Signaling Pathways in Liver Diseases 2nd ed

Signaling Pathways in Liver Diseases

Jean-Francois Dufour Pierre-Alain Clavien (Eds.)

Signaling Pathways in Liver Diseases Second edition

Prof. Jean-François Dufour University of Bern Institute of Clinical Pharmacology and Visceral Research Murtenstrasse 35 3010 Bern Switzerland [email protected]

Prof. Dr. Pierre-Alain Clavien Swiss HPB (Hepato-Pancreatico Biliary) Center University Hospital of Zurich Department of Visceral and Transplant Surgery Rämistrasse 100 8091 Zurich Switzerland [email protected]

ISBN: 978-3-642-00149-9     e-ISBN: 978-3-642-00150-5 DOI: 10.1007/978-3-642-00150-5 Springer Heidelberg Dordrecht London New York Library of Congress Control Number: 2009931701 © Springer-Verlag Berlin Heidelberg 2010 This work is subject to copyright. All rights are reserved, whether the whole or part of the material is ­concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilm or in any other way, and storage in data banks. Duplication of this publication or parts thereof is permitted only under the provisions of the German Copyright Law of September 9, 1965, in its current version, and permission for use must always be obtained from Springer. Violations are liable to prosecution under the German Copyright Law. The use of general descriptive names, registered names, trademarks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant ­protective laws and regulations and therefore free for general use. Product liability: The publishers cannot guarantee the accuracy of any information about dosage and application contained in this book. In every individual case the user must check such information by consulting the relevant literature. Cover design: eStudio Calamar, Figueres/Berlin Printed on acid-free paper Springer is part of Springer Science+Business Media (www.springer.com)

Preface

Recent advances in diagnostic and therapeutic approaches to liver disease have carried hepatology to new frontiers. The increasing frequency with which steatotic and cirrhotic livers undergo complex curative treatment strategies is a challenge to liver surgeons and hepatologists, who need to understand the molecular mechanisms at play in these situations. Comprehension of the signaling pathways participating in liver regeneration, hepatocellular apoptosis, and ischemia/reperfusion injury is essential. This book serves as a source of information to facilitate the reading of the literature and the planning of trials. Translational medicine implies knowledge of the molecular targets of novel therapeutic strategies. It is our goal to stimulate research that leads to exchanges between the laboratory, the clinical ward, and the operating room. Such a comprehensive insight including molecular and cellular events will pave the way for improvement of pharmacological and surgical interventions in complex liver disease. Bern, Switzerland Zurich, Switzerland

Jean-Francois Dufour Pierre-Alain Clavien

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Contents

Part I  The Cell Types and the Matrix . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

  1 Hepatocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Giuliano Ramadori and Pierluigi Ramadori

3

  2 Signaling Pathways in Biliary Epithelial Cells . . . . . . . . . . . . . . . . . . . . M. Fatima Leite, Viviane A. Andrade, and Michael H. Nathanson

25

  3 Stellate Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fabio Marra, Sara Galastri, Sara Aleffi, and Massimo Pinzani

41

  4 Signaling Pathways in Liver Diseases Kupffer Cells . . . . . . . . . . . . . . Christian J. Steib and Alexander L. Gerbes

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  5 Hepatic Sinusoidal Endothelial Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . Robert C. Huebert and Vijay H. Shah

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  6 Extracellular Matrix . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Scott L. Friedman

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  7 Platelets: A New Cell Type in Liver Physiology . . . . . . . . . . . . . . . . . . . 105 Mickael Lesurtel and Pierre-Alain Clavien   8 Immune Cell Communication and Signaling Systems in Liver Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 Ricky H. Bhogal and Simon C. Afford Part II  The Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 147   9 Toll-Like Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 149 Gyongyi Szabo and Pranoti Mandrekar 10 TNF/TNF Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161 Jörn M. Schattenberg and Mark J. Czaja 11 Fas/FasL . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 179 Maria Eugenia Guicciardi and Gregory J. Gores

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12 Interferon Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189 Markus H. Heim 13 NF-kB . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 201 Tom Luedde and Christian Trautwein 14 JNKs in liver diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 215 R. Schwabe 15 Insulin Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 229 Miran Kim and Jack R. Wands 16 Role of PKB/Akt in Liver Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 243 Elena Zhuravleva, Oliver Tschopp, and Brian A. Hemmings 17 Targeting mTOR Signaling Pathways in Liver Diseases . . . . . . . . . . . . 261 Hala E. Thomas and Sara C. Kozma 18 AMP-Activated Protein Kinase in Liver . . . . . . . . . . . . . . . . . . . . . . . . . 275 Louis Hue, Laurent Bultot, and Mark H. Rider 19 Er Stress Signaling in Hepatic Injury . . . . . . . . . . . . . . . . . . . . . . . . . . 287 Cheng Ji and Neil Kaplowitz 20 PPARa, A Key Regulator of Hepatic Energy Homeostasis in Health and Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 305 Nicolas Leuenberger and Walter Wahli 21 Bile Acids and Their Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 317 Thierry Claudel and Michael Trauner 22 Signaling Pathways in Liver Diseases: PXR and CAR . . . . . . . . . . . . . 333 Catherine A.M. Stedman, Michael Downes, and Christopher Liddle 23 p53 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 345 Wen-Wei Tsai and Michelle Craig Barton 24 The MYC Network and Cancer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 359 Snorri S. Thorgeirsson and Valentina M. Factor 25 The WNT/b-Catenin Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 367 Satdarshan P. S. Monga 26 Sonic Hedgehog Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 393 Alessia Omenetti and Anna Mae Diehl

Contents

Contents

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27 Hypoxia-Inducible Factor-1 Signaling System . . . . . . . . . . . . . . . . . . . . 403 Deborah Stroka and Daniel Candinas 28 VEGF Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 421 David Semela and Jean-François Dufour 29 Apoptosis and Mitochondria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 439 Jose C. Fernández-Checa and Carmen Garcia-Ruiz 30 Calcium Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 455 Thierry Tordjmann 31 HBV Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 465 Massimo Levrero and Laura Belloni 32 Hepatitis C Virus and Insulin Signaling . . . . . . . . . . . . . . . . . . . . . . . . . 483 Francesco Negro and Sophie Clément 33 MicroRNAs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 493 Onpan Cheung and Arun J. Sanyal 34 Hepatic Clocks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 501 Ueli Schibler, Gad Asher, Camille Saini, Jörg Morf, and Hans Reinke Answers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 513 Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 515

Contributors

Simon C. Afford  The Liver Research Group, School of Infection and Immunity, College of Medicine and Dentistry, University of Birmingham, UK Sara Aleffi  Dipartimento di Medicina Interna, Università degli Studi di Firenze, Firenze, Italy Viviane A. Andrade  Department of Biochemistry and Immunology, UFMG, Belo Horizonte, Brazil Gad Asher  Department of Molecular Biology and National Center of Competence in Research “Frontiers in Genetics”, Sciences III, University of Geneva, 30, Quai Ernest Ansermet, CH-1211, Geneva-4, Switzerland Michelle Craig Barton  Department of Biochemistry and Molecular Biology, University of Texas M.D. Anderson Cancer Center, 1515 Holcombe Blvd., Houston, TX 77030, USA Laura Belloni  Rome Oncogenomic Center, CRS-Regina Elena Cancer Center, Rome, Italy Ricky H. Bhogal  The Liver Research Group, School of Infection and Immunity, College of Medicine and Dentistry, University of Birmingham, UK Daniel Candinas  Clinic of Visceral Surgery and Medicine, Inselspital, University of Bern, Switzerland Thierry Claudel  Laboratory of Experimental and Molecular Hepatology, Division of Gastroenterology and Hepatology, Department of Medicine, Medical University, Graz, Austria Pierre-Alain Clavien  Swiss HPB Center, Department of Visceral and Transplant Surgery, University Hospital of Zurich, Rämistrasse 100, 8091 Zürich, Switzerland Onpan Cheung  Department of Internal Medicine, Division of Gastroenterology, Hepatology and Nutrition, Virginia Commonwealth University Medical Center, Richmond, VA 23298, USA Sophie Clément  Division of Clinical Pathology, University Hospitals, Geneva, Switzerland Mark J. Czaja  Marion Bessin Liver Research Center, Albert Einstein College of Medicine, 1300 Morris Park Avenue, Bronx, NY 10461, USA xi

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Anna Mae Diehl  Department of Medicine, Division of Gastroenterology, Duke University Medical Center, GSRBI, 595 LaSalle Street, Suite 1073, Box 3256, Durham, NC 27710, USA Michael Downes  Howard Hughes Medical Institute and Gene Expression Laboratory, The Salk Institute for Biological Studies, 10010 Torrey Pines Road, La Jolla, CA 92037, USA Jean-François Dufour  Institute of Clinical Pharmacology and Visceral Research, University of Bern, Murtenstrasse 35, 3010 Bern, Switzerland Valentina M. Factor  Laboratory of Experimental Carcinogenesis, Center for Cancer Research, National Cancer Institute, National Institutes of Health, 37 Convent Drive, Bethesda, MD 20892, USA Jose C. Fernández-Checa  Liver Unit and Centro de Investigaciones Biomédicas Esther Koplowitz, IMDiM, Hospital Clínic i Provincial and CIBEREHD, IDIBAPS, C/Villarroel 170, 08036-Barcelona, Spain Scott L. Friedman  Division of Liver Diseases, Mount Sinai School of Medicine, 1425 Madison Avenue, Box 1123, New York, NY 10029-6574, USA Sara Galastri  Dipartimento di Medicina Interna, Università degli Studi di Firenze, Firenze, Italy Carmen Garcia-Ruiz  Department of Cell Death and Proliferation, Instituto Investigaciones Biomédicas de Barcelona, Consejo Superior de Investigaciones Científicas, 08036 B arcelona, Spain Alexander L. Gerbes  Department of Medicine II, Ludwig-Maximilians-University, Klinikum Großhadern, Marchioninistraβe 15, 81377 Munich, Germany Gregory J. Gores  Mayo Clinic College of Medicine, 200 First Street SW, Rochester, MN 55905, USA Maria Eugenia Guicciardi  Miles and Shirley Fiterman Center for Digestive Diseases, Division of Gastroenterology and Hepatology, Mayo Clinic College of Medicine, 200 First Street SW, Rochester, MN 55905, USA Markus H. Heim  Division of Gastroenterology and Hepatology, University Hospital, Basel, 4031 Basel, Switzerland Brian A. Hemmings  Friedrich Miescher Institute for Biomedical Research, Maulbeerstrasse 66, 4058 Basel, Switzerland Louis Hue  Hormone and Metabolic Research Unit, Université Catholique de Louvain, and de Duve Institute Avenue, Hippocrate 75 UCL 7529, B-1200 Brussels, Belgium Robert C. Huebert  GI Research Unit, Guggenheim 10, Mayo Clinic, 200 First Street SW, Rochester, MN 55905, USA Cheng Ji  Keck School of Medicine, University of Southern California, 2011 Zonal Avenue, HMR 101, Los Angeles, CA 90033, USA

Contributors

Contributors

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Neil Kaplowitz  Department of Medicine, USC–UCLA Research Center for Alcoholic, and Pancreatic Diseases, and USC Research Center for Liver Diseases, Keck School of Medicine, University of Southern California, Los Angeles, CA, USA Miran Kim  The Liver Research Center, Alpert Medical School of Brown University, Providence, RI 02903, USA Sara C. Kozma  Genome Research Institute, 2180 E. Galbraith Road, University of Cincinnati, Cincinnati, OH 45237, USA M. Fatima Leite  Department of Physiology and Biophysics, UFMG, Belo Horizonte, Brazil Mickael Lesurtel  Swiss HPB (Hepato-Pancreatico- Biliary) Center, Department of Surgery, University Hospital of Zurich, Zurich, Switzerland Nicolas Leuenberger  Center for Integrative Genomics and National Research Center Frontiers in Genetics, University of Lausanne, Switzerland Massimo Levrero  Dipartimento di Medicina Interna, Sapienza Universita’ di Roma, Policlinico Umberto I, Viale del Policlinico 155, 0061 Rome, Italy Christopher Liddle  Department of Clinical Pharmacology, Storr Liver Unit, Westmead Millennium Institute and University of Sydney, Westmead Hospital, Westmead NSW 2145, Australia Tom Luedde  Medical Department III, University Hospital RWTH Aachen, Pauwelsstraβe 30, 52074 Aachen, Germany Pranoti Mandrekar  Department of Medicine, University of Massachusetts Medical School, Worcester, MA 01605, USA Fabio Marra  Dipartimento di Medicina Interna, Università di Firenze, Viale Morgagni, 85 50134 Firenze, Italy Satdarshan P.S. Monga  Division of Experimental Pathology, University of Pittsburgh, School of Medicine, Pittsburgh, PA 15261, USA Jörg Morf  Department of Molecular Biology and National Center of Competence in Research “Frontiers in Genetics”, Sciences III, University of Geneva, Quai Ernest Ansermet 30, 1211 Geneva-4, Switzerland Michael H. Nathanson  Department of Medicine, Yale University School of Medicine, 1 Gilbert Street, New Haven, CT 06520-8019, USA Francesco Negro  Departments of Internal Medicine and Pathology and Immunology, University of Geneva Medical Center, 1 Rue Michel-Servet, 1205 Geneva, Switzerland Alessia Omenetti  Department of Medicine, Division of Gastroenterology, Duke University Medical Center, Durham, NC, USA Massimo Pinzani  Dipartimento di Medicina Interna, Università degli Studi di Firenze, Firenze, Italy

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Giuliano Ramadori  Department of Internal Medicine, Section of Gastroenterology and Endocrinology, Georg-August-University Göttingen, Robert-Koch-Straβe 40, 37075 Göttingen, Germany Pierluigi Ramadori  Department of Internal Medicine, Section of Gastroenterology and Endocrinology, Georg-August-University Göttingen, Robert-Koch-Straβe 40, 37075 Göttingen, Germany Hans Reinke  Department of Molecular Biology and National Center of Competence in Research “Frontiers in Genetics”, Sciences III, University of Geneva, Quai Ernest Ansermet 30, 1211 Geneva-4, Switzerland Mark H. Rider  Université Catholique de Louvain, de Duve Institute, Brussels, Belgium Arun Sanyal  Department of Internal Medicine, Division of Gastroenterology, Hepatology and Nutrition, Virginia Commonwealth University Medical Center, Richmond, VA 23298, USA Jörn M. Schattenberg  I. Department of Medicine, Johannes Gutenberg University, 5501 Mainz, Germany Ueli Schibler  Department of Molecular Biology and National Center of Competence in Research “Frontiers in Genetics”, Sciences III, University of Geneva, Quai Ernest Ansermet 30, 1211, Geneva-4, Switzerland Robert Schwabe  Department of Medicine, Columbia University, Russ Berrie Pavilion, Room 415, 1150 St. Nicholas Avenue, New York, NY 10032, USA David Semela  Division of Gastroenterology and Hepatology, University Hospital Basel, Basel, Switzerland Vijay Shah  GI Research Unit, Guggenheim 10, Mayo Clinic, 200 First Street SW, Rochester, MN 55905, USA Catherine A.M. Stedman  Department of Gastroenterology, Christchurch Hospital and University of Otago, Christchurch, Private Bag 4710, Christchurch, New Zealand Christian Steib  Department of Medicine II, Ludwig-Maximilians-University, Klinikum Großhadern, Munich, Germany Deborah Stroka  Visceral Surgery Research Laboratory, Department of Clinical Research, University of Bern, Murtenstrasse 35, 3010 Bern, Switzerland Gyongyi Szabo  Department of Medicine, University of Massachusetts Medical School, 364 Plantation Street, Worcester, MA 01605, USA Hala E. Thomas  Genome Research Institute, 2180 E. Galbraith Road, University of Cincinnati, Cincinnati, OH 45237, USA Snorri S. Thorgeirsson  Laboratory of Experimental Carcinogenesis, Center for Cancer Research, National Cancer Institute, National Institutes of Health, 37 Convent Drive, Bethesda, MD 20892, USA

Contributors

Contributors

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Thierry Tordjmann  INSERM U757, Université Paris Sud, Bât. 443, 91405, Orsay, France Michael Trauner  Laboratory of Experimental and Molecular Hepatology, Division of Gastroenterology and Hepatology, Department of Internal Medicine, Medical University, Graz, Stiftingalstrasse 24, 8010 Graz, Austria Christian Trautwein  Medical Department III, University Hospital RWTH Aachen, Pauwelsstraβe 30, 52074 Aachen, Germany Wen-Wei Tsai  Department of Biochemistry and Molecular Biology, Program in Genes and Development, Graduate School of Biomedical Sciences, Center for Cancer Epigenetics, University of Texas M.D. Anderson Cancer Center, Houston, TX 77030, USA Oliver Tschopp  Clinic of Endocrinology, Diabetes and Clinical Nutrition, University Clinic Zurich, Rämistrasse 100, 8091 Zurich, Switzerland Walter Wahli  Center for Integrative Genomics, National Research Center Frontiers in Genetics, University of Lausanne, Genopode Building, 1015 Lausanne, Switzerland Jack R. Wands  The Liver Research Center, Alpert Medical School of Brown University, 55 Claverick Street, 4th Floor, Providence RI 02903, USA Elena Zhuravleva  Friedrich Miescher Institute for Biomedical Research, Maulbeerstrasse 66, 4058 Basel, Switzerland

Part The Cell Types and the Matrix

I

1

Hepatocytes Giuliano Ramadori and Pierluigi Ramadori

Introduction The liver is the largest organ of the body. Its weight (1.5–1.8  kg) represents about 2% of the total human body weight. The anatomical location is of course linked to its function. The liver function is comparable to that of the stomach, intestine, pancreas, and kidney together. In fact, all nutrients resulting from the digestion of the food are taken up by the intestine and then by the liver. Furthermore, the liver is responsible for the synthesis of most of the serum proteins and by this means for the oncotic pressure and the retention of water within the vessels. The liver stores nutrients and the energy derived from the oxidation of the nutrients. However, the liver is not only a power plant but also a cleaning device. In fact, the direct relationship with the intestine is not without danger. The large intestine despite the reabsorption of water contains an enormous number of bacteria and an enormous amount of their products. The bacteria and their products can reach the venous blood and the liver sinusoid where they are taken up and digested. Although the liver is made of several cell populations (Table 1.1), the most abundant cell type by mass and number is the hepatocyte. The human liver is made of 80 × 109 hepatocytes. To understand best the functions of the hepatocytes, it is useful to look at the laboratory findings of a 60-year old lady who developed jaundice followed by loss of appetite and was admitted

G. Ramadori () Department of Internal Medicine, Section of Gastroenterology and Endocrinology, Georg-August-University Göttingen, Robert-Koch-Strabe 40, 37075 Göttingen, Germany e-mail: [email protected]

to the university clinic because of reduced liver function. The following parameters describe her clinical situation well: serum bilirubin levels 27.7 mg/dl (normal <2), tromboplastin time 29% (normal >60%), SGPT of 2,850  U/l, and SGOT 2,553  U/l (normal <40  U/l for both) (Fig.  1.1). Further, microscopical analysis of liver biopsy showed massive areas of necrosis with increased deposition of connective tissue and presence of regenerative nodules with proliferating hepatocytes. An immunostaining for the proliferation marker Ki67 revealed that all the intact hepatocytes were in a proliferative phase. As can be seen from the follow-up study, regeneration succeeded and liver function recovered almost fully, 2 months later. The case is paradigmatic for the consequence of the lack of sufficient functional liver cell mass and the enormous capacity of hepatocytes to regenerate (Fig. 1.2).

Hepatocyte Development Recent studies on different mammalian species indicate that during embryonic development the hepatic genes are induced in a segment of the ventral endoderm through the activation of specific transcription factors (Foxa2, GATA-4, C/EBPb, Nf-1) forming a complex able to bind the chromatin upstream liver specific genes such as albumin [109]. The subsequent cell migration and organogenesis depend tightly on the orchestration of inductive signals between epithelial cells, mesenchymal cells, and endothelial cells. Experimental stimulation of in vitro embryo cultures suggests that fibroblast growth factor (Fgf) signaling from the cardiogenic mesoderm induces the liver budding in the ventral foregut endoderm [2]. Moreover, Bone morphogenetics proteins (Bmp-2 and Bmp-4)

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_1, © Springer-Verlag Berlin Heidelberg 2010

3

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G. Ramadori and P. Ramadori

Table 1.1  Liver resident cell types Parenchymal cells Resident immune cells

Hepatocytes Kuffer Cells NK-lymphocytes [57] T-Lymphocytes (e.g.,NK1.1 Ag+T cells) Sinusoidal endothelial cells Hepatic stellate cells liver-myofibroblasts portal fibroblasts

Specialized cells Mesenchymal cells with fibrogenic potential

produced by the septum transversum mesenchyme cells have been shown to be crucial for hepatogenesis [3]. The growth and the organization of the hepatic bud require dynamic processes of hepatoblast migration and cell–cell interaction with the continuous disruption and remodeling of the extra-cellular matrix (ECM) – steps that are regulated by important transcription factors such as Hex and Prox-1 [4]. A recent study illustrates how the angioblast, precursor of endothelial cells, also influences the liver bud outgrow providing an important growth stimulus prior to the formation of a local vasculature and proliferation of hepatocyte, attributing an organogenic role to the endothelial compartment too [5].

a

3000

The hepatoblast is considered to be the primitive hepatocyte precursor. However, it is not conclusively established whether there is only one type of hepatoblast or there is a hierarchy of lineage progression consisting of primitive hepatoblasts and stronger committed bipotential precursors. While the mesenchymal cells migrating from the septum transversum might give rise to the stromal cells of the liver, the hepatoblast is supposed to follow mainly a bipotential polarization: hepatocyte and biliary epithelial cell. Although this aspect requires more evidence, in experiments conducted with knockout mouse embryos, the absence of the transcription factor, hepatic nuclear factor 6 (HNF6), has been observed to be related with the absence of ALT and AST values in time course

ALT AST

U/l

2000

1000

0

Fig. 1.1  Biochemical parameters from clinical chemistry determined at different time points, before and during hospitalization. (a) Serum transaminases levels increased over 2,500 U/l within 2 months and declined to normal levels 20 days after. (b) Serum bilirubin levels and quick percentage describe very well the changes of the hepatic functional activity during damage and regeneration

b

110 100 90

26.02.2008

12.04.2008

14.04.2008

16.04.2008

28.04.2008

05.06.2008

Bilirubin and Quick values in time course Bilirubin (mg/dl) Quick (%)

80 70 60 50 40 30 20 10 0

26.02.2008 12.04.2008 14.04.2008 16.04.2008 28.04.2008 05.06.2008

1  Hepatocytes

the gall bladder and an excess of the biliary cell population, suggesting a regulatory role of this protein in the hepatoblast-­biliary shift [6]. A similar morphological picture was observed in conditionally homozygous hepatoblast for the transcription factor HNF-1b [7]. An important contribution to the biliary cells differentiation seems to be afforded by the Notch pathway as well. Mutations in the Notch pathway are in fact related in humans with the Alagille syndrome, a disease characterized by a reduced number of intrahepatic ducts [8]. A solid experimental evidence is given by observations on knockout Jag1 or Notch-2 mice, showing an analog phenotype at the birth; a more detailed analysis suggests an involvement of this pathway in the bile duct morphogenesis and remodeling rather than in the hepatoblast fate differentiation [9]. In humans, the progenitor cells are immunoreactive for cytokeratins 8, 18, 19, and 14. Most of the progen-itor cells develop to adult hepatocytes while losing cytokeratin 14 and 19; the latter is no longer detectable in human hepatoblasts at 20-week gestation [10, 11]. Using this marker together with the transcription factor Prox-1, as an hepatoblast marker, we detected CK-19+/Prox-1 small cells in cell cultures obtained from fetal rat liver at ED 14 [12], which indicate a possible different developmental pathway for biliary cells. The synthesis of alpha-fetoprotein (AFP) has the characteristic of the hepatoblast, which begins in the human liver as early as day 29. Although it is known that albumin gene expression starts later and increases in parallel with the decrease of AFP-gene expression which almost completely stops at birth, several reposts claim that albumin gene expression begins as early as AFP expression [13, 14]. Analysis with the use of more sensible methods such as in situ hybridization and biosynthetic labeling on rat embryos revealed that the albumin gene expression, together with its protein synthesis and the ­complete enzymatic secretory apparatus, is present and functional at the very early stages of liver development and in the ventral foregut endodermal cells, comparable with that of adult hepatocytes. Furthermore, although the ratio of albumin- and AFP-expressing cells to proliferating cells increases during embryonic stage, at the time of liver formation, both the genes and proteins are expressed in hepatoblast together with fibrinogen. In brief, during embryonic and fetal stages, 50% of the liver cells express secretory functions; while in the embryonic stage the liver growth depends mainly on cell proliferation, in the successive step, that

5

is, the fetal stage, the increase of cellular volume plays a more important role than cell division [15]. During fetal life the cell plates are three to five cells thick. At birth the plates are two cells thick and one cell-thick plate of the adult human liver is reached at 5  years of age. During fetal life hepatocytes exhibit considerable DNA-synthesis and replication. Two hours after birth DNA-synthesis rates are elevated with »18% of the hepatocytes incorporating 3H-thymidine into DNA. Three weeks later only 9% of the hepatocytes show evidence of DNA-synthesis. This activity declines continuously after birth until at 6 weeks of age only few hepatocytes (0.1–0.4%) show evidence of DNA synthesis [16]. Within the first 3 weeks after birth liver mass increases together with hepatocellular DNA-synthesis but no increase of mitosis in hepatocytes can be observed. This may mean that increased metabolic requirements induce a hypertrophy of the hepatocytes characterized by DNA-synthesis and enlargement of the hepatocytes and consequently of liver growth. A similar phenomenon can be observed in the adult animal after a certain period of fasting [17, 18]. In the fetal liver, the DNA-synthesis is quite strong and only mononuclear diploid cells are observed with the one third of the nuclei being in the S-phase in the suckling phase; in young adult animals, the DNAsynthesis strongly decreases and in  the young adult, the number of diploid nuclei decreases to »50%, the most being polyploidy (44% tetraploid) [19]. The hepatocyte polyploidy parallels increasing cell size and cytoplasmatic complexity correspond to the adaption of the cell to the increasing metabolic demand of the adult status. In the early life diurnal variability of the hepatic DNA-synthesis can be observed. This phenomenon is linked to feeding; in fact, reversal of food intake schedules alters the profiles of DNAsynthesis. In lower animals, food intake induces polyploidy in most gut cells.

Hepatocyte Structure and Renewal The hepatocyte belongs to the largest cells of the body. It has a size of 20–30 mm with a volume of 11,000 mm3 (information can vary between 10 and 60,000 mm3). The size however can vary quite strongly depending on age, location, and therefore on the blood

6

flow and metabolic load. The hepatocyte is an unbelievably complex system which has to fulfill several complex functions at the same time. These different functions are accomplished by means of very effectively functioning structures and organelles. A hepatocyte can be compared with the picture of the “Potsdamer Platz” in Berlin at the peak time of reconstruction or with the “big dig” in Boston where one could have difficulties to believe that a perfect synchronism between machine and human activity, give rise to a well organized end product. Hepatocytes are long-lived cells with little turnover in the absence of cell loss. Some people believe that liver tissue renews entirely approximately once a year and approximately one mitotic hepatocyte could be identified per 20,000 hepatocytes throughout the liver acinus [20]. For many years the hepatocytes response to the liver injury, in terms of a reparative proliferation through a compensatory hyperplasia, was based on a great number of studies of the liver after partial hepatectomy. This great capacity to divide in response to injury clearly emerged in contrast to a very limited pool of adult hepatocytes that lost the capacity to proliferate and differentiate. Even if there is no confirmed evidence of stem-cell involvement in normal hepatocytes turnover, in recent years the body of experimental data suggesting that hematopoietic cells could represent a possible source for replacement of hepatocytes in adult livers has been increasing [21]. On the other hand, the very low efficiency observed after cell transplantation, the possible errors in the interpretation of the results in terms of cell-fusion against cells amplification, low specificity of the methods and the proper characterization of distinct cell populations, give rise to a conspicuous number of questions that still requires concrete answers [22]. The hepatocyte is polyedric and possesses 5–12 facettes. Of these 1–3 are in contact with the sinusoidal blood whereas 4–9 are in contact with the biliary pool of the neighborhood-cell.

Plasmamembrane The hepatocyte is a polarized cell possessing three different specialized membrane domains (a) the basolateral or sinusoidal domain, (b) the canalicular domain, and (c) the lateral domain.

G. Ramadori and P. Ramadori

The Basolateral or Sinusoidal Domain Basolateral or sinusoidal domain faces the sinusoids and the perisinusoidal space of Disse. This domain is also called the vascular pole of the hepatocyte and constitutes 70% of total cell surface. It presents itself by 25–50 microvilli/mm, each of them measuring 0.5 mm in length and 0.1 mm in diameter. However, they are not uniformly distributed as there are clusters of thinner and longer microvilli on concavities existing on the basolateral domain that face the concavities on the surface of the opposite hepatocyte, that also contains these long, slender microvilli. The microvilli pervade the space of Disse and protrude the fenestrae of the sinusoidal endothelial cells into the sinusoids. For this reason they are thought to play a pivotal role in maintaining the integrity of the space of Disse [23]. However endo and exocytosis is the major function of the basolateral domain. For this reason the basolateral domain shows indentations or pits. Some of them represent exocytosis by secretory vesiculi, whereas others represent the so-called coated pits which are involved in receptor-­mediated endocytosis. The Na+/K+-adenosine triphosphate (ATPase) ion pump, the Na+/H+-exchanger as well as the Na+:HCO3 – cotransporter are located at the basolateral domain maintaining a substantial ion gradient and trans-membrane potential necessary for driving these transports across the cell membrane [24, 25].

The Canalicular or Apical Domain Canalicular or apical domain is also called the biliary pole of the hepatocyte. This domain constitutes 15% of the total hepatocyte surface and forms the bile canaliculus along with the canalicular domain of the opposite hepatocyte. The bile canaliculus is a half tubule cut into the hepatocyte surface. Laterally it is restricted by a smooth surface with junctional complexes. The diameter of the bile canaliculi changes with the site in the organ. In acinar zone 3, it is 0.5–1 mm and in acinar zone 1, it is 1–2.5 mm [23]. The canaliculus contains microvilli that are more abundant at the edges of the half tubulus. In the cytoplasm around the canaliculus there is also a network of contractile microfilaments driving the caliber of the canaliculus thereby regulating the bile flow. In the canalicular domain the apical bile acid transporter, organic ion transporters and

1  Hepatocytes

P-glycoproteins are located being responsible for the primary triphosphate (ATP)-dependent transport of organic components [26, 27].

The Lateral Domain The lateral domain of the hepatocyte ranges from the edge of the canalicular domain to the edge of the sinusoidal domain, representing about 15% of the total cell surface. The border between the lateral and the canalicular domains are represented by junctional complexes that include (a) tight junctions, (b) gap junctions, and (c) desmosomes.

Tight Junctions, Gap Junctions, and Desmosomes The tight junctions represent the barrier between the canaliculus and the rest of the intercellular space. They are composed of belt-like zones made up of three to five parallel strands, whereby the cohesiveness of the tight junctions depends on the number of strands. The gap junctions are patches in close approximation with adjacent membranes which are separated by a gap of 2–4 nm. The gap is bridged by trans-membrane protein particles which protrude from the membrane and contain a central pore. Two of those protrusions from opposite cells, both containing a central pore serve as a channel of intercellular communication [28]. In addition, the lateral surface also contains the so-called “press-stud” or “snap-fastener” types of intercellular junctions that consist of membrane protrusions that interact with membrane indentations on the opposite cell.

Organelles As noticed above the hepatocyte is one of the most metabolically active cell type of the body. This already suggests that it contains a large amount of organelles. The most abundant are the endocytoplasmatic reticulum (ER), the mitochondria, lysosomes, and the peroxisomes (Fig. 1.3).

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Endoplasmic Reticulum The ER represents 15% of total cell volume, however, its surface area is about 35-fold that of the cell membrane. The ER represents a complex system of membrane bound channels. Two different types of ER could be distinguished according to its appearance on electron microscopy – the rough ER (RER), so called because of its association with ribosomes that give a rough appearance and the smooth ER (SER) which consists of smooth membrane-bound channels and is less in number when compared to the RER. The relative amount of the two types in the hepatocyte is not constant but varies with location of the hepatocyte in the acinus and its physiologic state; e.g., the surface area of the ER in zone three is twice that in zone one. The RER appears as parallel profiles of flattened cisternae distributed randomly in the cytoplasm. It communicates on the one hand with the nuclear envelope, on the other hand with the SER. The SER in turn communicates with the RER and the Golgi complex but not with the nuclear envelope and consists of anastomosing and interlinked channels. It is noteworthy that neither the RER nor the SER communicates with the plasma membrane. The synthesis of proteins takes place in the polyribosomes that are attached to the RER. From here they finally reach the cisternae of the RER and are transported to the SER and finally to the Golgi complex. In the Golgi complex they are packed into the secretory Golgi vesicles. Many functions of the hepatocyte are accredited to the ER: synthesis of secretory proteins, synthesis of structural membrane proteins, metabolism of fatty acids, phospholipids and triglycerides, production and metabolism of cholesterol, xenobiotic metabolism, ascorbic acid synthesis and heme degradation. All this functions are localized to specialized domains of the ER. Structural proteins can also be synthesized by free ribosomes, especially during liver development and regeneration. While the RER is mainly involved in protein synthesis, in the SER, there are enzymes needed for drug metabolism, cholesterol biosynthesis, and conversion of cholesterol to bile acids. For example the well known cytochrome P450 enzyme system is located in the ER. When this system is induced, this leads to hypertrophy of the SER whose histological correlate is a “ground glass” appearance of the cytoplasm. Another very important enzyme, the Glucose-6-phosphatase is also associated with the ER. During glycogenesis and glycogenolysis the SER proliferates. As noticed above

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G. Ramadori and P. Ramadori

a

b

c

d

Fig. 1.2  Liver sections stained with the Goldner staining (connective tissue) (sections are a gift from Prof. L. Fuzesi, department of Pathology, University Hospital of Goettingen). (a) It is possible to appreciate collagen and reticular fibers disposition in the area of necrosis (blue stain). (b) Presence of areas of

proliferative hepatocytes and of necrotic areas with connective tissue deposition (arrowheads). (c) Detail of a regenerative nodule of proliferating hepatocytes (H/E). (d) Part of the same liver biopsy immuno-stained with the anti-Ki67-antibody, showing a massive number of cells in a proliferative phase

there is a close connection between the RER and the mitochondria and these complexes are important for synthesis of membranes and heme. Heme itself is an important component of the cytochromes [29, 30].

the ER and the vesicles from the ER transport proteins from the ER to this surface. The trans-surface is associated with vesicles containing osmophilic spheres corresponding to very low-density lipoproteins (VLDL) demonstrating that the Golgi complex of the hepatocytes is important for VLDL synthesis. In addition the Golgi complex is responsible for terminal glycosylation of secretory protein and recycling of membrane glycoprotein receptors [25, 31, 32].

Golgi Complex Two to four percent of the total cell volume is made by the Golgi complex which is located nearby the bile canaliculus and the nucleus. It is made up of about 50 interconnected complexes, each of them being composed of 3–5 parallel cisternae with associated vesicles and lysosomes. The surface of the cisternae can be divided into the cis-surface (convex surface) and the trans-surface (concave surface). The cis-surface faces

Mitochondria The mitochondria make up 13–20% of total hepatocyte cytoplasmic volume. About 1,000 mitochondria can be found in a hepatocyte. They are the power plant of

1  Hepatocytes

the hepatocyte generating the energy required for the metabolic functions of the hepatocyte e.g., fatty acid oxidation. They are able to change their shape, have the possibility to fuse and are able to move in the cytoplasm which is associated with microtubules. They consist of an outer and an inner membrane. The outer membrane has no enzymatic activity but contains a transport protein called porin. Porin is able to form channels that are permeable for proteins of <2 kDa. The inner membrane is highly convoluted and folded to the so called cristae that are closely associated with matrix granules. The inner membrane contains the enzymes of the respiratory chain, responsible for oxidative phosphorylation and generation of ATP. The matrix of the mitochondria contains a small circular DNA, ribosomes, and enzymes of the citric acid cycle and urea cycle and beta oxidation of fatty acids. In addition, calcium stores are found that form electron-dense granules. While the mitochondrial DNA encodes the mitochondrial proteins synthesized by the mitochondrial ribosomes, most of the mitochondrial proteins are imported from the nucleusderived transcription. Interestingly, the mitochondria are able to self-replicate and the half life is about 10 days [33–36].

Lysosomes The primary lysosomes are enveloped by a single membrane. On their inner side, they contain enzymes such as acid hydolases, acid phosphatase, aryl sulphatase, esterase, and b-glucuronidase. By these means they serve as storage granules and sequester enzymes produced by the ER and packed by the Golgi. The so called secondary lysosomes are formed when primary lysosomes fuse with other membrane bound vesicles containing endogenous cellular material or exogenous material destined for degradation. These are called autophagic vesicles. The secondary lysosomes are mainly located near the Golgi complex and the canalicular domain. In electron microscopy, they appear as the so called peribiliary dense bodies. The secondary lysosomes vary in size and number and contain a variety of materials. During starvation, regeneration and development they are particularly numerous. For example lysosomes accumulate lipofuscin, ferritin in iron overload states, copper in case of Wilson’s disease and glycogen during glycogenolysis. Coated endocytic vesicles, the so

9

called endosomes, are derived from receptor-­mediated endocytosis (after the internalization of receptor-ligand complexes on the basolateral domain). Such ligands are insulin, low-density lipoproteins, transferrin, immunoglobulins (IgA), and asialoglycoproteins. The ligands themselves may be processed before exocytosis, degraded (in autophagic vesicles made by fusion with lysosomes), or transported across the cell along microtubules. The latter process is called transcytosis.

Peroxysomes The peroxisomes are small, ovoid membrane bound granula of 0.1–0.2 mm diameter. Each hepatocyte contains 300–600 of them. They contain oxidases and catalases that are responsible for 20% of the total oxygen consumption of the hepatocyte. The oxygen is used to oxidize numerous substrates and to produce energy. The hydrogen peroxide formed during this process is hydrolyzed by the peroxisomal catalyze. Ethanol is also metabolized in the liver by the peroxisomal catalase. The importance of the peroxisomes becomes obvious by their proliferation during liver development and liver regeneration (e.g., recovery from partial hepatectomy and various liver diseases) as well as after administration of salicylate and clofibrate [1, 33, 37].

Nucleus/Polyploidy The onset of polyploidy in the liver is known for quite a long time and has been studied widely in hepatocytes. While most hepatocytes (see above) contain a single nucleus, up to 40–50% contain two and more nuclei. Multinucleated hepatocytes are thought to develop in response to completion of DNA synthesis and mitosis; however, failure of cell division of cytokinesis would normally generate daughter cells containing single nuclei. It has also been hypothesized that polyploidy cells originate from the fusion of multinucleated cells, especially because studies have shown that the prevalence of multinucleated cells declines in response to simultaneous induction of hepatic polyploidy. However, this has not been validated until now and other mechanisms resulting in loss of multinucleated cells have not yet been excluded e.g., cells containing

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two nuclei could divide into two mononuclear cells or undergo apoptosis [17, 38]. Another concept considers the fate of mammalian cells when mitotic progression is perturbated like in case of disruption of the centromere assembly. In this situation cells are able to continue to synthesize DNA and generate additional complements of chromosomes in the same nucleus with loss of cytokinesis ability [39]. The post natal liver growth is mainly due to hypertrophy, during which hepatocytes accumulate increased amounts of DNA, probably as an adaptive response to increasing metabolic requirements [40]. During postnatal liver growth, hepatocytes undergo polyploidization. Analysis of DNA content in the nuclei of hepatocytes demonstrated that DNA synthesis was prominent in the fetal liver with almost 1/3 of all nuclei being in the S-phase.This decreases fast after birth with fewer than 5% being in the S- phase. In young adults DNA synthesis was even less and only 1% of nuclei were in the S-phase. In contrast the liver of fetal and suckling animals contained only diploid cells whereas prevalence of diploid nuclei in young adults declined to 53% with remainder showing progression toward polyploidy. The ploidy class here is predominantly tetraploid (44%) [41]. Until now little is known about the function and fate of polyploidy hepatocytes. However some suggestion might be allowed. When diseased hepatocytes are replaced rapidly by proliferation of hepatocytes following self-limited liver injury, full recovery is expected, even though some cells undergo polyploidy. On the other hand, perpetuation of polyploidizing stimuli in the liver would lead to excessive appearance of polyploidy cells in the liver, which might be associated with accelerated rates of apoptosis and impaired capacity for organ restoration by proliferation in polyploidy cells. This could lead to two consequences. First, when proliferation of hepatocytes fails, liver failure will occur. Second, the emergence of cell clones resistant to ongoing disease processes or with genetic instabilities such as in chronic liver hepatitis, alcoholic liver disease, or genetic hemochromatosis, etc. could be associated with the development of liver cancer.

