The Liver: Biology and Pathobiology

The Liver The Liver Biology and Pathobiology Fifth Edition Editor Irwin M. Arias Senior Scientist, National Institut...

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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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2

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

20

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

22

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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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

32

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.

38

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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113. Schell, M.J., Maurice, M., Stieger, B. and Hubbard, A.L. (1992) 5 nucleotidase is sorted to the apical domain of hepatocytes via an indirect route. J Cell Biol , 119, 1173–82. 114. Ihrke, G., Neufeld, E.B., Meads, T., Shanks, M.R., Cassio, D., Laurent, M., Schroer, T.A., Pagano, R.E. and Hubbard, A.L. (1993) WIF-B cells: an in vitro model for studies of hepatocyte polarity. J Cell Biol , 123, 1761–75. 115. Wakabayashi, Y., Kipp, H. and Arias, I.M. (2006) Transporters on demand: intracellular reservoirs and cycling of bile canalicular ABC transporters. J Biol Chem, 281, 27669–73. 116. Berg, J.S., Powell, B.C. and Cheney, R.E. (2001) A millennial myosin census. Mol Biol Cell , 12, 780–94. 117. Stafford, W.F., Walker, M.L., Trinick, J.A. and Coluccio, L.M. (2005) Mammalian class I myosin, Myo1b, is monomeric and crosslinks actin ﬁlaments as determined by hydrodynamic studies and electron microscopy. Biophys J , 88, 384–91. 118. Balish, M.F., Moeller, E.F. III and Coluccio, L.M. (1999) Overlapping distribution of the 130- and 110-kDa myosin I isoforms on rat liver membranes. Arch Biochem Biophys, 370, 285–93. 119. Raposo, G., Cordonnier, M.N., Tenza, D., Menichi, B., Durrbach, A., Louvard, D. and Coudrier, E. (1999) Association of myosin I alpha with endosomes and lysosomes in mammalian cells. Mol Biol Cell , 10, 1477–94. 120. Salas-Cortes, L., Ye, F., Tenza, D., Wilhelm, C., Theos, A., Louvard, D., Raposo, G. and Coudrier, E. (2005) Myosin Ib modulates the morphology and the protein transport within multi-vesicular sorting endosomes. J Cell Sci , 118, 4823–32. 121. Valentijn, K., Valentijn, J.A. and Jamieson, J.D. (1999) Role of actin in regulated exocytosis and compensatory membrane retrieval: insights from an old acquaintance. Biochem Biophys Res Commun, 266, 652–61. 122. Chan, W., Calderon, G., Swift, A.L., Moseley, J., Li, S., Hosoya, H., Arias, I.M. and Ortiz, D.F. (2005) Myosin II regulatory light chain is required for trafﬁcking of bile salt export protein to the apical membrane in Madin–Darby canine kidney cells. J Biol Chem, 280, 23741–47. 123. 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. 124. Wu, X., Bowers, B., Rao, K., Wei, Q. and Hammer, J.A. III (1998) Visualization of melanosome dynamics within wild-type and dilute melanocytes suggests a paradigm for myosin V function in vivo. J Cell Biol , 143, 1899–918. 125. Takagishi, Y., Oda, S., Hayasaka, S., Dekker-Ohno, K., Shikata, T., Inouye, M. and Yamamura, H. (1996) The dilute-lethal (dl) gene attacks a Ca2+ store in the dendritic spine of Purkinje cells in mice. Neurosci Lett , 215, 169–72. 126. Jones, J.M., Huang, J.D., Mermall, V., Hamilton, B.A., Mooseker, M.S., Escayg, A., Copeland, N.G., Jenkins,

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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

4: MOLECULAR MOTORS

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

48

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

50

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

52

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.

REFERENCES 1. Satir, P. (2001) Motor molecules, in The Liver: Biology and Pathobiology, 4th edn, Chapter 4 (eds I.M. Arias et al.), Lippincott Williams & Wilkins, Philadelphia, pp. 47–58. 2. Novikoff, P., Cammer, M., Tao, L. et al. (1996) Three-dimensional organization of rat hepatocyte cytoskeleton: relation to the asialoglycoprotein endocytosis pathway. J Cell Sci , 109, 21–32. 3. Masyuk, A.I., Maysuk, T.V. and LaRusso, N.F. (2008) Cholangiocyte primary cilia in liver health and disease. Develop Dyn, 237, 2007–12. 4. Zhu, C., Zhao, J., Bibikova, M. et al. (2005) Functional analysis of human microtubule-based motor proteins, the kinesins and dyneins in mitosis/cytokinesis using RNA interference. Mol Biol Cell , 16, 3187–99.

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THE LIVER: REFERENCES

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25. Wang, H. and Satir, P. (1998) The 29 kDa light chain that regulates axonemal dynein activity binds to cytoplasmic dynein. Cell Motil Cytoskeleton, 39, 1–8. 26. Mizuno, N., Narita, A., Kon, T. et al. (2007) Three dimensional structure of cytoplasmic dynein bound to microtubules. Proc Natl Acad Sci USA, 104, 20832–37. 27. Howard, J. (1997) Molecular motors: structural adaptations to cellular functions. Nature, 389, 561–67. 28. Burgess, S.A., Walker, M.L., Sakakibara, H. et al. (2004) The structure of dynein-c by negative stain electron microscopy. J Struct Biol , 146, 205–16. 29. Imamula, K., Kon, T., Ohkura, R. and Sutoh, K. (2007) The coordination of cyclic microtubule association/dissociation and tail swing of cytoplasmic dynein. Proc Natl Acad Sci USA, 104, 16134–39. 30. Hirokawa, N. (1998) Kinesin and dynein superfamily proteins and the mechanism of organelle transport. Science, 279, 519–26. 31. Oleynikov, Y. and Singer, R.H. (1998) RNA localization: different zip codes, same postman? Trends Cell Biol, 8, 381–83. 32. Reck-Peterson, S.L., Yildiz, A., Carter, A.P. et al. (2006) Single molecule analysis of dynein processivity and stepping behavior. Cell , 126, 335–48. 33. Gennerich, A., Carter, A.P., Reck-Peterson, S.L. and Vale, R.D. (2007) Force-induced bidirectional stepping of cytoplasmic dynein. Cell , 131, 952–65. 34. Gill, S.R., Schroer, T.A., Szilak, I. et al. (1991) Dynactin, a conserved ubiquitously expressed component of an activator of vesicle motility mediated by cytoplasmic dynein. J Cell Biol , 115, 1639–50. 35. Schroer, T.A. (2004) Dynactin. Annu Rev Cell Dev Biol , 20, 759–79. 36. Holleran, E.A., Ligon, L.A., Tokito, M. et al. (2001) Beta III spectrin binds to the Arp1 subunit of dynactin. J Biol Chem, 276, 36598–605. 37. Muresan, V., Stankewich, M.C., Steffen, W. et al. (2001) Dynactin-dependent dynein-driven vesicle transport in the absence of membrane proteins: a role for spectrin and phospholipids. Mol Cell , 7, 173–83. 38. Paschal, B.M., Holzbaur, E.L., Pﬁster, K.K. et al. (1993) Characterization of a 50 kDa polypeptide in cytoplasmic dynein preparations reveals a complex with p150Glued and a novel actin. J Biol Chem, 268, 15318–23. 39. Burkhardt, J.K., Echeverri, C.J., Nilsson, T. et al. (1997) Overexpression of the dynamitin (p50) subunit of the dynactin complex disrupts dynein-dependent maintenance of membrane organelle distribution. J Cell Biol , 139, 469–84. 40. Melkonian, K.A., Maier, K.C., Godfrey, J.E. et al. (2007) Mechanism of dynamitin-mediated disruption of dynactin. J Biol Chem, 282, 19355–64. 41. Harada, A., Takai, Y., Kanai, Y. et al. (1998) Golgi vesiculation and lysosome dispersion in cells lacking cytoplasmic dynein. J Cell Biol , 141, 51–59. 42. Valletti, C., Wetzel, D.M., Schrader, M. et al. (1999) Role of dynactin in endocytic trafﬁc: effects of dynamitin overexpression and localization with CLIP-170. Mol Biol Cell , 10, 4107–20.

4: MOLECULAR MOTORS

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, g,A6L

V0

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

a

ATP

T L

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

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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

63

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

64

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

66

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),

75

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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THE LIVER: CELL SYSTEMS FOR STUDYING THE ITINERARIES OF PM PROTEINS

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.

6: HEPATOCYTE SURFACE POLARITY: ITS DYNAMIC MAINTENANCE AND ESTABLISHMENT

77

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.

78

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 -VAMP7

TGN DPP4 APN -VAMP8

79

2

DPP4 APN -VAMP7

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

80

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

BASOLATERAL

2

7

1 Multi-TMD: AQPS (g’on) -cAMP BSEP (TC) -Pl3-K -cAMP ATP7B (Cu1+)

Alb HA321 CE9

8

Multi-TMD: NPLC1

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 -PCSK9

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

82

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

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

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]

See above and [158, 238]

See above

[157, 231, 232]

[228]

[225]

[222]

[158, 217]

[214]

[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

Afadin (aka AF-6)

[205, 206]

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

Armadillo repeat motifs

[197]

Reviews

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

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

Role in polarity (reports)

Armadillo repeat motifs

Functional domains

β-catenin

Protein

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

Transcription factor

S/T kinase; farnesylated at C-terminus

S/T kinase heterotrimeric complex S/T kinase

HNF4α (hepatocyte nuclear factor)

LKB1 (aka PAR4)

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

SMADs (small mothers against deca-pentaplegic)

[300]

[301]

Regulator of translation; phosphorylates 4E-BP1 and p70 Sk kinase [307]; involved in neuronal polarity [308]

[299–302]

[290]

[285, 286]

[279–281]

[273]

[194, 258]

Reviews

Energy-dependent regulation of TJ assembly in MDCK cells [306]

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

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

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

91

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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THE LIVER: ESTABLISHING EPITHELIAL CELL POLARITY

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

6: HEPATOCYTE SURFACE POLARITY: ITS DYNAMIC MAINTENANCE AND ESTABLISHMENT

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

93

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

94

THE LIVER: REFERENCES

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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45. 46.

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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

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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

113

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

115

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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THE LIVER: REFERENCES

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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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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R ab 11

Basolateral Membrane

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,

131

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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67. Narula, N. and Stow, J.L. (1995) Distinct coated vesicles labeled for p200 bud from trans-Golgi network membranes. Proc Natl Acad Sci USA, 92, 2874–78. 68. Ikonen, E., de Almeid, J.B., Fath, K.R., Burgess, D.R., Ashman, K., Simons, K. and Stow, J.L. (1997) Myosin II is associated with Golgi membranes: identiﬁcation of p200 as nonmuscle myosin II on Golgi-derived vesicles. J Cell Sci , 110 (Pt 18), 2155–64. 69. Derby, M.C., van Vliet, C., Brown, D., Luke, M.R., Lu, L., Hong, W., Stow, J.L. and Gleeson, P.A. (2004) Mammalian GRIP domain proteins differ in their membrane binding properties and are recruited to distinct domains of the TGN. J Cell Sci , 117, 5865–74. 70. Gleeson, P.A., Anderson, T.J., Stow, J.L., Grifﬁths, G., Toh, B.H. and Matheson, F. (1996) p230 is associated with vesicles budding from the trans-Golgi network. J Cell Sci , 109 (Pt 12), 2811–21. 71. Derby, M.C., Lieu, Z.Z., Brown, D., Stow, J.L., Goud, B. and Gleeson, P.A. (2007) The trans-Golgi network golgin, GCC185, is required for endosome-to-Golgi transport and maintenance of Golgi structure. Trafﬁc, 8, 758–73. 72. Lieu, Z.Z., Derby, M.C., Teasdale, R.D., Hart, C., Gunn, P. and Gleeson, P.A. (2007) The golgin GCC88 is required for efﬁcient retrograde transport of cargo from the early endosomes to the trans-Golgi network. Mol Biol Cell , 18, 4979–91. 73. Henley, J.R. and McNiven, M.A. (1996) Association of a dynamin-like protein with the Golgi apparatus in mammalian cells. J Cell Biol , 133, 761–75. 74. Maier, O., Knoblich, M. and Westermann, P. (1996) Dynamin II binds to the trans-Golgi network. Biochem Biophys Res Commun, 223, 229–33. 75. Cao, H., Garcia, F. and McNiven, M.A. (1998) Differential distribution of dynamin isoforms in mammalian cells. Mol Biol Cell , 9, 2595–609. 76. Cao, H., Weller, S., Orth, J.D., Chen, J., Huang, B., Chen, J.L., Stamnes, M. and McNiven, M.A. (2005) Actin and Arf1-dependent recruitment of a cortactin-dynamin complex to the Golgi regulates post-Golgi transport. Nat Cell Biol , 7, 483–92. 77. Orth, J.D. and McNiven, M.A. (2003) Dynamin at the actin-membrane interface. Curr Opin Cell Biol , 15, 31–39. 78. Brown, H.A., Gutowski, S., Moomaw, C.R., Slaughter, C. and Sternweis, P.C. (1993) ADP-ribosylation factor, a small GTP-dependent regulatory protein, stimulates phospholipase D activity. Cell , 75, 1137–44. 79. Randazzo, P.A. and Kahn, R.A. (1994) GTP hydrolysis by ADP-ribosylation factor is dependent on both an ADP-ribosylation factor GTPase-activating protein and acid phospholipids. J Biol Chem, 269, 10758–63. 80. Ktistakis, N.T., Brown, H.A., Sternweis, P.C. and Roth, M.G. (1995) Phospholipase D is present on Golgi-enriched membranes and its activation by ADP ribosylation factor is sensitive to brefeldin A. Proc Natl Acad Sci USA, 92, 4952–56. 81. Ktistakis, N.T., Brown, H.A., Waters, M.G., Sternweis, P.C. and Roth, M.G. (1996) Evidence that phospholipase

