417, 434 , 459, 462 1126 as source of ECM, 435 as “suicidal” command, 416–417, 443 transcription factors up-regulated, 413, 413t vitamin A deficiency leading to, 398, 1122 Wingless (WNT) pathway, 414, 415 , 416t, 442 see also hepatic fibrosis; myofibroblasts adipogenic phenotype, 412, 413–414 collagen type I and III expression, 413, 417, 747 CREB function, 528 culture, stiff matrices, 462 cytokines produced, 410, 411, 412 embryogenesis, 5, 5 , 411 ethanol-induced increase in 2-AG, CB1 role, 1099, 1099 fibrosis, role in, 408, 411, 1121 functions, 408, 409–411 blood flow regulation, 409 growth/metabolism regulation, 409–410 immunoregulation, 411, 414, 437 vitamin A storage/retinoid metabolism, 408, 409 gap junctions between, 408 general description, 408–409 genes expressed, 410, 410t, 411, 417 growth factors produced, 409, 410, 412 heterogeneity, 417 isolation, 408 leptin and adiponectin expression, 744, 746 leptin as mitogen, 437, 746 microcirculation regulation, 704, 707 morphology, 10, 408, 409 NADPH oxidase role, 752 origin/embryology, 5, 5 , 411 phagocytosis, 411, 414 pluripotential cell features, 411 quiescent, 409, 435, 436 adipogenic phenotype, 412, 413–414 functions, 409–411, 435 regulation and activation, 412–413 transcription factors expressed, 413, 413t vitamin A storage, 408, 409, 1122 resistin expression, 747 ROS formation, 753 sinusoidal endothelial cell phenotype control, 374 structure, 10 targeting with vitamin A-liposome-siRNA HSP47, 1123–1124, 1124 thrombin as mitogen, 649 TIMP-1 as survival factor for, 418 transformation into myofibroblasts, 408, 408 vitamin A storage, 408, 409, 1122

1161 vitamin A uptake mechanism, 1122, 1124 hepatic stem cells see stem cell(s) hepatic uptake transporters see solute carriers (SLCs) hepatic vascular resistance, 703 cirrhosis, 704 portal hypertension pathogenesis, 704–708 sinusoidal contraction, 704 vasoactive agents affecting, 704–708, 705 hyporesponse to vasodilators, 705 , 705–707 increased response to vasoconstrictors, 705 , 707–708 hepatic (“central”) veins, 3, 7, 8 hepatic veno-occlusive disease, 377 see also sinusoidal obstruction syndrome (SOS) hepatic venous pressure gradients (HVPGs), hepatitis C, 924 hepatitis alcoholic see alcoholic hepatitis autoimmune see autoimmune hepatitis (AIH) cholestatic fibrosing post-transplant in hepatitis B, 926 post-transplant in hepatitis C, 922 food-borne, 823 fulminant, 823 historical report, 803 neonatal, 661, 665 viral, 807–834 acute, 807 common features of viruses, 807 historical background/future prospects, 803–805 see also individual hepatitis viruses hepatitis A, 803 hepatitis A virus (HAV), 804, 807, 808–810 2A protein, 808, 809 assembly and release, 809 clearance and hepatocyte destruction, 810 cytopathology and cytolytic strains, 809–810 epidemiology, 808 genome structure, 808 , 809 HuHAVcr-1 receptor, 808–809, 810 incubation period, 809, 810 infection, and pathogenesis of, 810 morphology, 808–809 non-cytolytic clearance mechanisms, 810 passive immunization against, 808 persistent infection with, 809, 881 polyprotein, 809 protein translation, 809 replication, 809–810 stability, 808 transmission and source, 808 vaccine, 808 VP1 and VP4, 808 Vpg protein, 809 hepatitis B, 803, 814–815, 859–860 acute, 859 early adaptive immune response, 836–837, 837 fulminant, post-transplant, 926 innate immune response, 836–837, 837 T cell response, 837–838

1162 hepatitis B (continued) biomarkers vs HCC development, 1002–1003 chronic, 814, 859 evolving, T cell response, 838 , 838–839 hepatic fibrosis, 434 IFN-α therapy, 904 prevention, 870 progression to, 814–815 T cell response, 839 therapy development, 805 cirrhosis, 859, 867, 871 management post-transplant, 927 epidemiology, 859–860 flares, 839 gene expression profiling of effects on liver, 1002 gene therapy, 967, 976 glomerulonephritis, 622 HCC and see hepatocellular carcinoma (HCC) liver transplantation, 805, 926–928 graft failure, 926 graft reinfection, 926 HBIG after, protective effect, 926, 927 HBIG and lamivudine, 927–928 immunization effect, 928 lamivudine before, 927, 928 nucleoside monotherapy, 927 recurrence prediction, 926–927, 926t, 928 requirement, oral therapy impact, 927, 928 pathogenesis, 814–815 recurrence, 926 post-transplant, 926–927, 926t, 928 therapy, 859, 861, 866–867, 900t assessing/monitoring, 899 combination, 904 current medications, 866–867 development, 804–805 general principles, 899–900 HBV polymerase/RT inhibitors, 861, 862, 866, 869t, 901, 902t, 903 IFN-α, 861, 866, 904 impact on transplant requirement, 927, 928 indications and duration, 867 long-term goal, 899 mechanisms of action, 901 molecular/pathophysiological basis, 900 , 900–901 nucleoside monotherapy post-transplant, 927 nucleoside/nucleotide analogs, 866–867, 901, 902t, 903 replication inhibition but no elimination, 903 resistance, 861, 867 “surrogate markers” of efficacy, 867 sustained virological response, 867 targets, HBV replication and, 860 , 860–861 see also lamivudine; other specific drugs transient, 814 transmission by donor liver, 926, 928 hepatitis B immune globulin (HBIG), 870, 926, 927 intramuscular vs intravenous, 927

INDEX lamivudine with, post-transplant, 927–928 hepatitis B virus (HBV), 115, 807, 810–816, 859–876 “a” antigenic determinant, 868, 870, 927 mutants, vaccines, 870 antibodies to, 839–840 defective production, HBV persistence, 840 epitope specificity, 870 antibody-escape mutants, 811, 816, 840 antigens, HBV persistence and immune response, 840, 871 assembly, 813, 901 basal core promoter (BCP) region mutants, 868 capsid protein, 862–863, 862t antibodies, 863 gene mutations, 868 see also hepatitis B virus (HBV), HBcAg carriers, 815, 860 HBeAg loss, 863 HCC in, 868, 870 cccDNA (covalently closed circular) 812 , 813, 860, 861, 900 , 901 intracellular amplification pathway, 864 loss/intracellular destruction, 814, 815 measurement, 864 pool size, 864, 901 regulation of copy number, 813 synthesis, 860–861 circular DNA (double-stranded), 811, 860, 860 classification, 810 clearance, 814 antibodies role, 839–840 core promoter mutations, 870 core protein see hepatitis B virus (HBV), HBcAg cytopathic mechanisms, 836 Dane particle, 804 DNA, 837, 864–866 clearance, 836–837 cytoplasmic forms, 866 detection, HBV recurrence post-transplant, 928 elevation, CD8 cell absence, 837 integration into host DNA, 815, 866 plasma levels, 836, 864, 899 replication see below DNA polymerase/reverse transcriptase (Pol), 811–812, 813, 860, 861–862, 862t, 900 binding to epsilon, 861, 862 domains, 861 as drug target, 861 functions, 901 inhibitors, 861, 862, 866, 869t, 901, 902t, 903 mutants, 868, 869 , 869t structure, 861, 862 variants resistant to inhibitors, 867, 868 drug resistance, 901, 903 detection, 903 lamivudine-resistant, 861, 868, 903, 927 mechanism, 900 , 900–901 reduction by combination therapy, 904 drug-resistant variants, 861, 862 , 867, 868, 901, 903, 903t

envelope, 862t, 863–864 antibodies to, 839–840 gene mutations, 868, 870 vaccines and, 870 see also hepatitis B virus (HBV), HBsAg epitopes, 868, 870 escape mutants, 811, 816, 840, 870 evolution, 815–816 fibrosing cholestatic hepatitis, 926 gene transcription, 812 , 813, 860–861 activator (X protein), 811, 814, 864 genome replication, 812 , 837, 860 , 860–861, 864–866, 865 cccDNA as intermediate, 811, 812 , 864, 865 initiation, 811, 812 , 813, 861 in situ priming of plus strand synthesis, 812 , 813, 815 site (nucleocapsids), 863, 866 genome structure, 811–812, 812 , 860 , 868, 900 open reading frame (ORF), 810, 811–812, 812 , 861 terminal redundancy of minus strand, 811, 813 genotypes, 867–868, 927 outcome and persistence, 840–841 half-life, 867 HBcAg (core antigen), 811, 814, 862–863 antibodies, 863 mutants, 868, 870 HBeAg, 814, 840, 863, 900 antibodies, 814 drug therapy indication, 867 HBV persistence and, 840 hepatic fibrosis and, 434 loss and mutants, 863, 868 seroconversion, 863 translation and detection, 863 viral load, and therapy, 867 HBeAg-negative disease, therapy, 900t HBeAg-positive disease, therapy, 900t HBsAg, 803–804, 811, 862t, 863–864, 900 antibodies, 863 carriers, 815 clearance, IFNα, 904 in HDV assembly, 820, 822 L, M and S proteins, 811, 812, 813, 863 L, M and S protein translation, 813, 863 morphogenesis and MHB glycoprotein, 863–864 M protein oncogenicity, 815 mutants, 868, 869 , 869t persistence, post-transplant, 926 production, integrated viral DNA, 815 stability of polymers, 864 see also hepatitis B virus (HBV), envelope HBsAg-negative liver donors, 926, 928 HBsAg-positive disease, treatment, 900t HCC and see hepatocellular carcinoma (HCC) HCV coinfection, 838, 887 HDV requirement for, 820, 822 historical background/future prospects, 803–805 HIV co-infection, 434