Physiology The liver exerts many functions that comprise storage, metabolism, production, and secretion. Besides the functions of the hepatocytes mentioned above, the

G. Ramadori and P. Ramadori

processing of absorbed nutrients and xenobiotics as well as the bile acid synthesis and bile formation, the hepatocytes are also responsible for maintenance of glucose, amino acid, ammonia, and bicarbonate homeostasis in the body, the synthesis of most plasma proteins, the storage and processing of signal molecules, and last but not least participate in the acute phase reaction (Fig. 1.3). At this point it is mandatory to keep in mind that the well organized physiological function of the hepatocyte largely depends on the proper function of hepatocyte nuclear factors (HNF) and their receptors [42] on one side but also on the proper function of the excretory apparatus.

Lipid/Lipoprotein, Cholesterol, and Bile Metabolism The liver plays a central role in lipoprotein metabolism. The hepatocytes, like the parenchymal cells of many other tissues possess the LDL-receptor (low density lipoprotein receptor) which is capable to take up cholesterol. In addition hepatocytes possess another receptor that binds and internalizes apo-E containing lipoproteins. Cholesterol is the basis for bile acid synthesis or is directly shifted through the cell and secreted into the bile. While the formation of primary bile acids such as cholic and chenodeoxcholic acids takes place exclusively in the liver, the formation of secondary bile acids (deoxycholic acid, lithocholic acid) occurs by metabolism by the intestinal bacteria in which the primary bile acids are conjugated with amino acids, sulfate or glucuronic acid. This leads to a decrease in toxicity, relieved biliary excretion and an increase in water solubility. The rate controlling step in bile acid synthesis is the microsomal P450 enzyme cholesterol7a-hydroxylase [36]. This as well as the hydroxymethylglutaryl-CoA reductase (HMGCoA reductase), which is the rate limiting enzyme of the hepatocyte cholesterol synthesis, underlie a feedback inhibition by bile acids at the levels of transcription and activity. The bile acids undergo an entero-hepatic circulation so that only small amounts of bile salts excreted by the liver derive from a de novo synthesis. Therefore an efficient uptake mechanism is required, realized by transporters described earlier in this chapter. Little is known about the intracellular transport of bile acids from the sinusoidal to the canalicular region in the hepatocyte. While diffusion may be involved, three

1  Hepatocytes

cytosolic bile-acid binding proteins which also function as proteins binding 3-a-hydroxysteroid dehydrogenase, glutathione-S-transferase, or fatty acids have been described. Their role in bile acid transport is, however, discussed controversially. Since electron microscopy visualizes vesicles that contain bile acids there might also be a vesicular bile acid transport. Bile acids could enter these microsomal or Golgi derived vesicles driven by the positive membrane potential. Although these findings support the concept of a vesicular bile acid transport, it could also be that these vesicles transport the canalicular bile acid carrier to the canalicular membrane domain and not primarily bile acids. Thus the vesicular and nonvesicular bile acid transport is still unclear. However transcytotic transport processes have been shown for the protein excretion into the bile [43]. Thereby, the newly synthesized membrane proteins, including those for the canalicular membrane are initially targeted from the Golgi complex and the ER to the sinusoidal membrane. Here, those destined to the canalicular membrane are resorted, endocytosed, and transported to the canalicular membrane via transcytosis. Other proteins and polysaccharides can also be taken up by receptor-mediated or fluid-phase endocytosis and also reach the canalicular membrane per transcytosis. A vesicular transport and exocytosis are also thought to be responsible for the excretion of cholesterol and phospholipids into the bile [44]. The canalicular secretion of bile acids and other organic anions or cations generate an osmotic gradient that drives the osmotic water flow into the bile canaliculi. It is however noteworthy that for bile acid excretion the basolateral uptake is not the rate liming step but the transport across the canalicular membrane[36]. Beyond the synthesis and secretion of bile acids, the hepatocyte represents an important center for the storage and the metabolism of lipids together with the adipose tissue. Fatty acids and triacylglycerols coming from different sources (lipogenesis or systemic circulation) can be stored in the cytosol or released in the circulation as constituents of the VLDL, based on the body energy requirement and on their plasmatic concentration. This storage function of the hepatocytes allows to metabolize the excess plasma levels of circulating nonesterified fatty acids directing them to the catabolic beta-oxidation or to be complexed with other component through phosphorylation and glycosilation. While the lipogenesis is controlled by hormonal and nutritional conditions at a cytosolic level through a series of

11

decarboxylative condensation reactions starting from the acetyl-CoA substrate, the catabolism of the fatty acids is a more compartimentalized process [46]. In fact, peroxisomal beta-oxidation is responsible for the metabolism of very long chain fatty acids together with the microsomal cytochrome P450 CYP4A, while through mitochondrial beta-oxidation the short and medium chain fatty acids are metabolized. The regulation of the set of genes coding enzymes involved in the synthesis and catabolism of fatty acids is under the control of transcription factors that constitutes receptors from the same acids or their derivatives, such as eicosanoids or oxidized low-density lipoproteins [47]. Liver X receptor (LXRs), Retinoid receptor X (RRX), and Peroxisome-proliferator-activated receptors (PPARs) represent transcription factors regulating not only several metabolic signaling pathways, but also inflammatory pathways in the immunity field. In ­particular, PPARs family has recently emerged as important repressors of proinflammatory signals and cytokines [48] (see Chapter on PPAR). PPAR-alpha is highly expressed in the liver where it regulates the expression of enzymes involved in the transport, in the oxidation and in the catabolism of fatty acids, and the loss or the lack of this protein leads to reduced energy burning with subsequent hepatic steatosis [49]. PPAR-gamma which is less expressed in the liver, much more in the adipose tissue, acts as a repressor of the fatty acids flux to the liver and inducing the expression of lipogenic enzymes (e.g., acetylCoA-carboxylase), increasing insulin sensitivity by up-regulating glucose transporter GLUT-4 and promoting fatty acids uptake into adipocytes [50]. PPARgamma deficient mice are protected against the development of steatosis [51]. The new prospective depicted by recent clinical and experimental studies regarding the link between metabolic aspects regulated by this family of transcription factors and their activity as repressors of gene involved in the immune-modulation and inflammation, offers nonalcoholic fresh insight for the development of new therapeutic approaches in the treatment of fatty liver disease.

Glucose Metabolism The liver plays a pivotal role in glucose metabolism of the organism regulating blood glucose level by removing glucose if it is present in excess by glycogen synthesis or glycolysis and lipogenesis or ­supplying glucose if

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G. Ramadori and P. Ramadori

it is needed by glycogenolysis or gluconeogenesis (Fig.  1.4). The glucose uptake and release across the hepatocellular plasma membrane is guaranteed by a bidirectional transporter, the Glut-2 transporter. On the other hand it has to be noticed that the plasma membrane transport of glucose is not the major regulatory site of glucose metabolism. Several factors are capable to control the reversible switch between glycogenolysis/gluconeogenesis and glycogen synthesis/ glycolysis such as substrate concentrations, hormone levels (insulin above all), hepatic nerves, the hepatocellular hydratation, and zonal hepatocyte heterogeneity [52, 53]. The glycogen synthesis and glycolysis are predominantly

regulated by the portal glucose concentration with insulin (about 80% of endogenously secreted and 50% of intravenously infused insulin is cleared in the hepatocytes [54]) and parasympathic nerves being auxiliary factors). Glycogenolysis and gluconeogenesis, on the other hand, are initiated by glucagons and sympathic nerves but inhibited by high portal glucose concentration. Three substrate cycles, the glucose/ glucose-6-P cycle, the fructose-6-P/fructose-1,6-bisphosphate cycle and the pyruvate/ phosphoenolpyrophate cycle are involved in regulating the hepatic gluconeogenesis/ glycolysis. The nature of these enzymes allows a rapid switch of flux through the pathways determining

Fig. 1.3  Hepatocyte metabolism (modified from Drenckhahn, 1994). SER smooth endocytoplasmic reticulum; RER rough endocytoplasmic reticulum; Nu nucleus (the nuclei in the chart  are small because of lack of space); GC Golgi complex; Po peroxisomes; VLDL very low lipoproteins; Li lipid droplets;

Lf lipofuscin. It is possible to note how all the physiological functions of the hepatocyte are regulated by a fine network of transcriptional regulators and their receptors with hepatocyte nuclear factors (HNF) acting as master regulators

1  Hepatocytes

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whether the liver becomes a producer or consumer of glucose [55]. The gene transcription of these gluconeogenic enzymes is controlled by hormones, mainly insulin, glucagon, and glucocorticoids. While insulin acts as an inhibitor of the gluconeogenesis by suppressing the expression of PEPCK and G6Pase, glucagon, and glucocorticoids stimulate hepatic glucose production by inducing the same genes. In the case of diabetes type II, reduced uptake of insulin by the hepatocyte may be one of the causes of hyperinsulinemia and hyperglycemia. Furthermore, clinical reports show that those patients develop a liver fat content 54% higher compared to the nondiabetic subjects and an insulin clearance 24% lower, indicating an inverse association between hepatic fat accumulation and insulin sensitivity, independent of intraabdominal fat [56]. In humans, the hepatocytes are

considered to be also the main source of insulin-like factor binding protein 1 (IGFBP-1), and insulin is supposed to be its main regulator. The analysis of liver fat content, through magnetic resonance spectroscopy together with the serum levels of IGFBP-1, was able to confirm the inverse relationship between insulin sensitivity and liver fat accumulation even in nondiabetic subjects [57]. In the context of the metabolic syndrome, characterized by NAFLD, obesity and the development of diabetes type II, the development of an inflammatory condition could be responsible for the activation in the hepatocytes of pathways of signals that trigger the induction of the family of inhibitory proteins, SOCS. It has been recently described that a member of these proteins (SOCS-3) might reduce insulin signaling through inhibition of the insulin receptor [58] (Fig.  1.5). The hepatic insulin resistance linked to the accumulation of

brain

minoacids actate

luc

co

os

ns

ulin

lu

e

uli

ns

n

eural mediators

muscle

se

pancreas liver

Insulin

dipo ines

Glucose lycerol

adipocytes

Fig. 1.4  the liver is a central organ of insulin and glucose physiology. More than 70% of the insulin reaching the liver from the pancreas it is taken up by the hepatocytes, as the case of the glucose absorbed in the intestine is metabolized or stored in the cells (thick arrows). Only a part of the insulin and glucose reach the

intestine

systemic circulation (dashed arrow). This amount might increase if the functional capacity of the liver is reduced (by reduction of the number of hepatocytes or by reduction of the functional capa­ city of the single hepatocyte)

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body fat is a central component in the physiopathology of the pluri-metabolic syndrome.

G. Ramadori and P. Ramadori

The liver also participates in the amino acid homeostasis of the body. An increased portal amino acid load leads to swelling of the hepatocytes and consecutively to an increase of the amino acid flux across the plasma membrane. This stimulates the amino acid breakdown and utilization for protein synthesis as well as glycogen synthesis and simultaneously inhibits amino acid generation from proteolysis. However, it also has to be noted that hepatic amino acid extraction is not equipotent for all amino acids. The highest amino acid extraction is observed for the gluconeogenic aminoacids alanine, serine, and threonine. Also most essential amino acids are extracted by the liver. On the other hand branched amino acids such as leucine, valine, and isoleucine which are only used for protein synthesis are not catabolized. The portal blood contains high concentrations of ammonia derived from the generation by the intestinal mucosa from glutamine and from intestinal microorganisms. However, ammonia is also produced by the hepatocytes during processing of amino acids. The detoxification of ammonia occurs by both the liver-specific urea synthesis and the glutamine synthesis. Failure of the hepatocellular function leads to increased serum levels of the neurotoxin ammonia and thereby to encephalopathy and neuropsychiatric disorder. A spillover of two enzymes into the blood involved in amino acid metabolism is the maker for hepatocellular necrosis. There are the AST (aspartate aminotransferase) and the ALT (alanine aminotransferase). Since AST has also a mitochondrial isoform, it is expressed in relevant quantities in muscles and is therefore less specific for liver damage than ALT.

[60] and probably C1q which are synthesized by macrophages [61]. Among these serum proteins are albumin, factors of the coagulation system (Factor I (fibrinogen), Factor II (prothrombin), Factor V, Factor VI, Factor IX, Factor X) as well as compounds of the complement cascade [62–64]. The major protein secreted by the hepatocytes is albumin (50% of the secreted proteins!), whose main function is to regulate the fluid homeostasis within the vessels. The individual hepatocyte however is not specialized on production of specific plasma protein. In fact it is able to synthesize the whole spectrum of proteins in comparable amounts. However, in vivo for some plasma proteins, a heterogeneity of production dependant on the zonal location of the hepatocyte could be observed e.g., for albumin. The albumin production by the hepatocytes is especially high. 12 g per day are produced in man and this amount could be increased up to fourfold upon albumin loss. Except for albumin and C-reactive protein all proteins produced by the hepatocytes are glycoproteins. On the other hand specific receptors for galactose- or mannose-terminated glycoproteins allow the hepatocytes to take part in the clearance of the glycoproteins [65]. The hepatocytes secrete proteins constitutively, meaning that they continuously produce and secrete proteins and contain no stores for the synthesized proteins. Therefore the rate of protein secretion depends on the protein synthesis. However, as the transport time of the synthesized proteins from the ER to the Golgi complex as well as the retention of the proteins in the Golgi complex differ, the secretion nevertheless is different for the diverse proteins. Examples for fast secreted proteins are albumin (albumin is secreted so fast that it is difficult to find it also in the intracellular space), fibronectin, or a1-protease inhibitor; examples for slow secreted proteins are fibrinogen or transferrin. However, protein synthesis of the hepatocytes underlies a certain control by amino acid concentration, cell swelling growth and thyroid hormones, glucagons, and vasopressin.

Protein Synthesis

Acute Phase Response

Apart from immunoglobulins the liver provides the most circulating plasma proteins which are synthesized by the hepatocytes, except von Willebrand factor which is produced by endothelial cells [59], IGFBP3

The liver is the site of synthesis of acute phase proteins. They comprise a heterogeneous group of plasma protein concentrations which rapidly change upon tissue injury in extrahepatic sites. The synthesis of these

Amino Acid and Ammonia Metabolism

1  Hepatocytes

proteins in the liver is mediated by cytokines (acute phase mediators) such as interleukin-6 (IL6), interleukin-1 (IL-1), tumor necrosis factor alpha (TNF-a), and ultimately interferon-gamma (IFN-g) [66], which are secreted by inflammatory cells at sites of injury. Among these IL-6 seems to be the most important regulator of inflammation. Its secretion by monocytes, macrophages and endothelial cells underlies the control of IL-1, TNF-a, endotoxin, or microorganisms (e.g., viruses). IL-6 possesses a specific receptor on the hepatocyte surface leading to an increased transcription of genes encoding for acute phase proteins. However it is also noteworthy that the presence of glucocorticoids is necessary for the cytokine induced synthesis of acute phase proteins. The increased synthesis of acute phase proteins leads to increased plasma levels that range from 1.5-fold (ceruloplasmin, complement C3) to up to several hundred fold (C-reactive protein (CRP), serum amyloid A) and also their effects which is of a large scale. Proteins e.g., such as a1antitrypsin and a1-antichymotrypsin are protease inhibitors thus restricting the protease activity of enzymes of inflammatory cells. Others are immunomodulators (e.g. a1-acid glycoprotein) or take part in the clearance of xenobiotic material and microorgamnisms (e.g. CRP, serum amyloid A) or in host defence (e.g. proteins of the complement system) [54, 72, 74]. Acute phase mediators not only influence the gene expression of secretory proteins (IGFBP-1, IGFBP-4, etc.) but also of intracellular proteins belonging to the defense apparatus of the hepatocytes, such as HO-1 [69]. Recently, it has been demonstrated that hepatocytes can control the changes occurring in the bone marrow during the acute phase by synthesizing cytokines such as IL-8 [66], a chemo-attractive mediator for polymorphonucleate granulocytes, thrombospondin, and thrombopoietin (TPO) involved in the control of platelet production [70].

Iron Metabolism The liver is the central organ of iron metabolism. It receives the iron contained in the heme of the erythrocytes which are eliminated by the Kupffer cells. The hepatocyte represents the major iron-storing cells of the body. It expresses both classic transferrin receptors (TfR1 and TfR2) and is thought to possess ferritin

15

receptors. Regulation of TfR1 is highly iron dependent but its role in liver iron uptake is unclear and this receptor is expressed in only small amounts on hepatocytes. The TfR2 however is highly liver specific. It occurs in two forms. As it is missing the iron-responsive elements (IRS) the expression of the a form is not regulated by iron. The b form is widely expressed at low level and may be a secreted protein. Upon binding the Tf-TfR-complex is internalized into endosomes. Here the Fe is then released from Tf by a reduction of endosomal pH. Once released the Fe is transported through the endosomal membrane and into the cell via Nramp2 or the divalent metal transporter 1 (DMT1). From this point Fe can be used for a variety of metabolic processes or stored within the protein ferritin. However excessive iron uptake e.g., in all nontransfusion iron overload diseases is due to the uptake of non-Tf-bound iron by the hepatocyte. Following this, the deposition of iron in the liver occurs in any case in which the iron binding capacity of Tf is exceeded. However, little is known of this transport system, e.g., it is unclear whether it recognizes Fe2+ and/or Fe3+. On the other hand the non-Tf-iron transport ­system seems to be always active and is not down-­ regulated by iron as it is for TfR1 [71, 72]. Hepatocytes express all the proteins involved in the iron metabolism described so far. They synthesize and secrete hepcidin, which is considered to be responsible for the changes in the uptake of iron in the intestinal cells and in macrophages. It has been shown that hepcidin behaves like a positive acute phase protein, while hemojuvelin, another key protein in the regulation of iron metabolism, results to be down-regulated [73].

Hepatocyte as an Endocrine Cell In response to stimulation with the growth hormone (GH), hepatocytes can synthesize insulin-like growth factor-1 (IGF-1), a hypoglycaemic hormone inducing GLUT-4 membrane expression stimulating glucose uptake in the muscle. IGF-1 as well as IGF-1 binding protein production, which is important for maintaining IGF-I in the circulation, is controlled by the cooperation of multiple cytokine pathways (e.g., MAPK-pathway) and cytokines including IL-6,

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insulin, TGF-a, and TGF-b [60, 74]. However, although it has long been suggested that circulating IGF-1 is responsible for body growth, newer studies clearly indicate that locally produced IGF-1 is involved in growth but only to a much lesser extent than the circulating IGF-1 [75]. On the other hand lack of the IGF acid labile subunit which is also produced by the hepatocytes and regulated by GH may be responsible for growth impairment [76]. Above that a recent publication suggests that the hepatocyte derived IGF-I has an important role in maintaining a fine balance between GH and insulin to promote normal carbohydrate and lipid metabolism [77]. In addition hepatocyte IGF-1 production not alone but in concert with hepatocyte growth factor (HGF) may be an important factor in progression of hepatocellular carcinoma (HCC) [78]. During embryonic development the liver represents the major center of erythropoiesis and the most important producer of erythropoietin (EPO) [79]. After birth, the kidney replaces the hepatic production of the erythropoietic hormone, whereas the hepatocytes take part to EPO synthesis during anemic or hypoxic conditions [80]. Furthermore hormones (e.g., GH, thyroid hormone) stimulation can induce an increase of EPO gene expression in the hepatocytes [81].

Transport-Systems The hepatocellular transport systems can be divided into basolateral and canalicular transport systems (for an overview see Fig. 1.6). The main basolateral transport systems comprise of the Na+ dependent bile salt uptake, the Na+ independent uptake of amphipathic substrates and the basolateral efflux pumps.

Na+ Dependent Bile Salt Uptake Studies on rat liver and hepatocytes as well as on basolateral plasma membrane vesicles indicated that more than 80% of taurocholate uptake and less than 50% of cholate uptake into hepatocytes are sodiumdependent. While unconjugated bile salts are uncharged molecules that can transverse membranes by passive nonionic diffusion, conjugation with glycine, or taurine

G. Ramadori and P. Ramadori

requires the presence of a specific carrier protein for hepatocellular uptake. The main uptake system for conjugated bile salts in mammalian liver is called the Na+-taurocholate cotransporting polypeptide (NTCP), which is exclusively expressed at the basolateral domain of the hepatocyte. The only nonbile salt substrates that are transported by NTCP are selected sulfated steroid conjugates such as estrone-3-sulfate and dehydroepiandrosterone sulfate (DHEA). NTCP is structurally related to the intestinal bile salt transporter (IBAT), which also mediates Na+ dependent uptake of bile salts; it is not only expressed in the ileum but also in the kidney and in cholangiocytes [82]. The study of the molecular mechanisms regulating components of NTCP is important to throw light on the physiopathology of several liver diseases, in which NTCP expression has been shown to be altered, such as pregnancy or cholestatic diseases (e.g., primary biliary cirrhosis). High levels of bile acids have been observed to suppress its gene expression, as also endotoxin and proinflammatory IL-1b [83], while glucocorticoids and PPAR-g ligands trigger to the activation of NCTP gene [84].

Na+ Independent Hepatic Uptake of Amphipatic Substrates: The Organic Anion Transporting Polypeptide Family (OATP) While uptake of conjugated bile salts into the liver is largely a Na+-dependent process mediated by the NTCP, numerous other endogenous, or xenobiotic compounds including nonbile salt organic anions and drugs are cleared from the sinusoidal blood by carriermediated uptake into the hepatocyte. Following hepatocellular uptake, many of these compounds are biotransformed in two phases. Phase I is mediated by cytochrome P450 enzymes and prepares the drug for conjugation by creating polar groups. Phase  II conjugates drugs with glucuronate, sulfate, glycin, or methyl group thereby representing a detoxification step. These conjugates can then be excreted into the bile or the urine. The Na+ independent hepatocellular uptake of bile salts and nonbile salt amphipathic compounds cannot be attributed to a single transport protein. In fact it is guaranteed by a family of transport proteins called the OATP. So far three members of the OATP family have been identified in rat hepatocytes called OATP1, OATP2, and OATP4. OATP1 is localized

1  Hepatocytes

at the basolateral membrane of the hepatocytes and the apical membranes of the kidney proximal tubular cells and the choroids plexus epithelial cells. Several lines of evidence ­suggest that OATP1 is a “multispecific bile acid ­transporter”. OATP1 has been shown to mediate heptocellular uptake of bile salts, bromosulphophathalin (BSP), conjugated steroids, thyroid hormones, leukotriene C4, bilirubin monoglucuronide, ouabain, ochratoxin A, the anionic magnetic resonance imaging agent gadoxetate, the angiotensin-converting enzyme inhibitors enalapril and temocaprilat, the HMG-CoA reductase inhibitor pravastatin and even oligopeptides including the thrombin inhibitor CRC-220, and opioid receptor antagonists. The driving force for OATP mediated substrate transport is yet not fully understood although it has been shown that  OATP1 can mediate bidirectional transport of BSP  and anion exchange of taurocholate/HCO3−. An important driving force for the OATP1 dependent organic anion uptake may be the counter transport of reduced glutathione (GSH). OATP2 is located at the basolateral membrane of the hepatocytes and has also been detected in the retina, in endothelial cells of the blood brain barrier and at the basolateral plasma membrane of the choroids plexus epithelial cells. OATP2 is a close homologue of OATP1 and transports bile salts, cardiac glycosides (e.g., digoxin) and cyclic peptides. The most important difference between OATP1 and OATP2 is their acinar location in the liver. While OATP1 is distributed homogenously within the liver acinus, OATP2 is  predominously expressed in the perivenous hepatocytes excluding the innermost two cell layers around the central vein. Interestingly, drugs such as phenobarbital or digoxin and thyroxin lead to a significant increase of OATP2 expression even in the innermost layer of the central vein. OATP4 can also mediate NA+-independent uptake of bile salts. It transports numerous organic anions including taurocholate, BSP, conjugated steroids, and thyroid hormones. In human liver at least four OATPs have been identified called OATP-A, OATP-B, OATPC, and OATP8. OATP-C and OATP8 are exclusively expressed on the basolateral domain of the hepatocyte and are 80% homologous. The closest homologue in the rat liver is OATP4 which is 64 and 66%. Therefore the substrate specificity of the human OATP-C and OATP8 and that of the rat OATP4 are very similar. Transport substrates of OATP-C are ­taurocholate, bilirubin monoglucuronide, DHEAS, pravastatin, and BSP.

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OATP8 has a similar substrate specificity but additionally transports digoxin and cholecystokinin. OATP-B is strongly expressed in the human liver and additionally expressed in spleen, placenta, lung, kidney, heart, ovary, small intestine, and brain. OATP-B has a limited substrate specificity for the organic anions BSP, estrone-3-sulfate, and DHEAS when compared to OATP-C and OATP8. OATP-A is expressed in relatively low levels in human hepatocytes. Although it was originally isolated from the human liver it is predominantly expressed in human cerebral endothelial cells. OATP-A transports bile salts, BSP, estrone-3sulfate DHEAS, opioid receptor antagonists, antihistamics and amphipathic organic cations. Thus in contrast to the preference of OATP-B, OATP-C, and OATP-8 for organic anions, OATP-A additionally transports amphipathic organic cations indicating that they might mediate substrate uptake independently from their charge. Taken together, the OATP family of transporters plays a central role in organic anion and drug clearance of the hepatocyte [82, 85]. Although so far no specific diseases result from an impaired function of the OATP transporters, alterations of their activity might interfere with the biotransformation or catabolism of certain drugs, modifying their therapeutic effects. The genetic control of these transporters has been shown to depend on the activity of the hepatic nuclear factors HNF-1a and HNF-4a [86].

Na+ Independent Hepatic Uptake of Hydrophilic Organic Cations and Anions: The Organic Ion Transporter Family (OAT/OCT) In addition to the NCTP and the OATPs a third family mediates the substrate uptake called the organic anion transporter family. This family comprises the organic anion transporter (OAT), the organic cation transporter (OCT) and the organic cation transporter novel type (OCTN)/ carnitine transporter families. Whereas Oat1 is expressed only in rat kidney, Oat2 is expressed exclusively and Oat3 predominantly in rat liver. In human liver only Oat2 has been found so far. Oat2 mediates the sodium independent transport of a ketoglutarate and salicylates, whereas Oat3 transports para aminhippurate (PAH), estrone-3-sulfate, and the cationic compound cimetidine. The organic cation transporter called OCT1 is expressed in rat kidney, small intestine enterocytes

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and the basolateral domain of the hepatocytes. In humans hOCT1 is exclusively expressed in the liver and mediates the hepatic clearance of type I cations such as dopamine, choline, tetraethylammonium, or N-methylnicotinamide [87–89].

Basolateral Efflux Pumps The basolateral membrane of the hepatocyte also possesses members of the multidrug resistance protein family (MRP) belonging to the superfamily of ATPbinding cassette (ABC) transporters. MRP1 mediates ATPdependent efflux of glutathion S-conjugates, leukotriene C4, steroid conjugates, or bile salt conjugates. MRP1 is normally expressed at low levels in hepatocytes. MRP3 mediates the basolateral efflux of the organic anions estradiol-17-b-D-glucuronide and S(2,4-dinitrophenyl) GSH, the anticancer drugs methotrexate and etoposide and even monovalent bile salts. MRP5 is suggested to be an anion transporter; however, its expression in adult liver is very low. MRP6 is localized at the lateral membrane of the hepatocytes and transports the cyclic pentapeptide and endothelin antagonist BQ-123 [90–92]. The canalicular transport systems manage one of the major functions of the liver, the bile salt excretion, the excretion of nonbile salt organic anions and the copper excretion. The importance of this function becomes obvious from the view of the endoscopist who finds the green color of the bile to be predominant in the many parts of the gut.

Bile Salt Excretion The canalicular excretion is the rate limiting step in the secretion of bile salts from blood into the bile. Whereas bile salt concentrations in the hepatocyte are in the micromolar range canalicular bile salt concentrations are 1,000-fold higher, which requires active transport across the canalicular hepatocyte membrane. The characterization of ATP-dependent taurocholate transport in canalicular membrane vesicles indicated the presence of a specific carrier system for monovalent bile salts. The main transport system that mediates excretion of monovalent bile salts is the so called “bile salt export pump” (BSEP). The sequence is more homologous with the MDR family than with the MRPs. BSEP is

G. Ramadori and P. Ramadori

expressed on the surface of canalicular microvilli as indicated by electron microscopic studies. Rat BSEP mediates the ATPdependent transport of taurocholate, glycocholate, taurochenodeoxycholate, and tauroursodeoxcholate. The human BSEP gene locus has been identified as the positional candidate for progressive familial intrahepatic cholestasis type 2 (PFIC2), a progressive liver disease characterized by low biliary bile salt concentrations. In PFIC2, BSEP is absent from the canalicular membrane and biliary salt concentrations are less than 1% of normal [93, 94]. Interestingly, a recent work with a tissue-specific selection of the transcription factor FOXA2 in hepatocyte has been shown to lead to cholestasis, exacerbated with a cholic acid supplemented diet [95]. Accumulation of hepatic bile acids in those mice was  explained by the authors by lowered hepatic mRNA and protein levels of several transporters of bile acids and their conjugates, including Mrp2, Mrp4, Oatp2, and Mrp3, but the levels of expression of Bsep were unchanged. The identification of FOXA2 as an important regulator of the main bile acids transporters could represent a starting point for the development of new therapeutic strategies.

Excretion of Nonbile Salt Organic Anions The excretion of nonbile salt organic anions into the bile is mediated by the canalicular multidrug resistance protein 2 (MRP2). Both rat and human MRP2 are expressed predominantly in the liver with exclusive localization in the canalicular membrane. The spectrum of organic anions is qualitatively similar to that of MRP1. It includes glutathion conjugates, glucuronides (conjugated bilirubin), leukotriene C4 and divalent bile salts, but not monovalent bile salts. A role for MRP2 in the canalicular excretion of reduced GSH, a major driving force for the maintenance of bile salt-independent bile flow, has also been demonstrated. In addition it is suggested that various structurally and functionally unrelated xenobiotics e.g., robenecid, glibenclamide, rifampicin inhibit excretion of organic anion excretion [92, 93].

Phospholipid Excretion The major lipid that is excreted together with cholesterol is phosphatidylcholine. The constant delivery of

1  Hepatocytes

phosphatidylcholine from the inner to the outer hemileaflet of the canalicular membrane is mediated by several ATP-dependent and ATP-independent phosphatidylcholine floppases. The ATP dependent flippase is a multidrug resistance glycoprotein (MDR) called Mdr2 in rodents or MDR3 in humans. Mdr2 and MDR3 are highly expressed in the canalicular membrane of the hepatocytes [96, 97].

Copper Excretion The liver is the central organ of copper homeostasis and has great capacity to store and excrete this metal. The degree of copper excretion is directly proportional to the size of the hepatic copper pool, suggesting that hepatocytes can sense the copper status in the cytoplasm and then regulates copper excretion into the bile. The biliary excretion of heavy metals such as copper is an important detoxifying mechanism of the liver. Copper excretion is mediated by a copper transporting P-type ATPase called ATP7B, predominantly expressed in the liver. It is localized to the trans-Golgi network where it mediates the incorporation of cop-per into cuproenzymes such as coeruloplasmin. An isoform of ATP7B is located in the mitochondria which might be a reason for the abnormalities of the mitochondria morphology in Wilson´s disease. The ATP7B was also found in the so called late endosomes. Copper incorporated into late endosomes is probably transported to lysosomes and subsequently excreted into bile by a process known as biliary lysosomal excretion. Copper is probably taken up into the hepatocytes via the copper transporters hCTR1 and hCTR2. As the copper concentration of the hepatocyte in-creases, ATP7B redistributes from the trans Golgi network to a cytoplasmic vesicular compartment and to pericanalicular vacuoles. After copper depletion, ATP7B returns to the trans-Golgi network. By these means, copper can induce redistribution of its own transporter from the trans-Golgi network to the apical membrane, where it may mediate biliary copper excretion. Copperinduced redistribution of ATP7B may provide a mechanism to preserve copper when it is scarce and to prevent copper toxicity when levels become too high [98, 99].

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Summary

›› Hepatocytes ›› ›› ›› ›› ›› ››

›› ›› ››

are arranged in plates between the portal space and the central vein. Hepatocytes are polarized cells with an apical membrane facing the bile canaliculus and a basal side facing the sinusoid. Hepatocytes in adults are frequently polyploid. Hepatocytes play a central role in lipid, cholesterol, and glucose metabolism. Hepatocytes are responsive to acute phase signals and massively increase the synthesis of specific proteins. Hepatocytes regulates iron metabolism and secrete hepcidin. Hepatocytes are not only targets for hormones such as insulin and glucagons, but synthesize hormones such as IGF-1 and, during the fetal life, EPO. Hepatocytes are equipped with numerous transporters. Hepatocytes excrete into the bile conjugated bile salts, conjugated bilirubin and phospholipids. Hepatocytes excrete into the bile copper.

Multiple Choice Questions 1. What is correct? (a) The canalicular membrane contains several ATP-dependent transporters (b) Conjugated bile salts are taken up by the basolateral NTCP transporter (c) The apical transporter MRP2 excretes conjugated bilirubin into the bile (d) Tight junctions are essential to seal the bile canaliculus (e) All the above statements are correct 2. Hepatocytes synthesize (a) Von Willebrand factor (b) Immunoglobulins (c) Complement C1q (d) Factor V (e) Growth hormone

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G. Ramadori and P. Ramadori Insulin

IL-6

TNF-alpha

IL-1 beta

IRS

Ins Rec

STAT3

MyD88

SOCS1 SOCS3 JNK IKKβ Inflammatory genes NFkB

ROS ER STRESS

AP-1

nucleus

Hyperlipidemia

Fig. 1.5  Possible mechanisms of hepatocytes insulin resistance (modified from Raddatz and Ramadori [106]). The increase of circulating fatty acids and the accumulation of triglycerides in the liver and in the adipose tissue induce the release of cytokines

Fig. 1.6  Diagram of hepatobiliary transport systems. Canalicular transporters function in an ATP-dependent manner. OATP organic anion transporting polypeptide family; NTCP Na+taurocholate cotransporting polypeptide; OCT organic cation transporter; OAT organic anion transporter; BSEP bile salt excretion protein; MDR multidrug resistance glycoprotein; MRP multidrug resistance protein family; ABC ATP-binding cassette transporters; GSH glutathione

and adipokines that increasing the cellular stress (oxidative stress and lipid peroxidation) triggers to an amplification of the inflammatory state and to an inhibition of sensitivity of the insulin receptor

1  Hepatocytes

3. What is correct? (a) Insulin induces the expression of PECK and G6Pase (b) Glucagon inhibits the expression of PECK and G6Pase (c) Glucocorticoids inhibit the expression of PECK and G6Pase (d) Plasma membrane transport of glucose is a major regulatory site (e) Hepatocytes use the glucose transporter Glut2 4. What is incorrect regarding SER? (a) It is involved in protein synthesis (b) It is involved in drug metabolism (c) It is involved in cholesterol metabolism (d) It is involved in bile acids synthesis (e) It contains cytochrome P450 enzymes 5. Which of these factors is not involved in the development of biliary epithelial cells? (a) Notch-2 (b) Jag1 (c) HNF-6 (d) HNF-1b (e) Prox-1

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1  Hepatocytes inflammation: involvement of interleukin-6. Lab Invest 85(3): 376–387 70. Burmester H, Wolber EM, Freitag P et al (2005) Thrombopoietin production in wild type and interleukin-6 knockout mice with acute inflammation. J Interferon Cytokine Res 25(7):407–413 71. Anderson GJ (2001) Ironing out disease: inherited disorders of iron homeostasis. IUBMB Life 51(1):11–17 72. Angelucci E, Brittenham GM, McLaren CE et al (2000) Hepatic iron concentration and total body iron stores in thalassemia major. N Engl J Med 343(5):327–331 73. Sheikh N, Dudas J, Ramadori G (2007) Changes of gene expression of iron regulatory proteins during turpentine oilinduced acute-phase response in the rat. Lab Invest 87(7): 713–725 74. Voci A, Arvigo M, Massajoli M et al (1999) IGF-I production by adult rat hepatocytes is stimulated by transforming growth factor-alpha and transforming growth factorbeta1. Eur J Endocrinol 140(6):577–582 75. Sjogren K, Liu JL, Blad K et al (1999) Liver-derived insulinlike growth factor I (IGF-I) is the principal source of IGF-I in blood but is not required for postnatal body growth in mice. Proc Natl Acad Sci U S A 96(12):7088–7092 76. Domene HM, Bengolea SV, Martinez AS et al (2004) Deficiency of the circulating insulin-like growth factor system associated with inactivation of the acid-labile subunit gene. N Engl J Med 350(6):570–577 77. Yakar S, Setser J, Zhao H et al (2004) Inhibition of growth hormone action improves insulin sensitivity in liver IGF-1deficient mice. J Clin Invest 113(1):96–105 78. Price JA, Kovach SJ, Johnson T et al (2002) Insulin-like growth factor I is a comitogen for hepatocyte growth factor in a rat model of hepatocellular carcinoma. Hepatology 36(5): 1089–1097 79. Maxwell PH, Osmond MK, Pugh CW et al (1997) Sites of erythropoietin production. Kidney Int. 51:393–401 80. Bondurant M, Koury M (1986) Anemia induces accumulation of erythropoietin mRNA in the kidney and liver. Mol Cell Biol 6:2731–2733 81. Sohmiya M, Kato Y (2005) Human growth hormone and insulin-like growth factor-I inhibit erythropoietin secretion from the kidneys of adult rats. Journal of Endocrinology 184:199–207 82. Kullak-Ublick GA, Stieger B, Meier PJ (2004) Enterohepatic bile salt transporters in normal physiology and liver disease. Gastroenterology 126(1):322–342 83. Denson LA, Auld KL, Schiek M et al (2000) Interleukin1beta suppresses retinoid transactivation of two hepatic transporter genes involved in bile formation. J Biol Chem 275:8835–8843 84. Eloranta JJ, Jung D, Kullak-Ublik GA (2006) The human Na+taurocholate cotransporting polypeptide gene is acti-vated by glucocorticoid receptor and peroxisome proliferator-­activated receptor-gamma coactivator-1alpha, and suppressed by bile acids via a small heterodimer partner-dependent mechanism. Mol Endocrinol 20:65–79 85. Kim RB (2003) Organic anion-transporting polypeptide (OATP) transporter family and drug disposition. Eur J Clin Invest 33(Suppl 2):1–5 86. Shih DQ, Bussen M, Sehayek M et al (2001) Hepatocyte nuclear factor-1alpha is an essential regulator of bile acid and plasma cholesterol metabolism. Nature Gen 27:375–382

23 87. Gorboulev V, Ulzheimer JC, Akhoundova A et al (1997) Cloning and characterization of two human polyspecific organic cation transporters. DNA Cell Biol 16(7):871–881 88. Grundemann D, Gorboulev V, Gambaryan S et al (1994) Drug excretion mediated by a new prototype of polyspecific transporter. Nature 372(6506):549–552 89. Michalopoulos G (1991) Control of hepatocyte proliferation in regeneration, augmentative hepatomegaly, and neoplasia. Prog Clin Biol Res 369:227–236 90. Kauffmann HM, Keppler D, Kartenbeck J et al (1997) Induction of cMrp/cMoat gene expression by cisplatin, 2-acetylaminofluorene, or cycloheximide in rat hepatocytes. Hepatology 26(4):980–985 91. Keppler D, Leier I, Jedlitschky G et al (1996) The function of the multidrug resistance proteins (MRP and cMRP) in drug conjugate transport and hepatobiliary excretion. Adv Enzyme Regul 36:17–29 92. Silverman JA (1999) Multidrug-resistance transporters. Pharm Biotechnol 12:353–386 93. Redinger RN (2003) The coming of age of our understanding of the enterohepatic circulation of bile salts. Am J Surg 185(2):168–172 94. Strazzabosco M, Boyer JL (1996) Regulation of intracellular pH in the hepatocyte. Mechanisms and physiological implications. J Hepatol 24(5):631–644   95. Bochkis IM, Rubins NE, White P et al (2008) Hepatocytespecific ablation of Foxa2 alters bile acid homeostasis and results in endoplasmic reticulum stress. Nature Med 14(8): 828–836   96. Elferink RP, Tytgat GN, Groen AK (1997) Hepatic canalicular membrane 1: the role of mdr2 P-glycoprotein in hepatobiliary lipid transport. FASEB J 11(1):19–28   97. Erlinger S (1996) Mechanisms of hepatic transport and bile secretion. Acta Gastroenterol Belg 59(2):159–162   98. Harada M, Kawaguchi T, Kumemura H et al (2003) Where is the site that ATP7B transports copper within hepatocytes? Gastroenterology 125(6):1911–1912   99. Schilsky ML, Irani AN, Gorla GR et al (2000) Biliary copper excretion capacity in intact animals: correlation between ATP7B function, hepatic mass, and biliary copper excretion. J Biochem Mol Toxicol 14(4):210–214 100. Besinger SJ, Tontonoz P (2008) Integration of metabolism and inflammation by lipid-activated nuclear receptor. Nature 454:470–477 101. Francanzani AL, Valenti L, Bugianesi E et al (2008) Risk of severe liver disease in nonalcoholic fatty liver disease with normal aminotransferase levels: a role for insulin resistance and diabetes. Hepatology 48:792–798 102. Konig J, Nies AT, Cui Y et al (1999) Conjugate export pumps of the multidrug resistance protein (MRP) family: localization, substrate specificity, and MRP2-mediated drug resistance. Biochim Biophys Acta 1461(2): 377–394 103. Lemaigre F, Zaret KS (2004) Liver development update: new embryo models, cell lineage control, and morphogenesis. Curr Opin Gen Dev 14:582–590 104. Nagel G, Volk C, Friedrich T et al (1997) A reevaluation of substrate specificity of the rat cation transporter rOCT1. J Biol Chem 272(51):31953–31956 105. Nguyen P, Leray V, Diez M et al (2008) Liver lipid metabolism. J Anim Physiol Anim Nutr 92:272–283

24 106. Raddatz D, Ramadori G (2007) Carbohydrate metabolism and the liver: actual aspects from physiology and disease. Z Gastroenterol 45:51–62 107. Seki S, Habu Y, Kawamura T et al (2000) The liver as a crucial organ in the first line of host defense: the roles of Kupffer cells, natural killer (NK) cells and NK1.1 Ag+ T cells in T helper 1 immune responses. Immunol Rev 174: 35–46

G. Ramadori and P. Ramadori 108. Van de Werve G, Lange A, Newgard C et al (2000) New lessons in the regulation of glucose metabolism taught by the glucose 6-phosphatase system. Eur J Biochem 267(6): 1533–1549 109. Watford M (1991) The urea cycle: a two-compartment system. Essays Biochem 26:49–58

2

Signaling Pathways in Biliary Epithelial Cells M. Fatima Leite, Viviane A. Andrade, and Michael H. Nathanson

Introduction Biliary epithelial cells, or cholangiocytes, line the lumen of the biliary tree. Like hepatocytes, cholangiocytes are a polarized epithelium with structural features that include well-defined apical and basolateral membrane domains. Cholangiocytes constitute approximately 5% of the mass of the liver and play an important role in the formation of bile by altering primary canalicular bile through a series of secretory and reabsorptive events. These events are regulated by peptide hormones, nucleotides, bile salts, growth factors, cytokines, and neurotransmitters that bind to and stimulate specific apical or basolateral surface membrane receptors, which in turn initiate intracellular signal transduction pathways that regulate cell function. In addition to their role in the modification of ductal bile, cholangiocytes participate in the detoxification of xenobiotics [1]. In the adult liver, cholangiocytes are mitotically dormant [2]. Cholangiocyte proliferation may include some combination of proliferation of preexisting ductules, progenitor cell activation, and appearance of intermediate hepatocytes. Proliferating cholangiocytes display enhanced secretory activity. This may serve to compensate for the impaired secretion of injured cells and maintain biliary mass and secretory function during disease states [2]. Cholangiocytes are the primary target of injury in a variety of cholestatic liver diseases, such as sclerosing cholangitis, primary biliary cirrhosis, cystic fibrosis, and biliary atresia. There is evidence

M. H. Nathanson () Department of Medicine, Yale University School of Medicine, 1 Gilbert Street, New Haven, CT 06520-8019, USA e-mail: [email protected]

that signaling pathways are altered in such disorders, which may contribute to the secretory defects that characterize these diseases. Cholestatic disorders represent the main indication for liver transplantation in pediatrics and are a common indication in adults [3, 4]. This chapter will systematically review the membrane receptors and associated intracellular signaling pathways that regulate cholangiocyte function, and will discuss the alterations that occur in these signaling pathways in cholangiopathic diseases.