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9

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

9: MITOCHONDRIA

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

9: MITOCHONDRIA

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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A. (2001) Does oxidative damage to DNA increase with age? Proc Natl Acad Sci U S A, 98, 10469–74. Kujoth, G.C., Hiona, A., Pugh, T.D., Someya, S., Panzer, K., Wohlgemuth, S.E., Hofer, T., Seo, A.Y., Sullivan, R., Jobling, W.A. et al. (2005) Mitochondrial DNA mutations, oxidative stress, and apoptosis in mammalian aging. Science, 309, 481–84. Trifunovic, A., Wredenberg, A., Falkenberg, M., Spelbrink, J.N., Rovio, A.T., Bruder, C.E., Bohlooly, Y.M., Gidlof, S., Oldfors, A., Wibom, R. et al. (2004) Premature ageing in mice expressing defective mitochondrial DNA polymerase. Nature, 429, 417–23. Orr, W.C. and Sohal, R.S. (1994) Extension of life-span by overexpression of superoxide dismutase and catalase in Drosophila melanogaster. Science, 263, 1128–30. Schriner, S.E., Linford, N.J., Martin, G.M., Treuting, P., Ogburn, C.E., Emond, M., Coskun, P.E., Ladiges, W., Wolf, N., Van Remmen, H. et al. (2005) Extension of murine life span by overexpression of catalase targeted to mitochondria. Science, 308, 1909–11. Martin, D.E. and Hall, M.N. (2005) The expanding TOR signaling network. Curr Opin Cell Biol , 17, 158–66. Warburg, O. (1956) On the origin of cancer cells. Science, 123, 309–14. Wang, G.L. and Semenza, G.L. (1993) General involvement of hypoxia-inducible factor 1 in transcriptional response to hypoxia. Proc Natl Acad Sci U S A, 90, 4304–8. Papandreou, I., Cairns, R.A., Fontana, L., Lim, A.L. and Denko, N.C. (2006) HIF-1 mediates adaptation to hypoxia by actively downregulating mitochondrial oxygen consumption. Cell Metab, 3, 187–97. Schulz, T.J., Thierbach, R., Voigt, A., Drewes, G., Mietzner, B., Steinberg, P., Pfeiffer, A.F. and Ristow, M. (2006) Induction of oxidative metabolism by mitochondrial frataxin inhibits cancer growth: Otto Warburg revisited. J Biol Chem, 281, 977–81. Matoba, S., Kang, J.G., Patino, W.D., Wragg, A., Boehm, M., Gavrilova, O., Hurley, P.J., Bunz, F. and Hwang, P.M. (2006) p53 regulates mitochondrial respiration. Science, 312, 1650–53. Bensaad, K., Tsuruta, A., Selak, M.A., Vidal, M.N., Nakano, K., Bartrons, R., Gottlieb, E. and Vousden, K.H. (2006) TIGAR, a p53-inducible regulator of glycolysis and apoptosis. Cell , 126, 107–20. DeBerardinis, R.J., Lum, J.J., Hatzivassiliou, G. and Thompson, C.B. (2008) The biology of cancer: metabolic reprogramming fuels cell growth and proliferation. Cell Metab, 7, 11–20. Mandal, S., Guptan, P., Owusu-Ansah, E. and Banerjee, U. (2005) Mitochondrial regulation of cell cycle progression during development as revealed by the tenured mutation in Drosophila. Dev Cell , 9, 843–54. Gupta, S. (2000) Hepatic polyploidy and liver growth control. Semin Cancer Biol , 10, 161–71.

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”

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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].

149

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].

REFERENCES 1. Weis, K. (2003) Regulating access to the genome: nucleocytoplasmic transport throughout the cell cycle. Cell , 112 (4), 441–51. 2. Tran, E.J. and Wente, S.R. (2006) Dynamic nuclear pore complexes: life on the edge. Cell , 125 (6), 1041–53. 3. Cronshaw, J.M. and Matunis, M.J. (2004) The nuclear pore complex: disease associations and functional correlations. Trends Endocrinol Metab, 15 (1), 34–39. 4. Rout, M.P., Aitchison, J.D., Suprapto, A., Hjertaas, K., Zhao, Y. and Chait, B.T. (2000) The yeast nuclear pore complex: composition, architecture, and transport mechanism. J Cell Biol , 148 (4), 635–51. 5. Cronshaw, J.M., Krutchinsky, A.N., Zhang, W., Chait, B.T. and Matunis, M.J. (2002) Proteomic analysis of the mammalian nuclear pore complex. J Cell Biol , 158 (5), 915–27. 6. Beck, M., Forster, F., Ecke, M., Plitzko, J.M., Melchior, F., Gerisch, G. et al. (2004) Nuclear pore complex

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structure and dynamics revealed by cryoelectron tomography. Science, 306 (5700), 1387–90. Beck, M., Lucic, V., Forster, F., Baumeister, W. and Medalia, O. (2007) Snapshots of nuclear pore complexes in action captured by cryo-electron tomography. Nature, 449 (7162), 611–15. Beck, M. and Medalia, O. (2008) Structural and functional insights into nucleocytoplasmic transport. Histol Histopathol , 23 (8), 1025–33. Alber, F., Dokudovskaya, S., Veenhoff, L.M., Zhang, W., Kipper, J., Devos, D. et al. (2007) The molecular architecture of the nuclear pore complex. Nature, 450 (7170), 695–701. Alber, F., Dokudovskaya, S., Veenhoff, L.M., Zhang, W., Kipper, J., Devos, D. et al. (2007) Determining the architectures of macromolecular assemblies. Nature, 450 (7170), 683–94. Schwartz, T.U. (2005) Modularity within the architecture of the nuclear pore complex. Curr Opin Struct Biol , 15 (2), 221–26. Hetzer, M.W., Walther, T.C. and Mattaj, I.W. (2005) Pushing the envelope: structure, function, and dynamics of the nuclear periphery. Annu Rev Cell Dev Biol , 21, 347–80. Belgareh, N., Rabut, G., Bai, S.W., van Overbeek, M., Beaudouin, J., Daigle, N. et al. (2001) An evolutionarily conserved NPC subcomplex, which redistributes in part to kinetochores in mammalian cells. J Cell Biol , 154 (6), 1147–60. Walther, T.C., Alves, A., Pickersgill, H., Loiodice, I., Hetzer, M., Galy, V. et al. (2003) The conserved Nup107-160 complex is critical for nuclear pore complex assembly. Cell , 113 (2), 195–206. Glavy, J.S., Krutchinsky, A.N., Cristea, I.M., Berke, I.C., Boehmer, T., Blobel, G. et al. (2007) Cell-cycledependent phosphorylation of the nuclear pore Nup107-160 subcomplex. Proc Natl Acad Sci USA, 104 (10), 3811–16. Lim, R.Y., Aebi, U. and Fahrenkrog, B. (2008) Towards reconciling structure and function in the nuclear pore complex. Histochem Cell Biol , 129 (2), 105–16. Fontoura, B.M., Blobel, G. and Matunis, M.J. (1999) A conserved biogenesis pathway for nucleoporins: proteolytic processing of a 186-kilodalton precursor generates Nup98 and the novel nucleoporin, Nup96. J Cell Biol , 144 (6), 1097–112. Loiodice, I., Alves, A., Rabut, G., Van Overbeek, M., Ellenberg, J., Sibarita, J.B. et al. (2004) The entire Nup107-160 complex, including three new members, is targeted as one entity to kinetochores in mitosis. Mol Biol Cell , 15 (7), 3333–44. Cristea, I.M., Williams, R., Chait, B.T. and Rout, M.P. (2005) Fluorescent proteins as proteomic probes. Mol Cell Proteomics, 4 (12), 1933–41. Boehmer, T., Enninga, J., Dales, S., Blobel, G. and Zhong, H. (2003) Depletion of a single nucleoporin, Nup107, prevents the assembly of a subset of nucleoporins into the nuclear pore complex. Proc Natl Acad Sci USA, 100 (3), 981–85.

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21. Boehmer, T., Jeudy, S., Berke, I.C. and Schwartz, T.U. (2008) Structural and functional studies of Nup107/Nup133 interaction and its implications for the architecture of the nuclear pore complex. Mol Cell , 30 (6), 721–31. 22. Devos, D., Dokudovskaya, S., Alber, F., Williams, R., Chait, B.T., Sali, A. et al. (2004) Components of coated vesicles and nuclear pore complexes share a common molecular architecture. PLoS Biol , 2 (12), e380. 23. Drin, G., Casella, J.F., Gautier, R., Boehmer, T., Schwartz, T.U. and Antonny, B. (2007) A general amphipathic alpha-helical motif for sensing membrane curvature. Nat Struct Mol Biol , 14 (2), 138–46. 24. Lutzmann, M., Kunze, R., Buerer, A., Aebi, U. and Hurt, E. (2002) Modular self-assembly of a Y-shaped multiprotein complex from seven nucleoporins. EMBO J , 21 (3), 387–97. 25. Berke, I.C., Boehmer, T., Blobel, G. and Schwartz, T.U. (2004) Structural and functional analysis of Nup133 domains reveals modular building blocks of the nuclear pore complex. J Cell Biol , 167 (4), 591–97. 26. Hsia, K.C., Stavropoulos, P., Blobel, G. and Hoelz, A. (2007) Architecture of a coat for the nuclear pore membrane. Cell , 131 (7), 1313–26. 27. Brohawn, S.G., Leksa, N.C., Spear, E.D., Rajashankar, K.R. and Schwartz, T.U. (2008) Structural evidence for common ancestry of the nuclear pore complex and vesicle coats. Science, 322 (5906), 1369–73. 28. Hawryluk-Gara, L.A., Shibuya, E.K. and Wozniak, R.W. (2005) Vertebrate Nup53 interacts with the nuclear lamina and is required for the assembly of a Nup93containing complex. Mol Biol Cell , 16 (5), 2382–94. 29. Jeudy, S. and Schwartz, T.U. (2007) Crystal structure of nucleoporin Nic96 reveals a novel, intricate helical domain architecture. J Biol Chem, 282 (48), 34904–12. 30. Antonin, W., Ellenberg, J. and Dultz, E. (2008) Nuclear pore complex assembly through the cell cycle: regulation and membrane organization. FEBS Lett , 582 (14), 2004–16. 31. Melcak, I., Hoelz, A. and Blobel, G. (2007) Structure of Nup58/45 suggests ﬂexible nuclear pore diameter by intermolecular sliding. Science, 315 (5819), 1729–32. 32. Lim, R.Y., Aebi, U. and Stofﬂer, D. (2006) From the trap to the basket: getting to the bottom of the nuclear pore complex. Chromosoma, 115 (1), 15–26. 33. Napetschnig, J., Blobel, G. and Hoelz, A. (2007) Crystal structure of the N-terminal domain of the human protooncogene Nup214/CAN. Proc Natl Acad Sci USA, 104 (6), 1783–88. 34. Enninga, J., Levy, D.E., Blobel, G. and Fontoura, B.M. (2002) Role of nucleoporin induction in releasing an mRNA nuclear export block. Science, 295 (5559), 1523–25. 35. Rosenblum, J.S. and Blobel, G. (1999) Autoproteolysis in nucleoporin biogenesis. Proc Natl Acad Sci USA, 96 (20), 11370–75. 36. Fontoura, B.M., Blobel, G. and Yaseen, N.R. (2000) The nucleoporin Nup98 is a site for GDP/GTP exchange on ran and termination of karyopherin beta 2-mediated nuclear import. J Biol Chem, 275 (40), 31289–96.

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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

INTRODUCTION

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

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,

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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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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

12: PROTEIN DEGRADATION AND THE LYSOSOMAL SYSTEM

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

Cargo in solution

Cargo interacts weakly with the plasma membrane

Cargo binds to specific receptors

Non-saturable

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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181

(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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THE LIVER: PEROXISOMAL PROTEIN IMPORT

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].

REFERENCES 1. Wanders, R.J. and Waterham, H.R. (2006) Peroxisomal disorders: the single peroxisomal enzyme deﬁciencies. Biochim Biophys Acta, 1763 (12), 1707–20.

2. Platta, H.W. and Erdmann, R. (2007) Peroxisomal dynamics. Trends Cell Biol, 17 (10), 474–84. 3. Jansen, G.A. and Wanders, R.J. (2006) Alpha-oxidation. Biochim Biophys Acta, 1763 (12), 1403–12. 4. Wanders, R.J. and Waterham, H.R. (2006) Biochemistry of mammalian peroxisomes revisited. Annu Rev Biochem, 75, 295–332. 5. Schrader, M. and Fahimi, H.D. (2006) Peroxisomes and oxidative stress. Biochim Biophys Acta, 1763 (12), 1755–66. 6. Steinberg, S.J. et al. (2006) Peroxisome biogenesis disorders. Biochim Biophys Acta, 1763 (12), 1733–48. 7. Erdmann, R. et al. (1989) Isolation of peroxisomedeﬁcient mutants of Saccharomyces cerevisiae. Proc Natl Acad Sci U S A, 86 (14), 5419–23. 8. Tsukamoto, T., Yokota, S. and Fujiki, Y. (1990) Isolation and characterization of Chinese hamster ovary cell mutants defective in assembly of peroxisomes. J Cell Biol , 110 (3), 651–60. 9. Titorenko, V.I. and Mullen, R.T. (2006) Peroxisome biogenesis: the peroxisomal endomembrane system and the role of the ER. J Cell Biol , 174 (1), 11–17. 10. Sakai, Y. et al. (2006) Pexophagy: autophagic degradation of peroxisomes. Biochim Biophys Acta, 1763 (12), 1767–75. 11. Gould, S.J. and Valle, D. (2000) Peroxisome biogenesis disorders: genetics and cell biology. Trends Genet , 16 (8), 340–45. 12. Steinberg, D. et al. (1967) Studies on the metabolic error in Refsum’s disease. J Clin Invest , 46 (3), 313–22. 13. Steinberg, D. et al. (1966) Effects of dietary phytol and phytanic acid in animals. J Lipid Res, 7 (5), 684–91. 14. Mihalik, S.J. et al. (1997) Identiﬁcation of PAHX, a Refsum disease gene. Nat Genet , 17 (2), 185–89. 15. Jansen, G.A. et al. (1997) Phytanoyl-CoA hydroxylase is not only deﬁcient in classical Refsum disease but also in rhizomelic chondrodysplasia punctata. J Inherit Metab Dis, 20 (3), 444–46. 16. van den Brink, D.M. et al. (2004) Identiﬁcation of fatty aldehyde dehydrogenase in the breakdown of phytol to phytanic acid. Mol Genet Metab, 82 (1), 33–37. 17. Poirier, Y. et al. (2006) Peroxisomal beta-oxidation—a metabolic pathway with multiple functions. Biochim Biophys Acta, 1763 (12), 1413–26. 18. Ferdinandusse, S. and Houten, S.M. (2006) Peroxisomes and bile acid biosynthesis. Biochim Biophys Acta, 1763 (12), 1427–40. 19. Brites, P., Waterham, H.R. and Wanders, R.J. (2004) Functions and biosynthesis of plasmalogens in health and disease. Biochim Biophys Acta, 1636 (2–3), 219–31. 20. Gorgas, K. et al. (2006) The ether lipid-deﬁcient mouse: tracking down plasmalogen functions. Biochim Biophys Acta, 1763 (12), 1511–26. 21. Novikoff, A.B. and Shin, W.Y. (1964) The endoplasmic reticulum in the Golgi zone and its relations to microbodies: Golgi-apparatus and autophagic vacuoles in rat liver cells. J Microsc, 3, 187–206. 22. Lazarow, P.B. and Fujiki, Y. (1985) Biogenesis of peroxisomes. Annu Rev Cell Biol , 1, 489–530.