INDEX host gene expression profiling, 1002–1003 host genes altered by, 1002 immune response, 836 , 836–841 acute infection, 837 , 837–838 CD4 cells, 837–838, 839 CD8 cells, 837, 839 clearance, non-cytopathic mechanisms, 836 , 837 early innate/adaptive responses, 836 , 836–837 HBV persistence mechanisms, 840–841, 871 humoral, 839–840 T cells in acute self-limited infection, 837–838 T cells in chronically evolving infection, 838 , 838–839 T cells in chronic infection, 839 see also antibodies; CD4 T cells; CD8 T cells infection see hepatitis B insertional mutagenesis of HERT (telomerase) by, 1113 lamivudine-resistant, 861, 868, 903, 927 latency, 838 life cycle, 860 , 860–861, 900 , 900–901 linear DNA (precursor), 811 mononuclear cell infection, 815 morphology, 811, 900 mRNAs and enhancers, 813, 861, 901 mRNA translation, 813, 814, 861, 901 mutants/mutations, 861, 863, 868 core/precore genes, 868 core promoter, 870 drug resistance and, 861, 867, 901, 903, 903t envelope genes, 868 escape mutants, 811, 816, 840, 870 HBsAg, Pol inhibitor resistance, 868 Pol, 868, 869 , 869t nucleocapsid protein, 813, 862–863, 901 number produced daily, 867 passive immunization, 870 persistence, 814, 838 CD4/CD8 cells and, 837–838, 871 mechanisms, 840–841 POL gene, 812, 813 see also hepatitis B virus (HBV), DNA polymerase post-transfusion hepatitis, 804, 810 precore protein, 814 gene mutations, 868 pregenome and degrading of, 813 pregenomic RNA (pgRNA), 860, 866, 901 pre-S mutants, 868 proteins, 811–812, 813, 861–864, 862t accessory functions, 814 pX protein, CREB binding, 529 quasi-species (sequence variants), 816 “reactivation”, 864 receptors, 115 replication, 812 , 813–814, 860–861, 864–866, 865 , 900 , 900–901 high levels, defective CD4/CD8 function, 839 inhibition by NK-T cells, 837 inhibitors (drugs), 861 regulation, 813, 837 time course, 837 reverse transcriptase and transcription, 813, 860, 861–862

RNA, 866, 901 ribozymes cleaving, therapy, 976 RNA polymerase, 811, 861–862 RNase H activity, 813 surface antigen see hepatitis B virus (HBV), HBsAg transient infection by, 814 uptake by clathrin-mediated endocytosis, 115–116 vaccine, 804, 810, 823, 839–840, 870–871 childhood, 871 coverage expansion and schedule, 870 liver transplant recipients, 928 waning immunity, 871 variants, 815, 867–870 viral load, 864, 867 viremia, 864 T cell response suppression, 839 X protein, 811, 814, 862t, 864, 900 mutations, 870 oncogenicity, 814, 815 structure and function, 864 transcription activator, 811, 814, 864 hepatitis C acute antibody response, 844, 881–882 B cell response, 844–845 T cell response, 842–844 acute progressing, 882 biomarkers vs HCC development, 1002–1003 chronic, 433–434, 882 antifibrotic management, 443 clinical features, 883 epidemiology, 882 factors associated with outcome, 885–887, 885t hepatic fibrosis due to, 433–434, 884, 888 natural history, 883 progression to cirrhosis, 883–884 rates and factors associated, 882–883 reversibility of fibrosis/cirrhosis, 443 T cell response, 843, 844 therapy development, 805 transfusion-associated, natural history, 888 see also hepatitis C virus (HCV), persistence cirrhosis, 888 in allografts, 921–922, 925 compensated HCV-related, 885 decompensated, 885, 922 outcome, 884–885 prevalence, progression rates, 883–884 retransplantation for, 925 coagulation disorders, 645, 646 diagnosis, 820 factors associated with outcome, 885–887, 885t environmental (and alcohol), 887 host factors, 885–886, 885t viral factors, 886–887 fibrosing cholestatic and cholestatic hepatitis, 922 gene expression profiling of effects on liver, 1002 gene therapy, 967, 976 glomerulonephritis and nephropathy, 622–623

1163 HCC and see hepatocellular carcinoma (HCC) hepatic fibrosis, 433–434, 884, 888, 922 CB2 effect, 1095–1096 progression rate, post-transplant, 923–924 hepatosteatosis associated, 886 SREBPs role, 631 liver transplantation, 805, 921–926 acute cellular rejection, 924–925 allograft failure, 922 antiviral therapy before, 922–923 diagnosis of recurrent disease, 923–924 optimal immunosuppressive regimen, 925–926 preemptive antiviral therapy, 923 recurrent HCV infection, 921–922 retransplantation, 925 treatment of recurrent disease, 923–924 treatment of recurrent HCV after, 922, 923t mortality, 884 NAFLD and insulin resistance, 886 oxidative stress effect on fibrinolysis, 646 persistent, clinical consequences, 882 see also hepatitis C virus (HCV), persistence recurrent diagnosis, 923–924 maintenance therapy, 924 natural history, 921–922 severe, risk factors for, 922, 922t treatment, 922, 923–924, 923t, 925 telomere shortening, 1109 therapy, 804, 904–913 algorithm, 925 antivirals pre-transplant, 922–923 cessation, hepatitis progression, 924 current guidelines, 907, 907 “difficult-to-treat” patients, 908–909 duration, 908 elevated doses (ribavirin), 908 general principles, 904–905 maintenance, 924 molecular/pathophysiological basis, 905–906 new approaches, 905, 909–913, 910 novel forms of IFN-α, 909 optimization (pegylated IFN-α–ribavirin), 907–909 preemptive after transplantation, 923 rapid/slow virological responses, 908 retreatment of non-responders, 908 specific inhibitors, mechanisms, 911–913 standard-of-care (pegylated IFN-α–ribavirin), 888, 906–907, 924 taribavirin, 909 hepatitis C immune globulins, 911 hepatitis C virus (HCV), 116, 807, 816–820, 877–897 adaptive mutations, 819, 845–846 angiogenic factors up-regulated by, 1002 antibodies to diagnosis/persistent infection detection, 820 evidence for/research methods, 844, 881 monoclonal, 881

1164 hepatitis C virus (HCV) (continued) neutralization of pseudo-particles, 844–845, 881, 882 neutralizing, 844–845, 877, 880, 881–882 persistent infections, 842, 881 undetectable after recovery, 883 assembly and release, 819, 910 , 911 inhibitors, 913 VLDL secretion, 819 classification, 816 clathrin-mediated endocytosis, 909–910 claudin-1 expression requirement, 818 clearance, rates of and factors associated, 882–883 co-receptors, 818 core protein, 819, 847 inhibition of T cells, 847 diseases associated, 818, 887–888 DNA ribonuclease action, 977 entry/uptake mechanism, 116, 217, 818, 909–910 inhibitors, 911 envelope proteins (E1 and E2), 816, 817, 819, 909 CD81 binding, 817, 818 E2, antibodies, 881 E2 and CD81 interaction, 842, 845, 909 E2 and NK activation inhibition, 842, 848 escape mutants, 817, 845 T cell inhibition by, 847 eradication, hepatocellular carcinoma despite, 888 “error catastrophe” induced by ribavirin, 906 ER stress, 788–789 escape mutants, 817, 845–846, 880–881 CD8 epitopes, 846 F protein, 910 genome structure, 816–817, 817 , 910 3′ -untranslated region (3′ -UTR), 817, 1037 5′ -untranslated region (5′ -UTR), 816, 817, 910, 976, 1037 ‘kissing-loop’, 3′ -UTR, 817, 819 oligonucleotides targeting 5′ -UTR, 911 open reading frame (ORF), 816, 817 genotypes, 816 current therapy guidelines, 907, 907 genotyping, 820 half-life, 845 HBV coinfection, 838, 887 HCC and see hepatocellular carcinoma (HCC) historical background/future prospects, 803–805 HIV coinfection, 434, 886–887 host gene expression profiling, 1002–1003 host genes altered by, 1002 immune escape (antibody-driven), 817, 845, 881–882 immune evasion strategies, 805, 841, 877–897 from B cell surveillance, 845–846, 881–882 dendritic cell function impairment, 846, 847–848 direct T cell inhibition, 846, 847