Membrane Receptors Several well-characterized families of membrane receptors have been identified in cholangiocytes. The receptors that associate with guanosine triphosphate (GTP)-binding regulatory proteins, or G proteins, constitute the largest family. Members of the G protein-coupled receptor family share structural and functional similarities, and contain seven hydrophobic membrane-spanning domains plus a cytoplasmic site for binding to G proteins. At least fifteen G proteins have been identified and each consists of a heterotrimeric complex consisting of alpha, beta, and gamma subunits. The alpha subunit has intrinsic GTPase activity at the guanine nucleotide binding site, plus a specific binding site for the receptor and effector proteins. When a ligand binds to the receptor, guanosine diphosphate (GDP) rapidly exchanges for GTP, which allows the G protein to dissociate from the receptor and the alpha subunit to dissociate from the beta–gamma subunits. The dissociated subunits then activate effector proteins such as adenylate cyclase, phospholipase C (PLC) or other hydrolyses, which in turn generates or activates signaling molecules such as cyclic adenosine

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_2, © Springer-Verlag Berlin Heidelberg 2010

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monophosphate (cAMP), cAMP-dependent protein kinase A (PKA), inositol 1,4,5-trisphosphate (InsP3), 1-2 diacylglycerol (DAG), cytoplasmic calcium (Cai2+), or protein kinase C (PKC). G protein-coupled receptors that have been identified in cholangiocytes include the secretin receptor, the bombesin receptor, the vasoactive intestinal peptide (VIP) receptor, the M3 muscarinic acetylcholine receptor, the gastrin receptor, the a1B and a2 adrenergic receptors, the somatostatin receptor, the type A and B endothelin receptors, the serotonin receptor, and several subtypes of the P2Y nucleotide receptor. Cholangiocytes also express all four subtypes of the histamine receptor. These subtypes are classified as H1R, H2R, H3R, and H4 and have been identified in both small and large cholangiocytes [5]. Under normal conditions, the predominant histamine receptors in cholangiocytes are H1R, which increases cytoplasmic Ca2+ via Gq, and H2R, which increases cAMP via Gs. Activation of H1R is associated with calmodulindependent stimulation of calmodulin-dependent protein kinase (CaMK) and activation of cAMP-response element binding protein (CREB), which results in cholangiocyte proliferation [6]. Following bile duct ligation, expression shifts to H3R and H4R, both of which inhibit cAMP formation [7]. This serves to inhibit PKA activation, which decreases activation of ERK 1/2 and Elk-1, which in turn retards proliferation of cholangiocytes [7]. Functional evidence suggests that cholangiocytes can signal via the endocannabinoid system as well, which consists of the G protein-coupled cannabinoid receptors Cb1 and Cb2 that are activated by anandamide (AEA). These receptors could be of relevance in the pathogenesis of liver fibrosis and portal hypertension [8]. For example, treatment of cholangiocytes with AEA impairs cholangiocyte proliferation after extrahepatic biliary obstruction [9]. Activation of this pathway involves the accumulation of reactive oxygen species and activation and nuclear translocation of thioredoxin 1 (TRX1), where it interacts with redox factor 1 (Ref1) to modulate the DNA-binding activity of the activator protein 1 (AP-1) transcription factor complex [9]. Activation of Cb receptors and of the endocannabinoid pathway thus plays a role in the suppression of cholangiocyte proliferation. Non-G protein-coupled receptors have also been identified in cholangiocytes. This broad group includes receptors for epidermal growth factor (EGF), nerve growth factor, insulin, interleukins, and lipopolysaccharides. These receptors typically have a single

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plasma membrane spanning domain and possess ligand-­activated tyrosine kinase activity. Such receptor tyrosine kinases stimulate several downstream signal transduction pathways, including those associated with phosphatidyl inositol metabolism and the mitogenactivated protein kinase (MAPK) cascade. The MAPK pathway requires several steps for activation, involving autophosphorylation of the tyrosine kinase receptor that provides recognition and binding sites for the src homology (SH2) domain of the adaptor molecule Grb2. This in turn causes the binding of another protein, SOS, at the SH3 domain of Grb2, required for activation of the ras proto-oncogene. Ras triggers the Ras/Raf/MEK/ MAPK cascade reaction [10]. The serine/threonine protein kinases of the Raf family (Raf-1, B-Raf, and A-Raf) play a key role in growth factor signaling by the ­phosphorylation and stimulation of MEK, then MAPK [11,  12] and subsequently, by the intranuclear activation of transcription factors, including Elk-1, myc, fos, and jun [13]. This signaling pathway induces a variety of biological responses, including cell proliferation, differentiation, and apoptosis. ERK signaling furthermore undergoes spatial control by Sef, an inhibitor that acts as a molecular switch for ERK by specifically inhibiting nuclear translocation of ERK without inhibiting its activity in the cytoplasm [14]. The Ras/Raf/ MEK/MAPK cascade reaction also can be modulated via cross talk with other intracellular signaling pathways. In some cells, including cholangiocytes, cAMP/ PKA signaling inhibits MAPK activity through the inhibition of Raf-1 and B-Raf activation [15, 16]. On the other hand, stimulation of cytokine receptor C-X-C motif chemokine receptor 4 (CXCR4) via the CXC chemokine ligand 12 (CXCL12) plays an important role in cholangiocarcinoma cell invasion by induction of the ERK1/2 and AKT pathways [17]. Inflammation signals mediated by the inflammatory cytokine IL-6 also induce the ERK and AKT pathways and enhance expression of the anti-apoptotic protein Mcl-1 in cholangiocarcinoma cells, thereby enhancing their survival [18]. Bile acids activate several membrane receptors in cholangiocytes. Bile acids activate the epidermal growth factor receptor (EGFR), which occurs through a TGFa-dependent mechanism [13, 19]. Bile acids also regulate the expression of death receptor 5/TRAILreceptor 2 via a c-Jun N-terminal kinase-dependent pathway and modulate the IGF1 system [20]. This effect depends upon the hydrophobicity of the bile salt, with deoxycholate exerting a maximal effect and

2  Signaling Pathways in Biliary Epithelial Cells

ursodeoxycholate exerting no measurable effect. This pathway protects cholangiocytes as well as hepatocytes against bile acid-induced cytotoxicity [20, 21]. Finally, there is an evidence that bile salts may activate the cystic fibrosis transmembrane conductance regulator (CFTR) Cl− channel [22]. CFTR and the apical sodium-dependent bile acid transporter (ASBT) colocalize on the apical membrane of large cholangiocytes [23] and evidence suggests that taurocholate directly modulates gating of the CFTR Cl− channel [22]. Cholangiocytes express not only cell membrane receptors, but also intracellular steroid hormone receptors. Steroid hormones enter cells by passive diffusion across the plasma membrane in order to bind to their intracellular receptor. Receptor binding results in a conformational change that increases the affinity of the receptor to bind to DNA. Specific binding sites are present on regulatory regions of genes, which serve to alter transcription and thus protein synthesis. Steroid receptors characterized in cholangiocytes include the a and b estrogen receptors (ERa and b, respectively) [24]. Cholangiocarcinomas express ERa and b, insulinlike growth factor 1 (IGF1) and IGF1 receptor (IGF1R). In these cells, estrogens cooperate with IGF1 and their receptor to modulate tumor growth [25]. Estrogens also induce the expression of vascular endothelial growth factor receptor (VEGFR) and may enhance cell proliferation on that basis as well [24–26].

Cyclic Adenosine 3’, 5’-Monophosphate (cAMP) Adenylate cyclase is a membrane-bound enzyme that can be regulated by alpha subunits of two G proteins, Gi and Gs. Gi inhibits adenylate cyclase activity, while Gs catalyzes the formation of cAMP from adenosine triphosphate (ATP). Cyclic AMP then activates PKA, which in turn phosphorylates a range of intracellular proteins, triggering a series of events that lead to specific cellular responses, such as ion secretion, absorption, and motility. Specific effects of cAMP in cholangiocytes include stimulation of exocytosis [27], opening of aquaporin water channels [28, 29], activation of the cystic fibrosis transmembrane conductance regulator (CFTR) Cl−channel [30, 31], and activation of Cl−/HCO3− exchange [32, 33]. Increases in cAMP in cholangiocytes, increase ductular bile flow in isolated

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bile duct units [32–35], but not in the intact, perfused rat liver [36]. This apparent inconsistency may be explained by the observation that ductular secretion in rats accounts for only ~10% of total bile flow [37]. Secretin receptors belong to the class of G proteincoupled receptors that activate adenylate cyclase to increase cAMP formation [38, 39]. Activation of the secretin receptor does not affect Cai2+ in isolated bile duct cells [40], nor does it affect cAMP production in hepatocytes [38]. Thus, in the liver these receptors are expressed only by cholangiocytes [38] and are localized to the basolateral membrane of these cells. Stimulation of secretin receptors induces ductal secretion by activation of CFTR. Among the different types of ion channels expressed by cholangiocytes, CFTR has been investigated most extensively and appears to be largely responsible for secretin-stimulated increases in apical Cl− secretion. Phosphorylation by PKA increases the open probability of plasma membrane CFTR channels, which is the mechanism by which cAMP increases apical Cl− secretion through this ion channel [41]. Activation of CFTR is also associated with ductular bicarbonate excretion, since forskolin, a direct activator of adenyl cyclase, alkalinizes the lumen of isolated bile duct units [35]. Studies in isolated cholangiocytes and bile duct units suggest that Cl− secretion via CFTR is linked to Cl−/HCO3− exchange [32, 34], and that this linkage leads to net secretion of HCO3− by bile duct epithelia [42]. However, work in the intact, perfused liver instead suggests that secretin-induced HCO3− secretion depends on the activation of Cl− channels, but not on Cl−/HCO3− exchange [36]. In fact, direct evidence in other systems similarly shows that CFTR can regulate bicarbonate efflux directly, rather than through activation of Cl−/HCO3− exchange [43–46]. It has been suggested that these apparent differences between in vitro and in vivo studies may be related to paracrine effects as well as vascular factors that alter signaling and secretion in vivo. Yet another possible mechanism for cAMP-mediated ductular HCO3− secretion has been delineated in microdissected, microperfused intrahepatic bile ductal units (IBDUs). In this experimental system, forskolin-induced alkalinization of the ductular lumen could be inhibited by knockdown of the apical, type III inositol 1,4,5-trisphosphate receptor (InsP3R) in cholangiocytes or by hydrolysis of luminal ATP, inhibition of apical P2Y nucleotide receptors, or buffering of cytosolic Ca2+ signals [47]. These findings suggest that cAMP-induced ductular

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HCO3− secretion depends on an autocrine signaling pathway that involves CFTR-mediated apical secretion of ATP, which leads to the stimulation of apical P2Y receptors, and then to the activation of apical, type III InsP3Rs [47]. Regardless of the mechanism, it is well accepted that increases in cAMP lead to HCO3− excretion in cholangiocytes. Water movement into the ductular lumen is increased by the increase in cAMP in cholangiocytes [34, 35]. Cholangiocytes contain secretory vesicles that are enriched in aquaporin-1, a water-selective channel protein, as well as CFTR, other Cl− channels and the Cl−/HCO3− exchanger [48]. Secretin or dibutyryl cAMP causes a rapid redistribution of these secretory vesicles from the cell interior to the apical membrane of cholangiocytes, leading to ion-driven water-transport and ductal bile secretion [27, 28, 48, 49]. Hormonal stimulation can serve not only to increase cAMP-mediated secretion in cholangiocytes, but also to decrease it. For instance, somatostatin acts on cholangiocytes via SSTR2 somatostatin receptors to increase cGMP, which inhibits secretin-stimulated cAMP synthesis, thereby decreasing bile formation through a combination of events that involve inhibition of ductal fluid secretion and stimulation of ductal fluid absorption [50]. Gastrin and endothelin-1, hormones that activate the PLC pathway, also inhibit secretinstimulated secretion. Like somatostatin, gastrin and endothelin-1 inhibit secretin-induced ductal bile secretion by binding to their specific cholangiocyte receptors, which links to decreased expression of the secretin receptor and a decrease in secretin-stimulated cAMP formation [49, 51–53]. Cyclic AMP has effects not only on secretion but also on proliferation. Administration of a cell-permeant form of cAMP enhances proliferation of cholangiocytes after bile duct ligation, whereas, the neuroendocrine hormone serotonin inhibits proliferation. Serotonin acts on cholangiocytes through both the 1A and 1B subtypes of its receptor. Animals subjected to bile duct ligation are treated with serotonin receptor agonists; there is a strong reduction in secretin-induced bile flow, HCO3− secretion, cAMP synthesis, and PKA activity [54]. Therefore, activation of the serotonin 1A and 1B receptor in cholangiocytes inhibits proliferation by reducing cAMP formation and subsequent activation of PKA. Bombesin, VIP, and ATP each stimulate cholangiocyte secretory responses similar to the effect of

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secretin, but unlike secretin, the effects of these agents do not appear to be mediated by cAMP [55–57]. The signaling pathways involved in such cAMP-independent secretory responses in cholangiocytes are not well characterized yet, but seems likely that Cl− channels other than CFTR are involved. In fact, a number of other types of Cl− channels have been identified in cholangiocytes [58], but the exact role of these channels in ductular secretion remains unclear.

Cytosolic Ca2+ Phosphatidylinositol 4,5-bisphosphate (PIP2) is a membrane lipid that is a substrate for PLC. Once it has been activated by a specific class of G protein-coupled receptors, PLC cleaves PIP2 into two signaling molecules, DAG and InsP3. DAG is lipophilic and remains at the membrane, where it activates PKC. InsP3 is a water-soluble molecule that diffuses through the cytosol to interact with the InsP3R, which stimulates the release of Ca2+ from intracellular Ca2+ stores into the cytosol. Three isoforms of the InsP3R have been identified and are termed type I, type II, and type III. Although the three isoforms have a high degree of sequence homology and behave as InsP3-gated intracellular Ca2+ release channels, they differ in sensitivity to InsP3, with the type II isoform being the most sensitive, followed by the type I, and then the type III [59]. The open probability of each isoform of InsP3R is further regulated by Ca2+ itself, but the effect of Ca2+ on each isoform appears to be distinct [60, 61]. InsP3R isoforms are expressed in relatively unique proportions in different tissues and have specific subcellular patterns of distribution as well. The behavior of the InsP3R is also altered by tissue-specific expression of various cofactors that interact with and modify the behavior of the InsP3R. For example, chromogranin A and chromogranin B each binds to the ER luminal aspect of the InsP3R and increases the open probability of the receptor [62, 63]. Chromogranin A is found in cholangiocytes but not hepatocytes [64], while chromogranin B is not found in the liver. The differences in behavior of each InsP3R isoform, along with tissuespecific patterns of distribution of the isoforms and their cofactors, suggest that these factors may permit tissue-specific patterns of Ca2+ signaling. Rat cholangiocytes express all three isoforms of InsP3R [65]. The

2  Signaling Pathways in Biliary Epithelial Cells

Fig. 2.1  The type III InsP3 receptor is concentrated in the apical region of cholangiocytes. Image shows confocal immunofluorescence labeling of a section through rat liver. The tissue was labeled with a monoclonal antibody directed against the N-terminal region of the type III InsP3R (image provided courtesy of Dr. Keiji Hirata)

type III InsP3R is expressed most heavily and is concentrated apically (Fig.  2.1) while type I and type II InsP3R are expressed to a similar extent and are distributed uniformly throughout the cytosol. Human cholangiocytes, like rat cholangiocytes, express all three InsP3R isoforms, and express the type III isoform most heavily [66]. NRC cells, a polarized rat cholangiocyte cell line, express the type III InsP3R almost exclusively, and this isoform is concentrated apically in these cells, just as in primary cholangiocytes [65]. The ryanodine receptor (RyR) is a separate intracellular Ca2+ release channel that plays a major role in cytosolic Ca2+ signaling. Like the InsP3R, the RyR has three isoforms, each of which displays distinct functional properties. It was believed initially that RyRs regulate Ca2+ signaling only in myocytes, whereas InsP3Rs regulate Ca2+ signaling in nonexcitable cells such as epithelia. However, many cell types express both RyR and InsP3R [67, 68] including a number of polarized epithelia [69, 70]. Each RyR isoform is activated by a process known as Ca2+-induced Ca2+ release (CICR). The type II and III RyRs also are sensitive to cyclic adenosine diphosphate (ADP)-ribose (cADPr) [71, 72]. Cholangiocytes from rat liver do not express RyR, but faint expression of type I RyR can be detected in NRC cells by RT-PCR [65]. Cytosolic Ca2+ signals are encoded through signaling patterns such as Ca2+ waves and Ca2+ oscillations, and

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both of these types of Ca2+ signals occur in cholangiocytes [40, 65]. The ability to form Ca2+ waves in cholangiocytes and other polarized epithelia is thought to depend on the pattern of expression of InsP3R isoforms [73]. Specifically, Ca2+ waves begin in the apical region of cholangiocytes, where the type III InsP3R is concentrated [65]. This association is observed in other epithelia as well [74, 75], and indeed the single-channel behavior of the type III receptor suggests that it may trigger Ca2+ waves [60]. Ca2+ signals can spread among neighboring cholangiocytes. This effect is mediated by gap junctions in most epithelia [76], including cholangiocytes [77]. The types of Ca2+ release channels in the basolateral region may also affect the formation of Ca2+ waves. For example, in pancreatic acinar cells all three InsP3R isoforms are concentrated in the apical region, whereas the RyR is basolateral [69]. In contrast, the type III isoform of the InsP3R predominates in the apical region while expression of the other isoforms is not limited to this region in both cholangiocytes and nonpigmented epithelial cells of the ocular ciliary body [65, 74]. Formation of polarized Cai2+ waves may be important for regulation of secretion in particular. Cai2+ waves regulate secretion in several ways, including activation of specific transporters and ion channels [78], and stimulation of vesicle fusion with the plasma membrane [79]. Vesicle fusion can in turn promote secretion either via exocytic release of vesicles contents [80], or by inserting additional transporters or channels into the plasma membrane, as occurs with canalicular transporters in the hepatocyte [81]. In cholangiocytes, ductular secretion depends upon apical insertion of water channels [28, 29] and other membrane fusion events [27, 82]. In these cells, Cai2+ regulates ductular secretion through pathways that differ from those activated by cAMP. Cai2+ directly activates apical Ca2+-dependent Cl− channels that are distinct from CFTR [31, 83], so that either Ca2+ or cAMP can mediate apical bicarbonate secretion [36, 84]. However, Ca2+ may also potentiate cAMP-mediated secretion. In particular, activation of M3 muscarinic receptors by acetylcholine enhances secretin-induced cAMP formation [85, 86] and accelerates secretin-induced Cl−/HCO3− exchange [32]. This is mediated by calcineurin and inhibited by cyclosporin [32]. This effect has been observed in the intact, perfused liver as well, but only if secretin is administered before cyclosporin [36]. Ca2+ oscillations can be induced in cholangiocytes by either ATP or acetylcholine. ATP-induced oscillations

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have been observed both in individual cholangiocytes within isolated bile duct units and in NRC cells [40, 65, 77]. This Cai2+ signaling pattern is concentration-dependent, since lower concentration of ATP predominantly induces Cai2+ oscillations, whereas, higher concentrations predominantly induce single increases in Cai2+ [40]. ATP is a signaling molecule in virtually all types of tissues, and at least 18 types of purinergic receptors have been identified. Cholangiocytes and biliary cell lines express P2X receptors [84, 87], which are ATP-gated cation channels, as well as P2Y receptors [84, 88], which are G protein-coupled receptors that increase Cai2+ via InsP3. However, ATP-induced Ca2+ signaling in cholangiocytes occurs principally via apical P2Y1, P2Y2, P2Y4, and P2Y6 subtypes of the P2Y receptor [89, 90]. Individual hepatocytes can release nucleotides in amounts sufficient to induce Cai2+ signals in cocultured bile duct cells [91], and measurable amounts of ATP can be detected in rodent and human bile [92, 93]. Thus, one possible mechanism for paracrine regulation of Ca2+ signaling in cholangiocytes may be for hepatocytes to signal to cholangiocytes through release of nucleotides into bile, followed by activation of apical P2Y receptors. Stimulation of cholangiocyte P2Y receptors in turn results in InsP3-mediated increases in cytosolic Ca2+, leading to ductular bicarbonate secretion [84]. Autocrine regulation of Ca2+ signaling in cholangiocytes may occur through a related mechanism in which cAMP induces CFTR-mediated release of ATP into bile [47], as described above. Cholangiocytes also express basolateral P2Y receptors [40, 84], which may mediate signaling from neural or vascular tissues. However, signaling via basolateral nucleotide receptors is attenuated by the expression of NTPDases by portal fibroblasts [94]. Cholangiocytes express multiple P2Y receptor subtypes [84]. P2Y12 receptors are expressed in the primary cilia on the apical surface of cholangiocytes. Cilia are sensory organelles which respond to chemical, osmotic, and mechanical stimuli [95]. Unlike most other P2Y receptors, the P2Y12 receptor is a G protein-coupled receptor associated with cAMP instead of Ca2+ signaling [96]. The expression of P2Y12 receptors in rat cholangiocytes links specifically to the chemo-sensory function of primary cilia [96]. Stimulation of P2Y12 receptors in rat cholangiocytes by either ADP or ATP-gS activates Gi and thus inhibits forskolin-induced formation of cAMP [96]. Cholangiocyte cilia also express TRPV4, a member of the transient receptor potential (TRP) superfamily of

M. F. Leite et al.

Ca2+ channels. Activation of this channel via hypotonic stimuli induces an increase in cytosolic Ca2+, as well as apical release of ATP and an increase in HCO 3− secretion [95]. Therefore, stimulation of primary cilia activates a variety of signaling pathways in cholangiocytes, which depends in part upon the type of stimulus [95]. Most studies of Ca2+ signaling in cholangiocytes have examined signals induced by hormones and other humoral factors. However, increasing evidence suggests that neurotransmitters regulate Ca2+ signals in bile ducts as well. Acetylcholine activates M3 muscarinic receptors [85] to induce Ca2+ waves and oscillations in isolated bile duct units [40, 65]. Unlike ATP, acetylcholine induces Ca2+ oscillations that have no clear concentration dependence [40]. Ca2+ signals induced by both ATP and acetylcholine mediate ductular bicarbonate secretion, however [36, 84]. Additional studies of the role of cholinergic innervation have been performed by examining the effect of vagotomy on cholangiocytes. Bile duct ligation induces proliferation of cholangiocytes and enhances their response to secretin, but vagotomy impairs this response, in part by enhancing apoptosis of cholangiocytes [86]. Vagotomy also eliminates the choleretic response to secretin that is induced by bile duct ligation [86]. Interestingly, this effect is reversed by forskolin, which may be consistent with the idea that cholinergic stimulation serves in part to potentiate formation of cAMP in cholangiocytes [85]. Adrenergic stimulation also increases Cai2+ in cholangiocytes. The a1-adrenergic agonist phenylephrine potentiates secretin-stimulated ductal secretion through a Ca2+- and PKC-dependent amplification of the adenylyl cyclase system [97] Cholangiocytes express dopamine receptors as well, and stimulation of these receptors inhibits rather than stimulates secretion induced by secretin and cAMP. This effect appears to be mediated by increases in cytosolic Ca2+ and Ca2+dependent activation of PKC [97]. Thus, distinct Ca2+ agonists may have opposing effects on ductal secretion. Although the basis for this is not known, one possibility would be that different agonists activate distinct isoforms of PKC, which may be a downstream effector for Ca2+ in the cholangiocyte. Ca2+ signaling in cholangiocytes also is involved in apoptosis. A number of pro- and antiapototic proteins exhibit their effects by modulating Ca2+ signals. A particularly important example is Mcl-1, a member of the Bcl-2 family which is the primary antiapoptotic protein in cholangiocytes [18]. Mcl-1 exerts its anti-apoptotic

2  Signaling Pathways in Biliary Epithelial Cells

Fig. 2.2  The antiapoptotic protein Mcl-1 is heavily expressed in human cholangiocarcinoma. Confocal immunofluorescence image was obtained from a surgical resection specimen. Mcl-1 labeling is in green, and labeling for the type III InsP3R is in red. Note that Mcl-1 is distributed diffusely throughout the cytosol, with some patchy areas of increased expression (image provided courtesy of Dr. Noritaka Minagawa)

activity in cholangiocytes as well as in the Mz-Cha-1 cholangiocarcinoma cell line through inhibition of mitochondrial Ca2+ signals [98]. Overexpression of this protein is thought to be important for development of cholangiocarcinoma (Fig. 2.2) [99].

Protein Kinase C DAG is formed along with InsP3 upon hydrolysis of PIP2 by PLC, and DAG acts to activate PKC. PKC isoforms have been grouped into three classes [100], which include conventional, nonconventional, and atypical isoforms. PKCa, PKCb, and PKCg are the conventional isoforms, and are activated by Cai2+ and DAG. These isoforms are involved in the regulation of gene expression, secretion, and modulation of ion channels, cell proliferation, and differentiation. For example, gastrin inhibits cholangiocyte proliferation and secretion following bile duct ligation by activation of PKCa [101]. Stimulation of a1-adrenergic receptors by phenylephrine induces membrane translocation of PKCa as well as b-II, and this has been associated with potentiation of secretin-stimulated

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ductal secretion [102]. This is thought to induce maximal bicarbonate excretion in proliferating ducts, which may provide a compensatory mechanism for the impaired secretion that occurs in ductular damage [102]. Cholangiocytes also express prolactin receptors, which stimulate growth by an autocrine mechanism involving phosphorylation of PKCb-I and dephosphorylation of PKCa [103]. Progesterone increases cholangiocyte proliferation by activation of nuclear (PR-A and PR-B) and plasma membrane (PRGMC1, PRGMC2, and mPR) progesterone receptors. The effects of this receptor have been examined in cholangiocytes from normal and bile duct ligated rats, as well as in the NRC cell line [104]. The therapeutic bile acids ursodeoxycholic acid (UDCA) and tauroursodeoxycholic acid (TUDCA) similarly inhibit cholangiocyte proliferation after bile duct ligation by increasing cytosolic Cai2+ and activating PKCa. Specifically, both UDCA and its taurine conjugate induce an immediate and sustained increase in Cai2+ [101, 105]. This is associated with redistribution of PKCa from the cytoplasm to the plasma membrane, which is required for activation of this kinase [101]. In contrast, both taurocholic acid (TCA) and taurolithocholic acid (TLCA) increase cholangiocyte proliferation after bile duct ligation. Neither of these bile acids increases Cai2+ or activates PKC in cholangiocytes. Like PKCa, PKCg appears to play an inhibitory role in cholangiocytes. For example, dopaminergic agonists inhibit secretin-stimulated ductal secretion by decreasing cAMP formation and inducing Ca2+-mediated activation of PKCg [97]. Thus, while certain Ca2+ agonists such as acetylcholine and ATP stimulate secretion in cholangiocytes, other Ca2+ agonists such as gastrin, dopamine, and certain bile acids instead are inhibitory. Evidence at present suggests that agonist-specific activation of various PKC isoforms may in part be responsible for these differential, agonist-specific effects of Ca2+.

MAPK Signaling The MAPK pathways are important for the normal and abnormal regulation of cell growth, and are often activated by stimulation of receptor tyrosine kinases. Certain inflammatory mediators can stimulate receptor tyrosine kinases. Lipopolysaccharide stimulates the release of IL-6, transforming growth factor (TGF-b), interleukin

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8 (IL-8), tumor necrosis factor (TNF-a), and plateletderived growth factor (PDGF). These factors can interact with cholangiocytes in an autocrine/paracrine fashion to regulate cholangiocyte intracellular responses [106]. Secreted IL-6 stimulates cholangiocyte IL-6 receptors in an autocrine fashion, inducing activation of MAPK as well as members of the STAT family of transcription factors [107]. IL-6 increases cholangiocyte proliferation via this mechanism, since, IL-6 activates the p44/p42 and p38 MAPK signaling pathway [107]. The p44/p42 MAPK signaling cascade can be activated by mitogenic stimulation of nonmalignant human cholangiocytes, although the p38 MAPK pathway is activated by mitogenic stimulation of malignant but not nonmalignant cholangiocytes. p38 MAPK signaling affects the growth of malignant cholangiocytes by dysregulation of the eukaryotic initiation factor, eIF-4E [108]. The eIF-4E is known to bind the cap structure of eukaryotic messenger RNAs, mediating the recruitment of ribosomes to messenger RNA, a rate-liming step for translation [108]. Thus, protein synthesis is decreased after stimulation with mitogens in cholangiocarcinoma cells with a functional impairment in p38 MAPK activation, due to impaired initiation of translation [108]. Because of the importance of translational regulation in promoting tumor growth, the translational apparatus could represent an attractive target for therapeutic intervention in the treatment or prevention of cholangiocarcinoma. Activation of p38 MAPK by IL-6 also plays a role in inhibiting apoptosis. Cholangiocarcinoma cells secrete IL-6, which upregulates expression of Mcl-1, thereby inhibiting apoptosis through STAT 3 and Akt [18, 109]. Mcl-1, furthermore, inhibits apoptotic Ca2+ signals in mitochondria, suggesting that it inhibits apoptosis in cholangiocytes through a range of complementary effects [98]. Cholangiocarcinoma growth is regulated in part by the sympathetic nervous system, although this may involve MAPK signaling as well. The a2 adrenoreceptor agonist UK-14304 inhibits growth in the Mz-ChA-1 and TFK-1 cholangiocarcinoma cell lines, which elevates cAMP, and in turn substantially inhibits Raf-1 and B-Raf-1 activity induced by EGF. This is associated with sustained inhibition of MAPK activity and decreased cholangiocyte proliferation [16]. Thus, a2adrenergic receptor stimulation inhibits cholangiocarcinoma growth through modulation of Raf-1 and B-Raf-1 activities [16].

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Cholangiocarcinoma growth may also be regulated in part by other inflammatory mediators. Activation of inducible nitric oxide (NO) synthase (iNOS) can result in the generation of NO in sufficient amounts to damage DNA. In addition, activation of iNOS promotes upregulation of COX-2 in immortalized mouse cholangiocytes, suggesting that COX-2 and COX-2-derived prostanoids play a key role in cholangiocarcinogenesis [110]. A subset of cholangiocarcinomas harbor activating mutations in the oncogenes K-ras and Braf, which potentiates activation of the ERK1/2 pathway. Disruption of the Ras/Raf/MAPK pathway through these mechanisms may play a crucial role in regulating development of cholangiocarcinoma [111]. Together, these studies demonstrate that a number of stimuli may converge to affect growth of cholangiocarcinomas via MAPK pathways. Although the fundamental role of MAPK signaling in growth of nonmalignant cells has been established in many types of cells and tissues, this topic has received little attention to date in the cholangiocyte.

PI3-Kinase Signaling Increases in cell volume are known to activate phosphotidylinositol 3-kinase (PI3-K) [112]. PI3-K is a heterodimer that phosphorylates phosphotidylinositol upon activation, producing distinct phospholipid second messengers. Studies in a number of cell types show that this kinase is involved in controlling cell proliferation, organization of the actin cytoskeleton, regulation of vesicle trafficking between intracellular organelles, and a range of secretion-related processes [113]. In hepatocytes, PI3-K is activated by certain bile acids and plays an important role in the choleretic and anti-apoptotic effects of hydrophilic bile acids in particular [113, 114]. In cholangiocyt es, activation of PI3-K is an early event in the modulation of cholangiocyte proliferation and secretion by bile acids [115]. The activation of PI3-K that occurs in proliferating cholangiocytes during cholestasis may result in part from activation of the glucagon-like peptide 1 (GLP-1) receptor [116]. TCA also increases DNA synthesis via PI3-K in cholangiocytes, since, TCA-induced DNA synthesis is abolished in cholangiocytes incubated with the PI3-K inhibitor wortmannin [112].

2  Signaling Pathways in Biliary Epithelial Cells

Experimental evidence further suggests that PI3-K plays a role in the regulation of ATP release from cholangiocytes [117]. ATP release appears necessary for cell volume regulation, so PI3-K may be a key mediator of this autocrine pathway [117].

Pathological Conditions Alterations in signaling have been implicated in two types of disorders in cholangiocytes: cholestatic secretory disorders and disorders associated with changes in cell proliferation. Cholestasis is one of the principal manifestations of liver disease and often reflects impaired ductular secretion due to cholangiocyte dysfunction [42]. In models of ductular cholestasis such as bile duct ligation, expression of the secretin receptor and cAMP formation is preserved and even enhanced, so that stimulation with secretin leads to a massive, bicarbonate-rich choleresis [38]. However, Ca2+ signaling pathways are severely impaired in this and other cholestatic models and disorders [118]. For example, there is a marked loss of each InsP3R isoform in cholangiocytes 2 weeks after bile duct ligation. In particular, expression of the type III InsP3R, which is the predominant isoform in cholangiocytes, is nearly absent. This loss of InsP3Rs is associated with impaired Cai2+ signaling and Ca2+-mediated bicarbonate secretion as well. InsP3R expression and inhibition of ductular secretion also occur after treatment with endotoxin, indicating that loss of InsP3R expression occurs in animal models of both acute and chronic cholestasis [118]. Human liver biopsy specimens similarly show that InsP3R expression is decreased in bile duct epithelia from a range of human cholestatic disorders, including primary biliary cirrhosis, sclerosing cholangitis, bile duct obstruction, and biliary atresia [118]. This is in contrast to patients with hepatitis C infection, in which there is portal inflammation without bile duct damage, and no loss in InsP3Rs is observed in cholangiocytes [118]. Thus loss of InsP3Rs appears to be a general feature of ductular cholestasis rather than of portal inflammation. This raises the hypothesis that Ca2+-mediated bicarbonate secretion is not just an alternative pathway to secretion via cAMP/CFTR, but may in fact be important for biliary bicarbonate secretion to occur under normal conditions. Recent studies have supported this hypothesis by demonstrating that knockdown of the

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type III InsP3R in microperfused intrahepatic bile duct segments impairs bicarbonate secretion [47]. However, further work will be needed to understand how ductular damage leads to loss of InsP3Rs in cholangiocytes. Regulation of signaling pathways may provide approaches to treat cholestasis. For example, the only medical therapy of proven benefit in certain cholestatic disorders is administration of UDCA, which promotes bile flow and biliary bicarbonate excretion [119, 120]. Several observations suggest that UDCA may act in part through a novel series of signaling events in the cholangiocyte. First, this bile acid induces hepatocytes [93] as well as cholangiocytes [121] to secrete ATP into bile. Biliary ATP in turn activates apical purinergic receptors on cholangiocytes, which induces Ca2+ signals and then stimulates Ca2+-dependent ductular Cl− and HCO3− secretion [47, 84, 93]. Another novel potential therapy for cholestasis is based on the sulfonylurea glybenclamide. Glybenclamide stimulates bile flow by up to 50% and stimulates bicarbonate excretion as well in the isolated perfused rat liver. This effect appears to be mediated at the level of the cholangiocyte rather than the hepatocyte. Moreover, glybenclamide stimulates ductular secretion by activation of Na+-K+-2Cl− cotransport, rather than via mechanisms involving cAMP or Cai2+ [122]. The cAMP signaling pathway is important for the regulation of cholangiocyte proliferation. Cholang­ iocytes undergo proliferation in response to events such as bile duct ligation and partial hepatoctomy [51, 123, 124]. Under such conditions, proliferation of cholangiocytes is associated with the increases in expression of the secretin receptor, secretin-stimulated cAMP levels, and ductal secretion [38, 51, 125, 126]. Moreover, upregulation of cAMP-related pathways by chronic administration of forskolin is sufficient to induce cholangiocyte hyperplasia similar to what is observed following bile duct ligation [127]. Other signaling pathways regulate bile duct cell growth as well. For example, chronic feeding of certain bile acids, such as TCA and TLCA, also induces proliferation of cholangiocytes [125]. Such bile acids enter cholangiocytes through the apical Na+dependent bile acid transporter (ASBT), and then alter cell proliferation by activating the PI3-K pathway in a cAMP-independent mechanism [115]. Interestingly, the bile acids UDCA and TUDCA decrease bile duct proliferation following bile duct ligation [115, 127]. Adminis­ tration of either of these bile acids is associated with decreased secretin receptor gene expression as well as decreased secretin-induced cAMP synthesis [115, 127].