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23. Titorenko, V.I. and Rachubinski, R.A. (1998) Mutants of the yeast Yarrowia lipolytica defective in protein exit from the endoplasmic reticulum are also defective in peroxisome biogenesis. Mol Cell Biol , 18 (5), 2789–803. 24. Mullen, R.T. et al. (1999) Peroxisomal membrane ascorbate peroxidase is sorted to a membranous network that resembles a subdomain of the endoplasmic reticulum. Plant Cell , 11 (11), 2167–85. 25. Geuze, H.J. et al. (2003) Involvement of the endoplasmic reticulum in peroxisome formation. Mol Biol Cell , 14 (7), 2900–7. 26. South, S.T. and Gould, S.J. (1999) Peroxisome synthesis in the absence of preexisting peroxisomes. J Cell Biol , 144 (2), 255–66. 27. Honsho, M., Hiroshige, T. and Fujiki, Y. (2002) The membrane biogenesis peroxin Pex16p. Topogenesis and functional roles in peroxisomal membrane assembly. J Biol Chem, 277 (46), 44513–24. 28. Hoepfner, D. et al. (2005) Contribution of the endoplasmic reticulum to peroxisome formation. Cell , 122 (1), 85–95. 29. Tam, Y.Y. et al. (2005) Pex3p initiates the formation of a preperoxisomal compartment from a subdomain of the endoplasmic reticulum in Saccharomyces cerevisiae. J Biol Chem, 280 (41), 34933–39. 30. Kragt, A. et al. (2005) Endoplasmic reticulum-directed Pex3p routes to peroxisomes and restores peroxisome formation in a Saccharomyces cerevisiae pex3Delta strain. J Biol Chem, 280 (40), 34350–57. 31. Kim, P.K. et al. (2006) The origin and maintenance of mammalian peroxisomes involves a de novo PEX16dependent pathway from the ER. J Cell Biol , 173 (4), 521–32. 32. Schrader, M. and Fahimi, H.D. (2008) The peroxisome: still a mysterious organelle. Histochem Cell Biol , 129, 421–40. 33. Li, X. et al. (2002) PEX11alpha is required for peroxisome proliferation in response to 4-phenylbutyrate but is dispensable for peroxisome proliferator-activated receptor alpha-mediated peroxisome proliferation. Mol Cell Biol , 22 (23), 8226–40. 34. Li, X. et al. (2002) PEX11 beta deﬁciency is lethal and impairs neuronal migration but does not abrogate peroxisome function. Mol Cell Biol , 22 (12), 4358–65. 35. Schrader, M. et al. (1998) Expression of PEX11beta mediates peroxisome proliferation in the absence of extracellular stimuli. J Biol Chem, 273 (45), 29607–14. 36. Koch, A. et al. (2003) Dynamin-like protein 1 is involved in peroxisomal ﬁssion. J Biol Chem, 278 (10), 8597–605. 37. Yu, T. et al. (2005) Regulation of mitochondrial ﬁssion and apoptosis by the mitochondrial outer membrane protein hFis1. J Cell Sci , 118 (Pt 18), 4141–51. 38. Fang, Y. et al. (2004) PEX3 functions as a PEX19 docking factor in the import of class I peroxisomal membrane proteins. J Cell Biol , 164 (6), 863–75.

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39. Kliewer, S.A. et al. (1992) Convergence of 9-cis retinoic acid and peroxisome proliferator signalling pathways through heterodimer formation of their receptors. Nature, 358 (6389), 771–74. 40. Wei, H. et al. (2000) Pharmacological induction of peroxisomes in peroxisome biogenesis disorders. Ann Neurol , 47 (3), 286–96. 41. Jones, J.M., Morrell, J.C. and Gould, S.J. (2004) PEX19 is a predominantly cytosolic chaperone and import receptor for class 1 peroxisomal membrane proteins. J Cell Biol , 164 (1), 57–67. 42. Brown, L.A. and Baker, A. (2008) Shuttles and cycles: transport of proteins into the peroxisome matrix (review). Mol Membr Biol, 25 (5), 363–75. 43. Gould, S.J. et al. (1989) A conserved tripeptide sorts proteins to peroxisomes. J Cell Biol , 108 (5), 1657–64. 44. Swinkels, B.W. et al. (1991) A novel, cleavable peroxisomal targeting signal at the amino-terminus of the rat 3-ketoacyl-CoA thiolase. EMBO J , 10 (11), 3255–62. 45. Williams, C. and Distel, B. (2006) Pex13p: docking or cargo handling protein? Biochim Biophys Acta, 1763 (12), 1585–91. 46. Thoms, S. and Erdmann, R. (2006) Peroxisomal matrix protein receptor ubiquitination and recycling. Biochim Biophys Acta, 1763 (12), 1620–28. 47. Leon, S., Goodman, J.M. and Subramani, S. (2006) Uniqueness of the mechanism of protein import into the peroxisome matrix: transport of folded, co-factor-bound and oligomeric proteins by shuttling receptors. Biochim Biophys Acta, 1763 (12), 1552–64. 48. Erdmann, R. and Schliebs, W. (2005) Peroxisomal matrix protein import: the transient pore model. Nat Rev Mol Cell Biol , 6 (9), 738–42. 49. McNew, J.A. and Goodman, J.M. (1994) An oligomeric protein is imported into peroxisomes in vivo. J Cell Biol , 127 (5), 1245–57. 50. Leon, S. et al. (2006) Dynamics of the peroxisomal import cycle of PpPex20p: ubiquitin-dependent localization and regulation. J Cell Biol , 172 (1), 67–78. 51. Platta, H.W. et al. (2007) Ubiquitination of the peroxisomal import receptor Pex5p is required for its recycling. J Cell Biol , 177 (2), 197–204. 52. Miyata, N. and Fujiki, Y. (2005) Shuttling mechanism of peroxisome targeting signal type 1 receptor Pex5: ATP-independent import and ATP-dependent export. Mol Cell Biol , 25 (24), 10822–32. 53. Platta, H.W. et al. (2005) Functional role of the AAA peroxins in dislocation of the cycling PTS1 receptor back to the cytosol. Nat Cell Biol , 7 (8), 817–22. 54. Platta, H.W., Girzalsky, W. and Erdmann, R. (2004) Ubiquitination of the peroxisomal import receptor Pex5p. Biochem J , 384 (Pt 1), 37–45. 55. Iwata, J. et al. (2006) Excess peroxisomes are degraded by autophagic machinery in mammals. J Biol Chem, 281 (7), 4035–41. 56. Bellu, A.R. et al. (2002) Removal of Pex3p is an important initial stage in selective peroxisome degrada-

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tion in Hansenula polymorpha. J Biol Chem, 277 (45), 42875–80. 57. Bellu, A.R. et al. (2001) Peroxisome biogenesis and selective degradation converge at Pex14p. J Biol Chem, 276 (48), 44570–74. 58. Farre, J.C. et al. (2008) PpAtg30 tags peroxisomes for turnover by selective autophagy. Dev Cell , 14 (3), 365–76.

59. Komatsu, M. et al. (2007) Homeostatic levels of p62 control cytoplasmic inclusion body formation in autophagy-deﬁcient mice. Cell , 131 (6), 1149–63. 60. Pankiv, S. et al. (2007) p62/SQSTM1 binds directly to Atg8/LC3 to facilitate degradation of ubiquitinated protein aggregates by autophagy. J Biol Chem, 282 (33), 24131–45.

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)

(d)

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.

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

Cytoplasm

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(c)

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

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

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(bacterial toxin receptor) Paracellular Loop 2 space

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PKC-z Cx26

Occludin PI3k pERK1

c-Yes

PPA2

cytosol PDZ

(c)

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ZO-1, –2, –3; PATJ; MUPPI

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.

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

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:

211

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)

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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

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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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(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

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].

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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83. Shen, L. and Turner, J.R. (2005) Actin depolymerization disrupts tight junctions via caveolae-mediated endocytosis. Mol Biol Cell , 16, 3919–36. 84. Ivanov, A.I., Nusrat, A. and Parkos, C.A. (2004) Endocytosis of epithelial apical junctional proteins by a clathrin-mediated pathway into a unique storage compartment. Mol Biol Cell , 15, 176–88. 85. Morimoto, S., Nishimura, N., Terai, T. et al. (2005) Rab13 mediates the continuous endocytic recycling of occludin to the cell surface. J Biol Chem, 280, 2220–28. 86. Yamada, A., Fujita, N., Sato, T. et al. (2006) Requirement of nectin, but not cadherin, for formation of claudin-based tight junctions in annexin II-knockdown MDCK cells. Oncogene, 25, 5085–102. 87. Matsuda, M., Kubo, A., Furuse, M. et al. (2004) A peculiar internalization of claudins, tight junction-speciﬁc adhesion molecules, during the intercellular movement of epithelial cells. J Cell Sci , 117, 1247–57. 88. Furuse, M., Hata, M., Furuse, K. et al. (2004) Claudin-1 gene mutations in neonatal sclerosing cholangitis associated with ichthyosis: a tight junction disease. Gastroenterology, 127, 1386–90. 89. Helle, F. and Dubuisson, J. (2008) Hepatitis C virus entry into host cells. Cell Mol Life Sci , 65, 100–112. 90. Meertens, L., Bertaux, C., Cukierman, L. et al. (2008) The tight junction proteins claudin-1, -6, and -9 are entry cofactors for hepatitis C virus. J Virol , 82, 3555–60. 91. Forrest, J.C., Campbell, J.A., Schelling, P., Stehle, T. and Dermondy, T.S. (2003) Structure-function analysis of reovirus binding to junctional adhesion molecule. 1. Implications for the mechanism of reovirus attachment. J Biol Chem, 278, 48243–44. 92. Prota, A.E., Campbell, J.A., Schelling, P. et al. (2003) Crystal structure of human junctional adhesion molecule 1: implications for reovirus binding. Proc Natl Acad Sci U S A, 100, 5366–71. 93. Cyone, C.B. and Bergelson, J.M. (2005) CAR: a virus receptor within the tight junction. Adv Drug Del Rev , 57, 869–82. 94. Schmelzer, E., Zhang, L., Bruce, A. et al. (2007) Human hepatic stem cells from fetal and postnatal donors. J Exp Med , 204, 1973–87. 95. Schmelzer, E., Wauthier, E. and Reid, L.M. (2006) The phenotypes of pluripotent human hepatic progenitors. Stem Cells, 24, 1852–58. 96. Kojima, T., Murata, M., Go, M. et al. (2007) Connexins induce and maintain tight junctions in epithelial cells. J Membr Biol , 217, 13–19.

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.

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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

226

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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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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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

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

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THE LIVER: THE ROLE OF TfR2, A SECOND TRANSFERRIN RECEPTOR HIGHLY EXPRESSED IN HEPATOCYTES

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 <105 M−1 respectively) [12]. Upon binding ferric Tf, a Tf–TfR complex forms and then internalizes within endosomes. As endosomes undergo acidiﬁcation, iron is released from the TfR–Tf complex. The released iron is still in the oxidized ferric form (Fe3+) and must undergo reduction by a reductase of the Steap family of proteins [13] prior to being transported from within the endosome to cytosol by a divalent metal-ion transporter (DMT) known as DMT1 [14]. When iron enters the cytosol, it can be incorporated into many enzymes and proteins that require iron for function. Iron can also be transported to subcellular locations such as mitochondria, which use an iron importer known as mitoferrin to facilitate mitochondrial iron uptake [15].

THE ROLE OF TfR2, A SECOND TRANSFERRIN RECEPTOR HIGHLY EXPRESSED IN HEPATOCYTES

Figure 16.1 Ferritin is a 24 subunit protein in which subunits of H (more abundant in heart) or L (more abundant in liver) chains coassemble to create a protein spherical structure with a central cavity. In this depiction, generated from coordinates of a reported ferritin crystal structure [10], three separate subunits on the anterior of the spherical protein are in dark gray, to show the arrangement of individual subunits. Pores in the structure allow entry of iron, which accumulates as a ferric precipitate within the hollow core. Each ferritin multimer can sequester up to 4500 iron atoms

Whereas in most tissues, TfR1 is the ﬁrst component of the major cellular iron uptake pathway, the liver instead expresses large amounts of a second, homologous TfR, TfR2 [16]. The fact that TfR2 has a lower afﬁnity for diferric Tf initially led to the supposition that TfR2 was not important for iron uptake. However, even though the afﬁnity of TfR2 for diferric Tf (27 nM) is 25-fold lower than that of TfR1 [16, 17], both TfRs have sufﬁcient afﬁnity to efﬁciently bind serum Tf, an abundant protein with a concentration of 9–28 nmol l−1 in serum, of which less than one-third of the iron binding sites are occupied with iron. Extrapolating from these measurements, the concentration of diferric Tf is estimated at about 8–12 µM [18], a concentration that would likely be sufﬁcient to saturate both TfRs. Thus, both TfR1 and TfR2 should be functional in iron uptake and it is possible that TfR2 is a major source of iron in hepatocytes, as the abundance

16: THE CENTRAL ROLE OF THE LIVER IN IRON STORAGE AND REGULATION OF SYSTEMIC IRON HOMEOSTASIS

and relative importance of the two receptors have not yet been quantitatively compared in hepatocytes.

TfR2 EXPRESSION IS REGULATED BY DIFFERENT MECHANISMS THAN TfR1 Aside from the fact that TfR2 is mainly expressed in hepatocytes, whereas TfR1 is expressed highly in erythroid progenitors and to varying degrees in most other tissues, the major difference between the two receptors is a difference in the mechanisms by which their expression is regulated. TfR1 is regulated at the transcriptional level by hypoxia [19, 20], but it is also potently regulated by a post-transcriptional regulatory system that consists of iron-regulatory proteins (IRPs), which bind to RNA stem-loops known as iron responsive elements (IREs) in transcripts such as ferritin and TfR1 (Figure 16.2). In cells that have cytosolic iron deﬁciency, IRPs bind to IREs, and IRP binding prevents translation of transcripts such as ferritin H and L chains, which have IREs located within the 5 UTR of transcripts (Figure 16.3a). Conversely, IRP binding to IREs in the 3 UTR of TfR1 stabilizes the transcript, leading to an increase in mRNA levels of TfR1, and a concomitant increase in protein levels (Figure 16.3b). In most cells that are iron-loaded, IRPs do not bind to

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IREs, and ferritin synthesis increases, whereas TfR1 expression decreases. Thus, iron loading in most cells leads to an increase in iron sequestration and to a decrease in TfR-mediated iron uptake (reviewed in [21, 22]). Hepatocytes differ from other cells because they express high levels of TfR2, and TfR2 mRNA levels remain high with iron loading [24]. In fact, binding of diferric Tf leads to increased TfR2 stability by changing TfR2 trafﬁcking [25]. One reason why hepatocytes are an important iron repository may thus be because TfR2-dependent iron uptake continues even when expression of TfR1 is diminished by iron overload in other cells throughout the body. The propensity to maintain iron uptake through non-IRP-regulated TfR expression may also be valuable for the hepatocyte in its role as a systemic iron regulator, as discussed below. Although TfR2 is implicated in a signaling pathway that responds to iron status (see below), the liver also has a strong capacity to take up non-transferrin-bound iron (NTBI).