INDEX IFN synthesis disruption, 841, 878 , 878–880 innate immunity evasion, 841–842, 878 , 878–880 negative regulation, T cell dysfunction, 846–847 PD-1 expression role, 847, 880 signaling disruption, 841–842, 878 , 878–880 from T cell surveillance, 845–846, 880–881 immune response, 841–848, 843 , 877 B cell response, 844–845, 881–882 CD4 cells, 843–844, 845, 846, 880 CD4 response impairment, 843, 844 CD8 cells, 837, 845, 880 CD8 cells dysfunction, 842–843, 844, 880 CD8 escape mutants, 845–846, 880 early, CD4 response, 843 , 843–844 early, CD8 response, 842–843, 843 evolution of chronic infection, 843 failure, mechanisms, 845–848, 880–882 innate, 841–842, 877 innate, signaling pathways, 878 , 878–880 T cells in chronic HCV, 844, 880 infection see hepatitis C interferon-α susceptibility/resistance, 819, 841 genes affected, 1002 internal ribosome entry site (IRES), 816, 817, 819, 910 drugs targeting, 911 JFH strain, 844 life cycle, 818–819, 909–911, 910 macroautophagy inhibition, 183 , 184 miR-122 relationship, 1037 morphology, 816 mutation rate, 845 non-structural (NS) proteins, 817, 819, 910 oncogenic, 887 see also specific NS proteins below NS2, 818, 910 NS3–4A (serine proteinase), 818, 910 inhibitors, 879, 911–912 innate immune evasion, 841, 878 , 878–879 NS3 protease/helicase, 818, 819, 887, 911 NS4A, 818, 819 NS4B, 818, 887, 911 NS5A, 818, 819, 879, 887, 910 domains I/II, hinge region, 818–819 gene expression alteration by, 1002 interferon-sensitivity-determining region, 819 NFκB interaction, 1002 NS5B (RNA polymerase), 818, 910 nucleocapsid, 816 p7, 817–818, 910 persistence, 877–897 B cell response/evasion, 845, 881–882 clinical consequences, 882 innate immunity evasion, 841–842, 878 , 878–880 mechanisms, 845–848, 880–882 rates of and factors associated, 882–883 T cell response/evasion, 845–856, 880–881

see also hepatitis C, chronic; hepatitis C virus (HCV), immune evasion strategies polyprotein and cleavage of, 817, 818, 819, 841, 910, 910 inhibitors, 911–912 post-transfusion hepatitis, 804, 888 post-translational processing, 817, 818, 819, 910, 910 inhibitors, 911–912 proteins oncogenicity, 887 structure and functions, 817–818, 910–911 pseudo-particles (retrovirus-HCV), 844–845, 881, 882 quasi-species, 804, 805, 816, 845, 881 escape from neutralizing antibodies, 845, 882 rapid spread, mechanisms, 845 receptors, 116, 217, 817, 818, 909 CD81, 116, 217, 909 cofactors, 217, 909 replication, 818–819, 910 , 910–911 high rate and mutants, 845, 848 inhibitors, 912–913 innate immune response and, 841 interferon-stimulated gene (ISG) effect, 880 miR-122 role/action, 1037 regulation, 819 site, 818–819 RNA, 816–817, 817 , 910–911, 921 intrahepatic, detection post-transplant, 924 quantitation, 820, 907, 921 ribozymes cleaving, therapy, 976 RNA polymerase, RNA-dependent (RdRp; NS5B), 818, 910 non-nucleoside inhibitors, 912–913 nucleoside inhibitors, 912 seroconversion, 844 species and tissue tropism, 818 spread, 804 sustained virological response (SVR), 888, 904–905, 907, 924 transgenic mice, oncogenicity, 887 translation of proteins, 816–817, 819, 910–911 inhibitors, 911 vaccine development, 804, 805, 845 viral peptidases (NS2 and NS3/4A), 910 see also above hepatitis D virus (HDV; hepatitis delta virus), 807, 820–823 antibodies, 823 assembly, 822 assembly and spread, 822 diagnosis, 823 genome structure, 820–821, 821 genotypes, 820, 822–823 HBV co-infection, hepatic fibrosis, 434 HDAg, 820 cytotoxicity, 823 large form (L-HDAg), 820, 822, 823 mRNA translation, 822 post-translational modification, 821, 822 RNA editing, 822 small form (S-HDAg), 820–821, 822 liver fibrosis, 823 persistent infection, 822

INDEX protection on HBV recurrence post-transplant, 926 quasi-species, 820 receptor, 821 replication, 820, 821 , 821–822 RNA replication (RNA-dependent), 821 ribozyme activity, 820, 821, 821 rolling-circle replication mechanism, 821, 821 RNA polymerases, 822 structure/morphology, 820–821 treatment and control, 823 hepatitis E virus (HEV), 807, 823–824 genome structure, 824, 824 genotypes, 823 polyprotein, 824 replication, 824, 824 RNA-dependent RNA polymerase, 824 structure/morphology, 823–824 hepatitis viruses historical background, 803–805 uptake by endocytosis, 115–116 see also individual hepatitis viruses hepatoblast–mesenchymal cell interactions, 454 hepatoblasts, 4 cholangiocyte development, 4, 20 development, 584, 584 differentiation, 4, 20, 22 miRNA profiling, 1039 , 1040 , 1040–1041, 1045 signals for, 21, 21 zebrafish, 1069–1070 gene expression pattern, HCC similarity, 1006 hepatocyte development, 4, 20, 22 liver cell-based therapy, 937 migration, liver bud development, 19, 584, 584 hepatocellular carcinoma (HCC), 871, 1083 ABC transporters, 1075 abnormal cell cycle control, 1021 agrin role, 459 animal models, 993–994 biochemical markers, 871, 999–1000 biomarkers challenges to application of, 1008 epigenetic signatures, 1004 tumor vs cirrhosis, 1003–1004 tumor vs non-tumor, 1002–1003 γ-carboxylation impairment, 650 chronic high-level proliferation of hepatocytes, 1021 classification, 991, 993, 1079 Cu Zn SOD gene deficiency (mice), 224 detection, 871–872 diagnosis, 999–1000 microarray study data, 1001–1004 drug resistance, 1075 drug sensitivity, solute carrier role, 1080–1081 epidemiology and etiology, 991, 1078–1079 extracellular matrix, 459, 463–464 genes up-regulated in, 1003 genetic losses/gains, 1003 genome-wide expression profiling, 991–997, 999–1013, 1079–1080 cross-species comparisons, 993–994 diagnostic signatures, 1001–1004 DNA copy number changes, 992, 994 gene expression signature, 994, 1003

prognostic signatures, 1004–1006 subtype identification, 993 techniques, 992 see also comparative genomic hybridization (CGH) hepatic stem cell signatures, 1006 hepatitis B, 815, 859, 866, 867, 871–872 carriers, 868, 870 detection, 871–872 gene expression profiling changes, 1002 hepatitis C, 887–888 cirrhosis duration, 884–885 gene expression profiling changes, 1002 risk and rates, 887, 888 hepatocyte nuclear factors role, 91–92 induction, telomere shortening role, 1112 invasive, genes associated, 994 liver transplantation, 1000 Mcl-1 overexpression, 787 metastases, 1004–1006 gene signatures, 1004–1005 hepatic microenvironment role, 1005–1006 intra-hepatic, 1000 organ distribution, 1005 miR-122 overexpression, 1035 miRNA as diagnostic marker, 1003, 1042, 1079 miR-21, 1043–1044 miRNA as prognostic marker, 1005, 1079–1080 miRNA profiling, 1041–1044, 1045, 1079–1080 multidrug resistance, 1077 , 1080–1082, 1083 ABC transporters role, 1081–1082 ABC transporter variations, 1082–1083 pharmacogenetics, 1082–1083 solute carriers (SLCs) role, 1080–1081 solute carriers (SLCs) variations, 1083 nodular (NHCDD), 1004 origin/development, 994 osteopontin (OPN), 463, 1004 p53 loss and, 1019, 1021, 1111, 1112 polarity protein expression link, 84 premalignant changes, gene expression, 1002 prognosis, 991, 1000 DNA copy number changes and, 994 poor, factors associated, 1000 stem-cell signatures, 1006 prognostic signatures, 1004–1006 metastasis/recurrence in non-tumor tissues, 1005–1006 metastasis/recurrence in tumor tissues, 1004–1005 telomerase activity, 1112–1113 recurrence, gene signatures, 1005 risk in cirrhosis, 871, 888, 1112 solitary (SLHCC), 1004 staging, lack of consensus, 1079–1080 subtype arising from hepatic progenitor cells, 994, 1006 subtypes, 991, 993, 1004, 1006 stem cell-like phenotypes, 1006 suppression, telomere shortening, 1110 survival rates, 1075 systems biology role, 1080, 1084 telomerase activation and, 1112–1113

1165 telomere shortening and, 1111–1112 treatment, 991, 1000, 1079 response prediction, 1081 target identification, 994–995 telomerase inhibition, 1113–1114 telomere destabilization, 1113–1114 hepatocellular encapsulation platforms, 942 hepatocyte(s) age (cell)-dependent differentiation, 11 albumin synthesis rate, 75 anchorage dependence, 938, 941 apoB transcription, 273, 273 , 274 apoptosis see apoptosis beneficial functions vs cancer drug resistance, 1075, 1083–1084 bile acid uptake see bile acid(s), hepatic uptake bone marrow stem cells generating, 585–586 calcein-loaded, 513–514 calcium signaling see calcium signaling cAMP level peaks, 528–529, 530 caveolae see caveolae cell cycle, and regeneration, 552 see also cell cycle; liver regeneration copper metabolism see copper coupling between, regulation, 210–212 see also gap junctions culture see culture, hepatic cytoskeleton, 32, 45, 50 damaged, reactive oxygen species release, 438 differentiation, 4, 21, 22, 584, 937 ECM role, 463 mesenchymal cell role, 21 miRNA profiling, 1039 , 1040 , 1040–1041, 1045 miRNA role, 1032 zebrafish, 1069–1070 differentiation into bile duct epithelial cells, 580 DNA replication, 550, 551–552 ECM remodeling enzymes, 461 efflux transporters see ABC transporters embryology, 4, 5 , 19, 21, 22 see also hepatocyte(s), differentiation endocytosis/endocytic trafficking, 35, 38, 50–51, 107 mechanism, 50 , 50–52 see also endocytosis endoreduplication, 552 ER abundance, 167 exocytosis, 35 fatty acid metabolism see fatty acid(s); lipid(s) functional heterogeneity, 11 functions, 125, 1075 gap junctions see gap junctions giant-cell changes, 662 hepatotoxic agents targeting, 438 hepcidin synthesis/secretin, 239–240 hypermetabolic state, 752 hyperplasia, 11, 550 see also liver regeneration hypertrophy, c-MYC role, 571 hypoxia, 377 iron storage/homeostasis see iron lifespan, 11 lipoprotein metabolism see lipoprotein(s) metabolic zonation, 10–11, 206 microencapsulation, 942 microtubule and actin filament organization, 32, 45, 50