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The mechanism of inhibition of these bile acids requires Ca2+-dependent activation of PKCa [115]. Biliary tract inflammation predisposes to the development of cholangiocarcinoma, and the signaling pathways involved in this have been investigated. Inflammation leads to activation of the p38 MAPK stress signaling pathway, which facilitates cell proliferation by translational regulation of protein synthesis. Activation of p38 MAPK signaling may thereby contribute to tumor growth. It has thus been suggested that gene silencing of cellular eIF-4E may be a useful strategy to limit tumor cell growth [108]. Based on such observations, gene therapy may become an approach to treat biliary tract malignancies, since cholangiocytes are accessible by percutaneous or endoscopic interventions, and the feasibility of introducing genes into cholangiocytes via retrograde biliary infusion has been shown in animal models [128].

Summary

›› Cholangiocytes are polarized and mitotically dormant.

›› Cholangiocytes express numerous G protein›› ›› ›› ›› ››

coupled receptors and non-G protein-coupled receptors. Calcium and cAMP are the principal second messengers of the cholangiocytes. Cholangiocytes are responsive to basolateral as well as apical signals. ATP is an important extracellular signaling molecule for cholangiocytes. Secretin is a major agonist for cAMP-mediated Cl− secretion. Cholestasis affects in particular calcium signaling.

2. Which statement is not correct. In cholangiocytes, (a) Endocannabinoids impair cholangiocyte prolif­ eration (b) CXC Chemokine ligand 12 induces ERK and Akt pathway (c) IL-6 induces ERK and Akt pathway (d) Estrogens induce the expression of VEGF (e) EGF inhibits the Ras/Raf/MAPK cascade 3. In cholangiocytes cAMP (a) Stimulates exocytosis (b) Close aquaporin water channels (c) Inhibits CL−/HCO−3 exchange channel (d) Blocks the effect of secretin (e) Is increased by somatostatin via the SSTR2 3. Regarding the effects of bile acids in cholangiocytes: (a) They activate EGFR through a TGF-a dependent mechanism (b) They Regulate the expression of death receptor 5/TRAIL-receptor 2 (c) They modulate the IGF1 system (d) They activate CFTR (e) All the above are correct 5. Which statement is correct: (a) Cholangiocytes have basolateral cilia (b) Cilia are sensory organelles (c) P2Y12 receptors link primary cilia to calcium signaling (d) TRPV4 are Cl− channels associated with cilia (e) Cilia are responsive only to mechanical stimuli Acknowledgments  This work was supported by NIH grants TW01451, DK61747, DK45710, DK34989, and DK57751 and by a grant from the Howard Hughes Medical Institute.

References

Multiple Choice Questions 1. Cholangiocytes express: (a) Secretin receptor (b) Histamin 2 receptor (c) Serotonin receptor (d) M3 muscarinic acethylcholine receptor (e) All the above

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3

Stellate Cells Fabio Marra, Sara Galastri, Sara Aleffi, and Massimo Pinzani

Hepatic stellate cells (HSC) are located in the space of Disse in close contact with hepatocytes and sinusoidal endothelial cells. In human liver, HSC are disposed along the sinusoids with a nucleus-to-nucleus distance of 40 µm, indicating that the sinusoids are equipped with HSC at certain fixed distances [1]. These observations suggest that, although the total number of HSC constitutes a small percentage of the total number of liver cells (approximately 5–8%), their spatial disposition and extension may be sufficient to cover the entire hepatic sinusoidal microcirculatory network. This cell type has received much attention in the past two decades, the reason being its potential involvement in the fibrogenic transformation of liver tissue following chronic injury. Because of their anatomical location, ultrastructural features, and similarities with pericytes regulating blood flow in other organs, HSC have been proposed to function as liver-specific pericytes. Branches of the autonomic nerve fibers coursing through the space of Disse come in contact with HSC [2], and the nerve endings containing substance P and vasoactive intestinal peptide have been demonstrated in the vicinity of HSC [3]. Other peculiar features that suggest a functional relationship between the autonomic nervous system and HSC are the expression of N-CAM, a typical central nervous system adhesion molecule detected in hepatic nerves, and the expression of glial fibrillary acidic protein (GFAP) are restricted, among liver cell types, to HSC [4]. These observations, while reinforcing a potential functional relationship between the autonomic nervous system and HSC, raise a current key issue concerning the origin of

F. Marra () Dipartimento di Medicina Interna, Università di Firenze, Viale Morgagni, 85 50134 Firenze, Italy e-mail: [email protected]

this cell type, previously considered to be of myogenic origin owing to the expression of desmin and smooth muscle a-actin (a-SMA). Along these lines, activated HSC express nestin, a class VI intermediate filament protein originally identified as a marker for neural stem cells [5]. Remark­ably, the expression of this cell marker appears to be restricted to HSC and pericytes of brain parenchyma vessels, among all organ-specific pericytes. Adding to the intriguing features of these cells, ­quiescent stellate cells also express epimorphin, a mesenchymal morphogenic protein, which is increased after partial hepatectomy, coincident with a decline in stellate cell expression of a-SMA [6]. The most evident ultrastructural feature of HSC in normal adult liver is the presence of cytoplasmic lipid droplets ranging in diameter 1–2 µm (i.e., “fat-storing cells” or “lipocytes”) [wake]. These lipid droplets are involved in the hepatic storage of retinyl esters owing to the key role of HSC in the metabolism and storage of retinoids [7]. Among other cell types potentially involved in the abnormal progressive deposition of fibrillar extracellular matrix (ECM), HSC have received much attention, also because of the possibility of isolating them from liver tissue with a relatively high purity. Consequently, most of the present knowledge on the cell and molecular biology of hepatic fibrosis derives from in vitro studies employing culture-activated HSC isolated from rat, mouse, or human liver. Regardless of this, it is now evident that distinct ECM-producing cells, each with a distinct localization and a characteristic immunohistochemical and/or electron microscopic phenotype, are likely to contribute to liver fibrosis [8]. These include fibroblasts and myofibroblasts of the portal tract, smooth muscle cells localized in vessel walls, and myofibroblasts localized around the centrolobular vein. It is also evident that the relative participation of these

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_3, © Springer-Verlag Berlin Heidelberg 2010

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different cell types is dependent on the development of distinct patterns of fibrosis. It is likely that all these different­ ECM-producing cell types undergo a process of activation in the conditions of chronic liver damage or, anyhow, in which the physiological homeostasis of the tissue is chronically perturbed. For the reasons previously­ mentioned, the process of HSC activation has been the object of several studies and consistent information is at present available. Following prolonged culture on plastic, HSC undergo a process of activation from the quiescent “storing” phenotype to the highly proliferative “myofibroblast-like” phenotype (Fig. 3.1). In addition to using uncoated plastic as a culture substratum, stellate cells grow well on a variety of extracellular matrices which can up- or downregulate their activation [9]. Even more striking is the preservation of a quiescent phenotype when stellate cells are maintained on a laminin-rich gel that mimics the effects of a basement membrane [10–12]. This quiescent phenotype can also be maintained if cell adherence is prevented by culture in suspension on a nonadherent surface [13]. A fascinating explanation for the effects of

Proliferation

a gel substratum has been given by recent studies implicating matrix stiffness as the key determinant of stellate cell activation in these systems [14]. Thus, the deformability of the substrate and its chemical composition may regulate stellate cell responsiveness. While the receptors that mediate these responses have not been clearly identified, integrins are strong candidates based on their important role in mediating cell-­substratum interactions in stellate cells and other mesenchymal cell types. The fact that matrix stiffness may be a regulator of stellate cell biology has to be put in the context of recent clinical studies showing the efficacy of a device that measures stiffness to determine the amount of fibrosis in liver tissue [15]. In vitro activation is still regarded as similar to that occurring in liver tissue following chronic damage, although this assumption likely represents an oversimplification. The activated phenotype is characterized by a dramatically increased synthesis of collagen types I and III, that appears predominant over the synthesis of collagen type IV (I > III » IV) and other ECM components. Studies performed in recent years have

Migration Fibrillar ECM

Activation Contraction Cross - talk with immune system Cross - talk with cancer cells

Chemoattraction Survival/Apoptosis

Cross - talk with biliary cells

Fig.  3.1  Activation and phenotypical modulation of hepatic stellate cells. Activation of hepatic stellate cells and transition to the so-called “myofibroblast-like” phenotype is associated with remarkable changes in their biology. Activated HSC become highly proliferative, motile and contractile. In addition they are responsible for the deposition of increasing amounts of fibrillar

extracellular matrix associated with a reduced capability towards its degradation and remodeling. Activated HSC are also responsible for the synthesis and secretion of several proinflammatory mediators, including chemokines, thus leading to further amplification of the inflammatory process

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emphasized some important aspects potentially related to the initiation of HSC activation. A first important element concerns the disruption of the normal ECM pattern that follows liver tissue injury and acute inflammation. A perturbation in the composition of the normal hepatic ECM and/or the cell–cell relationship between epithelial and mesenchymal cells present in liver tissue, typical of some cholestatic disorders (i.e., those characterized by bile duct proliferation and lobular invasion), could also be considered a potent stimulus for the activation and proliferation of HSC, as well as other ECM-producing cells. Indeed, loss of adhesion with the various elements constituting the basal membrane-like ECM of the space of Disse is likely to determine a marked increase of the proliferative and synthetic properties of HSC. This issue is becoming more and more important with the demonstration that the movement, shape, and proliferation of cells are greatly influenced by the cooperation of ECM components and cell adhesion molecules. Another study demonstrated that stellate cell activation in liver fibrosis is associated with a switch from E- to N-cadherin expression [16], raising the interesting prospect that stellate cells undergo epithelial to mesenchymal transition [17]. In addition, in cholestatic liver injury, portal fibroblasts may be a more important source of activated myofibroblasts than stellate cells around proliferating bile ducts [18]. Collectively, these findings reinforce earlier histochemical data highlighting the heterogeneity of stellate cells with respect to

both classical markers of stellate cell activation and even raise the possibility of transdifferentiation from epithelium. Several soluble factors, including growth factors, cytokines, chemokines, and oxidative stress products, play a role in the activation of HSC (Table 3.1) [19, 20]. It must be emphasized that, although most studies address the role and the biochemical features of individual factors, these mediators do not work alone but rather in a complex network of interactions with their cellular targets and the ECM. Thus, the response to single agonists on cultured HSC does not completely reflect the complexity of the in vivo situation. In the past decade, there has been significant elucidation of the cascades of intracellular signals downstream of cytokine-receptor binding. In addition, extensive modifications in the composition and organization of the ECM and parallel changes in the expression and function of cell membrane molecules have been identified. The different groups of cytokines can be grouped according to their class of receptors, which tend to generate similar intracellular signals within each group: (a) factors promoting HSC proliferation, migration, and survival (polypeptide growth factor ­receptors); (b) factors promoting fibrillar ECM accumulation, and particularly TGF-b1 (TGF-b receptor superfamily); (c) factors with a prevalent contractile effect on HSC, such as endothelins, angiotensin-II (A-II), vasopressin, and thrombin, although all these agents may also promote HSC proliferation (seven transmembrane

Table 3.1  Growth factors, cytokines and other soluble factors affecting HSC biology Injured hepatocytes T-lymphocytes Mononuclear/ Sinusoidal Platelets Kupffer endothelium ROI Reactive aldehydes IGF-1 VEGF

TNF-a IFN-g

PDGF-AB bFGF TGF-b TNF-a IL-1 PGs ROI

PDGF-BB bFGF VEGF IL-1 TGF-b IGF-1 PGs NO ET-1 ROI

PDGF-AB EGF/TGF-a TGF-b TX IGF-1 VEGF

Serum

Autocrine

Thrombin A-II AVP Curcumin 15d-PGJ2 Vitamin D Estrogens Paxillin NGF

MCP-1 PDGF TGF-b VEGF ET-1 Leptin Adiponectin Resistin Follistatin

Different cellular sources are responsible for the release of factors affecting HSC activation and their profibrogenic properties once activated A-II angiotensin-II; 15d-PGJ2 15-deoxy-prostaglandin J2; vitamin D 1,25-dihydroxyvitamin D3; EGF epidermal growth factor; ET-1 endothelin-1; bFGF basic fibroblast growth factor; IFN-g interferon-g; IGF-1 insulin-like growth factor 1; IL-1 interleukin 1; MCP-1 monocyte chemotactic protein 1; NGF nerve growth factor; NO nitric oxide; PDGF platelet-derived growth factor; PGs prostaglandins; ROI reactive oxygen species; TGF-a transforming growth factor-a; TGF-b transforming growth factor-b1; TNF-a tumor necrosis factor-a; TX thromboxane A2; VEGF vascular endothelial growth factor

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domain receptors); and (d) receptors for chemokines and proinflammatory cytokines. An important determinant of the biologic response to these ligands is the interaction between these substances and the rapidly evolving ECM microenvironment, where they bind and can be stored.

Polypeptide Growth Factor Receptors Platelet-derived growth factor (PDGF), a dimer of two polypeptide chains referred to as A- and B-chain, is the most potent mitogen for cultured HSC isolated from rat, mouse, or human liver [21–23]. Of the three possible dimeric forms of PDGF (-AA, -AB, and -BB), PDGF-BB is most potent in stimulating HSC growth/ chemotaxis and the relative intracellular signaling, in agreement with a predominant expression of PDGFreceptor b (or type B) subunits compared to PDGFreceptor a (or type A) subunits in activated HSC [22]. Importantly, codistribution of PDGF with cells expressing PDGF receptor subunits has been demonstrated following both acute and chronic liver tissue damage [24, 25], thereby confirming an active role of this growth factor in liver repair and fibrosis. In addition, PDGF is profibrogenic in conditions where inflammation is less evident such as experimental cholestatic liver injury [26, 27]. In cholestasis, PDGF synthesis and release is sustained by proliferating bile duct cells. Recent work has shown that PDGF, in addition to inducing proliferation and chemoattraction of HSC toward bile ducts, is able to mediate the myofibroblastic conversion of peribiliary ECM-producing cells distinct from HSC [28]. Owing to the relevance of this polypeptide growth factor in stellate cell growth and chemotaxis, considerable effort has been invested in understanding the intracellular signaling events elicited by the interaction of PDGF with its receptor. Recently, two additional PDGF polypeptide chains were discovered, namely PDGF-C and PDGF-D. The discovery of two additional ligands for the two PDGF receptors suggests that PDGFmediated signaling is more complex than previously anticipated [29]. Interestingly, in a mouse model with transgenic expression of PDGF-C, development of fibrosis and hepatocellular carcinoma (HCC) has been observed [30]. This model, where fibrosis precedes cancer, is one of the few resembling human liver disease with HCC arising in cirrhotic livers.

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Figure 3.2 illustrates the major signaling pathways elicited by the interaction of PDGF with its membrane receptors. This response is the net result of the activation of both positive and negative intracellular signals, and, accordingly, each pathway leading to a specific effect is often provided with an intrinsic autoregulation. PDGF receptors, which have intrinsic tyrosine kinase activity, dimerize and become auto-phosphorylated on tyrosine residues upon binding to their ligand [31–33]. Association of the PDGF receptor with the adapter protein Grb2 leads to recruitment of the exchange factor mSos with the consequent activation of Ras. This event is followed by the sequential activation of Raf-1, MEK, and extracellular-signal regulated kinase (ERK) [34, 35]. Nuclear translocation of ERK is associated to the phosphorylation of several transcription factors, including Elk-1 and SAP, and represents an absolute requirement for triggering a proliferative response [36]. In cultured human HSC, there is activation of the ERK pathway followed by increased expression of c-fos in response to PDGF [37–39]. The activation of this pathway is necessary for PDGF-induced cell proliferation and accordingly, the pharmacological blockade of signaling molecules upstream of ERK (e.g., MEK) leads to a dose-dependent inhibition of cell growth. This observation is supported by the reduction in the downstream activation of the proto-oncogene c-fos and of the AP-1 complex binding activity that follows the inhibition of PDGF-induced ERK activation [39]. Inhibition of ERK is associated with a reduction of STAT-1 activation induced by PDGF, although the potential role of the cross talk occurring between ERK and STAT-1 in PDGF-induced cell growth is presently unclear. Work by Reeves and coworkers indicate that PDGF-induced generation of the lipid second messenger phosphatidic acid (PA) contributes to ERK activation in rat HSC, an action attributed to a positive interaction of PA with signaling molecules upstream of ERK [40]. ERK activation in rat HSC occurs following in vivo liver injury induced by the acute administration of CCl4 [39]. In this model, increased ERK activity temporally precedes HSC proliferation and peaks at 48 h after the administration of the toxin. Remarkably, this time point is associated with maximal availability of PDGF in acutely injured liver tissue [24], and precedes HSC proliferation, which begins at 48 h and peaks at 72 h after liver damage. Studies which employ other drugs to interfere with pathways upstream of ERK provide additional evidence for a key role of ERK in mediating ­PDGF-induced cell

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PDGF-BB Integrin Receptor

β

Ca2+

RAS PIP

c-Src

PIP

Sos

Sos

Grb2

Grb2 P

P

FAK

[Ca2+]i

P GDP

+ P

GTP

p110

p85 P

P

PLC gγ

PI 3-K

+

P

AKT Grb2

IP3

Sos

PKC

Raf MEK ERK

CYTOSKELETAL TENSION

Nuclear Signaling Fig. 3.2  Major intracellular signaling pathways induced by PDGF and cross-talk with integrin receptors. Ras plays a key role in the cross-talk between PDGF-receptor and FAK in human HSC. Integrin dimerization leads to the phosphorylation of focal adhe-

sion kinase (FAK) and to the consequent formation of a functional FAK/c-Src complex. This complex, through the interaction with the signaling molecule Shc, leads to an enhanced GTP exchange on Ras, thus facilitating the activation of downstream effectors

growth in HSC. Preincubation of HSC with drugs increasing intracellular cAMP levels such as pentoxifylline, a phosphodiesterase inhibitor, leads to a remarkable­ reduction in the PDGF-induced ERK phosphorylation and activity, c-fos expression, and mitogenesis [41]. Other examples of upstream ERK antagonists are compounds that increase prostaglandin (PG) E2 synthesis, which act via an increase in intracellular cAMP [42–44]. Interestingly, stimulation of HSC with PDGF leads to increased synthesis and release of PGE2. This causes in turn an autocrine increase in intracellular cAMP leading to a selflimitation of PDGF mitogenic potential [42, 43]. Increased levels of intracellular cAMP may inhibit PDGF-induced cell growth with two main mechanisms: A. inhibition of Raf kinase, an upstream activator of ERK, occurring through phosphorylation of Raf-1 by cAMP-activated protein-kinase A (PKA) [45], and B. inhibition of STAT1 activation [46].

Phosphatidylinositol 3-kinase (PI 3-K), another molecule that is recruited by the activated PDGF receptor, is comprised of a 85  kDa regulatory subunit, equipped with two SH-2 domains, and a catalytic 110  kDa subunit [34]. PDGF stimulation leads to the association of PI 3-K with the activated receptor, and to tyrosine phosphorylation of p85 but not of p110. The downstream effectors of PI 3-K activation are only partially known and include protein kinase Cz, ribosomal S6 kinase, and protein kinase B (c-Akt). Nevertheless, this pathway is sufficient to transduce PDGF-dependent mitogenic signals [47] and to be necessary for cell chemotaxis [48]. Thus, in human HSC cultures, PI 3-K activation is necessary for both mitogenesis and chemo­ taxis induced by PDGF [49]. The in vivo relevance of this finding is suggested by the recruitment of the p85 subunits by the PDGF-receptor and activation of PI 3-K following acute CCl4-induced liver damage in the rat.

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Wortmannin, a fungal metabolite that binds and noncompetitively inhibits PI 3-K, induces a dose-dependent inhibition of PDGF-BB-induced PI 3-K activation in HSC with a maximal effect at 100 nM. This concentration, which does not affect either PDGF receptor autophosphorylation or the physical association between the PI 3-K p85 subunit and the receptor, virtually abolishes PDGF-induced mitogenesis and chemotaxis in HSC, indicating a functional involvement of this pathway. Similar observations have been made with other PI 3-K inhibitors such as LY294002 [50]. In addition, PI 3-K is involved in the activation of the Ras-ERK pathway in human HSC, although it is not strictly necessary, since both wortmannin and LY294002 inhibit ERK activation only by 40–50% [49, 50]. Therefore, in HSC, PI 3-K regulates PDGF-related mitogenesis and cell migration by pathways that are at least in part independent of ERK activation. In addition to the involvement in cell growth and migration, growth factor-induced PI 3-K activation may contribute to the downstream signaling that regulates cell survival. Current evidence suggests that the “survival” or antiapoptotic action of PI 3-K is mediated by the activation of c-Akt (also referred to as protein kinase B – PKB), a signaling protein whose activity is regulated by several upstream events, and particularly the generation of phoshoinositides by PI 3-K [51]. PDGF and insulin-like growth factor-I (IGF-I) provide examples of two contrasting paradigms of action. In human HSC, PDGF induces a tenfold increase in DNA synthesis and cell migration, whereas the effect of IGF-I is in general 1/5 of that of PDGF. Regardless, PDGF and IGF-I are equipotent in the activation of the Ras/ERK and the PI 3-k pathways, at least at early time points (10–15 min) after stimulation [50]. In addition, IGF-I acts as a survival rather than a mitogenic growth factor in this cell type. Gentilini and coworkers have shown that in human activated HSC, IGF-I can activate the c-Akt pathway and its downstream targets regulating cell survival [52]. Importantly, in these experiments, activation of the c-Akt pathway is a PI 3-K-dependent event that is reversed by PI 3-K inhibitors. In general, early signaling events (i.e., observed within 5–20 min after growth factor stimulation) may be responsible for one or more biologic effects occurring after several hours. However, it is increasingly evident that the biological effect of a growth factor may be dependent on the activation of one or more intracellular signals occurring with a cyclic and/or reiterated pattern between the interaction of the growth factor and its

F. Marra et al.

receptor (early signaling) and the completion of the biological effect (intermediate and late signaling). This modality has been extensively elucidated for the relationship between Ras/ERK activation and the progression of the cell cycle induced by PDGF [53] and likely applies to other pathways such as PI 3-K [54]. In particular, this holds true for human HSC, where PDGF triggers a biphasic activation of ERK, with a late peak at 15–24 h after the addition of the stimulus [55]. In addition to specific intracellular signaling pathways that involve protein phosphorylation, PDGFsignaling relies also on changes in [Ca2+]i and pH. In particular, in HSC and other cells sustained changes in [Ca2+]i and intracellular pH are necessary for the correct articulation of pathways involving protein phosphorylation. The mitogenic potential of different PDGF dimeric forms is proportional to their effects on [Ca2+]i in activated rat and human HSC [22, 56]. The increase in [Ca2+]i induced by PDGF in HSC is characterized by two main components: (1) a consistent and transient increase (peak increase), due to calcium release from intracellular stores following the activation of PLCg and the consequent PIP2 hydrolysis, and (2) a lower but longer lasting increase (plateau phase) due to an influx from the external medium. Induction of replicative competence by PDGF is dependent on the maintenance of sustained increase in [Ca2+]i due to calcium entry rather than from the release from intracellular stores [57, 58]. Extracellular calcium entry induced by PDGF was originally ascribed to the opening of low threshold voltage-gated calcium channels consistent with “T” type designation [59]. Subsequently, this channel has been better characterized and defined as a PDGFreceptor-operated nonselective cation channel controlled by the tyrosine kinase activity of the PDGF-R and, particularly, by the activation of Ras through Grb2-Sos [60]. Another protein that is directly affected by PDGF and calcium is the myristoylated alanine rich protein kinase C substrate (MARCKS). This major protein kinase C substrate protein regulated by calcium/calmodulin affects cell motility through its direct binding with the PDGF-B receptor. Upon PDGF stimulation, MARCKS becomes phosphorylated and detaches from the plasma membrane. As a consequence, MARCKS loses its filament actin-binding capacity and therefore favors cell migration [61]. Stimulation with PDGF increases the activity of the Na+/H+ exchanger in rat or human HSC with consequent sustained changes in intracellular pH [62–64].

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This increased activity appears to occur through calciumcalmodulin and protein kinase C dependent pathways. Inhibition of the activity of the Na+/H+ exchanger by pretreatment with amiloride inhibits PDGF-induced mitogenesis, thus indicating that changes in intracellular pH induced by this growth factor are essential for its full biologic activity [65]. In addition, PDGFinduced Na+/H+ exchanger activity is linked to the activation of PI 3-K, and is blocked by preincubation with PI 3-K inhibitors. Furthermore, inhibition of the Na+/H+ exchanger leads to the interruption of downstream signaling events essential for growth factor-mediated cytoskeletal reorganization such as PDGF-induced focal adhesion kinase (FAK) phosphorylation [66]. Several observations indicate that a complex interplay may occur among a PDGF signaling and products of oxidative stress in conditions characterized by chronic inflammation typical of liver disease. In this context, the relationship stirring up the activation of PDGF receptors and the action of reactive aldehydes (HAKs) appears of particular interest. HAKs, and particularly 4-hydroxy-2,3-nonenal (HNE), exert direct profibrogenic effects via an upregulation of the procollagen type I gene. This effect is mediated through a peculiar signal pathway based on activation and nuclear translocation of c-Jun NH2-terminal kinases (JNKs), upregulation of c-jun and increased AP-1 binding [67]. HNE, as well as other HAKs at the same low concentrations induce procollagen type I synthesis, abolish PDGF-BB mitogenic signaling in human HSC [68]. This occurs through specific inhibition of the intrinsic tyrosine kinase activity associated with the PDGF-b receptor subunit, while autophosphorylation of other receptors, such as PDGF-a receptor or EGF receptor is not affected. Extracellular signal-regulated kinase 5 (ERK5) [also known as mitogen-activated protein kinase 7 (MAPK7) or big mitogen-activated protein kinase 1 (BMK1)], an additional member of the MAPK family, has been recently found to be targeted by the PDGF receptor in HSC. EGF and PDGF-DD also activate ERK5. After ERK5 silencing by siRNA, PDGF-BBinduced cell proliferation and expression and activation of c-Jun were inhibited. In contrast, depletion of ERK5 was associated with significantly increased cell migration, both in the presence or absence of PDGFBB. This effect was associated with a redistribution of focal contacts, and with decreased phosphorylation of FAK, paxillin, and PAK [69]. Established “angiogenic” growth factors such as basic fibroblast growth factor (bFGF or FGF-2) and

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vascular endothelial growth factor (VEGF) exert important biological effects on HSC. Both bFGF and VEGF play a central role in angiogenesis in health and disease and in all conditions characterized by chronic wound healing. When compared to other mitogenic and chemoattractant growth factors for HSC, bFGF is second only to PDGF-BB in potency [21, 70]. In addition, the potential in vivo relevance of bFGF in hepatic fibrogenesis has been clearly delineated in animal models [71, 72]. No studies have examined the expression of bFGF receptors and the relative intracellular signaling in HSC. However, it is conceivable that activation of HSC is associated with an increased expression of receptors for this growth factor and studies addressing this specific issue are awaited, especially in view of recent advances in bFGF signaling [73]. The VEGF family of ligands and receptors has been the focus of attention in vascular biology for more than a decade [27]. Activation of HSC is associated with an increased expression of VEGF and VEGF receptors, and VEGF induces cell growth in this cell type [29, 74]. However, in later stages of activation, expression of VEGFR1 progressively increases, whereas VEGFR2 expression decreases. At this later stage of activation, stimulation with VEGF induces an attenuation of the contractile properties of HSC and of a-SMA expression [75]. In agreement with the available background information on VEGF intracellular signaling, stimulation of activated HSC with this growth factor induces activation of the ERK and PI 3-K pathways [76]. Importantly, several stimuli potentially relevant during chronic liver injury, including hypoxia, nitric oxide, and oxidative stress-related conditions, can upregulate the VEGFR1 [77, 78].

TGF-b Receptor Superfamily Expression of TGF-b is markedly increased in animal models of liver fibrosis and in patients with chronic liver disease [79, 80]. Overexpression of TGF-b is associated with increased deposition of matrix in the target tissue, and neutralization of the biological activity of this cytokine ameliorates experimental liver fibrosis [81, 82]. These data clearly establish a role for TGF-b in mediating the development of fibrosis during chronic liver injury. TGF-b is the most potent stimulus for production of fibrillar and nonfibrillar matrix by HSC, and it also induces qualitative changes in the

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matrix by differentially stimulating its components [83]. In addition, TGF-b has also effects on matrix degradation, that are characterized by mixed actions on matrix metalloproteinases, and by inhibition of tissue inhibitors of metalloproteinases (TIMPs) and plasminogen activator inhibitor (PAI). Quiescent HSC are poorly responsive to TGF-b when maintained in suspension, a condition that prevents transition to an activated state [13]. Accordingly, affi­ nity labeling studies have demonstrated little amounts of TbRIII in quiescent cells, while all three TGF-b receptors are present after activation. Nevertheless, in nonactivated cells, TbRII expression could be demonstrated by western blotting, and mRNA transcripts for TbRII and TbRIII (betaglycan) were actually greater than after activation on uncoated plastic [13]. Decreased mRNA levels for TbRII in activated HSC have been confirmed by Roulot et al. [84], suggesting that the ratio between type II and type I receptors may be critical to differentially mediate the biological effects of TGF-b, including matrix synthesis. The biological significance of reduced expression of TbRIII in activated vs. quiescent HSC remains to be established. Similar to other TGF-b-responsive cells, HSC show phosphorylation of Smad2 and Smad3 upon exposure to this cytokine [85]. Compared to early cultured HSC, fully activated myofibroblast-like cells show reduced Smad activation, and this finding is associated with lower efficacy of the cytokine in inducing biological actions in fully activated cells [85]. However, these changes are likely to be limited to fully activated, myo-fibroblast-like cells, because binding of TGF-b to its receptors, as evaluated by affinity binding, is not reduced in HSC isolated after 48 or 72  h after acute CCl4 intoxication in the rat [86]. Interaction between activated Smad proteins and other factors may be relevant for the cell specificity of TGF-b actions. Sp1 binding to the promoter of the a2(I) collagen gene mediates the increased expression by TGF-b in HSC [87], while in hepatocytes Sp3 binds the same element, but with little transactivating activity. In HSC, but not in hepatocytes, activated Smad3 physically interacts with Sp1, thus providing a link between TGF-b signaling and matrix upregulation in specific cell types [88]. Moreover, constitutive phosphorylation and nuclear localization of Smad3 was found in a clone of HSC exhibiting high levels of collagen and PAI-1 expression together with poor response to TGF-b, confirming the relevance of this pathway for the upregulation of ECM production in HSC [89].

F. Marra et al.

Recent findings suggest that altered Smad signaling may underlie the response of stellate cells to matrix stiffness [14]. Smad7 is an endogenous antagonist of Smad-2/3-mediated stellate cell activation [90, 91]. Antagonism of TGF-b signaling is an important and promising approach to antifibrotic therapy, either through administration of N-acetyl cysteine [92], Smad7 [90], soluble TGF-b receptors [81, 93], interferon-g (IFN-g) [94], bone morphogenic protein-7 [95], or a variety of other approaches [96]. Besides Smads, other signaling pathways are activated by TGF-b in HSC, with potential involvement in ECM synthesis and other activities. TGF-b1 induces activation of the ERK pathway through sequential activation of Ras, Raf-1, and MEK [97]. Interestingly, in the absence of TGF-b, the members of this pathway have divergent actions on collagen gene expression. While dominant negative ERK, which inhibits ERK signaling, reduced procollagen a1(I) transcription, dominant negative raf increased collagen reporter gene expression. These results indicate that activation of ras exerts negative signals on collagen transcription via a Raf-dependent, ERK-independent pathway, while ERK positively modulates expression of the collagen gene [98]. Other Smad-independent signaling pathways contribute to the biological effects of TGF-b in HSC, in particular the exposure of the cells to reactive oxygen intermediates. TGF-b-induction of procollagen a1(I) mRNA is mimicked by the addition of H2O2, and is prevented by the addition of the antioxidant PDTC, or of catalase to cultured HSC. These effects are mediated, at least in part, by activation of the C/EBPb transcription factor, which binds to a cis-acting element of the promoter [99]. It has been proposed that glutathione levels in HSC enable the cells to discriminate between exogenously produced H2O2, and therefore oxidative stress, and the H2O2 generated as a signaling molecule such as in response to TGF-b [100]. In particular, exposure to TGF-b would be associated with high intracellular levels of H2O2, which could be efficiently removed by catalase, yielding high levels of glutathione and allowing autocrine secretion of TGF-b. On the contrary, exogenously added H2O2, simulating a condition of oxidative stress, generates low intracellular levels of H2O2 which is catabolized by glutathione peroxidase, resulting in low glutathione levels and interruption of a TGF-b autocrine pathway­[100].

3  Stellate Cells

HSC also produce activin and respond to recombinant activin. Activin A increased mRNA levels for type I collagen, synergistically with TGF-b, thus establishing that activin receptors are functional [101]. Activin A also stimulates MCP-1 expression in human HSC [102]. A natural inhibitor of activin is follistatin, whose expression decreases during stellate cell activation, leading to more unopposed activin activity [103]. Nevertheless, the dynamics of activin receptor expression during HSC activation and the specific role of activin receptor-dependent signaling pathways have not yet been evaluated. Connective tissue growth factor (CTGF) (also known as CCN2) is a growth factor modulator protein that promotes fibrogenesis in skin, lung, and kidney [104–112]. CTGF is strongly expressed during hepatic fibrosis, and stellate cells appear to be a source of this cytokine in the liver [113, 114], although hepatocytes may be even more important [115]. A recent report indicates that its expression in stellate cells is TGF-b and Smad 2/3 independent, in contrast to hepatocytes where CTGF regulation is TGF-b dependent. These findings are particularly interesting in emphasizing that regulation of the same cytokine may be completely divergent between two resident liver cell populations, pointing to the potential of cell- and pathway-specific inhibition of a factor that is widely expressed [115].