NON-TRANSFERRIN-BOUND IRON UPTAKE AND HEPATIC IRON OVERLOAD In addition to the uptake of Tf-bound iron, the liver can very efﬁciently take up iron that is not sequestered by Tf (NTBI). NTBI becomes detectable in the plasma mainly

A Consensus Motif for the IRE G can also be A GU A G C X U/C/A N-N' N-N' 5 base pair stem N-N' N-N' N-N' Unpaired C Nucleoside − N-N' Usually Cytosine N-N' Lower stem N-N' (at least 3 base pair) 5'-N-N'-3' Consensus IRE

Functional IREs in 5′UTRs

Ferritin Mitochondrial aconitase Succinate dehydrogenase D. melanogaster eALASynthase Ferroportin-basolateral iron exporter HIF2 alpha

C G

C

5′

3′

Functional IREs in 3′UTRs

Transferrin receptor Divalent metal transporter (non-consensus) CDC14A

Figure 16.2 Iron responsive elements (IREs) are RNA stem-loops found in numerous proteins important in iron regulation. IREs consist of a base-paired stem, an unpaired cytosine that separates the upper and lower stems, and a six-membered loop with the sequence CAGUGX, in which X can be any residue except G and there is a base pair between A2 and G5 of the loop [23]

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THE LIVER: NON-TRANSFERRIN-BOUND IRON UPTAKE AND HEPATIC IRON OVERLOAD

One IRE in 5’UTR

Ferritin mRNA

5’

3’

AUG

+Fe

−Fe IRP

AAAAAA

40s

IRE occupied by IRP, inhibiting translation initiation IRE unoccupied, allowing polysome formation and increased ferritin synthesis

60s

(a) TfR mRNA

Five IREs in 3’UTR

5’

AAAAAA

Protein Coding

3’

−Fe +Fe

IRP

Protein Coding

Endonuclease Cleavage Site

AAAAAA

AAAAAA Protein Coding

One or more IREs occupied by IRP, protecting mRNA from rate-determining step mRNA degradation (b)

IRE unoccupied, rendering mRNA susceptible to an endonuclease

Figure 16.3 (a) Iron regulatory proteins 1 and 2 repress translation of transcripts that contain an IRE in the 5 UTR of the transcript by preventing binding of translation factors and ribosomes. (b) Binding of IRPs to IREs in the 3 UTR protects the TfR transcript from degradation, and likely also protects other transcripts that contain a 3 IRE by a similar mechanism

when Tf iron binding sites become fully saturated, as occurs in various diseases of systemic iron overload. It has been demonstrated that the plasma half-life of NTBI is less than 30 seconds, whereas Tf-bound iron has a half-life of approximately 50 minutes [26, 27]. Thus, the isolated perfused liver can remove 60–75% of NTBI iron from plasma when it ﬁrst passes through, whereas it removes less than 2% of Tf-bound iron at ﬁrst pass [26, 28, 29]. In addition, if plasma Tf is ﬁrst saturated with iron, the clearance of a subsequent dose of radioactive iron, given either orally or intravenously, is very rapid,

with the liver being the major site of iron deposition [27]. Perhaps the best demonstration of the capacity of the liver to acquire NTBI comes from studies of congenital hypotransferrinemia, where little Tf is synthesized in either a mouse model [30] or human patients [31]. In this condition, the liver and some other organs develop massive iron loading at a very young age. Such studies show that the role of Tf is not simply to deliver iron to tissues, but to allow the regulated delivery of the metal. Tf has such a high binding afﬁnity for iron under normal physiological conditions (afﬁnity constant 1023 M−1 ) [32] that

16: THE CENTRAL ROLE OF THE LIVER IN IRON STORAGE AND REGULATION OF SYSTEMIC IRON HOMEOSTASIS

the amount of circulating NTBI is extremely small and is essentially undetectable. Where NTBI does become important is in various pathological states, particularly iron loading disorders in which the capacity of plasma Tf to bind iron is exceeded. In HFE -associated hemochromatosis, for example, NTBI can reach 10–15 µM [33]. It has proved difﬁcult to precisely deﬁne the chemical nature of NTBI, but most of it may be chelated by small organic acids such as citrate and amino acids, and some can also be bound by proteins such as albumin [34]. The uptake pathway of NTBI has not been unequivocally deﬁned. Early studies suggested that NTBI uptake was mediated by a carrier protein [35], perhaps DMT1 [36], but when DMT1 is disrupted the liver is still able to accumulate NTBI, suggesting that other uptake pathways are also involved [37, 38]. Another candidate is Zrt- and Irt-like protein 14 or SLC39A14 (ZIP14), a zinc transporter that has the capacity to transport iron and is expressed in the liver [39]. Hepatoma cells are able to take up NTBI in a ZIP14-dependent manner. It has also been proposed that L-type voltage-dependent calcium channels play a role in NTBI uptake by some tissues, including the heart [40], as calcium channel antagonists block iron uptake. Whether such channels play a role in iron uptake by the liver is not known. Early studies on the kinetics of NTBI uptake indicated that the process was increased under iron loading conditions and it was suggested that this plays a protective role by removing potentially toxic “free” iron from the circulation [41].

THE RELEASE OF IRON FROM HEPATIC CELLS Although a great deal is known about the uptake of iron, the mechanisms by which the liver can release stored iron in times of need are less well understood. The capacity of the liver to relinquish stored iron is particularly well illustrated during phlebotomy therapy for primary iron-loading diseases such as HFE -associated hemochromatosis [42, 43]. In such diseases, the liver may contain 20 g or more of stored iron, but all of this can be mobilized for hemoglobin synthesis following phlebotomy. While in HFE -associated hemochromatosis most of the iron storage in the liver occurs in hepatocytes (see below), in transfusional iron overload the resident macrophages (the Kupffer cells) also can store much of the iron, and iron can be mobilized from Kupffer cells as well as hepatocytes by phlebotomy. In addition, Kupffer cells play an important role in the recycling of hemoglobin-derived iron from senescent red blood cells, and this iron must also be released into the circulation [44]. The protein that appears to be responsible for iron release from most body cells is ferroportin (FPN) [45–47]. FPN is a highly conserved ferrous iron export protein that contains 9 or 10 transmembrane domains. While FPN is most highly expressed in cell types that efﬂux

239

large amounts of iron, such as intestinal enterocytes and macrophages [48], it also performs an iron efﬂux role in all cells studied to date, including hepatocytes [49]. FPN mRNA contains an IRE in its 5 UTR, like the ferritin genes, and its expression in the liver increases with iron loading, suggesting that it is under iron-dependent translational control through the IRE/IRP system [47]. Since FPN levels increase with iron loading, it is feasible that this increase represents a protective mechanism to help the liver limit its iron accumulation. Efﬁcient export of iron from tissues requires an iron oxidase in addition to FPN. For most tissues the oxidase activity is supplied by circulating ceruloplasmin (Cp) [50]. Cp is the major copper-containing protein in the plasma, but it is a protein of iron metabolism rather than copper metabolism. In the absence of Cp (e.g. in rare cases of aceruloplasminemia), the plasma iron level is reduced and iron accumulates in the tissues, whereas copper homeostasis is unaffected [51, 52]. How Cp contributes to iron efﬂux is unclear, but a GPI-linked form of Cp can apparently stabilize FPN on the cell surface by oxidizing transported ferrous iron [53]. More plasma-membrane FPN means increased cellular iron release. Whether this is a general mechanism of Cp action remains to be determined. The liver is the principal site of synthesis of Cp and thus it plays an important role systemically in facilitating iron efﬂux.

HEPATOCYTES SYNTHESIZE AND SECRETE THE REGULATORY PEPTIDE HORMONE HEPCIDIN In systemic iron regulation, secretion of the peptide hormone hepcidin has an important role in determining how much iron is released from macrophages, and likely other body cells, and how much iron is absorbed through the duodenum [54]. Macrophages either release or store iron after they metabolize senescent red cells, and the recycled iron produced by macrophages is the major source of serum iron. Macrophages modulate the return of iron to the circulation to maintain Tf saturation levels of approximately 30%, and hepcidin is believed to play an important role in macrophage iron release. Hepcidin is related to other peptide hormones known as defensins that function in defense against bacterial infections. Hepcidin is synthesized as an 84 amino acid precursor, which undergoes cleavage and is released to the circulation as an active 25 amino acid peptide hormone. Forms of hepcidin that are truncated at the N-terminus and contain 22 or 20 amino acids are also present in serum, but these forms are not active [55]. (Figure 16.4 shows the hepcidin sequence and the positions of disulﬁde bonds). As a circulating peptide, hepcidin can interact with tissues and cells throughout the body. Hepcidin binds

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THE LIVER: TRANSCRIPTIONAL REGULATION OF HEPCIDIN EXPRESSION

Figure 16.4 The sequence of the peptide hormone hepcidin is depicted. The peptide contains eight cysteines and is believed to contain four disulﬁde bonds. Arcs depict the proposed cysteine pairs, and the central two cysteines are believed to form a disulﬁde bond that forms a hairpin in the hepcidin structure [56]

to the iron exporter FPN on macrophage membranes, where it forms a complex that internalizes and undergoes lysosomal degradation. Thus, hepcidin can reduce iron efﬂux from macrophages by causing degradation of FPN [57]. It is not yet clear if this mechanism of action can explain all of the effects of hepcidin (see below). In general, transcription of hepcidin is low when there is systemic iron depletion, whereas transcription levels are high when there is systemic iron overload. The mechanisms by which hepatocytes transduce information about systemic iron status into transcriptional activity of the hepcidin promoter are under active investigation.

TRANSCRIPTIONAL REGULATION OF HEPCIDIN EXPRESSION IS THE MAJOR DETERMINANT OF SYSTEMIC IRON HOMEOSTASIS Hepcidin is the central regulator of systemic iron homeostasis as it coordinates release of iron to serum from macrophages and also regulates intestinal iron absorption. Whereas it was once believed that sensing of systemic iron homeostasis occurred in the cells of the duodenal mucosa, numerous studies now indicate that the most important site of sensing is within hepatocytes [58, 59]. Signaling information converges at the hepcidin promoter. Important elements of the signaling pathway that affect hepcidin transcription in hepatocytes include serum proteins such as Tf and IL-6. Circulating proteins such as Tf and IL-6 are believed to directly or indirectly interact with receptors and associated proteins on the hepatic plasma membrane. These proteins include TfR2, HFE, which is the protein affected in the most common form of genetic hemochromatosis, and hemojuvelin, a member of the bone morphogenetic protein (BMP) receptor family [60]. IL-6 stimulates hepcidin transcription through a signaling pathway that does not involve either TfR2 or HFE [61]. Initially, it was not recognized that these signaling pathways are active in hepatocytes because work was performed on hepatic cell lines rather than fresh primary hepatocytes, which are needed to observe activity of

the hemojuvelin/BMP pathway [62]. Tf itself is strongly implicated in the hepcidin signaling pathway. The serum Tf saturation measurement integrates the removal and return of iron to the circulation from many tissue sources into one common value. Thus, the Tf saturation may be the principle clinically measurable indicator of iron status that conveys information about bodily iron status to multiple tissues [58, 61, 63, 64]. Importantly, atransferrinemic mice and hypotransferrinemic patients develop tissue iron overload, which attests to both the importance of transferrin in regulation of body iron status [65, 66] and the importance of NTBI in tissue iron overload. There is much that remains unknown about the signaling pathways that enable hepatocytes to accurately gauge systemic iron levels. Many insights were gained when genes that cause hereditary hemochromotosis were identiﬁed. The ﬁrst gene identiﬁed as a cause of hereditary hemochromatosis, HFE [67], was initially believed to function by binding to TfR1 and competitively inhibiting binding of Tf [68], but not by binding to TfR2 [68]. Recent studies indicate that TfR2 binds HFE when both proteins are overexpressed [69]. Importantly, HFE binding appears to increase the afﬁnity of TfR2 for diferric Tf [70]. Thus, data increasingly suggest that TfR2-dependent Tf binding plays a dominant role in hepatocyte iron sensing and systemic iron homeostasis. Another surface protein important in hepatocyte iron sensing was identiﬁed in family studies of patients that develop severe juvenile-onset hemochromatosis. The gene, which was accordingly named hemojuvelin (HJV ) [71], is now recognized as a member of the bone morphogenetic protein receptor (BMPR) family. The mechanisms by which TfR2, HFE, and HJV link to a change in hepcidin transcription are incompletely understood [72], but the discovery of hemojuvelin revealed that the well-known SMAD signal transduction pathway is important in this process. Although the mechanisms by which TfR2, HFE, and HJV link to a change in hepcidin transcription are incompletely understood [72], the signaling pathways of BMP receptors have been studied extensively in other physiological settings. In general, BMP receptors function by activating SMAD proteins, a family of transcriptional activators/repressors that function in numerous pathways to connect receptor-binding events to changes in nuclear transcription of target genes [73]. Notably, animals that lack hepatocyte SMAD4, a protein that combines with other members of the SMAD family to regulate transcription of target genes, develop signiﬁcant iron overload associated with a profound reduction in hepcidin expression [74]. In addition to responding to signaling through the BMP–SMAD pathway, hepcidin expression is also induced by hypoxia and inﬂammation [75]. LPS, interleukin-6, and interleukin-1 activate hepcidin expression

16: THE CENTRAL ROLE OF THE LIVER IN IRON STORAGE AND REGULATION OF SYSTEMIC IRON HOMEOSTASIS

[61] through a pathway that involves JAK–STAT signaling and a binding site for the transcription factor STAT3 [76, 77]. Thus it appears that hepatocytes sense Tf saturation levels and some inﬂammatory signals through distinct traditional receptors, and that these signals converge at the hepcidin promoter. This may allow crosstalk between inﬂammatory and iron-dependent signals, because binding sites for these independent signals are nearly adjacent to one another in the promoter [78]. Increases in hepcidin transcription result in changes in circulating hepcidin levels. Thus, it is important to understand how hepcidin regulates iron release from macrophages and enterocytes (Figure 16.5). Although several pathways that activate hepcidin transcription have been described above, only one pathway has thus far been identiﬁed that represses hepcidin expression. The transmembrane serine protease TMPRSS6 is part of a pathway that suppresses hepcidin expression, as revealed by Tmprss6 mutant mice [79] and some human patients with refractory anemia that carry TMPRSS6 mutations [80]. It is not yet known how this protease reduces hepcidin expression, but it may do so by cleaving one of the membrane proteins involved in the hepcidin signaling pathway, such as HJV or TfR2. Patients with TMPRSS6 mutations have high urinary and plasma hepcidin levels

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[80], which results in diminished intestinal iron absorption and diminished release of iron from macrophages.