1166 hepatocyte(s) (continued) cytoplasmic dynein as “-” end motor protein, 33, 38, 48, 48 kinesins, 37–38 mitogens, 11 morphology, 409 motor proteins, 37t, 45, 46t mechanism of action, 50 , 50–52 myosin 1b, 35, 47 myosin, 2 47 see also dynein; kinesins; myosin(s) nuclear receptors, 11, 12 plasma membrane see plasma membrane; polarity ploidy variations, 10 in liver regeneration, 551–552 polarity see polarity (epithelial cells) primary, plasma membrane protein research, 76 progenitors, “oval cells” as, 581 see also “oval cells”, proliferation, 7, 11, 1019 activation, 12 cAMP role in regulating, 528, 529 capacity, retention in animal models, 579 chronic high-level, HCC development, 1021 in culture, 936 Dicer knockdown model and miRNA role, 1034 liver regeneration, 551–552 regeneration, 11, 550–552 see also liver regeneration replacement, 12 secretory pathway see secretory pathway senescence, in cirrhosis, 1109 shape and implications for membranes, 74, 74 size and number, 9 S phase, 552 stem cell properties of, 580 structure, 9, 74–75, 83 surface polarity see polarity tight junctions see tight junctions transcytosis, 35 transferrin receptor TfR2 expression, 236–237, 243 transplantation see hepatocyte transplantation transporters see transporters upregulation of development, 11 uptake of proteins/lipids see endocytosis hepatocyte-derived FBG-related-protein 1 (HFREP-1), 458 hepatocyte–fibroblast co-cultures, 940, 942 hepatocyte growth factor (HGF) activators, 558 blocked by small hairpin RNA, 558 cytoprotection by, 795 functions, 558 produced by stellate cells, 409, 410 progression phase of liver regeneration, 552, 558, 561 receptor see c-Met signaling, 486 storage, 558 hepatocyte growth factor activator (HGFA), 558 hepatocyte nuclear factor(s) (HNFs), 91–92 bile acid transporter regulation in cholestasis, 684–685 HCC development, 91–92

INDEX HNF1 and HNF1α, 22 hnf1b, 1069 HNF4 see hepatocyte nuclear factor 4 (HNF4) HNF6, 22 hnf6 , 20, 1069, 1070 hepatocyte nuclear factor 4 (HNF4), 22, 325–327, 686t bile acid biosynthesis regulation, 297, 325–327, 326, 333 in cholestasis, 685, 689 CYP7A1 regulation by, 327, 333 HNF-1, hepatocytes in culture, 463 HNF-4α hepatocyte polarity, 88t, 91 hepatocytes in culture, 463 phosphorylation by AMPK, 541 role, 88t, 91, 541 NTCP expression regulation, 687, 688 upregulation, hepatocytes in culture, 463 hepatocyte transplantation, 11, 577, 936–937 animal models, 579 , 579–581, 580 , 936 clinical trials, 587–588, 589 early studies, 577–578 future prospects, 588–589 indications, 577–578, 587, 589 Crigler–Najjar syndrome, 577, 587 fulminant hepatic failure, 588 Wilson disease, 228 in vivo proliferation, 936 limited division of transplanted cells, 587 number, in gene therapy using viral vectors, 967 progenitor cells (“oval cells”), 581–582, 937 rationale, 577–578, 580, 936 regulation and microenvironmental signals, 937 xenorepopulation models, 558 , 588 see also liver repopulation; “oval cells”; stem cells hepatogenesis see embryology of liver hepatoma see hepatocellular carcinoma (HCC) hepatoma cell line, CREB phosphorylation and ICER expression, 528 hepatorenal syndrome (HRS), 619–622 hyperdynamic circulation, 620 pathogenesis, 619–620 precipitating factors, 619–620, 621 therapies, 620–622 type I/type 2 and classification, 619, 620, 620t, 621t hepatosteatosis see fatty liver disease; hepatic steatosis hepatotoxic drugs, 438 hepatic fibrosis due to, 435 inflammatory stress associated see drug(s); inflammatory stress hepcidin, 235, 754–755 binding to ferroportin (FPN), 239–240, 241 elevated, anemia of chronic disease, 244–245 functions, 239, 240 iron homeostasis regulation, 241–242, 242 mutations, in hemochromatosis, 243, 244 reduced production, 243 signaling pathways, 240–241, 241 structure, 239, 240

synthesis by monocytes, 245 synthesis/secretion by hepatocytes, 239–240 transcriptional regulation, 240–241 see also iron HepG2 cells, 76, 77t, 308 HepSear see adefovir dipivoxil (HepSera) heptanol, 207 1-HER, hydroxyethyl radicals, 753–754 herbal medicines antifibrotic effects, 445 chronic hepatitis C outcome, 887 hepatic fibrosis due to, 435 herceptin, 994 Hering, canals of, 9, 582–583, 583 herpes simplex virus -1 (HSV-1), gene therapy vector, 973 herpes simplex virus thymidine kinase (HSV-TK), 966, 970 heterophagy/heterophagic pathways, 178–179 see also endocytosis heteroplasmy, 139 Hex homeodomain factor, 19–20, 21–22 hexokinase, 2 472 HFE gene, 240, 243 HFREP-1, 458 HGF see hepatocyte growth factor (HGF) HIF1-alpha see hypoxia inducible factor-1α (HIF-1α) high-density lipoprotein (HDL), 271, 272t biosynthesis, 260 cholesterol delivery to liver, 278, 280 cholesterol efflux from cells, 278, 279 chylomicron/VLDL remnant formation, 275, 275 formation, 278 functions, 272 increased by fibrates, 282 intravascular maturation, 278 metabolism, 277–280 preβ-HDL (nascent), 278 receptor, 278 spherical, cholesterol efflux, 278 highly active antiretroviral therapy (HAART), 886 highly up-regulated in liver cancer (HULC) RNA, 1003 high-mobility group box 1 (HHMGB1) protein, 837 high molecular weight kininogen (HK), 639 high-throughput systems, 937, 995, 1000, 1001 Hip1 (huntingtin-interacting protein, 1) 111 histology of liver, 7–9 histone 3 (H3), 759 histone acetyltransferase (HAT), 525, 540, 540 histone deacetylases, class IIa (HDACs IIa), 542 histone modification, in HCC, 1004 HIV-1 infection chronic hepatitis C outcome, 886–887 glomerulopathies, 623–624, 623t HBV/HCV co-infection, hepatic fibrosis, 434 HIV-associated nephropathy (HIVAN), 623, 623t HIV-immune complex disease, 623, 623t HLA chronic hepatitis C outcome, 886 class I pathway

INDEX breast cancer and, 1082 HBV persistence and, 840 HCV persistence, 846, 880 HMG-CoA, 258 HMG-CoA reductase alcohol-related lipid alterations and, 744–745 cholesterol synthesis inhibition, 281, 538 hypoxia stimulating, 627 intracellular cholesterol inhibiting, 277 regulation by AMPK, 538, 744–745 role of protein quality control/degradation, 167 hMSH2 repair protein, 979, 981 hnf1b transcription factor, 1069 HNF1 HNF4, HNF6 see hepatocyte nuclear factor(s) (HNFs) hnf6 transcription factor, 20, 1069, 1070 homocysteine, ER stress, 759 homocystinemia, 789 homologous recombination, targeted gene modification, 975, 978 HOPS, 528, 529 hormones, calcium signaling initiation, 485–486 hormone-sensitive lipase (HSL), 258, 723, 723 H-Ras gene, 1044 Hrs (hepatocyte growth factor-regulated tyrosine kinase substrate), 112 hsc70 (cytosolic chaperone), 182 Hsp47 see heat shock protein 47 (HSP47, gp46) Hsp70/hsp70 family, 162, 182 Hsp90 see heat shock protein 90 (Hsp90) HU-308, CB2 agonist, 1100 Huh7 cell line, 979, 1037 Human Liver Proteome Project (HLPP), 962 Human Proteome Organization (HUPO), 962 human telomerase-associated protein 1 (hTEP1), 1113 humoral immunity HBV infection, 839–840 HCV persistence, 881–882 see also antibodies; B cells huntingtin, 958 Huntington’s disease, 958 hyaluronans (HAs), normal liver ECM, 455 hyaluronic acid, 460, 462 hydrogels, PEG-based, implantable constructs, 942, 943, 944 hydrogen peroxide, 511 formation, 751 stellate cell transdifferentiation, 411, 412 hydrogen sulfide, vascular resistance in cirrhosis, 707 hydrolases, lysosomal, 176 hydrophilic substances, calcium signaling initiation, 485 hydrophobic substances, calcium signaling initiation, 485 3b-hydroxy-C27-steroid oxidoreductase deficiency, 663t hydroxyethyl radicals, 753–754 7α-hydroxylation, bile acid synthesis, 280, 281, 290 hydroxyl radical, 511 formation, 515, 751 4-hydroxynonenal (HNE), 744, 755, 794 hyperammonemia, 602, 609