Seven Transmembrane Domain Receptors Several factors acting through receptors belonging to the “seven transmembrane domain” family are active in HSC. Due to similarities in biological actions and signaling, these receptors may be divided, for the purpose of this review, into two subgroups, namely receptors for “vasoconstrictors” and chemokine receptors. Endothelins, A-II, vasopressin, and thrombin, although generally referred to as “vasoconstrictors,” promote profibrogenic actions and are considered pleiotropic cytokines when viewed in the context of the chronic wound healing process. Endothelin-1, a potent vasoactive 21-amino-acid peptide secreted by endothelial as  well as other cell types, has been shown to exert a multifunctional role in a variety of tissues and cells [116–118], including the liver. Infusion of ET-1 in the isolated perfused rat liver causes a sustained and dose-

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dependent increase in portal pressure associated with increased glycogenolysis and oxygen consumption [119–122]. ET-1 stimulates glycogenolysis, phosphoinositide turnover and repetitive, sustained ­intracellular calcium transients in isolated rat hepatocytes [99, 123]. Other studies indicate that ET-1 may also have important interactions with liver nonparenchymal cells. Cultured sinusoidal endothelial cells isolated from rat liver have been shown to release ET-1 [124], and preferential binding sites for ET-1 have been identified, both in vivo and in vitro [125, 126], on HSC. ET-1 induces a dose-dependent increase in intracellular free calcium, coupled with cell contraction in this cell type. Importantly activated rat and human HSC have been shown to express preproET-1 mRNA [23, 127] and to release ET-1 in cell supernatants in response to agonists such as A-II, PDGF, TGF-b, and ET-1 itself [23], thus raising the possibility of a paracrine and autocrine action of ET-1 [128]. ET-1 synthesis in HSC is regulated through modulation of endothelin converting enzyme-1 (ECE-1), the enzyme that converts precursor ET-1 to the mature peptide, rather than by modulation of the precursor preproET-1 [129]. Recent evidence suggests that upregulation of 56- and 62-kDa ECE-1 3¢-untranslated region (UTR) mRNA binding proteins occurs in HSC after liver injury and during activation in vitro [130]. In addition, TGF-b1 stimulates ET-1 production by inducing ECE-1 mRNA stabilization. Overall, it is increasingly evident that the process of HSC activation and phenotypical modulation is characterized by a close and complex relationship with the ET system. The ability to synthesize and release ET-1 is associated with a progressive shift in the relative predominance of ETA and ETB receptors observed during serial subculture: ETA are predominant in the early phases of activation, whereas ETB become increasingly more abundant in “myofibroblast-like” cells [23]. The upregulation of the ETB receptor is prevented by the incubation of HSC with retinoic acid during the activation process [131], thus confirming that increased expression of this receptor is part of the phenotypical modulation of HSC towards the “myofibroblast-like” phenotype [132]. The shift in the relative ET receptor densities may be directed at differentiating the possible paracrine and autocrine effects of ET-1 on HSC during the activation process. Indeed, when HSC are provided with a majority of ETA receptors (early phases of activation), stimulation with ET-1 causes a dose-dependent increase in cell growth, ERK activity and expression of

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c-fos. These effects, likely related to the activation of the Ras-ERK pathway, are completely blocked by pretreatment with BQ-123, a specific ETA receptor antagonist [23], and are in agreement with studies performed in other vascular pericytes such as glomerular mesangial cells [133]. Conversely, in the later stages of activation, when the number of ETB receptors increase, ET-1 appears to induce a prevalent antiproliferative effect linked to the activation of this receptor subtype [134]. In this setting the activation of the ETB receptor stimulates the production of prostaglandins, leading to an increase in intracellular cAMP, which in turn reduces the activation of both ERK and JNK. In addition, both cAMP and prostaglandins upregulate ETB binding sites, thus suggesting the possibility of a positive feedback regulatory loop. In addition, recent studies have further defined the action of cAMP on the ET-1 receptor system. Cyclic AMP rapidly desensitizes ETA in activated HSC and shifts their ET-1 responsiveness from picomolar to nanomolar concentrations with respect to Ca(2+) signals and HSC contraction. ETA desensitization also occurs in response to prostaglandin E2, adenosine, or ETB stimulation [135]. Similarly to ET-1, A-II, vasopressin, and thrombin, although generally referred to as “vasoconstrictors,” promote profibrogenic actions and are considered pleiotropic cytokines when viewed in the context of the chronic wound healing process. Circulating levels of A-II, a powerful vasoconstrictor, are frequently increased in cirrhotic patients, and have been implicated in the circulatory disturbances typical of this clinical condition. However, A-II, in addition to its action as vasoconstrictor, is provided with biological properties potentially relevant for the progression of chronic fibrogenic disorders. These include increase in cell proliferation and cell hypertrophy, and accordingly A-II may be considered also as a pleiotropic cytokine. In a recent study, Bataller et al. [136] have reported that activated human HSC express A-II receptors of the AT1 subtype, and that an increased expression of this type of receptors may represent a feature of HSC activation. Stimulation with A-II elicits a marked dose-dependent increase in [Ca2+]i concentration associated with rapid cell contraction, in agreement with a previous report [137]. Moreover, A-II stimulates DNA synthesis and cell growth. The involvement of AT1 receptors in these effects of A-II is confirmed by their complete abrogation following preincubation with the AT1 receptor antagonist losartan. Moreover, infusion

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of A-II induces stellate cell activation and inflammation in rats [138]. The effects of angiotensin are mediated through NADPH oxidase, a multiprotein complex that generates reactive oxygen species [139, 140]. Within this complex, only Rac1 has been identified as a functionally active component [140], as underscored by a transgenic mouse model in which over expression of Rac1 in stellate cells amplifies injury and fibrosis [141]. Most importantly, antagonism of angiotensin signaling, either by angiotensin converting enzyme inhibitors, or by receptor antagonists, is antifibrotic in animal models [142–144]. In a retrospective human study of patients following liver transplantation, treatment with this class of drugs also appear to be beneficial [145]. In consideration of these promising data, controlled prospective trials of A-II type I receptor antagonists are underway. Analogous effects on HSC biology have been described for arginine vasopressin (AVP). Human activated­ HSC express V1 receptors, and stimulation with AVP elicits a dose-dependent increase in intracellular [Ca2+]i coupled with cell contraction. Moreover, AVP increases ERK activity, DNA synthesis, and cell growth [146]. The serine protease thrombin (THR) regulates platelet aggregation, endothelial cell activation and other important responses in vascular biology and in acute and chronic wound repair. Although THR is a protease, it acts as a traditional hormone or as a pleiotropic cytokine based on the nature of its receptors, the protease-activated receptors or PARs. PARs are G-protein-coupled receptors that use a fascinating mechanism to convert an extracellular proteolytic cleavage event into a transmembrane signal: these receptors carry their own ligands, which remain cryptic until unmasked by receptor cleavage [147]. Four PARs are known in mouse and human: human PAR1, PAR3, and PAR4 can be activated by THR, whereas PAR2 is activated by trypsin and tryptase as well as by coagulation factors VIIa and Xa, but not by THR. Studies by Marra and coworkers have shown that expression of PAR1 is markedly increased in chronic fibrogenic disorders involving the liver. In addition human HSC express PAR1, and this expression increases during HSC activation [148, 149]. Stimu­ lation of human HSC with THR induces cell contraction [137], proliferation [148], synthesis and release of chemokines such as MCP-1 [149], or platelet activating factor [150]. The signaling mechanisms specifically

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regulating THR action in HSC have not been reported thus far. In addition to ET-1, A-II, and thrombin, the potential involvement of other vasoconstrictors synthesized and released within the liver tissue has been suggested. Titos et al. [151] have recently reported that in cirrhotic rat liver there is an increased synthesis of cysteinyl leukotrienes (LTs). In this context, hepatocytes exhibit the highest ability to generate cysteinyl-LTs. Importantly, these compounds elicit a strong contractile response in activated HSC. These findings further reinforce the concept of an imbalance between vasocostrictor and vasodilator agents within the intrahepatic circulation of cirrhotic liver. Importantly, the concentration of vasocostrictors acting on the intrahepatic microvasculature of cirrhotic liver may increase as a consequence of clinical or subclinical events such as infections in the peritoneal cavity, which are clearly associated with a worsening of portal hypertension and with an increased incidence of variceal bleeding [152]. Appropriate use of drugs currently indicated for the treatment of portal hypertension [118] should be carefully reconsidered in light of the current knowledge of the cellular and molecular mechanisms of portal hypertension. The chemokine system is a major modulator of many critical functions both in physiologic and pathologic conditions, including inflammation, development, leukocyte trafficking, angiogenesis and cancer [153]. In HSC, secretion of several chemokines belonging to different subclasses regulates recruitment of inflammatory cells to the sites of damage [154]. These findings demonstrate a first line of interaction between the inflammatory and the reparative phases of the wound healing response. However, chemokines also directly regulate the behavior of cells involved in tissue repair. In fact, glomerular mesangial cells, or smooth muscle cells express chemokine receptors which mediate functional responses that are relevant for tissue fibrosis [52, 155]. In agreement with these observations, the CC chemokine monocyte chemoattractant protein-1 (MCP-1), induces concentration-dependent chemotaxis of HSCs [154]. Interestingly, HSC do not express the receptor CCR2, which mediates the biological responses of this chemokine in leukocytes. Nevertheless, incubation of stellate cells with MCP-1 is associated with a rise in intracellular calcium concentration and activation of intracellular signaling pathways, including protein tyrosine phosphorylation, and activation of PI 3-K, which are necessary to stimulate cell migration. The receptor(s) responsible for

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these effects of MCP-1 have not been yet identified, but a possible candidate could be CCR11, which can bind MCP-1 with high affinity [156]. The chemokine receptor CXCR3, which binds the ligands IP-10, Mig, and I-TAC, is the first chemokine receptor to be identified in HSC [55]. Its expression increases after a few days in culture on uncoated plastic, and receptor activation is associated with increased cell migration, establishing that CXCR3 is functional in HSC. Inter­ action of CXCR3 with its ligands leads to activation of the Ras/ERK cascade through a Src-dependent pathway and PI 3-K and its downstream kinase Akt. Interestingly, CXCR3 activation increases cell proliferation in glomerular mesangial cells but not in HSC. This discrepancy in the biological actions elicited by this receptor is accompanied by a different time course of ERK activation in the two cell types. Indeed, in HSC, ERK is activated only transiently, whereas in mesangial cells biphasic activation occurs, including a late peak at 15–24 h [55]. The molecular mechanisms underlying this different behavior in the two cell types have not yet been elucidated. Expressions of other functional chemokine receptors have been recently demonstrated in cultured HSC. CCR7, a receptor expressed by different T cell subtypes, is also present in activated HSC, which mediates cell migration and secretion of other chemokines [157]. Similarly, CCR5 have been found on the surface of HSC. Remarkably, HSC also express the CCR5 ligand, CCL5, indicating the existence of an autocrine loop involving these two molecules [158]. Exposure of HSC to recombinant CCL5 resulted in increased DNA synthesis and migration, providing additional evidence for a cross-talk between leukocytes and HSC during liver fibrosis. Collectively, these data support the view that the mechanisms which regulate leukocyte infiltration and the persistence of inflammation are also responsible for migration and proliferation of HSC to the same sites of liver injury, contributing to the pathogenesis of tissue repair and fibrogenesis.

TNF Receptor Superfamily The receptors for tumor necrosis factor (TNF) belong to a superfamily which includes several transmembrane molecules that bind cytokines or other ligands and orphan receptors, and are characterized by similar,

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cysteine-rich extracellular domains [159]. This family may be divided into two large subgroups depending on the presence or absence of a “death domain” which allows physical association with molecules containing the same domain. TNFR1, Fas, and the p75 receptor for nerve growth factor belong to the group of “death domain” receptors, while TNFR2 and CD40 lack this domain [160]. TNFR1 plays a major role in mediating the biological actions of TNF, because animals lacking this receptor, unlike those lacking TNFR2, are unable to mount inflammatory reactions in response to TNF [161]. TNF activates pathways that regulate gene transcription and inflammation and others leading to cell death. Binding of TNF induces homotrimerization of TNFR-1, which binds to the death domain containing protein TRADD [160]. Association of TRADD with RIP and TRAF2 is responsible for the downstream activation of NF-kB and JNK, respectively, leading to the regulation of gene transcription. Activation of other members of the MAPK family, including ERK1/2 and p38MAPK is also elicited by TNF. In contrast, cell death is mediated by interaction of TRADD with FADD, which initiates a cascade of proteases, known as caspases, leading to the apoptotic effect. It is important to notice that most cells become sensitive to TNFinduced apoptosis only when protein synthesis or RNA transcription is inhibited. TNF has many important effects on HSC relevant to the pathophysiology of liver fibrosis. TNF participates in the activation process [162], but has an inhibitory effect on expression and synthesis of type I collagen [163] and on proliferation of HSC [162, 164]. Remarkably, TNF is a critical factor for the “proinflammatory” role of HSC, because it upregulates expression and secretion of several cytokines and chemokines (for a review see [154]). Unlike other cytokine receptors, quiescent HSC express mRNA transcripts for TNFR1, and TNF efficiently binds to the cell surface [165]. However, the receptor expressed by quiescent cells seems to be only partially functional, because exposure of nonactivated cells to the ligand does not result in activation of NF-kB, due to the inability to degrade the inhibitory protein IkBa in quiescent cells. Accordingly, increased expression of NF-kB-regulated proteins, such as the adhesion molecule ICAM-1 is observed only in activated cells. Interestingly, the JNK pathway may be activated by TNF both in quiescent and activated HSC, indicating that the block in NF-kB activation in quiescent cells occurs at a post-receptor

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level. Activation of NF-kB plays a pivotal role in mediating the proinflammatory effects of TNF on HSC. Interference with NF-kB activation by proteasome degradation inhibitors or an IkB super-repressor, blocks the expression of several cytokines, chemokines, and adhesion molecules, including interleukin-6 (IL-6), MCP-1, CINC, MIP-2, and ICAM-1 [154, 166]. Interestingly, novel Rel-like proteins may contribute to the NF-kB DNA binding complex observed in activated HSC and upregulated by exposure to TNF [167]. NF-kB activation is also required to induce cyclo-oxygenase 2 (COX-2), which mediates the growth-inhibitory effect of TNF-a in these cells and contributes to chemokine expression [164, 166]. NF-kB is also an important mediator of cell survival [168], and exposure of HSC to TNF together with inhibition of NF-kB activation resulted in apoptosis [169]. However, when used alone, TNF actually protects HSC from apoptosis, via reduction of Fas-ligand expression [170]. On the other hand, activation of stress-activated protein kinases, such as JNK or p38, may be involved in the phenotypic transition from quiescent to activated HSC [171], and in the expression of matrix metalloproteinases [172]. The inhibition of type I collagen expression is mediated by a complex mechanism. Preincubation of HSC with pertussis toxin abolishes the inhibitory effects of TNF on procollagen a1(I) mRNA expression, and ceramide mimicks the effects of this cytokine, indicating the involvement of a pathway requiring a G protein and sphingomyelin/ceramide [173]. Moreover, several transcription factors and regulatory elements are implicated in the inhibitory effects of TNF, including a tissue-specific regulatory region, increased binding of p20C/EBPb and C/EBPd, and reduced binding of Sp1 [174–176]. An interesting signaling cross-talk involving TNF and the peroxisome proliferator-activated receptor-g (PPAR-g) has been recently described. PPAR-g is a nuclear hormone receptor that binds antidiabetic thiazolidinediones and downregulates the activated phenotype of HSC [177–180]. In HSC exposed to TNF, the transcriptional activity of PPAR-g was reduced, via phosphorylation of Ser(82) mediated by activation of ERK and JNK [180]. These data indicate an additional mechanism by which TNF may be implicated in the acquisition of an activated phenotype by HSC. Expression of Fas (CD95) by HSC was first reported by Saile et al. [181], who showed that this receptor is upregulated during activation, in parallel with the

3  Stellate Cells

appearance of spontaneous apoptosis. Remarkably, activated HSC also express Fas-ligand, indicating that HSC possess the two components of the Fas system necessary to induce apoptosis [170]. As indicated above, Fas ligand may be downregulated by either TNF-a or TGF-b [82]. In a different study, activation of Fas by soluble ligand or cross-linking antibodies required protein synthesis or transcription inhibitors to induce apoptosis in activated HSC [182]. The expression of receptors for TRAIL, another molecule related to TNF, has been recently reported in spontaneously immortalized human HSC (LX-2) [183]. Both TRAILR1/DR4 and TRAIL-R2/DR5 expression was detected in these cells, although the expression of TRAIL-R2/ DR5 was much higher. This process was activationdependent and resulted in sensibility of HSC to TRAILmediated apoptosis. The p75 nerve growth factor receptor also possesses a death domain and binds NGF with low affinity as compared to TrkA. In activated HSC, p75 was demonstrated in vivo and in culture, and exposure to NGF caused apoptosis [184]. Very recently, mRNA for other neurotrophin receptors has been detected in rat HSC [185]. The biological significance and signaling of these receptors merits further investigation.

Nuclear Receptor Family Nuclear receptors are members of the nuclear hormone receptor super family [186]. In the presence of their cognate ligands, they translocate to the nucleus and act as transcription factors. In HSC, information is available for almost the entire nuclear receptor family, in particular, peroxisome proliferator-activated receptors (PPARs) as well as the farnesoid X receptor (FXR). Retinoid receptors have been extensively explored in stellate cells given their important role in retinoid metabolism. However, no clear, coherent model for retinoid receptors in this cell type has emerged, and some of the data remain controversial. Stellate cells express retinoic acid receptors (RAR) a, b, and g [187, 188] as well as retinoid X receptors (RXR) a and b, but not g [186]. In culture, RXR-a is the dominant recep­ tor [186], but stellate cells express all six major ­isoforms (RAR-a, -b, -g and RXR-a, -b, -g) [189] and modulate a number of target genes, including cellular retinol binding protein (CRBP) [189] and collagen I [190].

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Natural RAs and synthetic RAR- or RXR-specific ligands exert differential effects on activated HSC. 9-cis-RA and synthetic RXR agonists reduce HSC proliferation and synthesis of procollagen type I and fibronectin. Both all-trans-RA and RAR agonists reduce the synthesis of ECM proteins, but do not affect HSC proliferation, indicating that retinoids inhibit proliferation independent of their interaction with RARs. RAR specific antagonists enhance HSC proliferation and demonstrate that RARs control proliferation in a negative way [191]. Moreover, inhibition of RAR-a blocks the formation of TGFb in a RA-treated stellate cell line [192]. Finally, RAR-specific antagonists induce stellate cell mitogenesis. In contrast to the unclear results concerning retinoid receptors in stellate cells, exciting advances have been made in understanding PPARs, a family of transcription factors also belonging to the nuclear receptor superfamily [193]. The peroxisome proliferator-activated receptors (PPARs) were identified 10 years ago, and serve as receptors for a number of endogenous lipid compounds, in addition to the peroxisome proliferators that originally led to their discovery. Three receptors, a, d(or b), and g have been reported in mammals. PPAR-a is the most abundant form found in the liver, with smaller amounts of the d and g forms. PPAR-g is  predominantly expressed in adipose tissue where it  regulates lipid metabolism and adipocyte differ­ entiation. Recent investigation suggests that PPAR-g receptors may be important in the control of the activation state of stellate cells, and their repression or inactivation may predispose to hepatic fibrosis. Quiescent HSC express PPAR-g isoform, and its abundance and activity are reduced during activation in culture [194, 195]. PPAR-g functions as a heterodimer with RXR, the 9-cis-RA receptor [194]. The combination of PPAR-g and RAR agonists demonstrated an additive effect in the inhibition of thioacetamide-induced hepatic fibrosis, due to inhibition of HSC proliferation and reduction of profibrotic TGF-b1 and proinflammatory TNF-a [196]. The first approved drug that specifically activates PPAR-g, troglitazone, has been withdrawn from the market due to serious liver injury. The PPAR-g ligands 15-deoxy-prostaglandin J2 (15d-PGJ2) and troglitazone dose-dependently inhibit HSC proliferation and chemotaxis induced by PDGF and inhibit a-SMA expression during stellate cell activation [195]. By

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increasing the expression of PPAR-g in activated HSCs, rosiglitazone, another thiazolidinedione agonist of PPAR-g, decreases a-SMA expression and type I collagen synthesis, inhibits cell proliferation, and induces cell apoptosis [197]. At least one mechanism of PPARg’s antifibrotic activity involves suppression of the proximal alpha1(I) collagen promoter via inhibition of p300-facilitated binding of the transcription factor NF-1 [198]. Curcumin, which possesses antioxidant and chemopreventive properties inhibits the proliferation and activation of HSCs, induces apoptosis of activated HSCs, and enhances the activities of MMP-2 and MMP-9. The effects of curcumin are mediated through activation of the PPAR-g signal transduction pathway and are associated with PPAR-g nuclear translocation/ redistribution [199]. Furthermore, prostaglandins produced by stellate cells through the upregulation of COX-2 expression may exert autocrine effects through PPAR-g, which are blocked by COX-2 antagonism [200]. These data suggest that reduced transcriptional activity of PPAR-g augments stellate cell activation and modulates mitogen-induced proliferation in activated cells. The observation that PPAR-g ligands are antifibrotic in cultured stellate cells [201, 202] and in animal models [203–205] is an additional factors advocating for their use for the treatment of nonalcoholic steatohepatitis associated with the metabolic syndrome [206–208]. Other PPARs, although less extensively studied than PPAR-g, are active in stellate cells [209]. In particular, PPAR-d (or -b) is an important signal-transducing factor [210] contributing to the hepatic stellate cell proliferation during acute and chronic liver inflammation [211]. PPAR-d regulates the expression of some genes involved in vitamin A metabolism in HSC undergoing activation [212]. During transdifferentiation, the expression of adipocyte markers decreases, with the reciprocal induction of PPAR-d, which is known to promote fatty acid oxidation, suggesting the antiadipogenic nature of HSC activation [213]. The FXR is an endogenous sensor for bile acids and inhibits bile acid synthesis by inducing a small heterodimer partner (SHP) gene expression [214]. Its therapeutic activation may become a major advance in the management of cholestasis [215]. The FXR is expressed by HSC, which exerts antifibrotic activit­ ies through upregulation of its target molecule SHP [214, 216]. The FXR-SHP regulatory cascade promotes

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the development of a quiescent phenotype and in­­ creases apoptosis in HSC, demonstrating that FXR ligands may represent a novel therapeutic option to treat liver fibrosis. The activated pregnane X receptor has also been identified in stellate cells, where it is transcriptionally functional, and its ligands inhibit transdifferentiation and proliferation. This receptor may therefore be an additional effective target for antifibrotic therapy [217]. Estrogens, in particular, 17 b-estradiol [218] and estradiol [219–221], are antifibrotic in liver, and this action may contribute to the decreased risk of fibrosis progression in females [222–224]. Estradiol suppresses hepatic fibrosis in experimental models, attenuates HSC activation in primary culture, and inhibits the activation of activator protein 1 (AP-1) and nuclear factor-kB (NF-kB) in cultured hepatocytes undergoing oxidative stress. In HSC, the actions of estradiol are mediated through ERb but not ERa, since HSC express only the b-isoform [219]. Within the liver, hepatocytes are most likely negative for the vitamin D receptor (VDR), whereas in Kupffer cells, HSC, and endothelial cells, VDR is fully functional. Accordingly, 1,25-dihydroxyvitamin D3 is mitogenic in cultured stellate cells [225]. Glucocorticoid receptor is also expressed by stellate cells [226], but its contribution to stellate cell behavior has not been specifically explored beside the observation that glucocorticoids modulate some biological activities of HSC [227]. Liver X receptor, a nuclear receptor that is a nutritional sensor of cholesterol metabolism and a major regulator of lipid metabolism, glucose homeostasis, and inflammation [228], has also been identified in stellate cells [229], underscoring the connection between this cell type and lipid physiology.

Other Cytokine Receptors The actions of IL-1 on HSC are remarkably similar to those elicited by TNF with regard to its effects on proinflammatory molecules, and these actions are mediated by activation of NF-kB [154]. Similarly, quiescent HSC express IL-1 receptors, but the activation of NF-kB, is restricted to activated cells [165]. IL-1 has also inhibitory effects on procollagen a1(I) expression, which occurs at a posttranscriptional level [163],

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and stimulates HSC proliferation [230]. However, the net effects of this cytokine are profibrogenic, because administration of IL-1 receptor antagonist reduces matrix deposition [231]. Receptors for IFN-g are ubiquitous, although no studies have examined their expression during HSC activation. A critical step in IFN-g signaling is the activation of the latent transcription factor STAT1, which upon dimerization migrates to the nucleus where it regulates transcription [232]. IFN-g downregulates activation, matrix synthesis, and proliferation of HSC in vitro and in vivo [57, 232–235]. In addition, particularly if used in combination with other cytokines or LPS, IFN-g stimulates the expression of inducible NO synthase [57]. However, the molecular mechanisms underlying these effects have not been studied. IFN-g upregulates the expression of chemokines such as MCP-1 in HSC [154], and this effect is differentially regulated as compared to other cytokines [124]. Sur­ prisingly, when human HSC are incubated with IFN-g for prolonged periods and exposed to mitogens such as serum or PDGF, an increase in cell proliferation is observed which is dependent on synergistic activation of STAT1a [38, 236]. HSC also respond to other cytokines, including IL-10, IL-6, and oncostatin M [237–239]. IL-10 inhibits procollagen a(1) expression at the transcriptional level, and may have important antifibrogenic properties [239]. Expression of the receptor for IL-10 has been recently shown in activated HSC [240]. HSC express P2Y receptors, linking extracellular ATP to inositol trisphosphate-mediated cytosolic calcium signaling [241, 242]. Stellate cells only express the type I inositol trisphosphate receptor, which shifts into the nucleus and cell extensions upon activation [242]. As activation of P2Y receptors in activated HSC regulates procollagen-I transcription, P2Y receptors may be an attractive target to prevent or treat liver fibrosis [241]. Other nucleotide receptors have also been identified on stellate cells [243]. Adenosine is a potent endogenous regulator of inflammation and tissue repair. Adenosine A (2A) receptors are expressed on rat and human hepatic stellate cell lines and its occupancy promotes collagen production by these cells. Adenosine reversibly inhibits Ca2+ fluxes and chemotaxis of HSCs and upregulates TGF-b and collagen-I mRNA, possibly providing a “stop signal” to HSCs when they reach sites of tissue injury following migration, and stimulate transdifferentiation of HSCs by

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upregulating collagen and TGF-b production [244]. Caffeine, a nonselective adenosine receptor antagonist, and ZM241385, a more selective antagonist of the adenosine A (2A) receptor, diminish hepatic fibrosis in wild-type mice exposed to either CCl4 or thioacetamide. Epidemiologic studies indicate that coffee and caffeine consumption are associated with a reduced risk of elevated alanine aminotransferase (ALT) activity in persons at high risk for liver injury. Therefore, hepatic adenosine A(2A) receptors, playing an active role in the pathogenesis of hepatic fibrosis, suggest a novel therapeutic target in the treatment and prevention of hepatic diseases [245, 246].

Adipokine Receptors Recently, the role of leptin as a mediator of liver fibrogenesis has been demonstrated by different groups. The expression and signaling of leptin receptors has been reported in primary HSC and in LX-2 cells. Incubation of HSC with recombinant leptin stimulates mRNA and protein expression of type I procollagen, potentiates the effects of TGF-b, and upregulates expression of the tissue inhibitor of metalloproteinase (TIMP)-1, thus blocking collagen degradation [247]. HSC express the long form of the receptor (ObRb), which is responsible for activation of protein tyrosine kinases of the Jak family and activation of Stat3 and the MAPK cascade [248]. These pathways are ultimately responsible for increased TIMP-1 expression and the resulting profibrogenic effect. Exposure of HSC to leptin also results in upregulation of monocyte chemoattractant protein 1 (MCP-1) expression, and increases gene expression of the proangiogenic cytokines VEGF and angiopoietin-1. Several additional signaling pathways are activated by leptin in HSC, including extracellular-signal-regulated kinase, Akt, and NF-kB, the latter being relevant for chemokine expression. Leptin also increases the abundance of hypoxia-inducible factor 1a (HIF-1a), which ­regulates angiogenic gene expression, in an Akt and PI3K dependent fashion. Leptin’s natural counterregulator, adiponectin, is also expressed by stellate cells [249] and limits the development of liver fibrosis [250–253]. Two adiponectin receptors, Adipo R1 e Adipo R2, have been identified, and the expression of both was found in

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transdifferentiated HSC with a predominance of Adipo R1 expression [254]. Globular adiponectin, which has a higher affinity for Adipo R1, effectively modulates HSC biology, activating adenosine monophosphateactivated protein kinase (AMPK), a sensor of cellular energy status. AMPK activation negatively modulates the myofibroblastic phenotype of human HSC, exerting a downregulatory action on the fibrogenic properties of these cells [254]. Resistin, which has been identified in rodents as the mediator linking obesity and type II diabetes[255], probably exerts different actions in humans [256, 257]. Recent data demonstrate an increased expression of resistin within human liver in conditions of severe damage and a proinflammatory action of this adipokine on cultured activated HSC, via a Ca2+/NF-kBdependent pathway [258].

Cooperation Between Growth Factor Receptor and Integrin Signaling Binding of cell to ECM is mediated, at least in part, by cell surface receptors belonging to the integrin family. In addition to mediating cell adhesion, integrins make transmembrane connections to the cytoskeleton and activate many intracellular signaling pathways. Integrins are heterodimeric transmembrane proteins that consist of an a subunit and b subunit. At present, 8a and 18b subunits and 24 possible distinct integrin dimers have been identified. Combination in dimers leads to the formation of integrin receptors with defined specificity for different ECM ligands. Expression of different dimers reflects the attitude of a given cell type to interact with a specific matrix microenvironment and this interaction has profound consequences on the biology of the cell type including its responsiveness to soluble factors. In addition to their roles in adhesion to ECM ligands or counterreceptors on adjacent cells, integrins serve as transmembrane mechanical links from those extracellular contacts to the cytoskeleton inside cells. For most integrins, the linkage is to the actin-based microfila­ ment system [259]. In part related to the integrin-mediated assembly of cytoskeletal linkages, ligation of integrins also triggers a large variety of signal transduction events that serve to modulate many aspects of cell behavior including proliferation, survival/apoptosis, shape, polarity, motility, gene expression, and

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differentiation. Protein phosphorylation is one of the earliest events detected in response to integrin stimulation, and in particular tyrosine phosphorylation is a common response to integrin engagement in many cell types [66, 260]. Several protein tyrosine kinases have been implicated in integrin-related signaling events. These signal transduction pathways are complex, like those emanating from receptors for soluble factors (e.g., G protein-coupled and kinase receptors). Indeed, many integrin-stimulated pathways are very similar to those triggered by growth factor receptors and are intimately coupled with them. In fact, many cellular responses to soluble growth factors, such as EGF, PDGF, LPA, and thrombin, etc., are dependent on the cell’s being adherent to a substrate via integrins. That is the essence of anchorage dependence of cell survival and proliferation and integrins lie at the basis of these phenomena. Along these lines, studies have begun to address the multiple potential interactions of cells with the microenvironment. The major advances reflect elucidation of the fine tuning occurring upon cell adhesion and the consequent cytoskeletal organization. Key elements in these responses are: (a) the specificity of integrin receptors and their downstream signaling; (b) the cross talk between integrin and cytokine signaling; (c) the relationship between the above mechanisms and the organization and tension of the cytoskeleton. Activated HSC express several integrin b1-­associated a subunits. A particularly high expression has been demonstrated for a1b1 and a2b1 [261]. FAK plays a ­central role in integrin-mediated signal transduction [262, 263]. In addition to its activation by integrins, FAK is also activated by several growth factors. FAK phosphorylation is stimulated by mitogenic neuropeptides such as bombesin, vasopressin, and by PDGF and ET-1. PI 3-K co-immunoprecipitates with tyrosinephosphorylated FAK in response to cell adhesion. Tyrosine phosphorylated PLCg may be a transducer molecule in integrin-mediated signaling pathways. Adhesion of human HSC to ECM proteins does not result in PLCg tyrosine phosphorylation [263]. Never­ theless, adhesion of HSC induces interactions between PLCg and cellular proteins undergoing tyrosine phosphorylation, one of which has been identified as FAK, suggesting that adhesion of HSC is followed by recruitment of PLCg to phosphorylated FAK. Since PLCg also physically associates with the PDGF-receptor, there may be cross-talk between this receptor and proteins of the focal adhesion complex. Indeed, stimulation of

3  Stellate Cells

HSC with PDGF-BB leads to clustering of PDGF-b receptor subunits in areas possibly corresponding to focal adhesion complexes. Along these lines upon autophosphorylation, PDGF-receptors are codistributed with FAK, thus suggesting a potential functional crosstalk between these signaling molecules [261]. In addition, experimental evidence indicates that Ras plays a key role in the cross-talk between PDGF-receptor and FAK in human HSC [60] (Fig. 3.2). In a recent study we evaluated the influence of cell adhesion on the major intracellular signaling pathways elicited by PDGF in activated human HSC [264]. PDGF signaling was investigated in an experimental condition characterized by lack of cell adhesion for different intervals of time. Prolonged lack of cell adhesion resulted in the abrogation of both basal and PDGF-induced FAK tyrosine phosphorylation. In these con­ditions ERK/MAP kinase activity correlated with FAK phosphorylation. Stimulation with PDGF was able to stimulate Ras-GTP loading only in adherent cells. The ability of PDGF to induce PI 3-kinase activity was abrogated in cells maintained in suspension. The activation of PKB/Akt was only marginally affected by the lack of cell adhesion. We then evaluated the association of FAK with c-Src. This association was found to be cell adhesion-dependent, and it did not appear to be dependent on phosphorylayed FAK. These changes in PDGF-induced intracellular signaling were associated with a remarkable reduction of PDGF proliferative potential in nonadherent cells, although no marked differences in the apoptotic rate were observed. Hence, to summarize, these results suggest that cell adhesion differentially regulate major signaling pathways activated by PDGF in human HSC. Therefore, it appears that multiple receptor systems can synergize with integrins to regulate biological phenomena such as cell proliferation, cell motility, and the signaling proteins activated by these synergistic agents are common to different receptor pathways. However, more recent advances indicate that cell adhesion and the consequent signaling events are necessary but not sufficient to provide a complete control of cell functions, and in particular their response to growth factors and cytokines. Indeed, a modern concept, defined in its complexity with the term “cellular tensegrity architecture” [265], implies that in the presence of growth ­factor stimulation and adequate adhesion to the substra­ tum, the cell is unable to progress into the cell cycle if

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restricted in its spreading. This view is supported by several observations indicating that lack of cytoskeletal tension, obtainable only with full cell spreading, is equivalent to cytoskeletal disruption and leads to the inability to enter the cell cycle. This general key mechanism appears relevant for the regulation of the molecular signaling cascades operating in the context of the structural and mechanical complexity of the living tissues and their pathophysiological alterations, including hepatic fibrogenesis.

HSC and Toll-like Receptors (TLRs) Toll-like receptors (TLRs) recognize pathogen-associated molecular patterns and are crucially involved in the regulation of innate immune responses. The normal liver shows no activation of TLR-signaling pathways. However, under pathologic conditions, Toll-like receptors promote proinflammatory signaling such as NF-kB, c-Jun-N-terminal kinase (JNK), p38, and interferon pathways in the liver and regulate antiviral and antibacterial responses, hepatic injury, and wound healing [266]. Toll-like receptors (TLRs) are expressed by activated stellate cells [267], indicating the capacity to interact with bacterial lipopolysaccharide (LPS), which in turn induces biological actions in these cells [268]. These effects are mediated, at least in part, by activation of IKK/NF-kB and JNK, leading to the expression of chemokines and adhesion molecules [267]. Activated HSC also respond to LPS, lipoteichoic acid and N-acetyl muramyl peptide with an upregulation of ERK phosphorylation [268]. In quiescent rat HSC, LPS stimulates the synthesis of TNF-a, IL-6, and IL-1 but not of transforming growth factor TGF-b [269]. These results suggest that bacterial wall products produce an inflammatory phenotype in stellate cells, but notably do not induce matrix deposition, since fibronectin and collagen transcripts were not increased. Signaling to stellate cells via TLR4 may function to enhance an adaptive immune response against bacterial pathogens. It is also possible that ligation of TLR4 is just the initial step of a series of sig­nals that are required for differentiation of stellate cells into a fully fibrogenic phenotype. Recent studies have indicated that signaling in stellate cells in response to lipopolysaccharide and possibly endogenous ligands of TLR4 may be more

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important than in Kupffer cells in eliciting a fibrogenic response. TLR4-activated HSCs produce chemokines (CCL2, CCL3, and CCL4) and express adhesion molecules (ICAM-1 and VCAM-1) that recruit Kupffer cells to the site of injury. Simultaneously, TLR4 signaling downregulates the TGF-b decoy receptor, Bambi, to boost TGF-b signaling, leading to hepatic fibrosis [270]. These findings converge with recent evidence indicating that specific single-nucleotide polymorphisms of TLR4 contribute to the rate of fibrosis progression in HCV infection [271] thereby linking a genetic risk marker to disease pathogenesis. Although TLR2 plays no significant role in hepatic fibrosis in vivo, TLR2 is dramatically upregulated in HSCs in response to LPS or TNF-a, implying a potential role of TLR2 ligands such as HCV core and NS3 protein in HSC activation [270, 272–274]. HSCs also express TLR9. Endogenous DNA from damaged hepatocytes activates HSCs to produce collagen, but suppresses HSC chemotaxis through TLR9 [275]. Although PDGF stimulation may drive cellular proliferation in parallel with fibrogenic stimulation in some settings, TLR9 activation blocks PDGF-mediated migration while provoking fibrogenesis [275], thereby providing a stop signal that allows activated cells to accumulate at sites of injury where they can deposit more scar. The injection of TLR3 ligand poly-I:C inhibits HSC activation by IFN-g from NK cells, which attenuate hepatic fibrosis [276]. Notably, chronic ethanol consumption abolishes the antifibrotic effects of TLR3, implicating a mechanism by which alcohol accelerates liver fibrosis [277].

Cannabinoids Receptors Cannabinoid system comprises CB1 and CB2 receptors, which specifically binds D9-tetra-hydrocannab­ inol (THC), the major psychotropic constituent of Cannabis sativa, as well as the endogenous cannabinoids anandamide and 2-arachidonoylglycerol. A marked upregulation of both receptors was found in cirrhotic livers [278, 279]. Stellate cells express cannabinoid receptors, and endogenous cannabinoids can provoke stellate cell death via necrosis involving mitochondrial reactive oxygen species [280–282]. Initial studies of the role of cannabinoids in hepatic fibrosis had yielded apparently paradoxical findings, which

F. Marra et al.

have largely been reconciled now that the divergent effects on liver fibrosis of the two cannabinoid G protein-coupled receptors (CB1 and CB2) have been clarified. In experimental models of liver injury, CB1 receptor is induced primarily in HSC as they activate into myofibroblasts during liver injury. CB1 receptors have been shown to exert profibrogenic actions [278, 279]. Antagonism of this receptor in a model of injury due to CCl4 or in isolated cells leads to decreased expression of TGF-b, a potent fibrogenic cytokine. In addition, CB1 antagonism reduces cellular proliferation and increases myofibroblast apoptosis, resulting in limitation of fibrosis. Moreover, CB1 KO mice or mice treated with the selective CB1 antagonist rimonabant develop less fibrosis following liver injury, compared to WT or nontreated mice, respectively [279]. In another experimental model of liver injury, lack of CB1 receptor is associated with reduced phosphorylation levels of ERK and Akt, in line with decreased cellular growth and survival, respectively [279]. On the contrary, emerging evidence identifies a protective role for CB2 receptors. CB2 activation inhibits proliferation and triggers apoptosis of cultured human myofibroblasts [278], and CB2 KO mice show enhanced fibrosis after CCl4 intoxication, compared to their WT counterparts [278].

Hedgehog Signal Pathway (See Also Chapter on this Pathway) The Wnt and Hedgehog (Hh) signaling pathways have long been known to direct growth and patterning during embryonic development. Recent evidence also implicates these pathways in the postembryonal regulation of stem-cell number in epithelia such as those of the skin and intestine [283]. This pathway was initially described in the development of Drosophila as a segment polarity gene required for embryonic patterning and is often reactivated in tumors [283, 284]. HSC have a complex phenotype that includes both neural and myofibroblastic features. The Hedgehog (Hh) pathway has been shown to direct the fate of neural and myofibroblastic cells during embryogenesis and during tissue remodeling in adults. Hh signaling regulates the fate of HSC in adults, inducing the activation and viability of HSC [285]. In addition, rat, mouse, and human HSC express proteins

3  Stellate Cells

that regulate Hh signaling, such as Sonic hedgehog (Shh) [286], Hh-interacting protein (HIP, the Hh pathway inhibitor), Hh receptor Patched (Ptc) [287], and Smoothened (SMO) transmembrane effector protein [285, 288]. When freshly isolated from adult livers, HSC produce some Shh but exhibit relatively little Hh signaling activity, due to relatively high levels of Hip. During HSC activation to the myofibroblastic phenotype, Hip expression is downregulated, Shh production increases, and expression of Gli-2, a Hh target gene, is induced [286]. Hh signaling is also required for HSC mitogens, such as PDGF-BB, to elicit their full mitogenic activity. Neutralizing Shh with anti-Shh antibodies or inhibiting Hh signaling downstream of Ptc by treatment with the Smo antagonist, cyclopamine, virtually abolishes PDGFinduced proliferation, inhibiting Akt activation [286]. These results identify Shh as an autocrine growth factor for HSC and suggest a role for Hh signaling in the pathogenesis of liver diseases. Hh signaling also contributes to the potential pluripotency of stellate cells, which express the stem/progenitor cell marker CD133 and exhibit properties of progenitor cells [289].

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Summary

›› Although the total number of HSC constitutes

›› ›› ›› ›› ›› ›› ››

Senescence of Activated Stellate Cells The hallmark of cellular senescence is an inability to progress through the cell cycle. Initially defined by the phenotype of human fibroblasts undergoing replicative exhaustion in culture [290], senescence can be triggered in many cell types in response to diverse forms of cellular damage or stress [291, 292]. Cells in cirrhotic livers of human patients express the senescence marker, SA-bgal [293]. The immunotype of senescent cells together with their location along the fibrotic scar strongly suggests that the majority of these arise from activated HSCs. In mice lacking key senescence regulators, stellate cells continue to proliferate, leading to excessive liver fibrosis. Moreover, senescent activated stellate cells exhibit gene expression profile consistent with cell-cycle exit, disabling the p53 and/or p16 Rb pathways, downregulating secretion of ECM components, and enhancing secretion of ECM degrading enzymes and immune surveillance [294]. Therefore, the senescence of HSCs limits the accumulation of fibrotic tissue following chronic liver damage and may facilitate the resolution of fibrosis upon withdrawal of the damaging agent.

››

››

a small percentage of the total number of liver cells (approximately 5–8%), their spatial disposition and spatial extension in the space of Dissé may be sufficient to cover the entire hepatic sinusoidal microcirculatory network. The most evident ultrastructural feature of HSC in normal adult liver is the presence of cytoplasmic droplets storing retinyl esters. HSC loose these droplets and acquire myofibroblasts phenotype during a process of activation. PDGF is the most potent mitogen for HSC. TGF-b1 is the most potent stimulus for production of fibrillar and nonfibrilar matrix by HSC. HSC are responsive to contractile signals such as endothelins, A-II, vasopressin via their respective G protein-coupled receptors. Stimulation of HSC with thrombin induces contraction, proliferation, and release of MCP-1. Nuclear receptors in HSC offer several antifibrotic therapeutic avenues such as for PPAR-g agonists, FXR ligands, and estradiol. In contrast to caffeine, a nonselective adenosine receptor antagonist which diminishes hepatic fibrosis, leptin potentiates the effects of TGF-b and stimulates secretion of collagen. TLR4 signaling downregulates the TGF-b decoy receptor Bambi to boost TGF-b signaling leading to hepatic fibrosis.