HEPCIDIN REGULATES SYSTEMIC IRON HOMEOSTASIS MAINLY THROUGH ITS EFFECTS ON MACROPHAGES AND THE DUODENAL MUCOSA In general, peptide hormones such as hepcidin effect changes by binding to speciﬁc receptors in target cell populations. The process of binding to a receptor often activates a process of signal transduction, in which binding of the ligand to the receptor activates a cascade of intracellular events that often change the transcription of various genes in the target cell population. Hepcidin signaling does not conform to this general pathway, because hepcidin binds to the iron exporter FPN rather than to a member of a previously recognized receptor family. Upon binding, FPN is internalized and subsequently degraded by lysosomes, which results in reduced export of iron because FPN is the only known iron

Figure 16.5 The hepcidin signaling pathway in hepatocytes. Diferric transferrin likely binds to TfR2, and TfR2 signals the nucleus information about iron status, aided by HFE and by signaling information from BMP–BMPR signaling, which is facilitated by HJV. IL-6 signals through a separate surface receptor. Transcription factors activated by this pathway activate hepcidin transcription by binding to a Stat3 binding site, whereas signals through the TfR2/HFE, BMPR pathway likely activate SMAD transcription factors. These sites are adjacent on the promoter and can affect one another. Ultimately, hepcidin (HAMP) transcription increases in hepatocytes that sense high systemic iron or inﬂammatory signals

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THE LIVER: MUTATIONS IN GENES INVOLVED IN HEPATOCYTE SENSING OF SYSTEMIC IRON LEVELS

Figure 16.6 A model for systemic iron homeostasis. Serum signals reﬂecting iron stores, erythropoiesis, and inﬂammation are transduced by receptors, and information converges on the hepcidin promoter, which determines how much hepcidin is synthesized. Hepcidin binds to ferroportin on the membrane surface of macrophages and enterocytes, leading to reduced ferroportin expression, and thereby enhancing macrophage iron sequestration and reducing duodenal iron absorption

exporter [81, 82]. Hepcidin binds to the N-terminus of FPN [55] and alters the phosphorylation of several cytosolic residues of the protein [57]. The present model for hepcidin function is that it binds to FPN on the membrane of macrophages, which leads to degradation of FPN, and the absence of functional FPN results in retention of iron by macrophages. The same process of hepcidin-induced degradation of FPN is also proposed to reduce export of iron from duodenal mucosal cells to the circulation, and thus to reduce the absorption of dietary iron. Although hepcidin-dependent degradation of FPN is accepted as a major mechanism by which intestinal iron uptake is decreased by hepcidin overexpression, some questions remain unsolved. For instance, FPN mRNA expression increases dramatically in iron deﬁciency, and it is not clear how this occurs [45]. Similarly, FPN is often observed in internal vesicles rather than on the basolateral membrane, and large transcriptional changes of FPN, DMT1, and Dcytb cannot be readily explained by degradation of FPN protein [83, 84]. The timing of the hepcidin effect on the duodenum also differs from its effects on macrophages, with the hepcidin effect on macrophages occurring before the duodenal mucosa is affected [85]. An overall potential model for hepcidin

activity is that low hepcidin levels result in high levels of FPN in duodenal mucosal cells, and these cells therefore become iron-deﬁcient, which then results in increased expression within duodenal mucosal cells of the iron importer DMT1, the ferric reductase Dcytb, and the iron exporter FPN (Figure 16.6).

MUTATIONS IN GENES INVOLVED IN HEPATOCYTE SENSING OF SYSTEMIC IRON LEVELS CAUSE HEMOCHROMATOSIS “Hemochromatosis” is the term for a group of inherited diseases characterized initially by increased serum iron levels, and later by parenchymal iron overload in the liver, heart, and endocrine organs [86]. Macrophages and the duodenal mucosa are the tissues most responsible for the abnormal increase in serum iron levels. Macrophages are normally the major repository of iron in the body, because they metabolize senescent red cells and store any iron derived from heme catabolism that

16: THE CENTRAL ROLE OF THE LIVER IN IRON STORAGE AND REGULATION OF SYSTEMIC IRON HOMEOSTASIS

is not released to plasma transferrin. Whereas the bone marrow macrophages of non-hemochromatosis patients are usually rich in stored iron, as assessed by Prussian blue iron stain, the macrophages of hemochromatosis patients have reduced ability to retain iron retrieved from erythrophagocytosis, and Prussian blue staining reveals that macrophages contain little iron in the spleen and liver. Moreover, despite high serum iron levels, duodenal iron absorption remains relatively high in hemochromatosis patients. As the disease progresses, increased serum iron levels cause parenchymal iron overload in the liver, heart, and endocrine organs. There are now four different genes recognized to cause the form of genetic hemochromatosis in which macrophage iron depletion is associated with parenchymal iron overload. These are HFE , TfR2 , HJV , and hepcidin, and it is likely that the list of disease genes will grow as research continues. All of these disorders are characterized by abnormally reduced hepcidin production in conjunction with elevated plasma transferrin saturation and increased parenchymal rather than macrophage iron stores. Fortunately, impaired function of these genes does not adversely affect hematopoiesis. As a result, iron balance can be restored by regular phlebotomy procedures to remove red cells rich in hemoglobin iron; these red cells are readily replaced by increased erythropoiesis.

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with TfR2, which is highly expressed in the liver where systemic iron sensing occurs [25, 70]. The role of HFE mutations in developing hemochromatosis is not as clear as was initially expected. Many homozygotes for the C282Y mutation have little evidence of iron overload and few symptoms [90]. A large study recently concluded that overt hemochromatosis was observed in 28% of males that were homozygous for the C282Y mutation, whereas only 1% of female homozygotes developed disease [91]. It is likely that there are genetic modiﬁer genes that determine whether HFE mutations will cause disease, and among the possible modiﬁers are other gene products involved in the same sensing pathway. Mutations in hepcidin can modify the iron loading in patients with C282Y alleles [92]. Numerous tests have been used to diagnose hemochromatosis in the past, including Tf saturation, serum ferritin levels, liver biopsy with evidence of parenchymal iron overload and cirrhosis, and, more recently, genetic tests for the C282Y mutation. However, recent data indicate that a single serum ferritin determination may be the most cost-effective approach to discovering patients with HFE mutations that are prone to developing complications [91, 93], and the best approach to diagnosis continues to evolve.

TfR2 AND HEMOCHROMATOSIS THE DISCOVERY OF HFE, THE FIRST IDENTIFIED CAUSE OF GENETIC HEMOCHROMATOSIS, AND THE ENIGMA OF ITS UNEXPECTEDLY LOW PENETRANCE Although early studies demonstrated that the commonest form of genetic hemochromatosis was caused by a gene on chromosome 6 based on linkage to the HLA region [87], the affected gene, HFE , was not identiﬁed until 1996, in part because the disease gene resided in a region of the chromosome that was not prone to the informative recombination events that facilitate positional cloning [67]. Initially, a single mutation in the coding region of HFE that results in substitution of a tyrosine for a cysteine at codon 282 (C282Y) was recognized, and a PCR test for the C282Y mutation conﬁrmed that it was widespread in populations of Northern European descent [88]. HFE did not belong to a known gene family, and insight into its potential function came from the observation that it could bind to TfR1 [89] and interfere with the binding of diferric transferrin [68]. However, an animal model indicated that Hfe does not exert its effects on phenotype through binding to TfR1 [72]. These results are consistent with the possibility that HFE functions in conjunction

Initially discovered as a second TfR with 45% homology to TfR1, TfR2 differs in that it is mainly expressed in liver, with some expression in brain, heart, and spleen, but little expression in other tissues [16]. Soon after its initial discovery, mutations in TfR2 were found to cause hemochromatosis similar to HFE hemochromatosis in both the histological pattern of iron overload (parenchymal loading with sparing of macrophages) and in disease severity. As we have learned more about TfR2-associated hemochromatosis, it has become apparent that it is more severe than HFE-related disease, but not as severe as juvenile hemochromatosis(JH). The disease usually presents in adults [94, 95], although it sometimes presents in juveniles [96]. Recapitulation of the essential features of the disease in a mouse model conﬁrmed that TfR2 was another cause of classical hemochromatosis [97]. Interestingly, TfR2 levels increase with binding of Tf, due to decreased degradation of TfR2 in lysosomes, and binding of Tf results in more recycling of TfR2 [25].

CAUSES OF JUVENILE HEMOCHROMATOSIS Among hemochromatosis patients, the severity of disease can vary substantially. For many years, investigators recognized a distinct type of hemochromatosis in which

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THE LIVER: ELEVATED HEPCIDIN EXPRESSION EXPLAINS THE ANEMIA OF CHRONIC DISEASE

severe iron overload and organ damage occurred before patients reached the age of 30. Thus, it was not surprising that the unusually severe clinical presentation correlated with mutations in other genes, including the gene that encodes hepcidin (HAMP ) and the gene that encodes hemojuvelin.

Hepcidin mutations cause severe early onset hemochromatosis Hepcidin is now acknowledged to be the most important regulator of systemic iron homeostasis through its capacity to regulate iron release from macrophages and from duodenal enterocytes, and it is not surprising that mutations in hepcidin itself cause a severe phenotype of hemochromatosis [98].

Hemojuvelin mutations cause severe early onset hemochromatosis Using positional cloning approaches to families with juvenile hemochromatosis, a gene not previously recognized as important in iron homeostasis was discovered to harbor mutations that cause disease [71, 99]. The gene product was given the name “hemojuvelin” because of its causal role in juvenile hemochromatosis, in which patients typically develop hypogonadotrophic hypogonadism, hepatic ﬁbrosis or cirrhosis, and cardiomyopathy before age 30. By sequence homology, HJV belongs in the family of repulsive guidance molecules (RGMs), proteins that bind to pairs of Type I and Type II serine/threonine kinase receptors that are important in transducing signals from proteins in the transforming growth factor β (TGFβ) superfamily. Upon ligand binding, these Type I and Type II receptor pairs activate the Smad proteins, which translocate to the nucleus to modulate transcription of various target genes when they have undergone receptor-dependent phosphorylation. BMPs are signaling molecules in the TGFβ superfamily that interact with a speciﬁc set of Type I and Type II receptor pairs known as BMP receptors (reviewed in [100]). For some receptors in the BMP signaling pathway, RGMs bind to the receptor-ligand complex and potentiate signaling [101]. It appears that HJV, which is required for the increase of hepcidin transcription that occurs in response to iron loading [102], is a co-receptor that facilitates BMP signaling in hepatocytes [103]. HJV is linked to membranes through a glycophosphatidyl–inositol (GPI) membrane linkage, and it speciﬁcally facilitates signaling of BMP2 and BMP4 ligands [62]. Mouse models in which the hemojuvelin gene is deleted (HJV–/–) develop parenchymal iron overload typical of hemochromatosis, but they develop relatively little organ damage, unlike humans [102, 104], an observation that remains unexplained.

MUTATIONS OF THE IRON EXPORTER FERROPORTIN CAUSE A GROUP OF DISTINCT IRON OVERLOAD SYNDROMES KNOWN AS FERROPORTIN DISEASE Mutations in the gene encoding the iron export protein FPN (SLC40A1 ) can also lead to iron loading [105], causing a group of syndromes often known as ferroportin disease. Patients with mutations in SLC40A1 fall into two classes, with distinct phenotypes depending on the particular mutation they possess [106]. The most common type of mutation leads to reduced levels of FPN on the plasma membrane or reduced iron transport capacity. Reduced iron export means that less iron enters the plasma and that iron accumulates in the tissues, and particularly in tissue macrophages, as internal iron recycling is disrupted. Thus the characteristic of this type of ferroportin disease is reduced plasma iron in the face of reticuloendothelial cell iron accumulation. In the liver, iron accumulation is seen in a panlobular distribution, which is distinct from the periportal hepatocyte-dominated distribution observed in the early stages of HFE -related hemochromatosis. The reduced plasma iron can also mean that a mild anemia develops. This in turn can stimulate iron absorption and increase the body iron load. In ferroportin disease, iron absorption is less efﬁcient due to functional FPN deﬁciency, but there still appears to be a net accretion of iron by the body. In the second form of ferroportin disease, missense mutations in SLC401 generate an FPN protein that has reduced responsiveness to hepcidin. In this situation, the iron loading is quite similar to that associated with disruption of HFE, TfR2, hemojuvelin, and hepcidin, with elevated transferrin saturation and periportal deposition of iron in the hepatocytes of the liver. The failure of FPN to respond appropriately to hepcidin means that the body’s normal feedback mechanism for limiting iron loading is dysfunctional.