hyperbilirubinemia, 251, 253, 254 conjugated, 251, 253, 663 unconjugated, 252 hypercholanemia, familial, 667t, 669–670 hypercholesterolemia, 323 hypercoagulation, 649 hyperdynamic circulation, 708 , 708–712 hepatorenal syndrome, 620 hyperfibrinolytic state, 650 hyperglucagonism, 709 hyperglycemia, 478 fasting, 477–478 mTORC1 hyperactivation, 544 hyperinsulinemia, 260–261, 473, 720 hyperlipidemia, drugs treating, 281–283 hypertonic saline, 611 hypertriglyceridemia CB1 receptor role, 1098 defenestration leading to, 393 mechanism, in insulin resistance, 475, 475 hyperventilation, 611 hypofibrinogenemia, 647 hyponatremia, 609–610 hypothermia, mild, induction, 611 hypoxanthine phosphoribosyl transferase, gene replacement, 976 hypoxia drug–inflammatory stress interaction, 777 hepatocyte, 377 hepcidin expression induction, 240–241 hypoxia inducible factor 1 (HIF-1), 416, 516 hypoxia inducible factor-1α (HIF-1α), 143 macrophage in adipose tissue, 722 , 723 in NAFLD and renal disease, 627 hypoxia inducible factor-2α (HIF-2α), in NAFLD and renal disease, 627

I ibuprofen, 611 ICER (inducible cAMP early repressor), 523 , 526 expression, liver regeneration, 528–529, 530 inducibility and functions, 526, 528 IDEAL trial, 907 idiopathic copper toxicosis, 229 IDN, 6556 796 IFT88 protein, 52 IκB, 555, 750, 791, 795, 878 IκB kinase complex (IKK), 555, 722, 791 IKKα (IKK1) and IKKβ (IKK2), 791 IKKγ (NEMO), 791, 792 inhibition, 732 as therapeutic target in NAFLD, 731–732 IL-6/STAT3 pathway see JAK/STAT pathway ileal bile acid binding protein (IBABP), 325, 329 ileum, bile acid transport system, 294 defects, 298, 299 imaging techniques, 1053–1066 acquisition modes, 1057 diffraction, 1055–1056, 1056 fluorescence resonance energy transfer (FRET), 1062 fluorescent dyes/proteins advances, 1053–1055 future prospects, 1064

1167 high-speed, 1062–1064 high-speed photomanipulation, 1062–1064 PALM and STORM, 1058, 1060, 1060 , 1061 , 1064 resolution, 1055, 1055 resolution increase, techniques, 1058 S/N ratio see signal-to-noise (S/N) ratio STED (stimulated emission depletion), 1058, 1060–1062, 1064 super-resolution see super-resolution microscopy see also microscopy imatinib, 1081 imino sugars, 913 immune response/system adaptive see antibodies; B cells; T cells HBV infections see hepatitis B virus (HBV) HCV infections see hepatitis C virus (HCV) innate see innate immune response liver-centered, 10 siRNA–HSP47 antifibrotic effect, 1126 viral infections, 835–836, 839–840, 882 immune tolerance, 944 to adenovirus vector antigens, 973 HBeAg role, 863 sinusoidal endothelial cell role, 376–377, 394 T cell, liver role, 840, 841 immunization see vaccination; vaccine immunocytes, of liver, 10 immunoglobulin A (IgA), HuHAVcr-1 (HCV) receptor, 808–809 immunomagnetic isolation, sinusoidal endothelial cells, 381 immunosuppressive mediators, sinusoidal endothelial cells, 377 immunosuppressive therapy, liver transplant in HCV infection, 925–926 immunotherapy, cancer, 966 IMP-1 (RNA-binding protein), 1044 implantable therapeutic constructs see under tissue engineering indomethacin, 611 “induced proximity model”, 783 infantile cholestasis, 662, 663t infantile cholestatic syndromes, 661–662 infants, liver development, 5 infections hepatic, effect on coagulation, 645–646 hepatic encephalopathy pathogenesis, 609 see also specific hepatitis viruses inferior vena cava, blood pressure in, 6 inflammation, 773–781, 774 definition, 773 drugs/xenobiotics associated, 774–776 see also inflammatory stress extracellular matrix, 454t, 455–458 hepatic encephalopathy pathogenesis, 609 management approaches, 611 hepcidin expression induction, 240–241 Kupffer cell role, 438 liver damage mechanism, 748, 749 NFκB activation, 792–793 persistent, lymphocyte accumulation, 748 stellate cell role, 437 inflammatory cell recruitment, ECM, 456, 457

1168 inflammatory mediators, 774, 775 TNF-α role, 749–750, 774, 777 inflammatory response, 773, 774 acute and chronic, 773 adipocytes, NASH and, 729–730 homeostasis alterations due to, 773 macrophage in adipose tissue, 722 , 723 inflammatory stress, 774 , 775, 775 drug hepatotoxicity, 774–775, 775 hemostatic system activation, 774, 777–778 idiosyncratic adverse drug reactions, 776 animal models, 776–777 mechanisms, 777–778, 778 influenza virus, 64 inhibitor of apoptosis (IAP), 750, 790 inhibitor of DNA binding 2 (ID2), 1004, 1005 ink4 family, 1017–1018 innate immune response activation, inflammation, 773 HBV infection, 836 , 836–837 HCV infection, 841–842, 877 evasion mechanisms, 841–842, 878 , 878–880 liver role, 551 in liver regeneration initiation, 552, 553 Toll-like receptor as activator, 554 signaling pathways, 841–842, 878 , 878–880 innexin proteins, 203 inosine monophosphate dehydrogenase (IMPDH), 906 inositol 1,4,5-triphosphate (IP3 ) Ca2+ release into nucleus, 495 calcium signaling, 486, 487 in cholangiocyte cilia, 361, 500 see also calcium signaling gap junction-mediated signaling, 207–208, 494 mitochondrial sensitivity heterogeneity, 491 inositol 1,4,5-triphosphate (IP3 ) receptor, 141, 486–489 binding affinity of InsP3 to, 488 calcium signaling, 486, 487 , 495 Ca2+ waves, 493 mechanism, 486–489 see also calcium signaling isoforms, 487 cells expressing, 488–489 cholangiocytes, 493–494 novel proteins interacting, 488 nuclear envelope, 495 phosphorylation, 488 regulation, 493, 495 structure and domains, 487 inositol-requiring protein 1 (IRE1), 265, 728, 788 insulin CYP7A1 regulation, 333 gluconeogenesis regulation, 478 glucose uptake, 720 lipid formation/storage, 720 macroautophagy regulation, 180–181 mechanism of action, 471, 474 , 720 in muscle, 471, 474 resistance see insulin resistance sensitivity AMPK regulating, 543 , 543–544 ER stress reducing, 727, 727 increased by resveratrol, 537, 539

INDEX PPARγ role, 263, 726–727 triglyceride storage affecting, 262 sensitizing agents, 478–480 adiponectin, 625 signaling alcoholic liver disease, 746 AMPK regulation of, 542–544 block, hepatic insulin resistance, 475 lipogenic inducer, 261 NAFLD, 722, 722 , 726 , 726–727 as therapeutic target in NAFLD, 731–732 insulin-induced genes, Insig-1 and Insig-2, 627, 628 insulin-like growth factor (IGF) IGF type 1, activated stellate cells, 436 IGF type, 2 1034 insulin-like growth factor binding protein (IGFBP), hepatocellular carcinoma, 1004 insulin-like growth factor binding protein 1 (IGFBP1), 788 insulin receptor substrate (IRS) function, model, 261, 261 IRS1, 261 IRS1, activation/phosphorylation, 471, 474 , 544, 722, 722 , 746 impairment, insulin resistance, 471, 472, 544 JNK signaling and, 722, 722 , 727 PI3 kinase activation, GLUT4 increase, 471, 474 protein kinase C θ activating, 473 IRS2, 261 block of PI3 kinase, insulin resistance, 476, 544 phosphorylation, 544 phosphorylation, mTORC1 role, 544 signaling, 261 insulin resistance, 257, 261, 471–483, 544, 720 adipose fat storage and, 261–263, 262 , 723 alcoholic liver disease, 745–746 chronic hepatitis C outcome, 886 endocannabinoids mediating, CB1 /CB2 receptors, 1098 endotoxin link, 722, 722 enhanced glucose production, 261, 477–478 ER stress-induced, 727, 727 evolutionary perspective, 719–720 fasting hyperglycemia, 261, 477–478 fat-induced in muscle, mechanism, 471–473, 474 intramyocellular lipid accumulation, 472 lipid accumulation in muscle, 472–473 muscle mitochondrial impairment, 472–473 Randle’s hypothesis, 471–472 hepatic, management, 478–480 obesity and, 745 hepatic, mechanisms, 475–477, 624, 624 , 745 acute high-fat feeding, 476 diacylglycerol role, 262, 476 fatless mouse, 476 fatty acid-induced, 262, 262 , 726, 726 FoxO role, 727 insulin signaling block, 261, 475 LPL over-expressing mouse, 475