Multiple Choice Questions 1. Which kinds of collagen are mainly produced by activated hepatic stellate cells? (a) Collagen I and IV (b) Collagen I and II (c) Collagen I and III (d) Collagen II and III (e) Collagen IV and V

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2. PDGF is known to regulate HSC proliferation and chemotaxis. Which of the following pathways is the major signaling cascade elicited by the interaction of PDGF with its membrane receptors? (a) Grb2–Ras–Raf 1–MEK–ERK (b) Grb2–Ras–Raf 1–MEK–JNK (c) PI3K–Akt/PKB–IKKs–NF-kB (d) PKC–DAG–PIP2–PLCg (e) PLCg–PIP2–DAG–PKC 3. TGF-b effects on matrix degradation are characterized by mixed actions on matrix metalloproteinases, including inhibition of tissue inhibitors of metalloproteinases (TIMPs). After (TIMP)1 inhibition: (a) Collagen degradation is upregulated (b) Collagen degradation is downregulated (c) Collagen degradation remains unchanged (d) HSC lose their activated phenotype (e) HSC go to apoptosis 4. The cannabinoid system comprises CB1 and CB2 receptors. CB1 receptors have been shown to exert profibrogenic actions. Antagonism of this receptor: (a) Decreases expression of TGF-b and reduces cellular proliferation (b) Reduces expression of TGF-b and myofibroblast apoptosis (c) Decreases expression of TGF-b and induces cellular proliferation (d) Induces expression of TGF-b and myofibroblast apoptosis (e) Induces cellular proliferation and myofibroblast apoptosis 5. Incubation of HSC with recombinant Leptin: (a) Reduces the effects of TGF-b and increases type I procollagen expression (b) Increases tissue inhibitor of metalloproteinase (TIMP)-1 expression and mimics Adiponectin effects (c) Reduces the effects of TGF-b and Resistin (d) Potentiates the effects of TGF-b and Adiponectin (e) Increases type I procollagen and tissue inhibitor of metalloproteinase (TIMP)-1 expression

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4

Signaling Pathways in Liver Diseases Kupffer Cells Christian J. Steib and Alexander L. Gerbes

Introduction Kupffer cells (KC) constitute 80–90% of tissue ­macrophages present in the body. These liver macrophages are named after the pathologist C. von Kupffer, who apparently first recognized this nonparenchymal cell type [1]. KC represents about 35% of the nonparenchymal liver cells in normal adult mice [2]. They reside within the lumen of the liver sinusoids, adherent to the endothelial cells that compose the blood vessel walls. KC, found in greatest number in the periportal area, constitute the first macrophage population of the body to come in contact with bacteria, bacterial endotoxins, and microbial debris derived from the gastrointestinal tract and transported to the liver via the portal vein [3]. Consequently, KC are constantly exposed to proinflammatory factors, e.g., bacterial endotoxins, known to activate macrophages. Upon activation, KC release various products including cytokines, prostanoides, nitric oxide, and reactive oxygen species [4]. These factors regulate the phenotype of the KC that produce them, as well as the phenotypes of neighboring cells, such as hepatocytes, stellate cells, and endothelial cells and other immune cells that traffic through the liver [5]. Therefore, KC are intimately involved in the liver’s response to infection, toxins, ischemia, resection, and various other stresses. This review will summarize established basic concepts of KC function as well as their role in the pathogenesis of various liver diseases. Due to the

A. L. Gerbes () Department of Medicine II, Ludwig-Maximilians-University, Klinikum Großhadern, Marchioninistraße 15, 81377 Munich, Germany e-mail: [email protected]

complexity of processes mediated by KC, this review focuses on selected aspects of the pathophysiology.

Molecular Mechanisms of Kupffer Cell Activation KC from mammalian livers can be obtained after perfusion with proteolytic enzymes (e.g., collagenase, pronase), density-gradient centrifugation, and centrifugal elutriation of the resulting liver cell suspension [4]. Isolated KC can be kept in primary culture for several days and can be used to study the mechanisms of activation. Activation of KC after exposure to various stimuli is characterized by a rapid change of the KC phenotype. Phagocytosable particles and several soluble substances are able to activate macrophages via binding to specific receptors on the plasma membrane. The most important activators of KC are the complement factors C3a and C5a [6], b-glucans from bacteria and fungi [4, 7], or lipopolysaccharides (LPS), the endotoxins of gram-negative intestinal bacteria [4, 8]. As illustrated in Fig. 4.1, LPS activate KC directly via toll-like receptor (TLR) signaling [8]. LPS trigger TLR4 signaling on KC which results in the production of TNF-a, IL-1b, IL-6, IL-12, IL-18, and IL-10 [9]. IL-12 and IL-18 interact with natural killer cells. Upon activation, they produce antimicrobial IFNg. KC also express TLR2, TLR3, and TLR9 [10]. In addition, high LPS concentrations can activate KC indirectly by triggering complement activation either in the portal or in the systemic circulation [11]. After complement activation, cleavage of C3 and C5 leads to the generation of the potent anaphylatoxins C3a and C5a and subsequent stimulation of their specific receptors C3aR and C5aR [12]. Thus, KC activation

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_4, © Springer-Verlag Berlin Heidelberg 2010

69

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C. J. Steib and A. L. Gerbes

complement activation

LPS

LBP

TLR4

LPS

MD2

CD14

C3

C3a

C5 C5a

C3aR G protein C5AR

MyD88

PLC

IRAK DAG TRAP-6

PKC

IP3

Ca2+cyt

Ca2+ER

PLA2

COX-I/II

NFkB i NOS TNFa

IL-1 IL-6

NO

NADPH oxidase H 2O 2

-

O 2-

O2

PGD2 PGE2

PGF2a TXA2

Fig. 4.1  Molecular mechanisms of Kupffer cell activation. C3 and C5 complement factor 3 and 5; C3a and C5a activated complement factor 3 and 5; CD14 CD14 receptor; COX-I/II cyclooxygenase-I/II; DAG diacylglycerol; H2O2 hydrogen peroxide; iNOS inducible nitric oxide synthase; IL-1 interleukin-1; IL-6 interleukin-6; IP3 inositol-3-phosphate; IRAK interleukin-1 receptor associated kinase; LPB LPS-binding protein; LPS

lipopolysaccharide; NFkB nuclear factor kB; NO nitric oxide; PGD2, PGE2, and PGF2a, Prostaglandin D2, Prostaglandin E2, Prostaglandin F2a; PKC protein kinase C; PLA2 phospholipase A2; PLC phospholipase C; TLR4 Toll-like receptor 4; TNFa tumor necrosis factor a; TRAP-6 tumor necrosis factor activated factor 6; TXA2 thromboxane A2

can be best described as a wide spectrum of gradually different alterations of the KC phenotype resulting from a complex interplay of various activators and signaling pathways. A dominant role in signal transduction from plasma membrane-associated C3a and C5a-receptor stimulation is attributed to G proteins which regulate phospholipase C as a major key factor of signal transduction in KC [4, 12]. The enhanced activity of this enzyme may lead to activation of protein kinase C (PKC) and to calcium mobilization from the endoplasmic reticulum and the extracellular space via opening of L-type calcium channels. Experimental data indicate that PKC is involved in the activation of NADPH oxidase while Ca2+-influx is necessary for phospholipase activation and eicosanoid synthesis [13]. The resulting formation of superoxide anions by NADPH oxidase helps to destroy phagocytosed organisms but may be toxic to neighboring cells 10, 36]. The effects of the prostanoids are manifold. In the hepatocyte, prostanoids increase

the glycogenolytic activity thereby supplying KC with glucose for the production of NADPH via the hexose monophosphate shunt [14]. Other prostanoids such as thromboxane A2 may induce vasoconstrictory effects [15] most likely due to contraction of hepatic stellate cells [16] while prostaglandin E2 (PGE2) plays an autoregulatory role in KC. PGE2 inhibits the synthesis of prostaglandins and TNF-a by KC [17, 18] which may explain its well known hepatoprotective effects [19]. Increasing attention has focused on the mechanisms by which LPS activate KC. This process seems to be mediated by LPS binding protein (LBP), CD14, and toll-like receptor 4 (TLR4) [8]. In blood, LPS binds to LBP, a 60-kDa acute-phase protein produced predominantly by the liver and secreted into the circulation [20]. Although LBP is not required for interactions of LPS with CD14, its presence significantly decreases the concentration of LPS required for KC activation [21]. Thus, the LBP-CD14 pathway is crucial at low concentrations of LPS found under physiological

4  Signaling Pathways in Liver Diseases Kupffer Cells

conditions. Due to their location in the liver sinusoids, KC are chronically exposed to higher concentrations of LPS than the circulating blood monocytes. Unlike monocytes, KC have relatively low baseline expression of CD14 [22]. However, expression of CD14 on KC can be upregulated by multiple stimuli including LPS [23]. Furthermore, CD14 expression in the liver is also increased in various liver diseases [8]. The physiological significance of these observed differences in KC CD14 expression is not entirely clear. However, it is tempting to hypothesize that changes in CD14 expression determine the liver’s sensitivity to LPS toxicity. Since CD14 is a glycosylphosphatidylinositol (GPI)anchored protein without a transmembrane component, downstream partners for this receptor have long been sought. Meanwhile there is substantial evidence that LPS signaling is mediated by TLR4 [8]. Signaling through TLR4 requires MD-2, a secreted protein closely associated with the extracellular domain of TLR4 [24]. Downstream of TLR4, signaling occurs via MyD88, which associates with interleukin-1 receptor-associated kinase (IRAK) and TNF-activated factor 6 (TRAF-6) [25]. TRAF-6 mediated signaling pathways activate NFkB which results in the production of proinflammatory cytokines [25]. Based on these molecular findings a very interesting new therapeutic tool has been developed. NFkB inhibiting decoy oligodeoxynucleotide loaded gelatin nanoparticles selectively inhibit NFkB activation in Kupffer cells [26]. The benefit of selective inhibition of NF-kappaB is a big advantage, but not for everything [27].

Kupffer Cell: Neutrophil Interaction in Host Defense, Immune Tolerance, and Liver Regeneration Host Defense The rapid clearance of bacteria from the blood stream has been attributed to fixed tissue macrophages, in particular to KC [28]. Recent experiments indicate that the actual mechanism is far more complicated than phagocytosis by KC alone. Rather, the rapid elimination of bacteria taken up by the liver is dependent on the complex interaction of KC and microbicidal neutrophils which immigrate rapidly in response

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to infection [29, 30]. In brief, most organisms taken up in the liver are bound extracellularly by KC [29]. Experimental data suggest that the binding is mediated in part by the interaction of the lectins expressed by KC and the carbohydrate residues expressed by the bacteria [31, 32]. In parallel complementary adhesion, molecules (i.e., CD11b/CD18 and CD54) facilitate the adherence of neutrophils to the KC [33, 34], which subsequently internalize and kill the organisms bound to the KC surface [35]. Clearance of infiltrating neutrophils from inflamed tissues is required for the resolution of inflammation. Immunohistochemical detection of KC positive for myeloperoxidase in sections of mouse and human livers supports the role of KC in neutrophil elimination [36–38]. Taken together, these findings suggest that KC play a critical role in eliminating activated neutrophils, thereby suppressing their production of toxic metabolic compounds and degradative enzymes. Furthermore, the ingestion of apoptotic neutrophils may enhance or abrogate the inflammatory response of macrophages depending on the receptors mediating the uptake. Phagocytosis of apoptotic neutrophils via the avb3integrin/CD36 complex suppresses the production of proinflammatory cytokines such as IL-1b, TNF-a as well as eicosanoides [39]. Uptake of neutrophils mediated by Fc-receptors, on the other hand, induces the production of these inflammatory mediators [39].

Immune Tolerance The ingestion of neutrophils by KC may also have profound implications for the development and expression of adaptive immunity in the liver. It has been suggested, that the liver is actively tolerogenic and plays the key role in preventing generalized inflammation by eliminating circulating CD8+ T-cells specific for systemically disseminated antigens [40, 41]. Interestingly, KC recovered from chronically accepted hepatic allografts have a greater ability to induce apoptosis of alloreactive T-cells whereas the administration of these cells significantly prolongs the survival of hepatic allografts in an acute rejection model [42]. Macrophages in vitro cross-present epitopes derived from ingested apoptotic cells via the vacuolar alternate MHC class I pathway [43]. In the absence of appropriate costimulatory molecules, cross-presentation of antigens by macrophages

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is more likely to promote tolerance than to induce adaptative immunity in naïve T-cells [43, 44]. Thus, apoptotic neutrophils ingested by KC may exert a significant influence on the development and expression of antigen-specific CD4+ and/or CD8+ T-cell-mediated immunity in the liver, where sensitized T-cells are activated and naïve T-cells are tolerized. Furthermore, KC could inhibit activation of T cells. As main mechanisms of KC-mediated T cell suppression of IL-10, nitric oxide and TGF-b were found. These data indicated that KC’s may play a critical role in regulating immune reactions within the liver and contributing to livermediated systemic immune tolerance [45].

Liver Regeneration The capacity to regenerate is critical for the successful outcome of liver resections and is especially important in the context of split liver and living donor liver transplantations. The liver’s unique ability to regenerate itself has been known since antiquity, but the underlying mechanisms have been explored only recently. Activation of KC is necessary for the optimal regenerative ability of ?

the liver, possibly through the release of TNF-a [46–48] and interleukin-6 [49]. These cytokines initiate hepatocyte proliferation in vivo, at least in part, through NFkB and STAT-3 translocation (Fig. 4.2). Interestingly, recent data suggest that these events are triggered by leukocyte–KC interaction mediated by the intracellular adhesion molecule ICAM-1 [50]. Livers from ICAM-1 deficient mice exhibited impaired regeneration after 70% hepatectomy which was associated with a dramatic decrease in leukocyte recruitment as well as tissue TNF-a and interleukin levels. Similar impairment of liver regeneration and cytokine production in neutropenic and KC-depleted animals suggest a novel pathway in liver regeneration, where KC and leukocytes trigger a local inflammatory response leading to KC-dependent release of TNF-a and interleukin-6. KC activation could be a direct consequence of leukocyte-KC interaction. In addition, this local inflammation may induce complement activation thereby liberating potent KC activators. This view is supported by a recent study demonstrating the essential role of complement components C3a and C5a for liver regeneration [51]. C3a and C5a deficiency as well as interception of C5a receptor signaling resulted in suppression of interleukin-6/TNFa induction and

complement activation

leukocyte CD11b/ CD18

ICAM-1

C3a

C5a

C3aR

C5aR

G protein

?

TNF-R1

NFκB

TNFα IL-6 IL-6R

Kupffer cell Fig.  4.2  Role of Kupffer cells in liver regeneration. C3a and C5a activated complement factor 3 and 5; C5aR complement receptors; CD11b/18 C-receptor type 3, also known as MAC-1

NFkB

STAT-3

hepatocyte or b2aM-integrin; ICAM-1 intercellular adhesion molecule 1; ­IL-6R interleukin-6 receptor; NFkB nuclear factor kB; TNFa tumor necrosis factor; TNF-R1 tumor necrosis factor receptor 1

4  Signaling Pathways in Liver Diseases Kupffer Cells

NFkB/STAT-3 activation after hepatectomy [51]. Altogether, these results indicate that KC-leukocyte interaction together with complement activation contribute to liver regeneration (Fig. 4.2).

Role of Kupffer Cells in Liver Injury Multiple lines of evidence indicate that KC contribute to the pathogenesis of different liver injuries, including alcoholic liver disease [52], nonalcoholic fatty liver [53], liver failure following acetaminophen-intoxication [54], iron- and copper toxicity [55], and ischemia-reperfusion injury during liver resection or transplantation [56, 57]. Furthermore, there is first evidence for a pathogeneic role of KC in carbon tetrachloride-induced hepatic fibrosis [58], galactosamine-induced liver injury [59], and portal hypertension [60, 61]. Several studies point to LPS as cofactor in the pathogenesis and exacerbation of liver injury [8, 62]. These findings are in line with the concept of LPS-mediated KC activation as key mechanism in the pathogenesis of various liver diseases. However, there is evidence that other mechanisms of KC activation as well as the cytotoxic mechanisms induced by KC may play a role depending on the underlying liver disease. This aspect may be illustrated by several examples discussed below.

Acetaminophen In overdose, the analgesic/antipyretic acetaminophen produces centrilobular hepatic necrosis which can lead to acute liver failure. Depletion of intracellular glutathione and the increased generation of reactive oxygen and nitrogen species have been considered as critical patho-mechanisms [54]. Recent work shows the central role of tyrosin nitration by peroxynitrite formed by the rapid reaction of nitric oxide (NO) and superoxide [63]. Surprisingly, acetaminophen toxicity and tyrosin nitration can be dramatically reduced by several KC inactivators and knockout of inducible nitric oxide synthase (iNOS) [54] while liver injury still occurs in NADPH oxidase knockout mice [64]. These findings suggest activation of KC, iNOS induction, and the

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subsequent formation of NO, but not of KC-derived superoxide, as major determinants of peroxynitrite formation and consequently, acetaminophen toxicity. Isolated KC can be activated by acetaminophen in the absence of LPS or complement factors [65] which points to alternative pathways of KC activation by hepatotoxins.

Ischemia-Reperfusion Injury Ischemia-reperfusion injury is the major factor responsible for the morbidity associated with liver resection under vascular exclusion (Pringlemaneuver) or after liver transplantation [56, 57]. The pathophysiology of hepatic ischemia-reperfusion injury includes a number of mechanisms which contribute to various degrees to the overall injury. An excessive inflammatory response by activated KC is clearly recognized as a key mechanism of injury during reperfusion [56, 66, 67]. Interestingly, ischemia alone induces activation of KC [68]. During reperfusion additional KC activation and cell injury may occur by accumulated LPS during the unhepatic phase of liver transplantation [69] through activation of complement factors [11, 70]. This example demonstrates the activation of KC by an uncommon still undefined pathway induced by ischemia and the subsequent activation by two common pathways. In contrast to acetaminophen toxicity, nitric oxide derived from KC or other sources may reduce ischemia-reperfusion injury due to its vasodilatory effects in line with the observation of only little peroxynitrite formation [66]. Moreover, the production of a vascular oxidant stress by the NADPH-oxidase of KC has been identified as the central pathomechanism of hepatic reperfusion injury which enhances NFkB and activator protein-1 (AP-1) mediated expression of genes such as TNF-a, chemokines, and adhesion molecules [71]. Subsequent studies indicated that glutathione (GSH) released by hepatocytes via the sinusoidal GSH transporter may partially counteract the vascular oxidant stress by KC [67]. This endogenous defense mechanism is limited by the capacity of the sinusoidal GSH transporter and the rapid elimination of plasma GSH as illustrated by the low concentration of GSH in the vascular space in contrast to the

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H 2O 2

O 2•

NADPH-Oxidase NADPH-Oxidase

+

GSH (10 000 µM) Hepatocyte

GSH (10 µM)

GSSG

KupfferCell O2

Sinusoid

Fig.  4.3  Vascular oxidant stress by activated Kupffer cells: endogenous defense by glutathione. GSH reduced glutathione; GSSG oxidized glutathione; H2O2 hydrogen peroxide

1,000-fold higher intracellular concentrations of GSH (Fig. 4.3). Accordingly, prevention of hepatic reperfusion injury can be achieved by increasing the extracellular antioxidant capacity around the KC by intravenous infusion of GSH [72–75]. Interestingly, reperfusion injury by activated KC could be attenuated not only by antioxidants but also by the hormone atrial natriuretic peptide (ANP) [76, 77], which had no relevant effect on reactive oxygen formation by KC [78]. Thus, we concluded that ANP increased the resistance of hepatocytes against the oxidant stress by KC. This may be due to an inhibition of the increase in cytosolic calcium following oxidant stress in hepatocytes [79]. Additionally, ANP may exert cytoprotective effects through the inhibition of TNF-a released from activated KC [80]. Recent data show the potency of ANP to specifically induce hemoxygenase-1 in KC independently of cGMP [81]. However, this increased expression of HO-1 seems not involved in hepatoprotection conveyed by ANP because inhibition of HO-1 did not abrogate the hepatoprotective effects of ANP [81].

Alcoholic Liver Disease Hepatic macrophages play an important role in the pathogenesis of alcoholic liver disease. Early alcoholic liver injury in the intragastric ethanol infusion model is attenuated by depletion of KC with gadolinium chloride [82]. The role of KC activation by

increased permeability of the gut to endotoxins [83] is also supported by studies showing hepatoprotection in ethanol-fed animals given polymyxin B and neomycin [84] or lactobacillus [85]. In fact, the administration of antibodies against TNF-a attenuates alcoholic liver injury [86], and the importance of TNF-a is confirmed by the absence of alcoholic liver injury in the TNF receptor-1 lock out mice [87]. Acute or chronic exposure to ethanol influences macrophage function in opposite ways: KC production of proinflammatory cytokines is inhibited by acute, but stimulated after chronic exposure to alcohol [88]. Consistent with these results, KC harvested from the livers of rats that were fed ethanol for several weeks demonstrated a ­time-dependent increase in TNF-a and interleukin-6 expression [89]. The studies discussed above implicate a role for LPS and intestinal bacteria. Indeed, TLR4 recognizes endotoxin, a trigger of inflammation in alcoholic liver disease. Interestingly, TLR4-induced and MyD88-independent pathways play an important role in the pathogenesis of alcoholic liver disease [90]. Additionally, there may be several other factors contributing to phenotypic alterations. Suppression of NFkB and TNF-a expression by antioxidants in monocytes from alcoholic hepatitis patients suggests dysregulation of TNF-a gene transcription driven by NFkB [91]. As shown previously increased iron storage by KC may prime NFkB activation in experimental alcoholic liver disease [92]. Another postulated molecular mechanism for modified TNF-a production is the Trif/IRF-3 pathway [93]. Furthermore, altered methionine metabolism, in particular deficiency of S-adenosylmethionine, may also play a role in dysregulation of TNF-a-gene expression [52].

Kupffer Cells and Portal Pressure Several studies have shown that infection is closely associated with variceal bleeding. Although the mechanisms were not clear the importance of KC to produce vasoconstrictors upon activation has been hypothesized earlier [62]. Recent studies found that KC activation indeed results in a massive increase of portal pressure. One of the important vasoconstrictor seems to be thromboxane A2 [94, 95].

4  Signaling Pathways in Liver Diseases Kupffer Cells

Summary

›› KC ›› ›› ›› ›› ›› ›› ››

constitute 80–90% tissue macrophages present in the body. They reside within the lumen of the liver sinusoids and come in contact with microbial debris derived from the GI tract. The most significant activators of KC are the complement factors C3a, C5a, b-glucans, and LPS. LPS-mediated KC activation is a key mechanism in the pathogenesis of various liver diseases. LPS activate TLR4 on KC. Ingestion of activated neutrophils by KC contributes to liver-mediated systemic immune tolerance. Activation of KC is necessary for the optimal regenerative ability of the liver, possibly through the release of TNF-a and interleukin-6. Activation of KC plays a pathogenic role in ischemia/reperfusion injury, alcoholic liver disease and portal hypertension.

Multiple Choice Questions 1. During liver regeneration, KC (a) Are activated by complement components (b) Release TNF and IL-6 (c) Are activated by interaction with neutrophils (d) Stimulate hepatocyte proliferation (e) All the above 2. CD14 (a) Is a G protein-coupled receptor (b) Is a tyrosine kinase receptor (c) Is a glycosylphosphatidylinositol (GPI) an-chored receptor (d) Does not signal through MyD88 (e) Has cAMP as second messenger 3. Acetaminophen toxicity: (a) Is reduced by KC activation (b) Is reduced in NADPH oxidase knockout mice

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(c) Is reduced by NO derived by KC (d) Is mediated by LPS activation of KC (e) All the above are wrong 4. Concerning the role of KC in ischemia/reperfusion injury, the following is correct: (a) ANP decreased the resistance of hepatocytes against the oxidant stress by KC (b) Ischemia alone cannot activate KC (c) Nitric oxide derived from KC aggravates ischemia/reperfusion injury (d) Oxidative stress induced by NADPH-oxidase of KC aggravates ischemia/reperfusion injury (e) Extracellular GSH is deleterious 5. Which is the correct statement? (a) KC production of proinflammatory cytokines is stimulated by acute exposure to alcohol (b) KC production of proinflammatory cytokines is inhibited by chronic exposure to alcohol (c) KC activation results in a decrease of portal pressure (d) KC activation by increased permeability of the gut to endotoxins is supported by studies showing hepatoprotection in ethanol-fed animals given nonresorbable antibiotics (e) Alcoholic liver injury is worse in TNF receptor-1 lock out mice

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77 dietary glycine. Am J Physiol Gastrointest Liver Physiol 281:G200–G207 59. Czaja MJ, Xy J, Yue J et al (1994) Lipopolysaccharideneutralizing antibody reduces hepatocyte injury from acute hepatotoxin administration. Hepatology 19:1282–1289 60. Graupera M, Garcia-Pagan JC, Titos E et al (2002) 5-lipoxygenase inhibition reduces intrahepatic vascular resistance of cirrhotic rat livers: a possibile role of cysteinyl-leukotrienes. Gastroenterology 122:387–393 61. Yokoyama Y, Xu H, Kresge N et al (2003) Role of thromboxane A2 in early BDL-induced portal hypertension. Am J Physiol Gastrointest Liver Physiol 284:G453–G460 62. Goulis J, Patch D, Burroughs AK (1999) Bacterial infection in the pathogenesis of variceal bleeding. Lancet 353: 139–142 63. Hinson JA, Pike SL, Pumford NR et al (1998) Nitrotyrosineprotein adducts in hepatic centrilobular areas following toxic doses of acetaminophen in mice. Chem Res Toxicol 11: 604–607 64. James LP, McCullough SS, Knight TR et al (2003) Acetaminophen toxicity in mice lacking NADPH oxidase activity: role of peroxynitrite formation and mitochondrial oxidant stress. Free Radical Res 37:1289–1297 65. Goldin RD, Ratnayka ID, Breach CS et al (1996) Role of macrophages in acetaminophen (paracetamol)-induced hepatotoxicity. J Pathol 179:432–435 66. Jaeschke H (2003) Role of reactive oxygen species in hepatic ischemia-reperfusion injury and preconditioning. J Invest Surg 16:127–140 67. Jaeschke H, Farhood A (1991) Neutrophil and Kupffer cellinduced oxidant stress and ischemia-reperfusion injury in rat liver in vivo. Am J Physiol Gastrointest Liver Physiol 260:G355–G362 68. Rymsa B, Wang JF, de Groot H (1991) O2-release by activated Kupffer cells upon hypoxia-reoxygenation. Am J Physiol Gastrointest Liver Physiol 261:G602–G607 69. Yokoyama I, Todo S, Miyata T et al (1989) Endotoxemia and human liver transplantation. Transplant Proc 21: 3833–3841 70. Jaeschke H, Farhood A, Bautista AP et al (1993) Complement activates Kupffer cells and neutrophils during reperfusion after hepatic ischemia. Am J Physiol Gastrointest Liver Physiol 264:G801–G809 71. Jaeschke H (2003) Molecular mechanisms of hepatic ischemia-reperfusion injury and preconditioning. Am J Physiol Gastrointest Liver Physiol 284:G15–G26 72. Bilzer M, Baron A, Schauer R et al (2002) Glutathione treatment protects the rat liver against injury after warm ischemia and Kupffer cell activation. Digestion 66:49–57 73. Bilzer M, Paumgartner G, Gerbes AL (1999) Glutathione protects the rat liver against reperfusion injury after hypothermic preservation. Gastroenterology 117:200–210 74. Liu P, Fisher MA, Farhood A et al (1995) Beneficial effects of extracellular glutathione against endotoxin-induced liver injury during ischemia and reperfusion. Shock 43:64–70 75. Schauer RJ, Gerbes AL, Vonier D et al (2004) Glutathione protects the rat liver against reperfusion injury after prolonged warm ischemia. Ann Surg 239:220–231 76. Bilzer M, Witthaut R, Paumgartner G et al (1994) Prevention of ischemia/reperfusion injury in the rat liver by atrial natriuretic peptide. Gastroenterology 106:143–151 77. Gerbes AL, Vollmar AM, Kiemer AK et al (1998) The guanylate cyclase-coupled natriuretic peptide receptor: a new

78 target for prevention of cold ischemia-reperfusion damage of the rat liver. Hepatology 28:1309–1317 78. Bilzer M, Jaeschke H, Vollmar AM et al (1999) Prevention of Kupffer cell-induced oxidant injury in rat liver by atrial natriuretic peptide. Am J Physiol 276:G1137–G1144 79. von Ruecker AA, Wild M, Rao GS et al (1989) Atrial natriuretic peptide protects hepatocytes against damage induced by hypoxia and reactive oxygen: possible role of intracellular free ionized calcium. J Clin Chem Clin Biochem 27: 531–537 80. Kiemer AK, Baron A, Gerbes AL et al (2002) The atrial natriuretic peptide as a regulator of Kupffer cell functions. Shock 17:365–371 81. Kiemer AK, Gerwig T, Gerbes AL et al (2003) Kupffer-cell specific induction of heme oxygenase 1 (hsp32) by the atrial natriuretic peptide-role of cGMP. J Hepatol 38:490–498 82. Adachi Y, Bradford BU, Gao W et al (1994) Inactivation of Kupffer cells prevents early alcohol-induced liver injury. Hepatology 20:453–460 83. Enomoto M, Ikejima K, Yamashima S et al (2001) Kupffer cell sensitization by alcohol involves increased permeability to gut-derived endotoxin. Alcohol Clin Exp Res 25:51–54 84. Adachi Y, Moore LE, Bradford BU et al (1995) Antibiotics prevent liver injury in rats following long-term exposure to ethanol. Gastroenterology 108:218–224 85. Nanji AA, Khettry U, Sadrzadeh SM (1994) Lactobacillus feeding reduces endotoxemia and severity of experimental alcoholic liver disease. Proc Soc Exp Boil Med 205: 243–247 86. Iimuro Y, Gallucci RM, Luster MI et al (1997) Antibodies to tumor necrosis factor alfa attenuate hepatic necrosis and inflammation caused by chronic exposure to ethanol in the rat. Hepatology 26:1530–1537

C. J. Steib and A. L. Gerbes 87. Yin M, Wheeler MD, Kono H et al (1999) Essential role of tumor necrosis factor alpha in alcohol-induced liver injury in mice. Gastroenterology 117:942–952 88. McClain CJ, Hill DB, Schmidt J et al (1993) Cytokines and alcoholic liver disease. Semin Liv Dis 13:170–182 89. Kamimura S, Tsukamoto H (1995) Cytokine gene expression by Kupffer cells in experimental alcoholic liver disease. Hepatology 22:1304–1309 90. Hritz I, Mandrekar P, Velayudham A, Catalano D, Dolganiuc A, Kodys K, Kurt-Jones E, Szabo G (2008) The critical role of TLR4 in alcoholic liver disease is independent of the common TLR adapter MyD88. Hepatology 48: 1224–1231 91. Hill DB, Devalaraja R, Joshi-Barve S et al (1999) Antioxidants attenuate nuclear factor-kappa B activation and tumor necrosis factor-alpha production in a alcoholic hepatitis patient monocytes and rat Kupffer cells, in vitro. Clin Biochem 32:563–570 92. Tsukamoto H, Lin M, Ohata M et al (1999) Iron primes hepatic macrophages for NF-kB activation in alcoholic liver injury. Am J Physiol 277:G1240–G1250 93. Zhao XJ, Dong Q, Bindas J, Piganelli JD, Magill A, Reiser J, Kolls JK (2008) TRIF and IRF-3 binding to the TNF promoter results in macrophage TNF dysregulation and steatosis induced by chronic ethanol. J Immunol 181: 3049–3056 94. Xu H, Korneszczuk K, Karaa A, Lin T, Clemens MG, Zhang JX (2005) Thromboxane A2 from Kupffer cells contributes to the hyperresponsiveness of hepatic portal circulation to endothelin-1 in endotoxemic rats. Am J Physiol Gastrointest Liver Physiol 288(2):G277–G283 95. Steib CJ, Gerbes AL, Bystron M, Op den Winkel M, Härtl J, Roggel F, Prüfer T, Göke B, Bilzer M (2007) Kupffer cell activation in normal and fibrotic livers increases portal pressure via thromboxane A(2). J Hepatol 47(2): 228–238

5

Hepatic Sinusoidal Endothelial Cells Robert C. Huebert and Vijay H. Shah

Development and Structure Hepatic sinusoidal endothelial cells (HSECs) are a morphologically distinct population of cells that form the lining of liver sinusoids. Features that distinguish HSEC from endothelial cells present in other organs and in larger liver vessels are the presence of multiple fenestrae throughout the cells and the lack of an underlying basement membrane [1–4]. The sinusoids are positioned between hepatocyte plates, and they initiate at the portal tract and terminate at the central vein. Sinusoids carry blood that converges in the liver from the portal venous supply as well as from the hepatic artery [5] (Fig. 5.1). Sinusoids are separated from adjacent hepatocytes by the perisinusoidal space of Disse. Due to their position, HSECs are the first cells that are in contact with blood flow into the sinusoids and serve to compartmentalize the vascular sinusoidal channels from the hepatic parenchyma [3, 4]. The hepatic sinusoids range in diameter from 4 mm near the portal triad to 5.5 mm near the central vein [6]. As this is smaller than the size of both red and white blood cells, there is distortion of both cells and the sinusoid during the passage of blood cells [6, 7]. This process has been referred to as an “endothelial massage” that allows efficient exchange of compounds from the blood through sinusoidal fenestrae into the space of Disse [5, 6]. Also residing in the sinusoidal space are hepatic macrophages (Kupffer cells), hepatic natural killer cells (pit cells), and liver-specific pericytes (hepatic stellate cells), each of which are covered in other chapters. V. h. Shah () GI Research Unit, Guggenheim 10, Mayo Clinic, 200 First Street SW, Rochester, MN 55905, USA e-mail: [email protected]

During organogenesis, the primordial liver is vascularized by capillaries that are continuous and have a basement membrane. Differentiation of these capillaries occurs from precursors located in the septum transversum, a mesenchymal structure located between the pericardium and the hepatic diverticulum [8, 9]. At 5 weeks of gestation, growing cords of hepatoblasts derived from the hepatic diverticulum surround the precursor vessels of the septum transversum. Intrahepatic capillaries at this early stage of gestation have the phenotype of typical fetal capillaries, containing cell–cell junctions and a basement membrane. Between 5 and 12 weeks of gestation, the vessels adjacent to hepatocytes develop their mature phenotype, marked by the development of fenestrae and the loss of both basement membrane and cell–cell junctions [10–12]. Interestingly, in some hepatic disease states, HSECs dedifferentiate from their specialized phenotype back toward a more typical endothelial cell phenotype [7, 12–14].

Cellular Functions Fenestration/Filtration The most prominent feature of sinusoidal endothelial cells is the presence of multiple fenestrae throughout the cell. Fenestrae have diameters of about 100 nm, and while the number of fenestrae increases as the sinusoids reach the central vein, their diameter decreases [1, 6, 15]. An intracellular cytoskeleton composed of microfilaments, intermediate filaments, and microtubules supports the dynamic fenestrae [2], which can reorganize to form additional fenestrae when stimulated [16]. The fenestrae of the HSEC serve as a mechanical sieve,

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Fig. 5.1  Vasculature architecture of the liver. Blood flow enters the liver via the portal vein (PVb) as well as the hepatic artery (HAb). While portal blood enters directly into the sinusoids (S), hepatic arterial blood perfuses into distinct anatomic locations prior to re-entering the sinusoids. Sinusoidal blood leaves the liver via the central veins (CV). Sinusoidal endothelial cells (En) form the fenestrated sinusoidal wall. Kupffer cells (K) are located within the sinusoids, whereas hepatic stellate cells, also

termed fat storing cells (FSCs), lie within the space of Disse (DS), adjacent to the single layer of hepatocytes (liver plate, LP). Bile canaliculi (BC) drain bile into the interlobular bile ducts (BDL) via the caniliculoductar junction (CD) in the opposite direction to flow in the vascular channels (Permission requested from Motta P, Muto M, Fujita T, (1978). “The Liver. An Atlas of Scanning Electron Microscopy.” Igaku-Shoin Medical Publishers, Inc)

which facilitates the transfer of nutrients and molecules from the sinusoidal space to the hepatic parenchyma [15, 17]. As HSECs lack a basement membrane, the presence of the fenestrae allow for steric selection of compounds that permeate the endothelial cell barrier and gain access to the space of Disse and the hepatic parenchymal cells [2, 15]. For example, the presence of fenestrae allows chylomicron remnants passing into the space of Disse to be recognized and affected by hepatocyte metabolism [2, 6]. Chylomicrons themselves, too large to pass through the HSEC fenestrae, remain in the

sinusoidal space and eventually pass from the liver. The effect on HSEC fenestrae by various disease states is felt to have major implications on the clinical manifestations of each disease state [14, 18].

Expression of Adhesion Molecules Leukocytes are attracted to and localized at sites of inflam­ mation throughout the body. This occurs because of

5  Hepatic Sinusoidal Endothelial Cells

their attachment to adhesion molecules expressed on the surface of endothelial cells [19, 20]. The liver is exposed to numerous foreign antigens that traverse the bowel mucosa and enter the portal circulation, thereby gaining access to the liver via the hepatic sinusoids [20]. Adhesion molecules on the surface of typical endothelial cells capture leukocytes in the passing blood flow, permitting recruitment of inflammatory cells to areas of invasion by foreign pathogens [21]. HSECs have been shown to rarely express selectins, a class of adhesion molecules commonly expressed in various other tissues [22]. However, other adhesion molecules, such as vascular adhesion protein (VAP-1), are expressed in HSEC and are responsible for the capture of rolling leukocytes through the sinusoids, thus recruiting them to areas of inflammation [20, 23]. In addition, other leukocyte adhesion molecules such as vascular cell adhesion molecule 1 (VCAM-1) are induced on HSEC in states of liver inflammation [19, 20, 22, 24]. VCAM-1 also mediates, in part, the adhesion of melanoma cells to hepatic endothelium, thereby playing a key role in the development of hepatic metastatic lesions in this disease [20]. Neutrophil attraction to HSEC is often mediated by intercellular adhesion molecule 1 (ICAM-1), which is constitutively expressed on HSEC and is upregulated in response to liver injury [19, 22, 25]. Recent evidence suggests that interaction between CD44 and hyaluronan may be another important mechanism for neutrophil sequestration in inflamed liver sinusoids [26]. Chemokines produced by inflamed tissues bind to receptors on leukocytes and prompt the subsequent firm adhesion of leukocytes to the endothelium. This process allows migration of leukocytes into inflamed tissues. The expression of multiple chemokines is upregulated in the inflamed liver [19]. T cells that infiltrate the liver have been demonstrated to express receptors for these chemokines [22]. After adhesion, mole­­cules expressed by tissue endothelial cells have captured a leukocyte; the leukocyte must transit through the endothelium to reach the inflamed tissue [27]. Many tissues have inter-endothelial-cell tight junctions that present a barrier to the migration of leukocytes. However, HSECs form a discontinuous barrier without tight junctions, which likely allows for unique means of leukocyte migration through the endothelium barrier and into the inflamed parenchyma [20].