ELEVATED HEPCIDIN EXPRESSION EXPLAINS THE ANEMIA OF CHRONIC DISEASE The anemia of chronic disease, better termed the anemia of chronic inﬂammation, is characterized by decreased production of erythrocytes by the bone marrow without underlying nutrient or hormonal deﬁciency and with no intrinsic abnormality of the bone marrow. This hypoproliferative anemia is marked by the presence of one of several chronic underlying inﬂammatory disorders, which can be of infectious, malignant, inﬂammatory, or traumatic origin. Typically the anemia is mild or moderate in character,

16: THE CENTRAL ROLE OF THE LIVER IN IRON STORAGE AND REGULATION OF SYSTEMIC IRON HOMEOSTASIS

the mean corpuscular volume is mildly reduced, the serum concentrations of iron and transferrin are decreased, and the serum ferritin concentration is in the upper end of the normal range or frankly elevated. Duodenal iron absorption is reduced and storage of iron in macrophages is increased. The changes in iron metabolism in the anemia of chronic inﬂammation are likely attributable to increased secretion of hepcidin by hepatocytes. Several experiments indicate that exogenous delivery of hepcidin or overexpression of hepcidin can cause chronic anemia. The injection of hepcidin in mice rapidly leads to a prolonged hypoferremia [107]. Mice that overexpress hepcidin under the control of a liver-speciﬁc promoter depend on parenteral iron injections after birth, suggesting that hepcidin is a negative regulator of iron transport in the small intestine and that it induces iron retention in macrophages that normally recycle iron from senescent erythrocytes [108]. Chronic overexpression of human hepcidin in mice leads to hypoferremia, increased hepatic iron, and anemia [109]. Patients with large hepatic adenomas that produce inappropriately high levels of hepcidin mRNA have iron refractory anemia similar to that observed in anemia of chronic disease, and this anemia resolves spontaneously after adenoma resection or liver transplantation [110]. Recent results suggest that, in addition to hepatocytes, monocytes also produce hepcidin and that such autocrine production may contribute to macrophage iron loading and the anemia of chronic inﬂammation [111]. Changes in iron metabolism probably do not fully explain the anemia of chronic disease. Additional factors may include suppression of erythroid precursors by inﬂammatory cytokines and some limitation to the production of or response to erythropoietin.

THE LIVER IS IMPORTANT IN CLEARING REACTIVE HEME FROM CIRCULATION Hepatocytes are the main site of synthesis of hemopexin, an abundant serum protein that binds free heme with high afﬁnity [112]. The structure of hemopexin enables it to sequester heme in a non-reactive form [113] and heme–hemopexin complexes are then metabolized by cells that have hemopexin receptors, including hepatocytes, macrophages, neurons, and syncytiotrophoblasts [114]. Increased hepatic uptake of hemopexin triggers an increase in hepatic heme oxygenase-1 expression, which enables hepatocytes to fully metabolize the newly absorbed heme. The hemopexin system is a major defense against tissue injury by free heme, which is important in the pathogenesis of ischemia-reperfusion and traumatic injuries. Mice engineered to lack hemopexin are more prone to developing renal failure after hemolysis than wild-type

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mice, indicating that hemopexin is important for protecting animals from the toxic effects of free heme in the circulation [115]. Haptoglobin is a tetrameric protein expressed by the liver that combines with free plasma hemoglobin and protects tissues, particularly the kidney, from hemoglobindependent damage [116]. Hemoglobin–haptoglobin complexes are cleared from the circulation by liver and spleen macrophages, which express the hemoglobin–haptoglobin receptor CD163 [117]. The liver protects the other organs against damage from hemolysis and trauma by synthesizing and secreting hemopexin and haptoglobin, and also by taking up heme–hemopexin and hemoglobin–haptoglobin complexes, and catabolizing the heme it acquires through these pathways using heme oxygenase.

HEPATIC IRON OVERLOAD AND THE RELATIONSHIP OF IRON OVERLOAD TO ALCOHOLIC LIVER DISEASE A third or more of alcoholics develop increased amounts of hepatic iron compared to controls [118, 119]. In rural Africa, there is a strong association among the consumption of a traditional fermented beverage with high ionic iron concentration, increased body iron stores, and liver toxicity and cirrhosis [120–124]. In experimental rats, dietary iron supplementation exacerbates alcohol-induced hepatocyte damage and promotes liver ﬁbrogenesis [125]. Similarly, the presence of elevated iron stores accentuates the hepatic toxicity of alcohol in the setting of HFE hemochromatosis in Caucasians [126]. Recent studies suggest that increased iron absorption associated with alcohol ingestion may be related to suppression of hepcidin production by hepatocytes [127–129].

ACKNOWLEDGMENTS We thank Helge Uhrigshardt for creating the ferritin structural representation.

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76. Wrighting, D.M. and Andrews, N.C. (2006) Interleukin-6 induces hepcidin expression through STAT3. Blood , 108, 3204–9. 77. Verga Falzacappa, M.V., Vujic Spasic, M., Kessler, R., Stolte, J., Hentze, M.W. and Muckenthaler, M.U. (2007) STAT3 mediates hepatic hepcidin expression and its inﬂammatory stimulation. Blood , 109, 353–58. 78. Verga Falzacappa, M.V., Casanovas, G., Hentze, M.W. and Muckenthaler, M.U. (2008) A bone morphogenetic protein (BMP)-responsive element in the hepcidin promoter controls Hfe2-mediated hepatic hepcidin expression and its response to IL-6 in cultured cells. J Mol Med , 86 (5), 531–40. 79. Du, X., She, E., Gelbart, T., Truksa, J., Lee, P., Xia, Y., Khovananth, K., Mudd, S., Mann, N., Moresco, E.M., Beutler, E. and Beutler, B. (2008) The serine protease TMPRSS6 is required to sense iron deﬁciency. Science, 320, 1088–92. 80. Finberg, K.E., Heeney, M.M., Campagna, D.R., Aydinok, Y., Pearson, H.A., Hartman, K.R., Mayo, M.M., Samuel, S.M., Strouse, J.J., Markianos, K., Andrews, N.C. and Fleming, M.D. (2008) Mutations in TMPRSS6 cause iron-refractory iron deﬁciency anemia (IRIDA). Nat Genet , 40, 569–71. 81. Nemeth, E., Tuttle, M.S., Powelson, J., Vaughn, M.B., Donovan, A., Ward, D.M., Ganz, T. and Kaplan, J. (2004) Hepcidin regulates cellular iron efﬂux by binding to ferroportin and inducing its internalization. Science, 306, 2090–93. 82. Delaby, C., Pilard, N., Goncalves, A.S., Beaumont, C. and Canonne-Hergaux, F. (2005) Presence of the iron exporter ferroportin at the plasma membrane of macrophages is enhanced by iron loading and down-regulated by hepcidin. Blood , 106, 3979–84. 83. Oates, P.S. (2007) The role of hepcidin and ferroportin in iron absorption. Histol Histopathol , 22, 791–804. 84. Laftah, A.H., Ramesh, B., Simpson, R.J., Solanky, N., Bahram, S., Schumann, K., Debnam, E.S. and Srai, S.K. (2004) Effect of hepcidin on intestinal iron absorption in mice. Blood , 103, 3940–44. 85. Chaston, T., Chung, B., Mascarenhas, M., Marks, J., Patel, B., Srai, S.K. and Sharp, P. (2008) Evidence for differential effects of hepcidin in macrophages and intestinal epithelial cells. Gut , 57, 374–82. 86. Pietrangelo, A. (2006) Hereditary hemochromatosis. Biochim Biophys Acta, 1763, 700–10. 87. Simon, M., Bourel, M., Fauchet, R. and Genetet, B. (1976) Association of HLA-A3 and HLA-B14 antigens with idiopathic haemochromatosis. Gut , 17, 332–34. 88. Merryweather-Clarke, A.T., Pointon, J.J., Shearman, J.D. and Robson, K.J. (1997) Global prevalence of putative haemochromatosis mutations. J Med Genet , 34, 275–78. 89. Lebron, J.A., Bennett, M.J., Vaughn, D.E., Chirino, A.J., Snow, P.M., Mintier, G.A., Feder, J.N. and Bjorkman, P.J. (1998) Crystal structure of the hemochromatosis protein HFE and characterization of its interaction with transferrin receptor. Cell , 93, 111–23.

16: THE CENTRAL ROLE OF THE LIVER IN IRON STORAGE AND REGULATION OF SYSTEMIC IRON HOMEOSTASIS

90. Waalen, J. and Beutler, E. (2006) Hereditary hemochromatosis: screening and management. Curr Hematol Rep, 5, 34–40. 91. Allen, K.J., Gurrin, L.C., Constantine, C.C., Osborne, N.J., Delatycki, M.B., Nicoll, A.J., McLaren, C.E., Bahlo, M., Nisselle, A.E., Vulpe, C.D., Anderson, G.J., Southey, M.C., Giles, G.G., English, D.R., Hopper, J.L., Olynyk, J.K., Powell, L.W. and Gertig, D.M. (2008) Iron-overload-related disease in HFE hereditary hemochromatosis. N Engl J Med , 358, 221–30. 92. Merryweather-Clarke, A.T., Cadet, E., Bomford, A., Capron, D., Viprakasit, V., Miller, A., McHugh, P.J., Chapman, R.W., Pointon, J.J., Wimhurst, V.L., Livesey, K.J., Tanphaichitr, V., Rochette, J. and Robson, K.J. (2003) Digenic inheritance of mutations in HAMP and HFE results in different types of haemochromatosis. Hum Mol Genet , 12, 2241–47. 93. Waalen, J., Felitti, V.J., Gelbart, T. and Beutler, E. (2008) Screening for hemochromatosis by measuring ferritin levels: a more effective approach. Blood , 111, 3373–76. 94. Camaschella, C., Roetto, A., Cali, A., De Gobbi, M., Garozzo, G., Carella, M., Majorano, N., Totaro, A. and Gasparini, P. (2000) The gene TFR2 is mutated in a new type of haemochromatosis mapping to 7q22. Nat Genet , 25, 14–15. 95. Roetto, A., Totaro, A., Piperno, A., Piga, A., Longo, F., Garozzo, G., Cali, A., De, G.M., Gasparini, P. and Camaschella, C. (2001) New mutations inactivating transferrin receptor 2 in hemochromatosis type 3. Blood , 97, 2555–60. 96. Loreal, O., Ropert, M., Mosser, A., Dehais, V., Deugnier, Y., David, V., Brissot, P. and Jouanolle, A.M. (2007) Pathophysiology and genetics of classic HFE (type 1) hemochromatosis. Presse Med , 36, 1271–77. 97. Fleming, R.E., Ahmann, J.R., Migas, M.C., Waheed, A., Koefﬂer, H.P., Kawabata, H., Britton, R.S., Bacon, B.R. and Sly, W.S. (2002) Targeted mutagenesis of the murine transferrin receptor-2 gene produces hemochromatosis. Proc Natl Acad Sci U S A, 99, 10653–58. 98. Roetto, A., Papanikolaou, G., Politou, M., Alberti, F., Girelli, D., Christakis, J., Loukopoulos, D. and Camaschella, C. (2003) Mutant antimicrobial peptide hepcidin is associated with severe juvenile hemochromatosis. Nat Genet , 33, 21–22. 99. Lanzara, C., Roetto, A., Daraio, F., Rivard, S., Ficarella, R., Simard, H., Cox, T., Cazzola, M., Piperno, A., Gimenez-Roqueplo, A.P., Grammatico, P., Volinia, S., Gasparini, P. and Camaschella, C. (2004) The spectrum of hemojuvelin gene mutations in 1q-linked juvenile hemochromatosis. Blood , 103, 4317–21. 100. Herpin, A. and Cunningham, C. (2007) Cross-talk between the bone morphogenetic protein pathway and other major signaling pathways results in tightly regulated cell-speciﬁc outcomes. FEBS J , 274, 2977–85. 101. Halbrooks, P.J., Ding, R., Wozney, J.M. and Bain, G. (2007) Role of RGM coreceptors in bone morphogenetic protein signaling. J Mol Signal , 2, 4. 102. Niederkoﬂer, V., Salie, R. and Arber, S. (2005) Hemojuvelin is essential for dietary iron sensing, and its

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17

Disorders of Bilirubin Metabolism Namita Roy Chowdhury and Jayanta Roy Chowdhury Departments of Medicine and Genetics and Marion Bessin Liver Research Center, Albert Einstein College of Medicine, Bronx, New York, NY, USA

INTRODUCTION

BILIRUBIN METABOLISM

Bilirubin is the end product of degradation of the heme moiety of hemoproteins. Hemoglobin, derived from senescent hepatocytes, is the major source of bilirubin. Significant fractions are also derived from other hemoproteins of liver and other organs. Historically, hyperbilirubinemia has attracted the attention of clinicians as a marker of liver dysfunction. Subsequently, the studies of bilirubin chemistry, synthesis, transport, metabolism, distribution, and excretion have provided important insights into the transport, metabolism, and excretion of biologically important organic anions, particularly those with limited aqueous solubility. Bilirubin is potentially toxic, but is normally rendered harmless by tight binding to albumin in the circulation, efﬁcient extraction and storage by hepatocytes, enzyme-catalyzed detoxiﬁcation, and subsequent active biliary excretion. Patients with very high levels of unconjugated hyperbilirubinemia are at risk for bilirubin encephalopathy (kernicterus). Kernicterus is found in some cases of severe neonatal jaundice and in inherited disorders associated with severe unconjugated hyperbilirubinemia. This chapter will provide a brief summary of bilirubin metabolism and its inherited disorders, with emphasis on some signiﬁcant development since the publication of the last edition of this book.