in NAFLD, 475–477, 722, 722 , 726–727 PKCε role, 476–477 steatosis link to resistance, 476–477, 722, 722 , 726–727 hepatic steatosis worsening, 726–727 increased de novo lipogenesis, 473, 475, 475 intestinal mechanisms, gut microbiota and, 722, 722 macroautophagy defect and, 184 in NAFLD see non-alcoholic fatty liver disease (NAFLD) partial, 726 peripheral tissues (muscle/fat), 471 hepatic steatosis association, 473–475, 474 , 475 , 726–727 hypertriglyceridemia mechanism, 475, 475 increased hepatic triglycerides, 475, 475 reduction AMPK activation, 544 CB1 receptor antagonists, 1098 metformin, 478–479, 544 Sirt1 activation, 733 thiazolidinediones, 479, 726–727 weight loss, 479–480 renal, cardiovascular and liver complications, pathogenesis, 624, 624 saturated fatty acid role, 262, 262 TNFα activity, 745 TRL4, endotoxin and inflammation leading to, 722, 722 VLDL secretion, 261 integrative systems genomics, HCC, 994–995 integrins, 796 activated stellate cells expressing, 437 β1 integrins liver development role, 92 polarity mediator, 88t, 92 cytoprotective effects, 796 maturation modification of ECM, 462 integromics, 995 intercellular adhesion complexes, 9 intercellular adhesion molecule-1 (ICAM-1), 748 interferon(s) (IFNs), 905 antiviral effectors produced, 905 receptors, 905 synthesis, signaling pathways, 841–842, 878 , 878–880, 905 disruption in HCV immune evasion, 841, 878 , 878–880 type I early HBV infection, 837 mechanisms of action, 905 interferon α (IFNα), 905 activity enhanced by ribavirin, 906 albumin-IFN-α2b, 909 antifibrotic effects, 443 chemokine upregulation, 905 HBV therapy, 861, 866, 904 HCV susceptibility/resistance, 819 HCV therapy, 888, 906–907 novel forms of IFN-α, 909 optimization, 907–909 immunomodulatory properties, 905 mechanism of actions, 905 pegylated, 905–906 HBV therapy, 866, 904

INDEX HCV combination therapy, 907–909, 911, 912 HCV therapy, 904–905, 906–907, 907–909 HCV therapy, post-transplant, 924 HCV therapy and rejection after transplant, 924–925 pegylated IFN-α2a (Pegasys), 905 pegylated IFN-α2b (PegIntron), 906 receptors (IFNAR-1 and IFNAR-2), 905 signaling by, 878, 878 , 879, 880, 905 siRNA–HSP47 antifibrotic effect not involving, 1126 synthesis induction in HCV infection, 841, 878, 878 , 879, 880 interferon β (IFNβ) HCV HS3–4A protease suppressing, 819–820 synthesis, HCV infection, 841, 878, 878 Toll-interleukin1 receptor domain-containing adapter protein inducing (TRIF; TICAM-1), 879 interferon β (IFNβ)-promoter stimulator (IPS-1), 841 interferon γ (IFNγ) acute HCV infection, 842–843, 848 chronic HCV infection, 844, 880 HBV infection, 837 interferon regulatory factor 3 (IRF3), 809, 841, 878, 878 interferon response elements (ISREs), HCV infection, 841, 878, 878 , 879 interferon sensitivity-determining region (ISDR), 819 interferon-stimulated genes (ISGs) HCV infection, 841, 878, 878 , 879 regulation by IFN induction, 880, 905 ISGF3, 841, 905 interleukin-1β (IL-1β), oatps/OATPs regulation, 688 interleukin-1 receptor-associated kinase (IRAK), 442 interleukin-2 (IL-2), deficiency, HCV infection, 842–843 interleukin-6 (IL-6) cytoprotective effects, 795 priming phase of liver regeneration, 554, 555–556 knockout mice, 555–556 production reduced by Myd88 deficiency, 554 produced by stellate cells, 410 released by adipose cells, and effects, 624t NAFLD, 723, 724 signaling by JAK/STAT pathway, 556, 556 regulation by SOCS, 556 , 556–557 transcriptional regulation of hepcidin, 240, 241 interleukin-10 (IL-10), 774 hepatic fibrogenesis, 438 produced by stellate cells, 410, 438 interleukin-12 (IL-12), 749 interleukin-13 (IL-13), 774 interleukin-15 (IL-15), 749 intermediate density lipoprotein (IDL), 272t, 276, 277 intermediate filament, 29, 45, 150 internal ribosome entry site (IRES), HCV, 816, 817, 819, 910, 911 intersectin, 110 interstitial matrix, 453

see also extracellular matrix (ECM) intestinal microbiota see gut microbiota intestine bile acid conservation, 294–295 bile acid secretion, 294, 296 insulin resistance development, 721–722, 722 “leaky”, 744 see also ileum intra-cellular trafficking, 48, 48 , 49 epithelial cells, 35 Vps18 role in zebrafish, 1071 see also vesicle transport/trafficking intracellular vaccination, 967 intracranial hypertension, 603–604, 605, 607–608 intraflagellar transport (IFT), 49, 52 intrahepatic biliary hypoplasia see Alagille syndrome intrahepatic cholestasis see cholestasis/cholestatic liver disease intrahepatic cholestasis of pregnancy (ICP), 663 Intralipid, 472 introns, miRNA located in, 1032 Invagination model, peroxisome protein translocation, 197 INVERSIN mutations, 662 in vitro hepatic culture models see culture, hepatic ion pumps, 57–71 see also ATP synthase; P-type ATPases; V-type ATPases ion transport P-type ATPases, 59–60 V-type ATPases, 63, 66–67 ion transporters canalicular, 693 regulation in cholestasis, 687, 693 Ire1, 167 IRE1 (inositol-requiring protein, 1) 265, 728, 788 IREs (iron responsive elements), 237, 237 , 239, 265 iron, 235–250 chelatable, 512 cytotoxicity after oxidative stress, 515, 515 , 516 mitochondrial uptake, 515, 516 oxidative stress and cell death, 511–512, 512 release from lysosomal/endosomal compartment, 513–515 chelators, 512, 513 depletion in macrophages, genes associated, 243 excess, toxicity, 512 free, 512 free radical generation by, 511, 512, 512 , 515, 517 homeostasis, 235, 242 , 754 regulation by hepcidin, 241–242, 242 , 754–755 signaling pathways, 240–241 transcriptional regulation of hepcidin for, 240–241 by transferrin receptor trafficking, 114–115 see also hepcidin hydroxyl radical formation, two-hit hypothesis, 515–516, 517 intracellular movement, 511–520 loading of cells, TfR role, 237

1169 metabolites, toxicity, 754–755 non-chelatable, 512 non-transferrin-bound (NTBI), 237–239 overload, 237–239 alcohol consumption and, 754 ferroportin disease, 244 hepatic, alcoholic liver disease and, 245 transfusional, 239 see also hemochromatosis oxidization, 236 release from hepatic cells, 239 release from macrophage/enterocytes, 235, 241–242 release into cytosol, 512–513 sensing, 240–241 mutations, hemochromatosis due to, 242–243 proteins involved, 240–241 storage/accumulation, 754 Kupffer cells, 239, 754 in liver, 235–236, 243, 754 macrophages, 239, 242–243, 245, 754 synergistic action with alcohol, 754 toxicity mechanism, 754 transferrin-bound, 236, 238 see also transferrin (Tf); transferrin receptors (TfR) transporter, 512 DMT1, 236, 239, 242, 512 uptake, 236 decrease, ferroportin degradation, 242 non-transferrin-bound, 237–239 receptor-mediated endocytosis, 512–513 reduced in anemia of chronic disease, 245 iron oxidase, 239 iron protoporphyrin (heme), 862 iron-regulatory proteins (IRPs), 237, 238 iron responsive elements (IREs), 237, 237 , 239, 265 ischemic-reperfusion injury, 380, 1100 iso (3β-hydroxy) bile acids, 295 ITMN-191, 911, 912

J JACOP (junction associated coiled-coil protein), 87t Jagged1 (JAG1) gene, 20, 666, 668 JAGGED1 mutations, 666, 1070 JAGGED1 signaling, liver development, 20, 1070 JAGGED/NOTCH2 , 662, 666, 668 JAGI, 666 JAK/STAT pathway hepcidin regulation, 241 IL-6 signaling, liver regeneration, 556, 556 innate immunity in HCV infection, 841, 878, 878 , 879 regulation by SOCS, 556 , 556–557 JAMA (junction adhesion molecule), 86t, 91 JAMC, 86t, 91 Janus kinases (JAKs), 556 HCV interference, 841 hepatic fibrosis, 442 Jak1, interferon type I signaling, 905 jaundice, 215 cholestatic, 661

1170 JNK (c-Jun N-terminal kinase) activation in apoptosis, 794 TNFR signaling, 750, 790–791, 791 cyclin D1 expression control, 1020 hepatic fibrosis, 441, 441 intrinsic apoptosis in cholestasis, 794 IRS activation, insulin resistance, 722, 722 , 727, 731 SOCS role in signaling, 726 as therapeutic target in NAFLD, 730, 731–732 JNK/c-Jun cascade, CYP7A1 regulation, 297, 333 jumonji gene expression, 21 junctional adhesion molecules (JAMs), 84, 205–206 JAM-A, virus binding/uptake, 217 in tight junctions, 204 trafficking, 216 junction proteins polarity establishment, 84, 85t–88t, 91 see also adherens junctions (AJ); gap junctions; tight junctions (TJ) jun oncogene, 529 JWH-133, CB2 agonist, 1100