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Signaling Specific signaling pathways including those in HSEC are covered in other chapters of this book. However, as nitric oxide (NO) is a key signaling pathway in HSEC, some discussion of NO production in HSEC is provided. HSECs produce NO via nitric oxide synthase. Three isoforms exist including the endothelial (eNOS), inducible (iNOS), and neuronal (nNOS) forms. eNOS is constitutively expressed by HSEC and is thought to be chiefly responsible for regulating sinusoidal vascular tone. Production of NO via eNOS is a characteristic that is exclusive to endothelial cells among hepatic cell types [20]. NO produced by the HSEC may serve to regulate sinusoidal blood flow through a paracrine action on perisinusoidal contractile cells [28]. Increase in NO production via eNOS is seen in HSEC subjected to shear stress, a characteristic that may serve to autoregulate blood flow through the liver [29]. By this mechanism, if blood flow through the hepatic sinusoid is high, eNOS mediated production of NO may serve locally to dilate the vascular bed, thus decreasing the resistance of the vascular bed [29]. In cirrhotic states, production of NO via eNOS is diminished [30]. One of the major phenotypic changes in HSEC during portal hypertension includes alteration of the NO generation system that results in a regional NO deficiency. Deficient NO generation, coupled with other changes that reduce its local effect, results in a myriad of downstream alterations. While eNOS protein levels appear to be unchanged, there is an increase in eNOS binding by the inhibitory protein caveolin. Additionally, Akt phosphorylation and activation of eNOS is impaired in cirrhosis. Other important agonists that promote eNOS derived NO generation include endothelin (via the ET-B receptor), VEGF, estrogen, and others [25, 28, 31, 32]. Every cell type within the liver has the capacity to generate NO via iNOS upon stimulation by liver injury or cytokine induction, though this does not occur during normal conditions [33, 34]. HSECs have been demonstrated to produce NO via induction of iNOS in response to cytokines such as IFN-g and LPS, agents that do not by themselves stimulate iNOS induction in hepatocytes [35]. In contrast, agents such as TNF-a and IL-1b that stimulate induction of iNOS in hepatocytes do not exert a similar effect in HSEC [35]. As HSECs are positioned adjacent to the blood supply of the liver, the majority of which is delivered from the

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splanchnic circulation, inflammatory cytokines may act more quickly on HSECs to induce a rise in NO production via iNOS than in other hepatic cell types.

Metabolism Numerous compounds and all orally ingested medications pass through the liver prior to distribution throughout the body to target sites of action. It has been demonstrated that HSECs possess only very low levels of cytochrome P450 enzymes; however, these cells do play a role in metabolism [36]. They possess low levels of oxidative enzymes, and possess high levels of the postoxidative enzymes epoxide hydrolase and glutathione transferase [37]. Therefore, although the ability to oxidize drugs is limited in HSEC, there is a relatively greater ability of the cells to conjugate or hydrolyze metabolized compounds.

Role in Blood Clearance The hepatic reticuloendothelial system contains two major scavenger cell types, including Kupffer cells and HSECs, which clear particulate matter and macromolecules from the blood, respectively. HSECs have welldeveloped apparatus for endocytotic functions, including lysosomes, endosomes, and pinocytotic vesicles. The capacity of HSEC to take up substances such as heparin, albumin, lipoproteins, and hyaluronate has been well described. For some substances such as heparin, endocytotic capacity of HSEC exceeds that of Kupffer cells [1]. The ability of HSEC to take up hyaluronate via endocytosis has been used as a functional marker of sinusoidal endothelial cells [38]. Rising serum hyaluronate levels have been demonstrated to correlate with decreased endocytotic capabilities of sinusoidal endothelial cells. A typical endocytosis pathway that has been described in HSEC involves the mannose receptor. The mannose receptor functions by binding to and internalizing compounds or antigens that contain terminal glycoproteins. The mannose receptor exists on the cell wall in coated pits and there is a large intracellular pool of additional receptors. When the receptor binds to ligand, rapid internalization occurs and the ligand is delivered to endosomes and lysosomes for degradation.

R. C. Huebert and V. H. Shah

New receptors, either from intracellular stores or recycled receptors, return to the cell surface to bind additional ligand [39]. The process of endocytosis through nonclathrin coated pits, such as caveolae, remains enigmatic in HSEC. Some studies have demonstrated the presence of caveolae or similar structures termed vesiculo-vacuolar organelles, which constitute grape-like clusters of caveolae in HSEC. However, protein levels of the caveolae coat protein caveolin, though upregulated in some disease states [40], seem very low in normal HSEC, and thus, it has been postulated that caveolae may not play an important role in HSEC endocytosis perhaps because of the availability of alternate endocytotic pathways.

Antigen Presentation HSECs have been demonstrated to be efficient presenters of bacterial antigens to T-cells, allowing for immune system activation. To this end, these cells have been demonstrated to carry major histocompatibility complex (MHC) class II molecules and to produce IL-1 after production and processing of bacterial antigens, both features common to antigen presenting cells [41]. The ability of HSECs to function as antigen presenting cells accounts for an additional mechanism by which they can attract T cells to the liver. In addition to nonspecific adhesion molecules, HSECs can recruit antigen-specific T cells to the liver by means of antigen presentation. The MHC class II possessing character of HSECs is shown to lead naïve CD4+ cells to differentiate into IL-4 and IL-10 expressing T cells [13, 42]. HSECs induce not only proliferation of CD4+ cells, but also the production of IFN-a [13, 43]. In addition, there is cross-presentation of antigen to CD8+ cells by HSEC that induces CD8+ cell proliferation and development of antigen tolerance. This is contrasted with the development of immunity, which is typical of CD8+ cell stimulation by dendritic cells [42]. This function may play a role in avoidance of unwanted autoimmune reactions against food antigens. Bacterial antigens, as well as endotoxin derived from the intestines, are commonly present in portal venous blood. Endotoxin has been shown to upregulate the functional activity of typical antigen presenting cells, such as dendritic cells and macrophages. In contrast to these cells, HSEC activity as antigen presenting

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cells is decreased in response to endotoxin [44]. The effect of endotoxin on HSEC is not via a decreased ability of the cells to endocytose bacterial antigens, but rather by affecting intracellular means of processing the bacterial antigens. The pH of endosomes and lysosomes is raised in HSEC in response to endotoxin, therefore decreasing the ability of these cells to process antigen and to activate naïve and memory T cells [44]. The significance of this effect is that bacterial antigens can be efficiently cleared by HSEC without inducing hepatic inflammation in response to the near constant presence of endotoxin in portal venous blood. This allows for hepatocytes to function in a relatively immunologically quiescent state.

Pathobiology Alcoholic Liver Disease The role of alcohol as a cause of chronic liver disease has been well described. It has recently been described that the earliest changes in the liver due to alcohol occur in the sinusoidal endothelium. Alcohol intake has been demonstrated to dilate the HSEC fenestrae [45], though chronic, excessive intake of alcohol has been shown paradoxically to lead to capillarization of the endothelium with closure of the fenestrae and the development of an endothelial basement membrane. Both the size and number of fenestrae are decreased with chronic alcohol ingestion [46]. These changes of HSEC fenestrae have been shown to be reversible upon abstinence from alcohol [6]. In addition, the scavenging activity of HSEC has been shown to be dysfunctional after only short periods of alcohol ingestion. These changes in the morphology and functional characteristics of HSEC precede the effects of alcohol on hepatic parenchymal cells. The effect of alcohol on HSEC has been demonstrated to be due primarily to intermediates produced by neighboring Kupffer cells rather than the alcohol itself. The effects of alcohol on sinusoidal endothelial cells can be eliminated by administration of compounds that inactivate Kupffer cells prior to the treatment on experimental animals with alcohol [8, 47, 48]. Chronic alcohol ingestion has also been demonstrated to affect the endocytotic ability of HSECs [48].

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Receptor mediated uptake of ligands is slower in HSEC from animals fed on ethanol chronically. In addition, the ability of HSEC to degrade ligands that had been imported by endocytosis is also diminished after alcohol ingestion [49]. The implications of these findings are that bacterial antigens may not be as efficiently removed by HSEC-mediated endocytosis in states of chronic ethanol ingestion. This may lead to prolonged antigen presence and immune stimulation, resulting in increased inflammation in the liver [49].

Nonalcoholic Fatty Liver Disease Nonalcoholic fatty liver disease (NAFLD) is a term encompassing a spectrum of liver damage including steatosis, steatohepatitis, and fibrosis/cirrhosis. The syndrome and pathological findings are often indistinguishable from those in alcoholic liver disease. Important risk factors include obesity, diabetes mellitus, and hyperlipidemia [50]. While much of the pathology in this disorder is attributable to the deposition of fat droplets within the hepatic parenchyma, important changes occur within the hepatic microcirculation as well. In the steatotic liver, the sinusoidal lumen is narrowed and distorted resulting in reduced sinusoidal volume and impaired microvascular blood flow [51]. Additionally, there is evidence of increased HSEC expression of cellular adhesion molecules, such as ICAM-1 and VCAM-1. This is thought to be related to upregulation of NF-kB and a significant increase in the number of adherent leukocytes [52]. This exaggerated peri-vascular inflammatory response may serve to potentiate further liver damage due to reduced tolerance for ischemia-reperfusion injury and increased sensitivity to hepatic toxins.

Portal Hypertension/Cirrhosis Chronic liver disease and its associated complications are increasing in frequency, due in a large part to rising rates of obesity and the hepatitis C epidemic. Cirrhosis is the final common pathway in chronic liver disease and a syndrome of increased portal pressure is the final common pathway in all etiologies of cirrhosis. Cirrhosis is a disease characterized by extensive

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scarring throughout the liver, destruction of the normal hepatic architecture, and high vascular resistance through the hepatic circulation. Endothelial dysfunction and impaired endothelium-dependant relaxation is thought to be an important early event in the pathogenesis of cirrhosis. In cirrhotic states, NO production from HSEC is diminished [30]. This most likely occurs through defects in posttranslational processing of eNOS including enhanced expression of the NOS inhibitory protein, caveolin, as well as decreased eNOS phosphorylation at Serine 1179 [32, 34]. Furthermore, production of the constrictor ET-1 is increased, in part through HSEC production [28]. HSEC fenestrae shrink and disappear in the cirrhotic state, a change that precedes the development of an endothelial basement membrane and capillarization of the sinusoids [6, 18, 53] (Fig. 5.2). Studies of liver biopsy specimens in patients with primary biliary cirrho­sis have demonstrated that as histological degree of disease

advances, the sinusoidal endothelium develops the capillarized phenotype [14, 54]. In addition, the functional marker of HSEC dysfunction, rising serum hyaluronate levels, is seen in the histological progression of primary biliary cirrhosis, and levels of hyaluronate appear to correlate with disease stage [54]. Activation of hepatic stellate cells into a phenotype that produces extracellular matrix, which leads to fibrosis, is central to the development of cirrhosis. HSECs produce growth factors that induce stellate cell proliferation and production of extracellular matrix [14]. Some studies have demonstrated that HSECs are also capable of producing extracellular matrix components such as fibronectin, laminin, and type IV collagen when stimulated with TGF-b [55]. This suggests that in cirrhotic or inflammatory states, basement membranes and extracellular matrix deposition may derive not only from perisinusoidal stellate cells, but also from the sinusoidal endothelial cells themselves [12, 55].

Fig. 5.2  Scanning electron microscopy of rat hepatic sinusoids in zone 3: (a) control and (b) after 4 weeks, (c) 6 weeks, and (d) 12 weeks of intraperitoneal injections of thioacetamide, an exper-

imental model of cirrhosis (Permission requested from Mori T, Okanoue T, Sawa Y, et al, (1993). “Defenestration of the sinusoidal endothelial cell in a rat model.” Mosby-Year Book, Inc.)

5  Hepatic Sinusoidal Endothelial Cells

An additional change seen in HSEC in cirrhotic states is the expression of numerous adhesion proteins not expressed in normal HSEC. Several integrins that serve as cell-matrix adhesion proteins are not seen in normal HSEC, but are produced in sinusoidal endothelial cells of the cirrhotic liver [11]. Integrins function to anchor the endothelial cells to the basement membrane that develops as described in cirrhotic states. Furthermore, von Willebrand factor expression also appears to increase in HSEC in cirrhosis [54]. Platelet-endothelial cell adhesion molecule-1 is present on HSECs of cirrhotic livers, but has been found to be similarly expressed in the HSECs of normal livers as well [56].

Angiogenesis/Hepatic Malignancies Hepatocellular carcinoma (HCC) is a primary liver tumor that is characterized by blood supply derived exclusively from the hepatic artery. During the development of small HCC, the blood supply to the tumor transitions from a dual blood supply to an exclusively arterialized supply. During this transition, the hepatic sinusoids undergo alterations of capillarization, with loss of fenestrae and development of a basement membrane [14, 57]. These changes may occur to preserve the sinusoidal structure in the setting of increased intrasinusoidal pressures resulting from the arterial blood flow. The liver is also a common site to find evidence of tumor metastasis. The sequence of events required to establish metastatic tumor includes tumor cell arrest by binding to endothelial adhesion molecules, migration through the endothelial barrier, migration into the subendothelial space, and ultimately proliferation  [58]. Melanoma cells have been found, in experimental conditions, to use mannose receptors and the adhesion molecule VCAM-1 to affect adherence to sinusoidal endothelium [59, 60]. The binding of these melanoma cells to HSEC was found to be dependent on IL-1, which induces expression of VCAM-1 [59–62]. This is consistent with other studies that have demonstrated enhanced binding of tumor cells to endothelium that has been “primed” by inflammation. HCC cells and other liver cancers release angiogenic factors to recruit new vessels to the growing tumor including VEGF, angiopoietins, epidermal growth factor, platelet-derived endothelial cell growth

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factor, and others [63]. In addition to angiogenesis, which involves increased tumor vasculature derived from neighboring endothelium, there is evidence suggesting that vasculogenesis, or de novo formation of new vessels, is an important contributor to tumor vascularity. This process is thought to be mediated by bone marrow-derived endothelial progenitor cells (EPCs), which are mobilized from the bone marrow by tumor-derived cytokines and hone to the site of the tumor via the bloodstream. Specific cell surface markers such as CD31 and VEGFR2 can identify these cells and some studies suggest that the contribution of EPCs to tumor neovascularization may be as high as 35–45% [64]. Higher levels of circulating EPCs have been detected in patients with HCC than in healthy controls [65]. Furthermore, levels are higher in advanced unresectable HCC versus patients with resectable HCC or cirrhosis alone, suggesting use for these cells as a prognostic marker [66]. With increasing evidence that antiangiogenic approaches may have efficacy in tumor growth and ongoing clinical trials related to antiangiogenic therapy in HCC, the processes of angiogenesis and vasculogenesis remain interesting and important potential targets in the treatment of both primary and metastatic hepatic malignancies.

Drug Toxicity One of the earliest findings of hepatic toxicity due to acetaminophen is the development of large pores in HSECs with separation of HSECs from the underlying hepatocytes, thereby widening the space of Disse [67]. Collapse of the sinusoidal lumen eventually occurs, likely secondary to enlargement of the space of Disse. Changes in HSEC due to acetaminophen were found to be due to primary effects of the drug on the endothelial cells in one mouse strain, and also due to metabolites of acetaminophen by adjacent cells in another mouse strain and in rats [67, 68]. In addition, depletion of cellular glutathione in both HSECs and adjacent cells was found to be crucial to the pathogenesis of acetaminopheninduced toxicity, as well as the toxicity of other drugs, with abrogation of toxic effects after treatment with exogenous glutathione [69–71]. Acetaminophen toxicity has been shown to be exacerbated by concomitant alcohol use, and its toxic effects on HSEC are exaggerated in experimental animals treated with ethanol [52].

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Veno-occlusive disease (VOD) is a well-described hepatotoxin induced condition characterized by hepatomegaly, jaundice, and ascites within 10–20 days after starting a chemotherapeutic regimen [72]. VOD has been most commonly associated with chemotherapy regimens used for marrow ablation in bone marrow transplants. Histological features include fibrosis of the liver sinusoids and necrosis of zone 3 hepatocytes [72, 73]. It has been demonstrated that the earliest microscopic changes of VOD are seen in HSEC [74]. The earliest histologic changes seen in the liver in VOD are dilatation and engorgement of the hepatic sinusoids with extravasation of red blood cells into the space of Disse [72, 73]. Electron microscopy demonstrates closure of fenestrae in HSEC and accumulation of collagen in the pores of sinusoids [74]. Animal models of VOD have demonstrated early morphologic changes in the HSEC such as loss of fenestration and the appearance of gaps in the junctions between HSEC [73, 74]. It has also been seen that HSECs undergo further morphologic changes such as cellular rounding and sloughing from the lining of the hepatic sinusoid. In experimental models of VOD, embolization of sloughed HSECs, as well as Kupffer cells can cause downstream occlusion of the hepatic sinusoids and resultant portal hypertension [75]. Acti­ vation of matrix metalloproteinases and diminished NO generation have also been implicated in this syndrome in the experimental models. Administration of an NO donor or MMP-2/MMP-9 inhibitors minimizes endothelial injury and red cell penetration into the space of Disse. Furthermore, inhibition of eNOS exacerbates, and the inhibition of iNOS reduces the endothelial injury suggesting that constitutive, HSEC-derived, NO is protective. The changes induced in HSEC, which can be abolished by administration of glutathione, precede pathological changes in hepatic parenchymal cells [71]. Thus, the disruption of the hepatic circulation by HSEC damage is the cause of hepatic parenchymal damage, rather than parenchymal damage being the cause of hepatic circulatory dysfunction [71].

Cellular Rejection Rejection of the transplanted organ is a major concern after orthotopic liver transplantation. One of the earliest signs of acute liver rejection is the presence of infiltrating immune cells within the sinusoids. There is also

R. C. Huebert and V. H. Shah

evidence of functional abnormalities of HSEC during episodes of acute rejection, such as increases of serum hyaluronate levels [76], reflecting diminished endocytotic capabilities and damage of HSEC. HSECs also play an important role in the inflammation of nonacute graft rejection. Leukocytes are arrested in sinusoids by binding to adhesion molecules that are induced by inflammatory cytokines such as TNF-a and IL-1b [24]. HSECs display a different pattern of adhesion molecules during graft rejection than do portal vein endothelial cells [24]. The lack of adhesion molecules such as selectins on HSEC may be necessary to prevent the firm adherence and microvascular thrombosis of the sinusoids by leukocytes in conditions such as rejection [24]. The difference in patterns of adhesion molecule expression by different types of endothelial cells also may explain the recruitment of different cell types to different tissues. At sites of the portal vein acute rejection, eosinophils are typically seen while in the liver sinusoid, natural killer cells are more typically seen [24].

Ischemia-Reperfusion Injury HSECs are the most sensitive cells in the liver to cold preservation damage. Studies of rat livers perfused with University of Wisconsin solution for varying times demonstrate that morphologic changes occurred after as few as 8  h of perfusion [77] earlier than changes seen in hepatocytes. The upper limit of time that graft tissue can be kept in a state of cold ischemia is dictated by the ability of HSEC to survive [78, 79]. Morphologic changes noted during periods of cold ischemia are small blebs and changes of surface texture including pits [77, 80]. Fenestrae are widened during periods of cold ischemia, prior to reperfusion. However, it is at the time of reperfusion that the most marked destructive changes in ­sinusoidal endothelial cells are seen [78, 81]. Upon reperfusion, lethal injury, rounding of the cells, denudation of sinusoids, and condensation of HSEC nuclei, all herald the loss of viability of the tissue graft [78, 82]. The changes that occur in HSEC upon reperfusion are probably apoptotic rather than necrotic [79, 81, 83, 84]. The changes of reperfusion injury to HSEC are most marked in periportal areas compared to pericentral areas [84]. Interestingly, apoptosis of rat HSEC is inhibited by ­fasting the donor animal prior to organ harvest.

5  Hepatic Sinusoidal Endothelial Cells

The mechanism of this is unclear [79]. Several signaling mediators are thought to be involved in ischemiareperfusion injury including cytokines, reactive oxygen species (ROS), activated complement, and calcium. Cytokines such as IL-1, IL-6, and TNF-a may induce neutrophil recruitment. ROS such as superoxide, NO, and peroxynitrite may produce cytotoxic effects via nitrosylation of iron–sulfur groups and tyrosine residues, inactivation of heme groups, and lipid peroxidation. Activated complement may act directly by the formation of membrane attack complexes and indirectly by production of pro-inflammatory cytokines and neutrophil recruitment. Calcium has been implicated in ischemia-reperfusion injury and is essential for activation of calcium-dependent phospholipases, nucleases, and proteases [85].

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Summary

›› The presence of multiple fenestrae throughout

›› ›› ›› ›› ››

Aging Process The fenestrations seen in HSEC allow transfer of macromolecules from the hepatic blood supply to the hepatic parenchyma. Due to the lack of a charged basement membrane, size is essentially the only factor that dictates which compounds will traverse the sieve-like fenestrae [15]. With aging, changes in the number of fenestrae in the HSEC are seen [86]. This aging process resembles the capillarization seen in other pathologic states of the liver because with the loss of fenestrae, the sinusoidal endothelium more closely resembles the capillary beds in other vascular tissues. There is thickening and defenestration of HSEC as well as increased levels of von Willebrand factor and ICAM-a. These changes have been termed pseudocapillarization [87]. The etiology of these age-related changes is uncertain, though chronic exposure to alcohol and oxidants have been shown to induce changes in HSEC [6, 88]. The implications of hepatic sinusoidal capillarization are uncertain, but there appears to be reduced sinusoidal perfusion and impaired clearance of substrates [87]. It has been postulated that this process may lead to disordered lipid metabolism [86]. Chylomicron remnants, rich in triglycerides, are unable to traverse the fenestrae of HSEC that have undergone capillarization due to aging, a process that may lead to postprandial hypertriglyceridemia [89]. It has been proposed that the ability of the liver to remove dietary cholesterol via the sieve function of the sinusoidal fenestrae plays a key role in the pathogenesis of atherosclerosis [18].

››

the cells and the lack of an underlying basement membrane are features distinguishing HSEC from other endothelial cells. Several adhesion molecules expressed at the surface of the HSECs capture leukocytes in the passing blood. Production of NO by the constitutive expression of eNOS regulates the sinusoidal vascular tone. HSECs are efficient presenters of bacterial antigens to T cells, this activity is decreased in response to endo­toxin. HSECs have a high capacity for endocytosis of macromolecules. In cirrhotic states, HSECs change; they have less fenestrae, secrete ECM components and produce less NO. HSECs are particularly sensitive to ischemia/ reperfusion injury.

Multiple Choice Questions 1. Which statement about the fenestrae of the HSEC is correct? (a) The chylomicrons are too large to passé through the fenestrae (b) The number of fenestrae increases as the sinusoids reach the central vein (c) The diameter of the fenestra decreases as the sinusoids reach the centra vein (d) The fenestrae decreases with aging (e) All the above are true 2. HSEC have been shown to (a) Express numerous selectins (b) Express less VCAM-1 in case of inflamma­ tion (c) Lack ICAM-1 (d) Express adhesion molecules such as VAP-1 (e) Have no expression of MHC class II 3. Which statement is true. (a) In cirrhotic states, production of NO via eNOS is diminished (b) Caveolin stimulates eNOS to produce NO

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(c) Every cell type in the liver can generate NO via eNOS (d) IFN-g impairs the induction of iNOS in HSEC (e) TBF-a induces iNOS in HSEC 4. Regarding the pathophysiology of HSEC, what is true: (a) Rising serum hyaluronate levels correlate with decreased endocytotic capabilities of HSEC (b) HSEC activity as antigen presenting cells is increased in response to endotoxin (c) Both size and number of fenestrae increase with chronic alcohol ingestion. (d) Bacterial antigens are more efficiently removed by HSEC mediated endocytosis in states of chronic ethanol ingestion (e) Serum hyaluronate levels are low during acute rejection 5. During ischemia/reperfusion injury (a) HSEC are particularly resistant cells (b) Changes that occurs during reperfusion are more apoptotic than necrotic (c) HSEC in the pericentral areas are more affected (d) Calcium seems protective (e) Fasting may increase HSEC apoptosis due to ischemia/reperfusion

Selected Readings   1. Wisse E, Braet F, Luo D, et al (1996) Structure and function of sinusoidal lining cells in the liver. Toxicol Pathol 24:100–111. this review summarizes the biology and function of HSEC and its neighboring cell types.   2. Selzner N, Rudiger H, Graf R, Clavien PA (2003) Protective strategies against ischemic injury of the liver. Gastroenterology 125:917–936. this review outlines approaches to protect the hepatic microcirculation from ischemic injury.   3. Shah V (2001) Cellular and molecular basis of portal hypertension. Clin Liver Dis 5:629–644. this review outlines mechanisms of endothelial cell pathobiology relevant to portal hypertension.   4. Shah V, Kamath PS (2003) Nitric oxide in liver transplantation: pathobiology and clinical implications. Liver Transplant 9:1–11. this review article outlines NO signaling as it relates to biologic processes relevant to liver transplantation.

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R. C. Huebert and V. H. Shah ment for hepatocyte activation in different mouse strains. Biochem Pharmacol 53(9):1339–1345 68. Laskin DL, Gardner CR, Price VF, Jollow DJ (1995) Modulation of macrophage functioning abrogates the acute hepatotoxicity of acetaminophen. Hepatology 21(4): 1045–1050 69. DeLeve LD (1994) Dacarbazine toxicity in murine liver cells: a model of hepatic endothelial injury and glutathione defense. J Pharmacol Exp Ther 268(3):1261–1270 70. DeLeve LD (1996) Cellular target of cyclophosphamide toxicity in the murine liver: role of glutathione and site of metabolic activation. Hepatology 24(4):830–837 71. DeLeve LD, Wang X, Kuhlenkamp JF, Kaplowitz N (1996) Toxicity of azathioprine and monocrotaline in murine sinusoidal endothelial cells and hepatocytes: the role of glutathione and relevance to hepatic venoocclusive disease. Hepatology 23(3):589–599 72. Shulman HM, Fisher LB, Schoch HG, Henne KW, McDonald GB (1994) Veno-occlusive disease of the liver after marrow transplantation: histological correlates of clinical signs and symptoms. Hepatology 19(5):1171–1181 73. DeLeve LD, McCuskey RS, Wang X et al (1999) Char­ acterization of a reproducible rat model of hepatic venoocclusive disease. Hepatology 29(6):1779–1791 74. DeLeve LD, Shulman HM, McDonald GB (2002) Toxic injury to hepatic sinusoids: sinusoidal obstruction syndrome (veno-occlusive disease). Semin Liver Dis 22(1):27–42 75. DeLeve LD, Ito Y, Bethea NW, McCuskey MK, Wang X, McCuskey RS (2003) Embolization by sinusoidal lining cells obstructs the microcirculation in rat sinusoidal obstruction syndrome. Am J Physiol Gastrointest Liver Physiol 284(6):G1045–G1052 76. Yokoi Y, Nakamura S, Muro H, Baba S (1994) Functional abnormalities of sinusoidal endothelial cells in rats with acute liver rejection. Transplantation 57(1):27–31 77. Okouchi Y, Sasaki K, Tamaki T (1994) Ultrastructural changes in hepatocytes, sinusoidal endothelial cells and macrophages in hypothermic preservation of the rat liver with University of Wisconsin solution. Virchows Arch 424(5):477–484 78. Caldwell-Kenkel JC, Currin RT, Tanaka Y, Thurman RG, Lemasters JJ (1989) Reperfusion injury to endothelial cells following cold ischemic storage of rat livers. Hepatology 10(3):292–299 79. Sun X, Kimura T, Kobayashi T et al (2001) Viability of liver grafts from fasted donor rats: relationship to sinusoidal endothelial cell apoptosis. J Hepatobiliary Pancreat Surg 8(3):268–273 80. McKeown CM, Edwards V, Phillips MJ, Harvey PR, Petrunka CN, Strasberg SM (1988) Sinusoidal lining cell damage: the critical injury in cold preservation of liver allografts in the rat. Transplantation 46(2):178–191 81. Clavien PA (1998) Sinusoidal endothelial cell injury during hepatic preservation and reperfusion. Hepatology 28(2): 281–285 82. Caldwell-Kenkel JC, Currin RT, Tanaka Y, Thurman RG, Lemasters JJ (1991) Kupffer cell activation and endothelial cell damage after storage of rat livers: effects of reperfusion. Hepatology 13(1):83–95 83. Gao W, Bentley RC, Madden JF, Clavien PA (1998) Apoptosis of sinusoidal endothelial cells is a critical mecha-

5  Hepatic Sinusoidal Endothelial Cells nism of preservation injury in rat liver transplantation. Hepatology 27(6):1652–1660 84. Kohli V, Selzner M, Madden JF, Bentley RC, Clavien PA (1999) Endothelial cell and hepatocyte deaths occur by apoptosis after ischemia-reperfusion injury in the rat liver. Transplantation 67(8):1099–1105 85. Montalvo-Jave EE, Escalante-Tattersfield T, OrtegaSalgado  JA, Pina E, Geller DA (2008) Factors in the pathophysiology of the liver ischemia-reperfusion injury. J Surg Res 147(1): 153–159

91 86. Le Couteur DG, Fraser R, Cogger VC, McLean AJ (2002) Hepatic pseudocapillarisation and atherosclerosis in ageing. Lancet 359(9317):1612–1615 87. Le Couteur DG, Warren A, Cogger VC et al (2008) Old age and the hepatic sinusoid. Anat Rec (Hoboken) 291(6): 672–683 88. Cogger VC, Mross PE, Hosie MJ, Ansselin AD, McLean AJ, Le Couteur DG (2001) The effect of acute oxidative stress on the ultrastructure of the perfused rat liver. Pharmacol Toxicol 89(6):306–311

6

Extracellular Matrix Scott L. Friedman

Introduction The hepatic extracellular matrix (ECM) is a complex network of macromolecules that not only provides cells with an extracellular scaffold but also plays an important role in the regulation of cellular activities [1, 2]. In a normal liver, the ECM comprises less than 3% of the relative area on a tissue section and approximately 0.5% of the wet weight [3]. In addition to Glisson’s capsule, ECM is found mainly in the portal tracts and the central veins. Small amounts of ECM, the perisinusoidal matrix, are also found in the subendothelial space of Disse. The sinusoids are lined by fenestrated endothelial cells which lack an electrondense basement membrane (BM), which facilitates the bidirectional flow of plasma between sinusoidal lumen and the hepatocytes. The strategic position of the perisinusoidal matrix at the interface between blood and the epithelial components of the liver explains why quantitative or qualitative change of ECM may significantly influence hepatic function [4]. Greater understanding of the structure and function of the ECM in the liver is vital not only for defining new therapeutic targets, but also for replicating functions of liver ex vivo using tissue engineering approaches in the hope of developing liver assist devices [5–7]. Generation of ECM, or fibrogenesis, occurs in response to different injuries to the liver. This is a wound-healing response that is reversible by ECM degradation upon elimination of the primary insult

[8–10]. Transition of a normal to a fibrotic liver involves both quantitative and compositional changes in the ECM [11]. Intense research over the past 20 years in clarify cellular sources ECM has established that hepatic stellate cells play a central role in the process [12] (see Part 1, Chap. 3), along with other resident and recruited fibrogenic cell types, including fibrocytes, epithelial mesenchymal transition, and resident portal fibroblasts [13, 14] (see Part 1, Chap. 3). In contrast, other cell types including hepatocytes and hepatic sinusoidal cells have only a modest contribution to the overall production of ECM. While the ECM has long been considered a relatively static scaffold, it is now increasingly recognized as a dynamic substratum that guides cellular and organ behavior. Moreover, the physical properties conferred by fibrotic ECM, specifically its stiffness [15], have been exploited in the development of noninvasive technologies to determine the stage of fibrosis in patients with chronic liver diseases [16]; thus, its relevance is now emerging in the clinical realm [17]. Particularly, the transient elastography [18] and magnetic resonance elastography [19] are appealing noninvasive approaches that correlate matrix stiffness with the extent of ECM accumulation, which may specifically reflect pericellular fibrosis [16]. Because acute hepatitis and edema can also increase stiffness in the absence of ECM deposition [20, 21], these technologies are only valuable when there is chronic but not acute liver injury.

Components of the ECM in Liver S. L. Friedman Division of Liver Diseases, Mount Sinai School of Medicine, 1425 Madison Avenue, Box 1123, New York, NY 10029-6574, USA e-mail: [email protected]

The components of ECM in liver include collagens, non­collagenous glycoproteins, glycosaminoglycans (GAGs), proteoglycans, matrix-bound growth factors,

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_6, © Springer-Verlag Berlin Heidelberg 2010

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Fig. 6.1  Molecules of the hepatic extracellular matrix. The hepatic ECM consists of collagens, noncollagenous glycoproteins, elastin, GAGs, and proteoglycans. ECM-bound molecules include fibrin, plasmin, urokinase type plasminogen activator (upa), plasminogen activator inhibitor (PAI)-1, tissue transglutaminase, lysyl oxidase, growth factors/cytokines, metalloproteinases (MMPs), and tissue inhibitors of metalloproteinase (TIMP)-3. In addition, transmembrane proteoglycans may serve as cell surface receptors are cleaved by proteases, becoming ECM-bound, including CTGF connective

tissue growth factor; a/bFGF acidic/basic fibroblast growth factor; GM-CSF granulocyte macrophage colony stimulating factor; IFN interferon; IL interleukin; HGF hepatocyte growth factor; KGF keratinocyte growth factor; OSM oncostatin M; PDGF plateletderived growth factor; SPARC secreted protein acidic and rich in cysteine (synonymous with osteonectin or BM-40); TGF transforming growth factor; TSP thrombospondin; VEGF vascular endothelial growth factor; BM basement membrane; and S sulfate. Reproduced from [1], with permission from the author and publisher

and matricellular proteins (Fig. 6.1). In the normal liver, the dense, interstitial ECM is largely confined to the capsule, around large vessels and in the portal triad. The perisinusoidal matrix, on the other hand, is composed of both an interstitial and a BM-like low-density ECM.

molecular classes: the relatively homogeneous group of fibril-forming collagens (collagens I, II, III, V, and XI) and the rather heterogeneous group of nonfibrillar collagens [22, 23]. In addition, noncollagenous proteins including fibronectin (FN) may be essential nucleators that promote the assembly of collagen fibrils [23]. The fibril-forming collagens, which consist of a triple helix of approximately 300  nm in length and 1.5  nm in diameter, self-assemble into fibrils in the extracellular space through the cleavage of terminal procollagen peptides by C-propeptidase and N-propeptidase. Types I, III, and V are the main

The Collagen Scaffold More than 20 genetically distinct collagens have been identified, and are grouped into two main

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components in the dense interstitial ECM in portal tract and central vein wall of normal liver. Among the nonfibrillar collagens, types IV, VI, VIII, XIV, XIX, XV, and XVIII are found in the liver with different locations and functions. Both fibril-forming and nonfibrillar collagens are found in the perisinusoidal matrix. These include fibrillar types I, III, and V, microfibrillar collagen VI, BM collagens IV and XVIII, and FACIT (fibril-associated collagens with interrupted triple helices) collagen [1].

Proteoglycans Proteoglycans belong to a distinct subset of noncollagenous glycoproteins that contain GAG side chains. They interact with other ECM molecules via specific GAG-binding domains in these molecules. By virtue of such properties, they regulate matrix architecture and spatial arrangement of structural polymers. They bind cytokines and growth factors and thus control their availability and biological activities. Proteoglycans identified in liver include aggrecan, fibromodulin, decorin, biglycan, perlecan, betaglycan, glypicans, and syndecan-1, -2, -3, and -4. Aggrecan belongs to the family of proteoglycans characterized by an N-terminal globular domain that interacts with hyaluronan and a C-terminal selectin domain. Fibromodulin, decorin, and biglycan are characterized by a protein core composed of leucine-rich repeats. These provide a horseshoe-like structure, which favors protein–protein interaction. In fact, these small proteoglycans bind transforming growth factor b-1 (TGF b-1), a potent fibrogenic cytokine, to stellate cells. In a normal liver tissue, biglycan and decorin are detected in the space of Disse, while in the liver of patients with chronic hepatitis, they are also found in fibrotic areas (Fig. 6.2). Betaglycan, syndecans, and glypicans are membraneanchored heparin sulfate proteoglycans. Betaglycan is the type III TGF b receptor while syndecans may function as coreceptors of cytokines. They are transmembrane proteins with an amino-terminal extracellular domain, a single transmembrane domain, and a short cytoplasmic tail. Glypicans are integral membrane proteoglycans that are anchored via glycosyl phosphatidylinositol. Overexpression of glypican-3 is seen in hepatocellular carcinoma, which lends evidence to the suggestion that glypican-3 can regulate cell growth [24].

Laminin Laminin is a noncollagenous glycoprotein, which, together with perlecan, nidogen, and collagen IV, is one of the main components of the BM. It is composed of three disulphide linked chains (a, b, and g) with a characteristic cross shape. A number of homologues of these chains have been discovered – five a chains, three b chains, and three g chains. Not all possible combinations of the three chains are used, and so far 12 distinct laminin isoforms have been identified. Among them, at least four may be found in human liver. Laminin is important not only in its structural role in the BM but also in its range of effects on cellular activities, namely cell adhesion, cell migration, and cell differentiation. It mediates the cell–matrix interaction via binding to the integrin receptors [25].

Fibronectin FN is a multifunctional glycoprotein that plays crucial roles in many cellular functions. It is a major component of normal and fibrotic hepatic ECM. FN molecules found in ECM are insoluble (“cellular fibronectin,” cFN) while “plasma fibronectin” (pFN), are soluble [1]. FN has a domain structure, consisting of three internally homologous repeats, termed types I, II, and III. The repeats are assembled into different functional domains that bind to various ligands such as collagen, heparin, fibrin, and integrin. FN mRNA is posttranscriptionally modified by alternative splicing at three variable regions of type III homology, EIIIA, EIIIB, and EIIICS. Two mRNA isoforms are generated by either inclusion or exclusion of EIIIA and EIIIB, respectively. The IIICS region has three subdomains (CS1, CS5, and the portion between these) and five isoforms may arise by exon subdivisions within this region. Alternative spliced variants have different biological properties. For example, only cFN has EIIIA and EIIIB regions. In vivo, FN forms fibrils stabilized by intermolecular disulfide bridges. The polymerization process is driven by cell surface receptors, especially by integrin a5b1. Hepatocytes are the major source of pFN while cFN is produced by hepatocytes, activated stellate cells, and sinusoidal endothelial cells [26]. In rats with normal

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NORMAL LIVER “Portal Triad” Hepatocytes

Sinusoidal space of Disse Bile duct

Stellate cell

Portal vein

Sinusoid

Central vein

Sinusoidal endothelial cells

Kupffer cell

Hepatic arteriole

FIBROTIC LIVER Stellate cell activation and proliferation

Loss of hepatocyte microvilli

Distortion of veins

Loss of endothelial fenestrations

Increase in fibrilforming collagen in Space of Disse KEY Fibril-forming collagens (Types I,III,V) Basement membrance collagens (Types IV,VI) Glycoconjugates (laminin, FN, glycosaminoglycans, tenascin)

Fig. 6.2  Extracellular matrix and cellular alterations in hepatic fibrosis. The normal liver (top) has modest amounts of fibrilforming collagen throughout the acinus, with basement membrane components concentrated in the subendothelial space of Disse. In fibrotic liver (bottom) fibril-forming matrix largely

replaces basement membrane matrix. This is associated with stellate cell activation and results in distortion of central and/or hepatic veins and loss of microvilli and endothelial fenestrations. Reprinted from [95]. Used with permission

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liver, none of the FN isoforms is present in quiescent stellate cells. In response to injury, sinusoidal endothelial cells express EIIIA fibronectin. This is a critical early event as the EIIIA segment provokes activation of stellate cells. The activated stellate cells in turn synthesize an EIIIA-containing FN themselves [26].

In the liver, expression of SPARC is upregulated during fibrogenesis and hepatocarcinogenesis. Stellate cells are the main source of SPARC as shown both in vivo and in culture [29, 30]. Tenascin and osteopontin have both been found in normal and fibrotic livers and hepatocellular carcinoma [31–33].