Bilirubin is derived from the breakdown of hemoglobin released from senescent erythrocytes and other hemoproteins, particularly cytochromes and several enzymes. Of the 250–400 mg of bilirubin produced daily in normal adults, ∼80% is derived from the hemoglobin [1], while the remainder comes from more rapidly turning-over hemoproteins, primarily in the liver. Cleavage of heme at the α-methene bridge is catalyzed by microsomal heme oxygenase by a reaction requiring oxygen and a reducing agent, such as NADPH. Heme oxygenase activity is rate-limiting in bilirubin production. Inhibition of heme oxygenase by nonmetabolized “dead-end” inhibitors, such as tin-protoporphyrin or tin-mesoporphyrin, reduces serum bilirubin levels in neonates [2, 3]. The reaction opens the tetrapyrrole structure of heme, giving rise to a linear tetrapyrrole, biliverdin, and releasing 1 mol each of CO and iron. Normally, oxidation of the α-chain carbon of heme accounts for the great majority of the body’s endogenous CO production, only a small fraction being contributed by intestinal bacteria [4]. Therefore, measurement of CO production by breath analysis can be used to quantify bilirubin production and heme breakdown, which, at a steady state, equals heme synthesis. Of the three heme oxygenase isoforms, heme oxygenase 1 is ubiquitous and inducible. Biliverdin, which is subsequently reduced to

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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bilirubin by the action of cytosolic biliverdin reductase(s), has signiﬁcant antioxidant activity. The heme oxygenase proteins and their products, biliverdin and bilirubin, are thought to provide tissue protection against reactive oxygen species [5, 6], especially when the levels of other antioxidants or free-radical scavengers are low. CO, the other product of heme oxygenase, is a potent vasodilator that regulates the vascular tone in the liver and in other organs, such as the heart, under conditions of stress. With a few exceptions, in most vertebrates biliverdin is reduced to bilirubin by the action of biliverdin reductases. Bilirubin is less polar than biliverdin and requires energy-consuming metabolic steps for excretion into bile. Despite this, bilirubin has been the end product of heme catabolism in most species since the evolution of early vertebrates, such as teleost and elasmobranch ﬁsh [7]. The stronger antioxidant activity of bilirubin may be particularly important during the neonatal period, when concentrations of other antioxidants are low in body ﬂuids. Another possible advantage of bilirubin formation may be that, being more non-polar, bilirubin is more efﬁciently extracted by the placenta in intrauterine life. The mechanism by which the placenta removes bilirubin from the fetal circulation has not been elucidated fully. A few women with high serum bilirubin levels due to an inherited deﬁciency of enzymatic bilirubin glucuronidation (see below) have had successful pregnancy. In these cases, serum bilirubin levels of the newborns were much lower than those of the mothers, suggesting a mechanism by which the placenta can transport bilirubin from the fetal to the maternal circulation against a steep concentration gradient. As heme oxygenase acts speciﬁcally at the α-bridge of heme, physiologically-generated biliverdin and bilirubin are designated as biliverdin IXα and bilirubin IXα, respectively. The two dipyrrolic halves are joined by a central methane bridge. Each half has a propionic acid side chain. X-ray diffraction crystallography and nuclear magnetic resonance studies have revealed that the propionic acid side chains of bilirubin are internally hydrogen-bonded to the pyrrolic and lactam sites on the opposite half of the molecule [8]. The internal hydrogen bonding engages all polar groups and “buries” the central methane bridge that joins the two dipyrrolic halves of the molecule. Physiologically, the hydrogen bonds can be disrupted by enzyme-catalyzed conjugation of one or both propionic acid side chains, forming bilirubin mono- and diglucuronides, respectively. In the van den Bergh reaction [9] that is commonly employed in clinical analysis of serum bilirubin levels, the diazo reagents, which attack the central methane bridge, act rapidly on the non-hydrogen-bonded conjugated bilirubin (“direct”-reacting bilirubin), whereas the unconjugated fraction reacts rapidly only when the hydrogen bonds are disrupted by a chemical accelerator (“total” bilirubin). The hydrogen bonds can also be disrupted transiently by conﬁgurational isomerization of bilirubin, which occurs upon

exposure to light. These bilirubin photoproducts can be excreted into bile without conjugation, which explains the efﬁcacy of phototherapy in reducing serum unconjugated bilirubin levels. Although clinicians are generally concerned with serum bilirubin levels as an indicator of liver function and disease, and the toxicity of bilirubin, within a near-physiological range of plasma concentrations, the antioxidative action of bilirubin may provide beneﬁcial effects. Serum bilirubin levels were found to be inversely related to the risk of ischemic coronary artery disease in middle-aged men in Britain [10]. An inverse relationship between serum bilirubin levels and cancer mortality was reported [11]. The study of a large number of subjects in the USA revealed that the odds ratio for the history of colorectal cancer was reduced by 0.295 in men and 0.186 in women per 1 mg dl−1 increment in serum bilirubin concentrations [12]. These studies suggest, but do not prove, a causative role of bilirubin in reducing the incidence of several common diseases in the population.

DISORDERS OF BILIRUBIN METABOLISM At higher concentrations, unconjugated bilirubin is toxic to many cells and organelles. Since albumin binding abrogates the toxic effect of bilirubin, the harmful effects occur when unconjugated bilirubin is present in a molar excess over albumin. Such conditions are usually limited to neonates with a high level of unconjugated hyperbilirubinemia and subjects with inherited disorders of bilirubin conjugation. Markedly elevated serum unconjugated bilirubin concentrations (usually over 20 mg dl−1 ) in the newborn may result in clinical evidence of brain damage, ranging from subtle neurological abnormalities to severe encephalopathy or permanent neurologic damage (kernicterus) [13]. The serum concentration at which clinical signs of neurotoxicity occur is variable. Reduction of plasma albumin concentrations due to intercurrent illnesses or release of bilirubin from albumin binding by drugs such as sulfonamides and warfarin can precipitate kernicterus even when bilirubin is not at a molar excess over albumin. Although the immaturity of the blood–brain barrier in neonates is often held responsible for increased susceptibility of neonates to kernicterus, there is little ﬁrm evidence to support this concept. The entry of the non-albumin-bound (free) fraction of bilirubin into the brain is probably independent of the intactness of the blood–brain barrier. Damaged and edematous brain may bind bilirubin avidly, and therefore be unable to clear it rapidly, increasing the susceptibility to bilirubin encephalopathy [14]. Normally, bilirubin toxicity is prevented by tight albumin binding in circulation, rapid clearance by the liver, enzymatic detoxiﬁcation within the hepatocytes, and efﬁcient excretion in the bile. These processes are brieﬂy summarized below.

17: DISORDERS OF BILIRUBIN METABOLISM

Bilirubin circulates in plasma bound to albumin. Albumin binding keeps unconjugated bilirubin in solution and prevents its diffusion into tissues and all its toxic effects. Unconjugated bilirubin binds more tightly to albumin than does conjugated bilirubin. Therefore, in the absence of proteinuria, unconjugated bilirubin does not undergo glomerular ﬁltration signiﬁcantly. Normally, unconjugated bilirubin constitutes the great majority of plasma bile pigments. Therefore, very little bilirubin appears in the urine. When conjugated bilirubin accumulates in plasma because of acquired or inherited liver diseases, a signiﬁcant amount of conjugated bilirubin is excreted in urine. In the presence of prolonged accumulation of conjugated bilirubin in plasma, a signiﬁcant fraction of the conjugated bilirubin becomes covalently bound to albumin, producing delta-bilirubin, which is not excreted in bile or urine. This may lead to prolonged persistence of conjugated hyperbilirubinemia even after biliary obstruction or intrahepatic cholestasis is relieved. In the liver, bilirubin dissociates from albumin and is internalized by hepatocytes via facilitated diffusion. Although the transport protein SLC21A6 (OATP-2) was implicated in bilirubin internalization, this could not be conﬁrmed in subsequent studies [15]. Within the hepatocyte, binding of bilirubin to GSTs inhibits its efﬂux, thereby increasing the net uptake. Microsomal uridinediphosphoglucuronate glucuronosyltransferase type 1 (UGT1A1) catalyzes the transfer of glucuronic acid from UDP-glucuronate to bilirubin, forming monoand diglucuronides. Glucuronidation makes bilirubin water soluble, reduces its toxicity, and promotes its Bilirubin-Albumin

253

secretion into bile. Glucuronide conjugation is critical in biliary excretion of bilirubin. Signiﬁcant reduction of hepatic bilirubin glucuronidating activity results in the accumulation of unconjugated bilirubin in plasma (see below). Finally, the bilirubin glucuronides are transported into the bile canaliculi by an energy-consuming process mediated by ABCC2 (also termed MRP2) (Figure 17.1). Among the many isoforms of uridinediphosphoglucuronate glucuronosyltransferase (UGT), UGT1A1 is the only one that contributes signiﬁcantly to bilirubin glucuronidation. UGT1A1 [16] also accepts estradiol and several drugs as substrates. UGT1A1 is expressed from an unusually organized gene that expresses eight UGT1 isoforms from eight different promoters, each next to a unique exon encoding the amino terminal half of a speciﬁc isoform. The transcript initiating from each promoter is spliced to four common region exons (exons 2–5) that encode the identical carboxy-terminal half of all UGT1A isoforms [17]. The presence of a different promoter for each isoform permits its independent regulation. Mutations in any of the ﬁve exons (1A1–5) encoding UGT1A1 can cause complete or partial deﬁciency of bilirubin glucuronidation, resulting in Crigler-Najjar syndrome type 1 (CN-1) or Crigler-Najjar syndrome type 2 (CN-2), which are characterized by hyperbilirubinemia with potential cerebral injury [18, 19]. Gilbert syndrome, a milder and much more common type of inherited hyperbilirubinemia, results from a variant TATA element within the UGT1A1 promoter. This results in reduced synthesis of normal UGT1A1 protein [20].

Bilirubin + Albumin Sinusoidal surface

1 2 3 GST's

Bilirubin UDPGA

4 UGT1A1

Contiguous membrane

UDP

5

ABCC2 (MRP2)

Bilirubin monoglucuronide Bilirubin diglucuronide Canalicular surface

Figure 17.1 Summary of hepatic metabolism of bilirubin. Bilirubin is strongly bound to albumin in the circulation (1). At the sinusoidal surface of the hepatocyte, this complex dissociates, and bilirubin enters hepatocytes by fascilitated diffusion (2). This process is non-adenosin triphosphate (ATP)-dependent and bidirectional. Within the hepatocyte, bilirubin binds to a group of cytosolic proteins, mainly to glutathione-s- transferases (GSTs) (3). GST binding inhibits the efﬂux of bilirubin from the cell, thereby increasing the net uptake. A speciﬁc form of uridine diphosphoglucuronate glucuronosyltransferase, UGT1A1, located in the endoplasmic reticulum, catalyzes the transfer of the glucuronic acid moity from UDP-glucuronic acid (UDPGA) to bilirubin, forming bilirubin diglucuronide and bilirubin monoglucuronide (4). Glucuronidation is necessary for efﬁcient excretion of bilirubin in bile (5). Canalicular excretion of bilirubin and other organic anions (except most bile acids) is primarily an energy-dependent process, mediated by the ATP-utilizing transporter ABCC2, also known as multidrug-resistance-related proteins (MRP2)

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Finally, the bilirubin glucuronides are transported into the bile canaliculi by the ATP-hydrolyzing canalicular pump, ABCC2. This energy-consuming pump transports bilirubin across the bile canaliculus against a steep concentration gradient. In Dubin-Johnson syndrome (DJS), the lack of ABCC2 activity results in the accumulation of conjugated bilirubin and many other organic anions, with the exception of most bile salts. ABCC2 is also downregulated in acquired biliary obstruction and intrahepatic cholestasis. In these situations, other MRPs are expressed on the sinusoidal surface of hepatocytes, leading to transport of conjugated bilirubin from hepatocytes into the plasma, and hence to conjugated hyperbilirubinemia. Bilirubin is degraded by intestinal bacteria into a series of urobilinogen and related products [21]. Most of the urobilinogen reabsorbed from the intestine is excreted in bile, but a small fraction is excreted in urine. Absence of urobilinogen in stool and urine indicates complete obstruction of the bile duct. In liver disease and states of increased bilirubin production, urinary urobilinogen excretion is increased. Urobilinogen is colorless; its oxidation product, urobilin, contributes to the color of normal urine and stool. Normally, the capacity of the liver for bilirubin uptake, conjugation, and excretion is closely balanced, so that reduction of any of these processes can limit the rate of bilirubin throughput by the liver. On the other hand, increase of the bilirubin handling capacity of the liver, for example, in response to increased bilirubin load, requires that all these pathways are coordinately upregulated. Several nuclear receptor proteins, such as CAR and PXR, may regulate such coordinated regulation [22–24]. Predominantly unconjugated hyperbilirubin can result from increased bilirubin production resulting from hemolytic disorders or ineffective erythropoiesis. In the presence of normal liver function, increased bilirubin production does not lead to plasma levels of bilirubin exceeding 5 mg dl−1 . On the other hand, complete CN1 or partial CN2 inherited deﬁciency of UGT1A1 is associated with serum bilirubin concentrations of 20–40 mg dl−1 and 5–20 mg dl−1 , respectively. In CN2, serum bilirubin levels are reduced by at least 25% after seven days of phenobarbital therapy, which induces microsomal UGT1A1. CN1 and CN2 are rare autosomal recessive disorders caused by genetic lesions of any one of the ﬁve exons that constitute the coding region of UGT1A1 or their ﬂanking intronic sequences [18, 19]. In contrast, Gilbert syndrome is one of the most common inherited disorders in humans, in which a promoter polymorphism of UGT1A1 leads to reduced expression (∼30% of normal) of structurally normal UGT1A1 [20]. In addition to mild and usually intermittent hyperbilirubinemia, which is exacerbated on fasting, subjects with Gilbert syndrome may have increased susceptibility to several drugs, such as irinotecan and acetaminophen. In CN2 and Gilbert syndrome, the proportion of bilirubin monoglucuronide in bile (normally >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

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

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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

FIT DGA

FIT DGAT

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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THE LIVER: PATHOBIOLOGY OF HEPATIC TRIGLYCERIDE

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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THE LIVER: ADIPOSE FAT STORAGE AND HEPATIC INSULIN RESISTANCE

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

263

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

264

THE LIVER: METABOLIC FATE OF “OLD” AND “NEW” FAT

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

FAS

D

PPARα

"NEW" FAT

GNEO

N

PPARα

(De novo) CHOL

"OLD" FAT

PERIPHERAL MOBILIZATION

FAO

FATTY LIVER

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

18: HEPATIC FATTY ACID METABOLISM AND DYSFUNCTION

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

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en

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Regulate gene expression for lipid synthesis

XBP-1

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n me

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lu

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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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THE LIVER: REFERENCES

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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and reduced fat storage in mice lacking acetyl-CoA carboxylase 2. Science, 291, 2613–16. Chirala, S.S., Chang, H., Matzuk, M., Abu-Elheiga, L., Mao, J., Mahon, K., Finegold, M. and Wakil, S.J. (2003) Fatty acid synthesis is essential in embryonic development: fatty acid synthase null mutants and most of the heterozygotes die in utero. Proc Natl Acad Sci U S A, 100, 6358–63. Abu-Elheiga, L., Matzuk, M.M., Kordari, P., Oh, W., Shaikenov, T., Gu, Z. and Wakil, S.J. (2005) Mutant mice lacking acetyl-CoA carboxylase 1 are embryonically lethal. Proc Natl Acad Sci U S A, 102, 12011–16. Harada, N., Oda, Z., Hara, Y., Fujinami, K., Okawa, M., Ohbuchi, K., Yonemoto, M., Ikeda, Y., Ohwaki, K., Aragane, K. et al. (2007) Hepatic de novo lipogenesis is present in liver-speciﬁc ACC1-deﬁcient mice. Mol Cell Biol , 27, 1881–88. Mao, J., DeMayo, F.J., Li, H., Abu-Elheiga, L., Gu, Z., Shaikenov, T.E., Kordari, P., Chirala, S.S., Heird, W.C. and Wakil, S.J. (2006) Liver-speciﬁc deletion of acetyl-CoA carboxylase 1 reduces hepatic triglyceride accumulation without affecting glucose homeostasis. Proc Natl Acad Sci U S A, 103, 8552–57. Savage, D.B., Choi, C.S., Samuel, V.T., Liu, Z.X., Zhang, D., Wang, A., Zhang, X.M., Cline, G.W., Yu, X.X., Geisler, J.G. et al. (2006) Reversal of dietinduced hepatic steatosis and hepatic insulin resistance by antisense oligonucleotide inhibitors of acetyl-CoA carboxylases 1 and 2. J Clin Invest , 116, 817–24. Yu, S., Rao, S. and Reddy, J.K. (2003) Peroxisome proliferator-activated receptors, fatty acid oxidation, steatohepatitis and hepatocarcinogenesis. Curr Mol Med , 3, 561–72. Rodriguez, J.C., Gil-Gomez, G., Hegardt, F.G. and Haro, D. (1994) Peroxisome proliferator-activated receptor mediates induction of the mitochondrial 3-hydroxy-3-methylglutaryl-CoA synthase gene by fatty acids. J Biol Chem, 269, 18767–72. Koo, S.H., Satoh, H., Herzig, S., Lee, C.H., Hedrick, S., Kulkarni, R., Evans, R.M., Olefsky, J. and Montminy, M. (2004) PGC-1 promotes insulin resistance in liver through PPAR-alpha-dependent induction of TRB-3. Nat Med , 10, 530–34.