K karyopherins (kaps), 149, 150, 151 Kayser-Fleischer rings, 228 KDEL receptor, 126 Keap1, 1133–1134, 1134 mutant, 1134–1135 regulatory regions, 1134 turnover rate and Nrf2 ubiquitination, 1134–1135 Kelch domain, Keap1, 1134 keratin 8 mutants, 21 kernicterus, 251, 252 ketogenesis, 260 α-ketoglutarate dehydrogenase, 604 ketone bodies, formation, 258, 260 KFERQ, 182 K+ gradients, 58 kidney in liver disease, 619–638, 620t acute renal injury/failure, 619 hepatorenal syndrome see hepatorenal syndrome impaired renal function, 619 in NAFLD see non-alcoholic fatty liver disease (NAFLD) viral infections affecting, 622–624 Kiernan lobule, 7–8 kinase inducible domain, 524 kinesins, 32–33, 33–34, 37t, 45, 49–50 catalytic head domain (catalytic core), 33, 49 direction of transport by, 33, 38, 49 discovery, 32–33 double-headed, 33–34 in epithelial cells, 37–38, 49–50 functions, 33, 46, 49 inhibition by AMP-PNP, 50, 51 KIF1A, 34 KIF1B (kinesin, 3) 50 KIF3A (kinesin, 2) 49–50, 51, 52 KIF3B, 49 KIF5 (kinesin 1 family), 49 KIF5B, 33, 37t, 38, 49, 51 properties, 47t KIF17, 49–50 KIFC1 and KIFC2 (kinesin, 14) 33, 37t, 38, 50, 51

INDEX KIF genes, 49 knockout mutant, 52 light chains (KLC), 49 mitochondrial distribution, role, 141 structure, 33, 49 kinetochores, 46 KIR 2DL3, 842 knockout (KO) animals see specific genes/functions Kruppel-like transcription factors, 413t Kupffer cells, 375, 375 activation endotoxin role, 749 in inflammation, 773–774 chelatable iron, effect, 516 CYP2E1 expression, 743 DNA replication, 551 free radical formation, 751 function, 10, 376 hepatic fibrosis and, 749 iron release from, hepcidin role, 239 iron storage, 239, 754 liver inflammation, role, 438 mediators released, 753 phagocytosis, 179 reactive oxygen species formation, 753 structure, 10 see also macrophage(s)

L “label-retaining cells”, BrdU, stem cells, 583 lacrimal gland acinar cells, 34, 35, 36, 38 lactate, brain accumulation, 604, 605 lactitol, 610 lactoferrin, 756 lactulose, 610 lacunae, 7 lacZ plasmid system, 978, 979 lamellipodia, 34 laminin distribution in liver, antibody staining, 455, 456 ECM and bile duct epithelium development, 454 lamins, 150 lamivudine (Epivir), 866, 901, 902t, 903 combination therapy in hepatitis B, 904 HBIG with, 927–928 post-transplant, 927 HBV resistance, 861, 868 pre-transplant in hepatitis B, 927, 928 resistance to, 861, 868, 903, 927 latrunculin, fenestrae regulation, 395t, 396 LC8 protein, 48 LCMV model, T cell regulation, 847, 882 LDL receptor-related protein (LRP), 277 Leber hereditary optic neuropathy, 139 lecithin:cholesterol acyltransferase (LCAT), 278 lectin-based chaperones, 162, 163 lentiviruses, gene therapy, 969–970 leptin, 415 AMPK activation, 537, 539 hepatic fibrosis pathogenesis, 439, 745, 746 as mitogen for stellate cells, 437, 746 released by adipose cells, and effects, 624t resistance, endocannabinoids mediating via CB1 receptors, 1098 stellate cells expressing, 744, 746

stellate cells transdifferentiation, 412, 436, 437 TIMP1 up-regulation, 746 leptospiral protein (Lsa21), 456 Let-7a family, 1037–1039 fetal liver and hepatocyte differentiation, 1039 , 1040 hepatocarcinogenesis, 1044 Let-7/let-7 gene, 1029, 1032 Lethal Giant Larvae (LGL) protein, 84, 85t leucine-zipper domain CREB, 523 transcription factors in xenobiotic detoxification, 1135, 1135 see also bZIP transcription factors leukemia acute lymphoid (ALL), 1082 acute myeloid (AML), 1081 chronic myeloid (CML), 1081 nucleoporins (Nups) role, 152 leukemia inhibitory factor (LIF) receptor, 557 leukocytes see individual leukocyte types levofloxacin, 776 Lhx2, LIM -homeobox gene, 21 lifespan of liver cells, 11 LIF receptor, 557 ligand/receptor-mediated delivery, gene therapy, 973–974 LIM domains, 525 lipase hepatic, 277, 280 hormone-sensitive (HSL), 258, 723, 723 lipoprotein see lipoprotein lipase (LPL) monoglyceride (MGL), 1093 lipid(s), 271 absorption, 273 gut microbiota role, 721 , 721–722 accumulation in liver, 721–727 see also under non-alcoholic fatty liver disease (NAFLD) accumulation in muscle, mechanism, 472–473 biliary secretion, 280, 298 in caveolae, 113 endocytosis regulation, 114 fate of “old” (stored) vs “new” (dietary-derived) fat, 264 , 264–265 insulin resistance and, 261–263, 262 intrahepatic, reduced by weight loss, 479–480 intramyocellular accumulation, 471, 474 insulin resistance due to, 472, 473 insulin resistance vs insulin sensitivity, 472 shifting to adipocytes, thiazolidinedione action, 479 metabolism, 257–270, 744 adipose tissue role, 262 alcoholic liver disease, 744–745 AMPK regulating, 539, 543–544 biosynthesis vs oxidation balance, 263–264, 725, 725 defects, PED-6 for screening, 1072 gene expression, after PH, 560 HIF-1α role, 627 insulin role, 720 mTOR effects on, 543–544 unfolded protein response relationship, 265 , 265–266 see also AMPK (AMP-activated protein kinase); lipogenesis

INDEX oxidation, 472, 725 , 725–726 see also fatty acid(s), oxidation peroxidation, 728, 743, 754 alcoholic liver disease, 754 end products, 755 hepatic ischemic–reperfusion injury, 1100 regulation of protein trafficking, 114, 131 storage adipocyte size/number increase, 723 adipose fat, insulin resistance and, 261–263, 262 gut microbiota role, 721 , 721–722 insulin role, 720 PPArγ role/functions, 263 translocator types in canalicular membrane, 341–342, 342 utilization during fasting, AMPK role, 542 vesicle biogenesis in secretory pathway, 131 see also fatty acid(s); specific fatty acids; specific lipids lipid-based delivery systems, gene therapy, 973, 974 lipid bilayers, membranes, 343–344 lipid droplets (LDs), 258 biogenesis, 259, 259 storage of triglycerides, 258–259 structure/composition, 258 lipid peroxides, stellate cell activation, 436 lipid peroxyl radical, 754 “lipid rafts”, 82, 113, 343–344 lipin, 258 lipogenesis enzymes, regulation, 261, 541 by AMPK, 538–539, 539 increased CB1 receptor activation, 1096 , 1096–1098 in insulin resistance, 473, 475, 475 inhibition by AMPK, 541 regulation/regulators, 261, 265, 265 transcriptional control, 261, 629, 631 unfolded protein response (UPR) and, 265 , 265–266 lipolysis, 258, 259, 725 apo-B-containing lipoproteins, 275 , 275–276 lipoprotein lipase-mediated, 275 , 275–276, 278, 721 rates, 276 lipopolyplex delivery systems, gene therapy, 973 lipopolysaccharide (LPS) (endotoxin), 645 alcoholic liver disease, 749–750 coagulation increased, 777 effects/actions, 750 gut microbiota and hepatic fat accumulation, 722, 722 hepatic microcirculation changes, 750 inflammation associated, 749, 773 insulin resistance pathogenesis and, 722, 722 liver damage, 456, 774 liver regeneration, 554 oxidant formation by NADPH oxidase, 751 signaling system, 555 TNFα elevation, 749, 759, 777, 778 lipopolysaccharide binding protein (LBP), 750

lipoprotein(s), 271–285 apo-B containing, 272–277 assembly, 273–275, 274 intravascular metabolism, 275 , 275–276 receptor-mediated clearance, 276 , 276–277 classes, and characteristics, 271, 272t fenestrations regulating deposition, 376, 393 high-density see high-density lipoprotein (HDL) intermediate-density, 272t, 276, 277 lipid membrane, 271–272 low-density see low-density lipoprotein (LDL) metabolism, 271–281 endogenous pathway, 273, 274 , 275 exogenous pathway, 273, 274 remnant receptor pathways, 276 , 276–277 structure, 271–272, 272 VLDL see very low-density lipoproteins (VLDL) lipoprotein lipase (LPL), 275 , 275–276, 278, 721 functions, 722–723, 723 increased activity, insulin resistance, 472, 721 overexpression (mice), insulin signaling block, 475 liposomes, vitamin A-coupled see vitamin A-coupled liposomes lipotoxicity, 262 5-lipoxygenase (5-LO), 707–708 LIRKO mouse, 261 lithocholic acid (LCA), 292, 294, 295 calcium signaling induced by, 498 detoxification, 295 “littoral cells”, DNA replication, 551 liver anatomy, 5–6 architecture see liver organization as ecosystem, 407 elasticity, 455, 455 functions, 3, 4, 11, 17, 935–936 mass/volume, 5, 6 size, 4, 5 regulator, YAP, 578 weight, 5 liver biopsy HCC, 871 recurrent HCV infection, 923 liver bud, 4, 19 formation from hepatic endoderm, 4, 19–20, 584, 584 liver cancer see hepatocellular carcinoma (HCC) liver/cell engineering see tissue engineering liver cells, 3, 9–10 lifespan, 11 proliferation during “regeneration”, 12 in repair of liver, 11, 12 structure, 9–10 turnover, 11 types see individual cell types liver disease, kidney disease in see kidney in liver disease liver-enriched transcriptional activator protein (LAP), 478 liver-enriched transcriptional inhibitory protein (LIP), 478