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 Editing

Intestine

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]

274

THE LIVER: BIOCHEMISTRY AND PHYSIOLOGY OF CHOLESTEROL AND LIPOPROTEIN METABOLISM

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

276

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

277

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

278

THE LIVER: BIOCHEMISTRY AND PHYSIOLOGY OF CHOLESTEROL AND LIPOPROTEIN METABOLISM

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].

19: LIPOPROTEIN METABOLISM AND CHOLESTEROL BALANCE

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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59. Grundy, S.M. and Metzger, A.L. (1972) A physiological method for estimation of hepatic secretion of biliary lipids in man. Gastroenterology, 62, 1200–17. 60. Grundy, S.M. (1983) Absorption and metabolism of dietary cholesterol. Annu Rev Nutr, 3, 71–96. 61. Reuben, A., Maton, P.N., Murphy, G.M. and Dowling, R.H. (1985) Bile lipid secretion in obese and non-obese individuals with and without gallstones. Clin Sci , 69, 71–79. 62. Northﬁeld, T.C. and Hofmann, A.F. (1975) Biliary lipid output during three meals and an overnight fast. I. Relationship to bile acid pool size and cholesterol saturation of bile in gallstone and control subjects. Gut , 16, 1–11. 63. Cohen, D.E. (2008) Balancing cholesterol synthesis and absorption in the gastrointestinal tract. J Clin Lipidol , 2, S1–S3. 64. Sehayek, E., Nath, C., Heinemann, T., McGee, M., Seidman, C.E., Samuel, P. and Breslow, J.L. (1998) U-shape relationship between change in dietary cholesterol absorption and plasma lipoprotein responsiveness and evidence for extreme interindividual variation in dietary cholesterol absorption in humans. J Lipid Res, 39, 2415–22. 65. Bosner, M.S., Lange, L.G., Stenson, W.F. and Ostlund, R.E. (1999) Percent cholesterol absorption in normal women and men quantiﬁed with dual stable isotopic tracers and negative ion mass spectrometry. J Lipid Res, 40, 302–8. 66. Expert Panel on Detection, Evaluation, and Treatment of High Blood Cholesterol in Adults (2001) Executive summary of the third report of The National Cholesterol Education Program (NCEP) expert panel on detection, evaluation, and treatment of high blood cholesterol in adults (adult treatment panel III). JAMA, 285, 2486–97. 67. Grundy, S.M., Cleeman, J.I., Merz, C.N., Brewer, H.B. Jr., Clark, L.T., Hunninghake, D.B., Pasternak, R.C., Smith, S.C. Jr. and Stone, N.J. (2004) Implications of recent clinical trials for the National Cholesterol Education Program Adult Treatment Panel III Guidelines. J Am Coll Cardiol , 44, 720–32. 68. Einarsson, K., Ericsson, S., Ewerth, S., Reihner, E., Rudling, M., Stahlberg, D. and Angelin, B. (1991) Bile acid sequestrants: mechanisms of action on bile acid and cholesterol metabolism. Eur J Clin Pharmacol , 40 (Suppl 1), S53–58. 69. Watanabe, M., Houten, S.M., Wang, L., Moschetta, A., Mangelsdorf, D.J., Heyman, R.A., Moore, D.D. and Auwerx, J. (2004) Bile acids lower triglyceride levels via a pathway involving FXR, SHP, and SREBP-1c. J Clin Invest , 113, 1408–18. 70. Turley, S.D. and Dietschy, J.M. (2003) The intestinal absorption of biliary and dietary cholesterol as a drug target for lowering the plasma cholesterol level. Prev Cardiol 6, 29–33, 64. 71. Armstrong, M.J. and Carey, M.C. (1987) Thermodynamic and molecular determinants of sterol solubilities in bile salt micelles. J Lipid Res, 28, 1144–55. 72. Davis, H.R. Jr., Zhu, L.J., Hoos, L.M., Tetzloff, G., Maguire, M., Liu, J., Yao, X., Iyer, S.P., Lam, M.H., Lund, E.G., Detmers, P.A., Graziano, M.P. and Altmann,

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S.W. (2004) Niemann-Pick C1 Like 1 (NPC1L1) is the intestinal phytosterol and cholesterol transporter and a key modulator of whole-body cholesterol homeostasis. J Biol Chem, 279, 33586–92. Zandbergen, F. and Plutzky, J. (2007) PPARalpha in atherosclerosis and inﬂammation. Biochim Biophys Acta, 1771, 972–82. Mandard, S., Muller, M. and Kersten, S. (2004) Peroxisome proliferator-activated receptor alpha target genes. Cell Mol Life Sci , 61, 393–416. Sacks, F.M. (2006) The apolipoprotein story. Atheroscler Suppl , 7, 23–27. Aalto-Setala, K., Fisher, E.A., Chen, X., Chajek-Shaul, T., Hayek, T., Zechner, R., Walsh, A., Ramakrishnan, R., Ginsberg, H.N. and Breslow, J.L. (1992) Mechanism of hypertriglyceridemia in human apolipoprotein (apo) CIII transgenic mice. Diminished very low density lipoprotein fractional catabolic rate associated with increased apo CIII and reduced apo E on the particles. J Clin Invest , 90, 1889–900. Duval, C., Muller, M. and Kersten, S. (2007) PPAR alpha and dyslipidemia. Biochim Biophys Acta, 1771, 961–71. Millar, J.S., Duffy, D., Gadi, R., Bloedon, L.T., Dunbar, R.L., Wolfe, M.L., Movva, R., Shah, A., Fuki, I.V., McCoy, M., Harris, C.J., Wang, M.D., Howey, D.C. and Rader, D.J. (2009) Potent and selective PPAR-{alpha} agonist LY518674 upregulates both ApoA-I production and catabolism in human subjects with the metabolic syndrome. Arterioscler Thromb Vasc Biol , 29, 140–6. Kamanna, V.S. and Kashyap, M.L. (2008) Mechanism of action of niacin. Am J Cardiol , 101, 20B–26B. Tunaru, S., Kero, J., Schaub, A., Wufka, C., Blaukat, A., Pfeffer, K. and Offermanns, S. (2003) PUMA-G and HM74 are receptors for nicotinic acid and mediate its anti-lipolytic effect. Nat Med , 9, 352–55. Shepherd, J., Packard, C.J., Patsch, J.R., Gotto, A.M. Jr. and Taunton, O.D. (1979) Effects of nicotinic acid therapy on plasma high density lipoprotein subfraction distribution and composition and on apolipoprotein A metabolism. J Clin Invest , 63, 858–67. Bays, H. (2006) Clinical overview of Omacor: a concentrated formulation of omega-3 polyunsaturated fatty acids. Am J Cardiol , 98, 71i–76i. Davidson, M.H. (2006) Mechanisms for the hypotriglyceridemic effect of marine omega-3 fatty acids. Am J Cardiol , 98, 27i–33i. Scappa, M.C., Kanno, K. and Cohen, D.E. (2007) Lipoprotein metabolism, In: The Textbook of Hepatology: From Basic Science to Clinical Practice, 3rd Edition (eds J. Rod´es, J.P. Benhamou, M.A. Rizzetto, J. Reichen and A. Blei), Blackwell, Oxford. Cohen, D.E. and Armstrong, E.J. (2007) Pharmacology of Cholesterol and Lipoprotein Metabolism, In: Principles of Pharmacology: The Pathophysiologic Basis of Drug Therapy (eds D.E. Golan, A.H. Tashjian, E.J. Armstrong, J.M. Galanter, A.W. Armstrong, R.A. Arnaout and H.S. Rose), 2nd edn, Lippincott, Williams and Wilkins, Philadelphia.

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

291

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.

20: BILE ACIDS AND THE ENTEROHEPATIC CIRCULATION

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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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.

20: BILE ACIDS AND THE ENTEROHEPATIC CIRCULATION

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

1. Nair, P.P. and Kritchevsky, D. (1971, 1988) The Bile Acids: Chemistry, Physiology, and Metabolism, Vols. 1–4, Plenum Press, New York. 2. Danielsson, H., Sj¨ovall, J., Neuberger, A. and Van Deenen, L.L.M. (eds) (1985) Sterols And Bile Acids: New Comprehensive Biochemistry, Elsevier, Amsterdam, p. 447. 3. Dawson, P.A., Shneider, B.L. and Hofmann, A.F. (2006) Bile formation and the enterohepatic circulation, in Physiology of the Gastrointestinal Tract , 4th edn, Vol. 2 (eds K.E. Barrett, F.K. Ghishan, J.L. Merchant, H.M. Said, J.D. Wood and L.R. Johnson), Elsevier Academic Press, San Diego, pp. 1437–62. 4. Hofmann, A.F. (2008) The enterohepatic circulation of bile acids in mammals: form and functions. Front Biosci , 14, 2584–98. 5. Hofmann, A.F. and Hagey, L.R. (2008) Bile acids: chemistry, pathochemistry, biology, pathobiology, and therapeutics. Cell Mol Life Sci , 65, 2461–83. 6. Keppler, D., Beuers, U., Leuschner, U., Stiehl, A., Trauner, M. and Paumgartner, G. (eds) (2007) Bile Acids: Biological Actions and Clinical Relevance, Springer, Dordrecht, p. 265.

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7. Russell, D.W. (2003) The enzymes, regulation, and genetics of bile acid synthesis. Annu Rev Biochem, 72, 137–74. 8. Norlin, M. and Wikwall, K. (2007) Enzymes in the conversion of cholesterol into bile acids. Curr Mol Med , 7, 199–218. 9. Hofmann, A.F. and Hoffman, N.E. (1974) Measurement of bile acid kinetics by isotope dilution in man. Gastroenterology, 67, 314–23. 10. Hofmann, A.F. and Mysels, K.J. (1988) Bile salts as biological surfactants. Colloids Surf , 39, 145–73. 11. Hofmann, A. and Mysels, K.J. (1992) Bile acid solubility and precipitation in vitro and in vivo: the role of conjugation, pH, and Ca2+ ions. J Lipid Res, 33, 617–26. 12. Ridlon, J.M., Kang, D.J. and Hylemon, P.B. (2006) Bile salt biotransformations by human intestinal bacteria. J Lipid Res, 47, 241–59. 13. Hamilton, J.P., Xie, G., Raufman, J.-P., Hugan, S., Grifﬁn, T.L., Packard, C.A., Chatﬁeld, D.A., Hagey, L.R., Steinbach, J.H. and Hofmann, A.F. (2007) Bile acids of human cecal contents: concentration and spectrum. Am J Physiol , 293, 256–63. 14. Hofmann, A.F. and Eckmann, L. (2006) How bile acids confer gut mucosal protection against bacteria. Proc Natl Acad Sci U S A, 103, 4333–34. 15. LaRusso, N.F., Hoffman, N.E., Hofmann, A.F., Northﬁeld, T.C. and Thistle, J.L. (1975) Effect of primary bile acid ingestion on bile acid metabolism and biliary lipid secretion in gallstone patients. Gastroenterology, 69, 1301–14. 16. Rossi, S.S., Converse, J.L. and Hofmann, A.F. (1987) High pressure liquid chromatographic analysis of conjugated bile acids in human bile: simultaneous resolution of sulfated and unsulfated lithocholyl amidates and the common conjugated bile acids. J Lipid Res, 28, 589–95. 17. Amelsberg, A., Schteingart, C.D. and Ton-Nu, H.-T. (1966) Hofmann AF Carrier-mediated jejunal absorption of conjugated bile acids in the guinea pig. Gastroenterology, 110, 1098–106. 18. Dawson, P.A., Haywood, J., Craddock, A.L., Wilson, M., Tietjen, M., Kluckman, K., Maeda, N. and Parks, J.S. (2003) Targeted deletion of the ileal bile acid transporter eliminates enterohepatic cycling of bile acids in mice. J Biol Chem, 278, 33920–27. 19. Oelkers, P., Kirby, L.C., Heubi, J.E. and Dawson, P.A. (1997) Primary bile acid malabsorption caused by mutations in the ileal sodium-dependent bile acid transporter gene (SLC10A2). J Clin Invest , 99, 1880–87. 20. Dawson, P.A., Hubbert, M., Haywood, J., Craddock, A.L., Zeranque, N., Christian, W.V. and Ballatori, N. (2005) The heteromeric organic solute transporter alpha-beta OSTalpha-OSTbeta is an ileal basolateral bile acid transporter. J Biol Chem, 280, 6960–68. 21. Hofmann, A.F., Molino, G., Milanese, M. and Belforte, G. (1983) Description and simulation of a physiological pharmacokinetic model for the metabolism and enterohepatic circulation of bile acids in man. Cholic acid in healthy man. J Clin Invest , 71, 1003–22. 22. Hofmann, A.F., Loening-Baucke, V., Lavine, J.E., Hagey, L.R., Steinbach, J.H., Packard, C.A., Grifﬁn,

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21

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

INTRODUCTION

excretion of a number of typical organic anions, including bile acids, for which the liver plays an essential role.

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)

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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

THE LIVER: HEPATOCYTE UPTAKE OF BILE ACIDS PI3K PIP2

activates

PIP3

LY294002

motile vesicles

PKCz

PKCzPS (a)

Motility of ntcp-containing vesicles (% of total MT-bound)

35

(289)

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(234)

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(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

(570) 30 25

* (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].

21: HEPATOCYTE BASOLATERAL MEMBRANE ORGANIC ANION TRANSPORTERS

Vector

311

ntcp-GFP (a)

70

(161)

(171)

60 (224)

ntcp-GFP vesicles

% motile vesicles

50

Late ASOR vesicles 40 (323)

30 20

(241) 10 (174)

0 0

25

50

75

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[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