1171 liver failure acute see acute liver failure (ALF) ammonia and glutamine trafficking, 602, 603 cell-based therapies, 935–937 chronic, 935 consequences, 602 fulminant, 588, 935 hyperammonemia, 602–603 mortality, 935 subacute, 457 liver fibrosis see hepatic fibrosis liver injury acute coagulation factors reduced, 646 healing and regeneration, 550 alcohol-induced epigenetic changes, 759 mechanisms, 744 inflammation associated, 748, 749 ischemic-reperfusion injury, 380, 1100 “oval cells” proliferation induction, 581 stellate cell transdifferentiation, 411 liver irradiation, hepatocyte transplantation model, 580–581 liver kinase B1 see LKB1 (liver kinase B1) liver organization, 3–15, 74 , 74–75 functional/structural heterogeneity, 10–11 liver cells, 3, 9–10, 74–75 principles, 3–4 zonation, 11 see also hemodynamics, liver liver perfusion, impaired, adaptation, 380–381 liver receptor homolog-1 (LRH-1; FTF), 325–327, 686t in cholestasis, 685 CYP7A1 regulation, 326–327 mechanism, and FXR–SHP–FTF cascade, 297, 328–329, 329 liver regeneration, 12, 549–565, 553 accelerated by TNF, 552, 553 after repeated hepatectomies, 550, 551 at all developmental stages, 22 basic requirements, 550, 578 bone marrow cells, role, 586–587 calcium signals role in cell proliferation, 499 cAMP role in cell proliferation, 528–529, 529 , 530 capacity/extent, 550, 560 cell cycle control and cyclins, 1019–1021 cell transplantation, 936 as compensatory growth process, 550 DNA replication, 550, 551–552, 558, 559 ECM remodeling and, 462–463 embryologic control, 21–22 gap junction expression, 208, 210 Cx32 mutants and, 210 gene expression, global patterns, 560 with hepatocyte hyperplasia, 11, 550 hepatocyte ploidy, 551–552 historical background, 549 inhibition, NK and NKT cells, 551 liver cell transplantation, 578, 935 Ntcp expression, bile acid uptake and, 311 organic anion transport, by oatp1a1, 314–315 parenchymal regeneration, 11–12

1172 liver regeneration (continued) priming phase (G0 to G1), 552–553, 553 , 560, 1019 cyclin gene up-regulation, 1019, 1020 cytokines regulating, 407, 552–553, 553 , 553–557, 560 IL-6 system, 555–556 NFκB system, 555, 555 termination by SOCS3, 557, 557 TNF and lymphotoxins, 553, 554, 554 toll-like receptor system, 554 transcription factor activation, 553, 555, 556, 560 transition to progression phase, 557, 559 progression phase (G1, S, G2), 552, 557–560 EGFR ligands, 559 growth factors regulating, 407, 552, 553, 561 HGF role, 552, 558, 561 TGF-β inhibiting replication, 559 research goal, 1021 ribosomal biogenesis role, 567, 571–573, 572 size/number of lobules, 551 stages, 552–553 stem cell role, 1021 telomere shortening and, 1107–1108 time course, 550, 560, 578 transcription factors, global expression, 560, 1020 trigger, portal blood flow, 12 weight regained, 550, 578 without hepatocyte hyperplasia, 552 zebrafish model, 1071 liver repopulation, 577–595 animal models, 579 , 579–581, 580 experimental conditions, 580–581 by extrahepatic/embryonic stem cells, 585–586 by fetal liver stem/progenitor cells, 584–585 by hepatocyte transplant see hepatocyte transplantation induced pluripotent stem cells and, 586 by “oval cells” (rats), 581–582 xenorepopulation models, 558 , 588 see also liver regeneration; stem cells “liver sieve”, 390, 393 liver-specific insulin receptor knockout (LIRKO) mouse, 261 liver stem cells see stem cells liver transplantation cell transplantation see hepatocyte transplantation donor shortage, 935 hepatic fibrosis management, 443 in hepatocellular carcinoma, 1000 in hepatorenal syndrome, 620–621 large livers, size reduction after, 550, 578 liver cell repletion, 11 in liver failure, 935 living donors, 228 non-alcoholic steatohepatitis recurrence, 720 partial, 935 reperfusion injury, sinusoidal endothelial cell damage, 380 small for size graft, 380, 550, 578 for virus-induced end-stage liver disease, 921–931

INDEX HBV, 926–928 HCV, 921–926 see also hepatitis B; hepatitis C Wilson disease, 228 liver X receptor (LXR) see LXR (liver X receptor) LKB1 (liver kinase B1) AMPK regulation, 537–538, 539 , 540, 540 , 545 mouse models, 541t polarity mediator, 88t, 92, 93 transcription regulation, 540, 540 LLC-PK1 cell line, plasma membrane protein/traffic research, 76, 77t lobes of liver, 6, 550 lobules, liver, 4, 7, 74 “hexagonal”, 8–9 increase in size/number during regeneration, 551 primary, 8 losartan, 730 low-density lipoprotein (LDL) apoB100, 277, 277 characteristics, 272t cholesterol decrease by bile acid transport inhibitors, 300 by fibrates or niacin, 282 by plant sterols/stanols and ezetimibe, 282 clearance from plasma, 277 formation, 277, 277 half-life, 277 receptor-mediated endocytosis, 277 low-density lipoprotein (LDL) receptor (LDLr), 277 biosynthetic pathway, 83 defect, hepatocyte transplantation for, 587 HCV receptor/uptake, 116 LDL clearance, 277 upregulation, 282 low-density lipoprotein (LDL) receptor-related protein (LRP), 646, 648 low-phospholipid associated cholelithiasis (LPAC), 666 LRH-1 see liver receptor homolog-1 (LRH-1; FTF) LST-1 see OATP1B1 (LST-1;SLCO1B1 ) LXR (liver X receptor), 260, 323, 326, 686t actions, 325 bile acid transporter regulation in cholestasis, 685, 686t CYP7A1 regulation, 297, 326, 327 FXR interactions, 328, 328 dietary cholesterol effect on, 327 lipid/glucose homeostasis, role, 724 , 725 LXRα, 325, 327, 328 Sirt1 deactylation of, 957 lymph, hepatic, 6, 394 lymphedema–cholestasis syndrome, 668t, 671 lymphocyte(s) alcoholic liver disease, 748 recruitment, 748 role in hepatic fibrosis, 748 stellate cells transdifferentiation, 414–415, 438 structure, 10 see also B cells; T cells lymphocyte function-associated antigen-1 (LFA-1), 748–749

lymphocytic choriomeningitis virus (LCMV) model, 847, 882 lymphotoxins, 553 , 554 LTα and LTβ, 554, 554 lysophosphatidic acid (LPA), 258, 412 lysophosphatidylcholine, 278 lysosomal apyrase-like protein, 177 lysosomal integral membrane protein, LIMP1 and LIMP2, 177 lysosomal storage disorders (LSDs), 176–177 lysosomal system, 176–183 proteolytic pathways, 178–183 autophagic pathways see autophagy/autophagic pathways heterophagic pathways, 178–179 in liver disease and aging, 183 , 183–184 see also lysosome(s) lysosome(s), 176 accumulation of products in, 177 apoptosis pathway, steatosis progression to steatohepatitis, 728 autophagosome delivery, 180 breakdown, chelatable iron role, 513 chelatable iron release, 513–515 endosome maturation to, 179 enzymes, 176–177 defective, 176–177 location at membrane, 177 synthesis, tagging and sorting, 176 transport, clathrin-coated vesicles derived from TGN, 130 functions, 176 in hepatocytes, 74 , 74–75 membrane lipids, age-related changes, 185 membrane proteins, 177–178 oxidative stress effect, 513, 515, 517 secondary, 176, 181 targeting of cargo to, 112, 179, 182 transporters at membrane, 177, 182 V-type ATPases, 64, 177 lysosome-associated membrane proteins (LAMPs), 177 LAMP-1, 177 LAMP-2, 177 deficiency, 178 isoforms, 177, 182 LAMP-2A receptor changes with age, 177 lysosome-related compartments, 176, 179 see also endosomes lysyl hydroxylases, 461 lysyl oxidases, 461

M macroautophagy, 179–182, 180 age-related changes, 184–185 alpha-1-antitrypsin deficiency, 184 carcinogenesis and, 183 , 184 conjugation events/membrane formation, 170–180 defects in fatty liver, 183 , 184 functions, 179, 184 hormonal regulation (insulin), 180–181, 184 inhibition by HCV, 183 , 184 initiation complex, 170, 180 negative regulatory complex, 170, 180 role in intracellular quality control, 181–182

INDEX Sirt1 control, 957 in starvation, 182–183 macropexophagy, 197, 198 macrophage(s) activated increased in obesity, 626 , 626–627 activation, by NFκB, 792–793 in adipose tissue, inflammatory response, 722 , 723, 746 chemotaxis, osteopontin role, 457 endocannabinoid generation, 1093–1094 hepatic fibrosis and, 749 hepatic wound healing role, 438 inflammatory, activation, 626 , 626–627 iron depletion, genes associated, 243 iron release from, hepcidin role, 239, 2