Matricellular Proteins

Changes in ECM from Normal to Fibrotic Liver

Matricellular proteins are a group of matrix proteins that modulate cell–matrix interaction and cell function but do not contribute directly to the formation of structural elements [27, 28]. Members of this group of distinct molecules include SPARC (secreted protein acidic and rich in cysteine, also termed osteonectin), thrombospondins (TSP1 and TSP2), osteopontin, tenascin-C, tenascin-X, and CCN family of proteins (CYR61, CTGF [connective tissue growth factor]). Studies in various cell types demonstrate that these molecules are capable of sequestering growth factors (e.g., SPARC and PDGF), binding ions (e.g., osteonectin and Ca2+), inhibiting (TSP1) or clearing proteases (TSP2) and activating cytokines (e.g., TSP2 and TGF-b1). They regulate cell adhesion, migration, chemotaxis, proliferation, and apoptosis. Furthermore, complex effects are also exerted on ECM synthesis and collagen assembly. a

Fig.  6.3  Changes in ECM and stellate cells during hepatic fibrosis. (a) In normal sinusoids stellate cells, located between sinusoidal endothelium and hepatocytes, are quiescent and contain vitamin A droplets. (b) In chronic liver injury, a fibrillar

Hepatic fibrosis is associated with a significant change in both the quantity and composition of the ECM. Total collagen content increases by three to tenfold [34]. The perisinusoidal low-density ECM is gradually replaced by a high-density ECM with accumulation of fibrillar collagens (types I and III) and an electron-dense BM [35]. There is also an increase in glycoproteins, proteoglycans, and GAGs and a shift from BM-type proteoglycans (heparan sulfate) to interstitial-type (dermatan sulfates and chondroitin sulfates) [36]. Changes in ECM are associated with disappearance of endothelial fenestrations, a process termed “sinusoid capillarization.” In vitro studies suggest that interstitial matrix induces loss of fenestrations whereas physiologically derived BM maintains them [37] (Fig. 6.3). Complex interactions exist between the cellular components of liver and the ECM. With the b

matrix accumulates in the subendothelial space, produced primarily by activated stellate cells. This matrix leads to loss of hepatocyte microvilli and reduced sinusoidal porosity. Reprinted from [96] with permission

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modification of the ECM microenvironment, cellular functions and phenotypes are inevitably affected [13]. This is evidenced by the loss of microvilli in hepatocytes in the fibrotic liver and compromised synthetic activity of hepatocytes when deprived of BM matrix [13]. Meanwhile, the high-density matrix activates stellate cells, which further perpetuates the process of fibrogenesis [38]. On the other hand, quiescent stellate cells cultured on BM-like matrix derived from Englebreth– Holm-Sarcoma remain nonproliferative and nonfibrogenic [38, 39]. A recent study further showed that BM-like matrix (Matrigel) can induce quiescence of activated stellate cells, suggesting that restoration of normal ECM in the liver might down regulate fibrogenesis by restoring stellate cells to a quiescent state as well [40]. Interactions between stellate cells and the microenvironment may be further modulated by the activity of metalloproteinases, which can remodel the microenvironment [41]. Specific components of the ECM can regulate cellular activities [42]. One example is EIIIA segment of fibronectin as described above. Another molecule is microfibrillar collagen VI, the expression of which is upregulated in liver fibrosis. Collagen VI stimulates DNA synthesis and inhibits apoptotic cell death in stellate cells in vitro [1].

Pathways of Cell–Matrix Interaction The Integrin Family The ECM interacts with cells via matrix receptors on the cell membrane; these cell membrane adhesion complexes link ECM to cell function [43, 44]. The best-characterized matrix-membrane integrators are the integrin family of heterodimeric transmembrane receptors [45–48]. Integrin receptors are composed of a and b subunits, and at least 18 a and 8 b subunits are currently known. Structural analysis of integrins has greatly aided efforts to define therapeutic targets [49]. The various combinations result in over 20 functional integrin dimers with different specificities. Their globular head domain binds to ligands which include components of the ECM and cell adhesion molecules. Most integrin ligands contain an Arg–Gly–Asp (RGD) trip-

S. L. Friedman

eptide sequence, which is necessary but is not sufficient for signaling. The integrins are important not only in their adhesive function but also their roles in modulating signal transduction pathways downstream of other receptors [50]. Modulation of signaling pathways takes place via a number of mechanisms. Cell adhesion may result in change in shape and tension of the cell and nucleus via the cytoskeleton, which in turn influences gene expression [51]. Meanwhile, integrins may affect signal transduction via parallel activation of pathways that synergize at the level of phosphorylation of proteins. In hepatic stellate cells, the key fibrogenic cells in liver injury, integrin linked kinase plays a vital role [52]. Signals generated by other receptors are also enhanced due to clustering of ECM-bound integrins in the plane of the membrane. The activated integrins recruit signaling molecules that form complexes called focal adhesion complexes [53]. Examples of such molecules are caveolin, paxillin, and tyrosine kinases such as fyn and focal adhesion kinase (FAK). Further downstream, the complexes are associated with other kinases and adaptor molecules. Clustering of these proteins results in an amplification of the extracellular signal. Lastly, integrins may cluster and transactivate signaling pathways involving receptor tyrosine kinases. An example of such functional cooperation is that between the PDGF pathway and integrin signaling as demonstrated by clustering of ligand-activated PDGF-b receptors in areas corresponding to focal adhesion complexes [54]. A complex web of crosstalk thus exists between the integrin pathways and signaling mechanisms of growth factor receptors. Normal adult human hepatocytes express low levels of three integrin dimers: a collagen and laminin receptor, a1b1; a FN receptor, a5b1; and a tenascin receptor, a9b1. On the other hand, receptors a1b1, a2b1, a5b1, and a6b4 have been identified in cultured stellate cells [55, 56]. In experimental liver fibrosis, there is upregulation of laminin-binding integrins a6b1, a2b1, and aVb8 [57] and FN-binding a5b1 [58]. Functionally, integrin antagonism by the soluble integrin recognition sequence pentapeptide GRGDS in rat stellate cells disturbs actin stress fiber formation and tyrosine phosphorylation of FAK caused by adhesion to ECM. Similarly, blockade of the vitronectin receptor aVb3 abrogates PDGF mediated migration of activated stellate cells [59].

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Recent studies have specifically identified upregulation of aVb6 integrin in biliary fibrosis models and cholestatic human disease [60, 61], and thus efforts to block this integrin have important therapeutic potential [62]. Involvement of this integrin molecule in liver fibrosis is particularly appealing since a similar role has been uncovered in models of pulmonary fibrosis, where it activates latent TGFb1 [63]. Thus, the fibrotic response of the liver to biliary injury may be functionally and structurally distinct from parenchymal injury, involving not only a unique matrix receptor expression, but also a larger dependence on epithelial mesenchymal transition as a source of fibrogenic cells, driven by [64] hedgehog signaling [65]. Integrins are also integral to cancer biology [45, 66], including hepa­ tocarcinogenesis.

ADAM Molecules A relatively new family of ECM membrane receptors, ADAM (“a disintegrin and metalloproteinase domain”) [67] also contribute to hepatic fibrosis [68]. Based on its components, the molecules can provide both protease activity as well as adhesive functions. In the liver, the ADAMSTS-1 gene has been identified in endothelial cells [69] whereas ADAMSTS13 is expressed by hepatic stellate cells [70, 71]. Intracellular links to membrane bound ADAM molecules are now being uncovered, for example RACK1, a receptor for activated protein kinase C, which binds to ADAM12 [72].

Discoidin Domain Receptors Another matrix receptor with a potential role in liver fibrosis is discoidin domain receptor-2 (DDR2) [73, 74]. DDR2 is a tyrosine kinase receptor that responds to ECM ligands but not to soluble peptide factors. It is activated primarily by collagen type I, and to a lesser extent, by collagen types II, III, and V. DDR2 is induced during stellate cell activation, and the phosphorylated receptor mediates growth stimulation and MMP2 production in response to type I collagen; thus DDR2 can stimulate degradation of normal liver ECM via MMP2, while it is further upregulated by accumulating

interstitial collagen, thereby establishing a positive feedback loop. The importance of DDR2 in liver fibrosis is substantiated by a recent study showing its presence at elevated levels in the small bile ducts of patients suffering from primary biliary cirrhosis [75]. DDRs also play a role in epithelial-mesenchymal transition [76].

Growth Factors in ECM In addition to its structural role and direct interaction with cells, ECM also regulates cell function indirectly via modulation of the availability and activity of growth factors including PDGF, TGF-b, CTGF [77] vascular endothelial growth factor [78], and HGF [1,  79] (Fig. 6.4). Proteoglycans such as decorin, biglycan, fibromodulin, and GAGs are the main ECM components that bind growth factors and cytokines. Proteoglycans can interact with growth factors either via their core proteins or via their GAG side chains. For example, decorin or biglycan binds TGF-b by their protein cores but interacts with HGF through the heparan sulfate. Other ECM components such as FN and laminin bind tumor necrosis factor-a (TNF-a), while collagen binds PDGF, HGF, and interleukin-2 (IL-2) [1]. Binding of survival factors by interstitial matrix may prevent apoptosis of hepatocytes in liver that have acquired DNA damage, thereby perpetuating the expansion of cells with mutations and genomic instability. This observation may explain why cancer is more likely in livers that are cirrhotic, particularly in patients with hepatitis C. While protecting such factors from proteolysis, the ECM also controls their release through the actions of proteases and their inhibitors, resulting in further modulation of their activities.

ECM and the Stem Cell Niche An intriguing body of evidence has begun to suggest that the ECM may provide a vital link to stem cell growth and activity in normal and injured liver. Anatomically, hepatic stellate cells, which are the key

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chemotaxis& proliferation of epithelial cells angiogenesis

collagen & heparan sulfate

PDGF

HGF KGF

chemotaxis& proliferation of ECM-producing cells

heparan sulfate

collagen

chemotaxis& differentiation of inflammatory cells

collagen

bFGF PDGF

VEGF CTGF

induction activation

heparan sulfate MCP-1 RANTES

OsM IL-2

M-CSF

IL-3 IFN-γ

GM-CSF

TGF-β

ECM production

Fig. 6.4  Binding of growth factors by ECM. The ECM can localize and store growth factors (see also Fig. 6.1), sequestering them for release by controlled proteolysis. Reproduced from [1], with permission from the author and publisher

source of hepatic ECM, surround putative stem cells in injured liver [80, 81]. Moreover, stellate cells may themselves have pluripotent potential in which the ECM could play a regulatory role [82]. Related studies have indicated that stellate cells provide vital growth factors that support hepatoblast growth [83]. A molecular link to ECM has been provided by evidence that an isoform of CTFG with a novel FN binding site may be particularly vital for expansion of oval cells in rodent liver [84], which are an epithelial progenitor cell type. The intersection of stellate cells, ECM, and stem cells is likely to be an important area of investigation in the coming years.

Metalloproteinases and Their Inhibitors Since the ECM components are highly stabilized and have cross-linked molecules, they can only be broken down by a specific family of enzymes, the matrix metalloproteinases (MMP). The MMP family comprises 25 different calcium and zinc-dependent enzymes divided into five broad categories: interstitial collagenases,

gelatinases, stromelysins, membrane-type MMP (MT-MMP), and metalloelastase. This functional classification is somewhat arbitrary as there is overlap in activities among categories. The activity of MMPs is antagonized by a group of proteins, the tissue inhibitors of the MMP family (TIMP) [85, 86]. The net activity of MMPs and TIMPs determines the rate of ECM degradation. Stellate cells are one key source of MMPs in the liver. In early primary culture, stellate cells express MMP-3 (a stromelysin) and MMP-2 but not TIMP [87, 88]. Although this matrix-degrading phenotype could be a result of the isolation process, it might also reflect the early phase of “pathological” matrix degradation in vivo after injury, during which the normal subendothelial matrix and BM are damaged. With prolonged culture of stellate cells, MMP-1/MMP-13 is downregulated while the expression of TIMP-1, TIMP-2, MMP-2, and MMP-14 is enhanced [85]. In vivo studies in rodent and human specimens have concurred with these findings by demonstrating increased TIMP expression associated with fibrosis [87]. According to one proposed model, increased MMP-2 and MMP-14 production by stellate cells causes

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degradation of the pericellular matrix. This results in an altered stellate cell–matrix interaction and further cellular activation [87]. Meanwhile, as the expression of TIMP is increased, degradation of newly synthesized collagen is inhibited, with consequent accumulation of ECM. Resolution of fibrosis, on the other hand, is associated with degradation of fibrillar ECM and restoration of normal hepatic architecture. In carbon tetrachloridetreated rats, liver fibrosis regresses after cessation of treatment. The process is associated with marked reduction in TIMP activity and an approximately fivefold increase in hepatic collagenase activity [87]. MMP-1/MMP-13 derived either from stellate or Kupffer cells, together with MMP-14 and MMP-2, contribute to the fibrolysis leading to restoration of normal histology [89, 90]. MMP-9 has additionally emerged as a key regulator of stellate cell functions [41].

Summary

›› ECM is both an important structural scaffold ››

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›› Conclusions The hepatic ECM can no longer be seen as an inert structural element of the liver. Dynamic changes in its composition and quantity take place in response to external stimuli. This plasticity and responsiveness serve important physiologic functions, as typified by the wound-healing response of fibrogenesis and associated angiogenesis [91–94]. Stellate cells and related fibrogenic cells are the predominant cells responsible for producing the components of ECM as well as the enzymes that break down the ECM. Injurious stimuli directly or indirectly through hepatocytes or Kupffer cells activate stellate cells via various signaling pathways, resulting in the synthesis of these components. Yet, interaction between ECM and their surrounding cells is bidirectional. The biological activities contained in the ECM components in addition to the growth factors sequestered regulate cellular functions in a multitude of ways. Further research and better understanding of this complex web of interactions and its molecular basis may eventually facilitate the development of new therapies for chronic liver diseases and carcinoma.

›› ›› ››

and a dynamic structure regulating cell and organ function. Key components include a large number of collagens (both fibrillar and nonfibrillar), matricellular proteins proteoglycans, and glycoproteins, including FN and laminin. Marked qualitative and quantitative changes in ECM accompany progressive injury and fibrosis. These changes favor loss of hepatocyte function and activation of hepatic stellate cells, the key resident fibrogenic cell in liver. Signals from the ECM are transduced through the cell membrane by receptors, the best characterized of which are the integrin family. Integrins are widely divergent in their ECM specificity, localization in normal and fibrotic liver, and downstream intracellular pathways. ECM is also an important reservoir of bound growth factors, whose activity on cells may be induced by release into the pericellular milieu through the activity of matrix degrading proteases. Protease activity in turn is regulated in part by the relative concentration and interaction with specific inhibitors known as TIMPs. Recent data implicate ECM in the regulation of the stem cell niche, with more data likely to emerge. Greater understanding of the structure, interactions and cellular effects of ECM is leading to important clinical advances in diagnosis and therapy of liver disease.

Multiple Choice Questions 1. Extracellular matrix (ECM) is composed primarily of: (a) Collagens (b) Proteoglycans (c) Glycoproteins (d) All of the above

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2. Which of the following features of ECM is NOT true: (a) A scaffold to support cells (b) Fixed in composition at the time of birth (c) A dynamic regulator of cell function (d) Increased accumulation may lead to increased liver stiffness 3. Which one answer is true about integrins: (a) There is one major type in fibrotic liver (b) They consist of a triple helix (c) They may be targets for therapies (d) They signal unidirectionally from ECM to the cell 4. Metalloproteinases are: (a) Highly insoluble aggregates (b) A family of matrix degrading enzymes (c) Stably expressed in health and disease (d) Too complicated to understand 5. Which of the following can enhance fibrogenesis by activated stellate cells: (a) EIIIA fibronectin (b) TIMP1 (c) Transient elastography (d) Extraterrestrial life forms

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S. L. Friedman 10. Friedman SL, Bansal MB (2006) Reversal of hepatic fibro­ sis – fact or fantasy? Hepatology 43(2 Suppl 1):S82–S88 11. Guo J, Friedman SL (2007) Hepatic fibrogenesis. Semin Liver Dis 27(4):413–426 12. Friedman SL (2008) Hepatic stellate cells–protean, multifunctional, and enigmatic cells of the liver. Physiol Rev 88(1):125–172 13. Friedman SL (2008) Mechanisms of hepatic fibrogenesis. Gastroenterology 134(6):1655–1669 14. Schuppan D, Afdhal NH (2008) Liver cirrhosis. Lancet 371(9615):838–851 15. Wells RG (2008) The role of matrix stiffness in regulating cell behavior. Hepatology 47(4):1394–1400 16. Wong GL et al (2008) Assessment of fibrosis by transient elastography compared with liver biopsy and morphometry in chronic liver diseases. Clin Gastroenterol Hepatol 6(9): 1027–1035 17. Kettaneh A et al (2007) Features associated with success rate and performance of fibroscan measurements for the diagnosis of cirrhosis in HCV patients: a prospective study of 935 patients. J Hepatol 46(4):628–634 18. Castera L et al (2005) Prospective comparison of transient elastography, fibrotest, APRI, and liver biopsy for the assessment of fibrosis in chronic hepatitis C. Gastroenterology 128(2):343–350 19. Talwalkar JA et al (2008) Magnetic resonance imaging of hepatic fibrosis: emerging clinical applications. Hepatology 47(1):332–342 20. Arena U et al (2008) Acute viral hepatitis increases liver stiffness values measured by transient elastography. Hepatology 47(2):380–384 21. Coco B et al (2007) Transient elastography: a new surrogate marker of liver fibrosis influenced by major changes of transaminases. J Viral Hepat 14(5):360–369 22. Kavitha O, Thampan RV (2008) Factors influencing collagen biosynthesis. J Cell Biochem 104(4):1150–1160 23. Kadler KE, Hill A, Canty-Laird EG (2008) Collagen fibrillogenesis: fibronectin, integrins, and minor collagens as organizers and nucleators. Curr Opin Cell Biol 20(5):495–501 24. Zhu ZW et al (2001) Enhanced glypican-3 expression differentiates the majority of hepatocellular carcinomas from benign hepatic disorders. Gut 48(4):558–564 25. Bosman FT, Stamenkovic I (2003) Functional structure and composition of the extracellular matrix. J Pathol 200(4): 423–428 26. Jarnagin WR et al (1994) Expression of variant fibronectins in wound healing: cellular source and biological activity of the EIIIA segment in rat hepatic fibrogenesis. J Cell Biol 127(6 Pt 2):2037–2048 27. Bornstein P (2002) Cell–matrix interactions: the view from the outside. Methods Cell Biol 69:7–11 28. Bornstein P, Sage EH (2002) Matricellular proteins: extracellular modulators of cell function. Curr Opin Cell Biol 14(5):608–616 29. Frizell E et al (1995) Expression of SPARC in normal and fibrotic livers. Hepatology 21(3):847–854 30. Nakatani K et al (2002) Expression of SPARC by activated hepatic stellate cells and its correlation with the stages of fibrogenesis in human chronic hepatitis. Virchows Arch 441(5):466–474 31. Tokairin T et al (2008) Osteopontin expression in the liver with severe perisinusoidal fibrosis: autopsy case of Down

6  Extracellular Matrix  103 syndrome with transient myeloproliferative disorder. Pathol Int 58(1):64–68 32. El-Karef A et al (2007) Expression of large tenascin-C splice variants by hepatic stellate cells/myofibroblasts in chronic hepatitis C. J Hepatol 46(4):664–673 33. Lee SH et al (2004) Effects and regulation of osteopontin in rat hepatic stellate cells. Biochem Pharmacol 68(12): 2367–2378 34. Rojkind M, Giambrone MA, Biempica L (1979) Collagen types in normal and cirrhotic liver. Gastroenterology 76(4):710–719 35. Hahn E et al (1980) Distribution of basement membrane proteins in normal and fibrotic human liver: collagen type IV, laminin, and fibronectin. Gut 21(1):63–71 36. Gressner AM, Bachem MG (1990) Cellular sources of noncollagenous matrix proteins: role of fat-storing cells in fibrogenesis. Semin Liver Dis 10(1):30–46 37. McGuire RF et al (1992) Role of extracellular matrix in regulating fenestrations of sinusoidal endothelial cells isolated from normal rat liver. Hepatology 15(6):989–997 38. Friedman SL et al (1989) Maintenance of differentiated phenotype of cultured rat hepatic lipocytes by basement membrane matrix. J Biol Chem 264(18):10756–10762 39. Sohara N et al (2002) Reversal of activation of human myofibroblast-like cells by culture on a basement membrane-like substrate. J Hepatol 37(2):214–221 40. Gaca MD et al (2003) Basement membrane-like matrix inhibits proliferation and collagen synthesis by activated rat hepatic stellate cells: evidence for matrix-dependent deactivation of stellate cells. Matrix Biol 22(3): 229–239 41. Han YP et al (2007) A matrix metalloproteinase-9 activation cascade by hepatic stellate cells in trans-differentiation in the three-dimensional extracellular matrix. J Biol Chem 282(17):12928–12939 42. Somasundaram R, Schuppan D (1996) Type I, II, III, IV, V, and VI collagens serve as extracellular ligands for the isoforms of platelet-derived growth factor (AA, BB, and AB). J Biol Chem 271(43):26884–26891 43. Berrier AL, Yamada KM (2007) Cell–matrix adhesion. J  Cell Physiol 213(3):565–573 44. Lock JG, Wehrle-Haller B, Stromblad S (2008) Cell–matrix adhesion complexes: master control machinery of cell migration. Semin Cancer Biol 18(1):65–76 45. Danen EH (2005) Integrins: regulators of tissue function and cancer progression. Curr Pharm Des 11(7):881–891 46. McCall-Culbreath KD, Zutter MM (2008) Collagen receptor integrins: rising to the challenge. Curr Drug Targets 9(2): 139–149 47. Avraamides CJ, Garmy-Susini B, Varner JA (2008) Integrins in angiogenesis and lymphangiogenesis. Nat Rev Cancer 8(8):604–617 48. Silva R et al (2008) Integrins: the keys to unlocking angiogenesis. Arterioscler Thromb Vasc Biol 28(10):1703–1713 49. Xiong JP, Goodman SL, Arnaout MA (2007) Purification, analysis, and crystal structure of integrins. Methods Enzymol 426:307–336 50. Stupack DG (2007) The biology of integrins. Oncology 21(9 Suppl 3):6–12 51. Lelievre SA et al (1998) Tissue phenotype depends on reciprocal interactions between the extracellular matrix and the structural organization of the nucleus. Proc Natl Acad Sci U S A 95(25):14711–14716 52. Shafiei MS, Rockey DC (2006) The role of integrin-linked kinase in liver wound healing. J Biol Chem 281(34): 24863–24872

53. Melton AC et al (2007) Focal adhesion disassembly is an essential early event in hepatic stellate cell chemotaxis. Am J Physiol Gastrointest Liver Physiol 293(6):G1272–G1280 54. Carloni V et al (2000) Tyrosine phosphorylation of focal adhesion kinase by PDGF is dependent on ras in human hepatic stellate cells. Hepatology 31(1):131–140 55. Carloni V et al (1996) Expression and function of integrin receptors for collagen and laminin in cultured human hepatic stellate cells. Gastroenterology 110(4):1127–1136 56. Pinzani M, Marra F, Carloni V (1998) Signal transduction in hepatic stellate cells. Liver 18:2–13 57. Levine D et al (2000) Expression of the integrin a8b1 during pulmonary and hepatic fibrosis. Am J Pathol 156(6):1927–1935 58. Znoyko I, Trojanowska M, Reuben A (2006) Collagen binding a2b1 and a1b1 integrins play contrasting roles in regulation of Ets-1 expression in human liver myofibroblasts. Mol Cell Biochem 282(1–2):89–99 59. Patsenker E et al (2007) Pharmacological inhibition of the vitronectin receptor abrogates PDGF-BB-induced hepatic stellate cell migration and activation in vitro. J Hepatol 46(5):878–887 60. Wang B et al (2007) Role of avb6 integrin in acute biliary fibrosis. Hepatology 46(5):1404–1412 61. Popov Y et al (2008) Integrin avb6 is a marker of the progression of biliary and portal liver fibrosis and a novel target for antifibrotic therapies. J Hepatol 48(3):453–464 62. Patsenker E et al (2008) Inhibition of integrin avb6 on cholangiocytes blocks transforming growth factor-b activation and retards biliary fibrosis progression. Gastroenterology 135(2):660–670 63. Munger JS et al (1999) The integrin avb6 binds and activates latent TGF b1: a mechanism for regulating pulmonary inflammation and fibrosis. Cell 96(3):319–328 64. Omenetti A et al (2008) Hedgehog signaling regulates epithelial-mesenchymal transition during biliary fibrosis in rodents and humans. J Clin Invest 118(10):3331–3342 65. Omenetti A et al (2008) The hedgehog pathway regulates remodelling responses to biliary obstruction in rats. Gut 57(9):1275–1282 66. Hehlgans S, Haase M, Cordes N (2007) Signalling via integrins: implications for cell survival and anticancer strategies. Biochim Biophys Acta 1775(1):163–180 67. Primakoff P, Myles DG (2000) The ADAM gene family: surface proteins with adhesion and protease activity. Trends Genet 16(2):83–87 68. Kesteloot F et al (2007) ADAM metallopeptidase with thrombospondin type 1 motif 2 inactivation reduces the extent and stability of carbon tetrachloride-induced hepatic fibrosis in mice. Hepatology 46(5):1620–1631 69. Diamantis I et al (2000) Cloning of the rat ADAMTS-1 gene and its down regulation in endothelial cells in cirrhotic rats. Liver 20(2):165–172 70. Zhou W et al (2005) ADAMTS13 is expressed in hepatic stellate cells. Lab Invest 85(6):780–788 71. Niiya M et al (2006) Increased ADAMTS-13 proteolytic activity in rat hepatic stellate cells upon activation in vitro and in vivo. J Thromb Haemost 4(5):1063–1070 72. Bourd-Boittin K et al (2008) RACK1, a new ADAM12 interacting protein. Contribution to liver fibrogenesis. J Biol Chem 283(38):26000–26009 73. Labrador JP et al (2001) The collagen receptor DDR2 regulates proliferation and its elimination leads to dwarfism. EMBO Rep 2(5):446–452

104 74. Olaso E et al (2001) DDR2 receptor promotes MMP-2mediated proliferation and invasion by hepatic stellate cells. J Clin Invest 108(9):1369–1378 75. Mao TK et al (2002) Elevated expression of tyrosine kinase DDR2 in primary biliary cirrhosis. Autoimmunity 35(8): 521–529 76. Maeyama M et al (2008) Switching in discoid domain receptor expressions in SLUG-induced epithelial–mesenchymal transition. Cancer 113(10):2823–2831 77. Gressner OA, Gressner AM (2008) Connective tissue growth factor: a fibrogenic master switch in fibrotic liver diseases. Liver Int 28(8):1065–1079 78. Yoshiji H et al (2003) Vascular endothelial growth factor and receptor interaction is a prerequisite for murine hepatic fibrogenesis. Gut 52(9):1347–1354 79. Asano Y et al (2007) Hepatocyte growth factor promotes remodeling of murine liver fibrosis, accelerating recruitment of bone marrow-derived cells into the liver. Hepatol Res 37(12):1080–1094 80. Roskams T (2006) Different types of liver progenitor cells and their niches. J Hepatol 45(1):1–4 81. Roskams T (2008) Relationships among stellate cell activation, progenitor cells, and hepatic regeneration. Clin Liver Dis 12(4):853–860; ix 82. Kordes C et al (2007) CD133+ hepatic stellate cells are progenitor cells. Biochem Biophys Res Commun 352(2):410–417 83. Kubota H, Yao HL, Reid LM (2007) Identification and ­characterization of vitamin A-storing cells in fetal liver: implications for functional importance of hepatic stellate cells in liver development and hematopoiesis. Stem Cells 25(9): 2339–2349 84. Pi L et al (2008) Connective tissue growth factor with a novel fibronectin binding site promotes cell adhesion and migration during rat oval cell activation. Hepatology 47(3): 996–1004

S. L. Friedman 85. Benyon RC, Arthur MJ (2001) Extracellular matrix degradation and the role of hepatic stellate cells. Semin Liver Dis 21(3):373–384 86. Han YP (2006) Matrix metalloproteinases, the pros and cons, in liver fibrosis. J Gastroenterol Hepatol 21(Suppl 3): S88–S91 87. Iredale JP (2001) Hepatic stellate cell behavior during resolution of liver injury. Semin Liver Dis 21(3):427–436 88. Iredale JP (2007) Models of liver fibrosis: exploring the dynamic nature of inflammation and repair in a solid organ. J Clin Invest 117(3):539–548 89. Duffield JS et al (2005) Selective depletion of macrophages reveals distinct, opposing roles during liver injury and repair. J Clin Invest 115(1):56–65 90. Fallowfield JA et al (2007) Scar-associated macrophages are a major source of hepatic matrix metalloproteinase-13 and facilitate the resolution of murine hepatic fibrosis. J Immunol 178(8):5288–5295 91. Lee JS et al (2007) Sinusoidal remodeling and angiogenesis: a new function for the liver-specific pericyte? Hepatology 45(3):817–825 92. Semela D et al (2008) Platelet-derived growth factor signaling through ephrin-b2 regulates hepatic vascular structure and function. Gastroenterology 135(2): 671–679 93. Taura K et al (2008) Hepatic stellate cells secrete angiopoietin 1 that induces angiogenesis in liver fibrosis. Gastroenterology 135(5):1729–1738 94. Yang L et al (2008) Sonic hedgehog is an autocrine viability factor for myofibroblastic hepatic stellate cells. J Hepatol 48(1):98–106 95. Friedman SL, Millward-Sadler H, Arthur MJP (1992) Liver fibrosis and cirrhosis. Wright’s liver and biliary disease, 3rd edn. WB Saunders, London 96. Friedman SL, Arthur MJP (2002) Reversing hepatic fibrosis. Sci Med 8(4):194–205

7

Platelets: A New Cell Type in Liver Physiology Mickael Lesurtel and Pierre-Alain Clavien

List of Abbreviations 5-HT 5-HIAA 5-HTP CTL EGF HGF HSC I/R NASH PDGF SERT TGF-b TPO TPH

5-hydroxytryptamine = serotonin 5-hydroxyindol acetic acid 5 hydroxytryptophan cytotoxic T lymphocytes response epidermal growth factor hepatocyte growth factor hepatic stellate cells ischemia/reperfusion nonalcoholic steatohepatitis platelet-derived growth factor serotonin reuptake transporter transforming growth factor-b thrombopoietin tryptophan hydroxylase

What Are Platelets? Platelets are the smallest type of blood cells, which are only fragments of bone marrow megakaryocyte cytoplasm and are biconvex discs, approximately 3 mm in diameter. The development of megakaryocytes and production of platelets are unique processes. Megakaryocyte maturation involves nuclear duplication without cell division, resulting in giant cells. Cytoplasmic organelles are organized into domains representing nascent platelets, demarcated by a

P.-A. Clavien () Swiss HPB Center, Department of Visceral and Transplant Surgery, University Hospital of Zurich, Rämistrasse, 100, 8091, Zürich, Switzerland e-mail: [email protected]

network of invaginated plasma membranes. Within the marrow, megakaryocytes are localized next to the sinusoidal walls and this facilitates the exit of large segments of cytoplasm into the circulation. The fragmentation of megakaryocyte cytoplasm into individual platelets then results from the shear forces of circulating blood [1]. Thrombopoietin (TPO) is the dominant hormone controlling megakaryocyte development, but many cytokines and hormones take part, including interleukins 3, 6, and 11 [2]. Platelets have no nucleus, but do have several important organelles. These include the open canalicular ­system, a complex arrangement of membranes within the platelet cytoplasm communicating with the extracellular space [1]. This provides a membrane store for rapid mobilization to the surface during morphological changes (Fig.  7.1). This system also promotes rapid discharge of granule contents from the platelet. There are two classes of secretary granules. The first type consists of a-granules, approximately 80 per platelet, and contain many important molecules within their lumen and membrane wall [3]. On activation, they fuse with the open canalicular system and surface membrane, discharging their contents. The second type consists of dense bodies (around five per platelet) which are additional storage organelles whose contents are also released on activation. Table 7.1 summarizes the content of platelet a-granules and dense bodies. Among the numerous molecules stored in platelets, serotonin has emerged as a crucial molecule involved in the pathophysiology of the liver. Though serotonin is present in the diet, most of it is metabolized before entering the bloodstream. In enterochromaffin cells and neurons, but not in platelets, serotonin is synthesized from the essential amino acid tryptophan by two enzymatic steps. First, hydroxylation of tryptophan by  the enzyme tryptophan hydroxylase (TPH) (the

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Fig. 7.1  Resting platelets are smooth and disc shape (left); activated platelets have an irregular shape with many protruding pseudopodia (right)

r­ ate-limiting step) produces 5 hydroxytryptophan (5-HTP). The second enzymatic step is decarboxylation of 5-HTP by the enzyme l-aromatic amino acid decarboxylase producing serotonin [4]. Platelets (and neurons) possess a high affinity serotonin uptake mechanism, and platelets become loaded with serotonin as they pass through the intestinal circulation, where the local concentration is relatively high. Therefore, about 95% of serotonin found in blood is stored in platelets. Serotonin in tissues can be very rapidly metabolized, mainly as a result of the activity of monoamine oxidase. In the kidney and the liver, the enzymes, monoamine oxidase and aldehyde dehydrogenase convert serotonin to 5-hydroxyindole acetic acid (5-HIAA), which is excreted in the urine. There are two isoforms of TPH [5]. TPH1 is present in the periphery, especially in the duodenum, while TPH2 is present exclusively in the brain coded by a different gene [6]. Serotonin does not have the ability to cross the membrane lipid bilayer; it has therefore to be bidirectionally transported. The serotonin reuptake transporter (SERT) is the major protein responsible for the uptake and release of serotonin. Serotonin is present in high concentration in platelets, where it accumulates from the plasma via the active transport system SERT. In laminar blood flow, platelets circulate at the periphery of the vessel lumen and can rapidly detect endothelial injury. In response, structural and biochemical changes occur, promoting thrombus formation and vessel wall repair. Approximately 15–20% of daily platelet turnover (7,000 platelets per microliter per day) is consumed maintaining vascular integrity and the mean platelet survival is 9 days [1]. The spleen continually but transiently sequesters about a third of

the circulating platelets. Splenomegaly, particularly when caused by passive congestion due to increased portal venous pressure, greatly increases the fraction of platelets retained in splenic sinusoids, without decreasing overall platelet survival time. This retention causes the mild thrombocytopenia associated with liver cirrhosis and portal hypertension. Most platelets are removed from circulation after senescence, but a constant small fraction is continually removed by involvement in the maintenance of vascular integrity. Although the normal platelet count is 150–300 × 109 per liter, a count of just 10 × 109 per liter is usually sufficient to prevent bleeding [7]. Does this discrepancy merely reflect great redundancy in this homoeostatic system or do platelets have other important roles? Apart from their well-known action on haemostasis, there is increasing evidence that platelets are also involved in many other mechanisms such as inflammation [8], atherosclerosis [9], antimicrobial defense [10], angiogenesis [11], tissue repair, and tumor cell growth [12]. This chapter focuses on the role of platelets that has recently emerged in the physiopathology of the liver.

Ischemia/Reperfusion Injury Hepatic ischemia/reperfusion (I/R) injury is the major source of morbidity associated with liver resection under vascular occlusion, i.e., Pringle maneuver, or after liver transplantation [13, 14]. The liver can be subjected to three forms of ischemia, namely cold (or hypothermic), warm (or normothermic), and rewarming [15]. Cold ischemia occurs almost exclusively in

7  Platelets: A New Cell Type in Liver Physiology Table 7.1  Contents of platelet a-granules and dense bodies Dense bodies a-Granules Serotonin Chemokines Platelet factor 4 b-Thromboglobulin Connective tissue activating protein 3 Neutrophil activating peptide 2 Regulated up on activation, normal T cell expressed and presumably secreted (RANTES) Macrophage inflammatory protein 1a Adhesive proteins

Adrenaline Noradrenaline Dopamine Histamine Bivalent cations, e.g., Ca2+ and Mg2+ Adenosine 5¢-diphosphate Adenosine 5¢-triphosphate Guanosine 5¢-diphosphate Guanosine 5¢-triphosphate P-selectin in the membrane

Thrombospondin Fibrinogen Fibronectin Vitronectin Growth factors Platelet-derived growth factor Transforming growth factor a (TGF-a) TGF-b Epidermal growth factor Insulin-like growth factor Vascular endothelial growth factor A Vascular endothelial growth factor C Basic fibroblast growth factor Hepatocyte growth factor Interleukin 8 Immunoglobulins IgG, IgM, IgA, IgE Cationic proteins P-selectin von Willebrand factor Coagulation factors Factors V and VIII Glycoprotein IIb–IIIa Plasminogen activator  inhibitor 1

the transplant setting where it is applied intentionally to reduce metabolic activities of the graft while the organ awaits implantation. Warm ischemia occurs in a variety of situations including transplantation, trauma, shock, and liver surgery, when hepatic inflow occlusion

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(Pringle maneuver) or inflow and outflow (total vascular exclusion) are induced to minimize blood loss while dividing the liver parenchyma. Rewarming ischemia typically occurs during manipulation of the graft (e.g., ex situ split liver preparation) or during the period of implantation of the graft when the cold liver is subjected to room or body temperature while performing vascular reconstruction. A point to note is that injury to the liver cells after any type of ischemia is detected mainly after reperfusion when oxygen supply and blood elements are restored. While inhibition of platelet activity reduces ischemic tissue injury in the heart [16], lung [17] and pancreas [18], little is known about the impact of platelets and serotonin on I/R injury of the liver.

Cold Ischemia During liver transplantation, platelets are rapidly sequestrated in the liver graft after reperfusion. In this process, some platelets adhere to the sinusoidal endothelium, which has been activated as a result of cold and warm ischemia. The extent of platelet activation to the activated endothelium has been shown to correlate with organ function, both in animal models and in human transplant patients [19–21]. The direct contribution of platelets to I/R injury was first suggested by Cywes et  al. [19] in experimental ­studies using isolated perfused rat livers. A positive correlation was seen among the duration of cold ischemia, degree of platelet adhesion to the activated liver endothelium, and injury of the perfused rat liver. When livers were ex vivo perfused with activated platelets, hepatic injury was increased in comparison to reperfusion with unactivated platelets, and this indicated that platelet activation is directly responsible for injury to the liver and that the correlation between the extent of platelet deposition and organ damage is not just reflecting enhanced organ damage as a result of cold ischemia. The interaction of platelets with activated endothelium resulting in organ damage is mediated by adhesion molecules such as selectins and integrins, which are highly expressed on activated platelets and endothelial cells. In agreement with this observation, it has been demonstrated that platelets induce apoptosis of sinusoidal endothelial cells, especially upon reperfusion [21]. Platelets seem to act in concert with leukocytes and Kupffer cells, and a triangular interaction between

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these cell types has been demonstrated in the mechanisms of reperfusion injury [22]. Kupffer cells, the resident liver macrophages, interact with circulating blood cells. These cells rapidly activate after reperfusion of the ischemic liver and mediate injury in interactions with leukocytes and platelets. Platelets and leukocytes need functional Kupffer cells to mediate injury, and Kupffer cells are much less harmful in the absence of platelets and leukocytes [22, 23].

Warm Ischemia In contrast to cold ischemia, warm ischemia is tolerated poorly and rapidly leads to the death of hepatocytes [24]. This severe injury of the hepatocytes probably is preceded by massive death of endothelial cells [25]. The role of Kupffer cells, adherent leukocytes and platelets remains an area of active investigation in the warm ischemic liver. Mice deficient for P-selectin, an adhesion molecule critical to the postischemic platelet-endothelial cell interaction display reduced platelet and neu­ trophil sequestration and a better survival following  warm ischemia [26]. In addition, the inhibition of  platelet adhesion by the administration of anti-­ fibrinogen antibody decreases short-term liver injury after ischemia [27]. However, these experimental approaches also inhibit leukocyte-mediated effects; thus, the specific contribution of platelets to warm I/R injury of the liver could not be known. Recently, our team studied the role of platelets in warm I/R injury using models of impaired platelet function and immune thrombocytopenia (leukocyte and erythrocyte counts unaffected) [28]. Neither abrogation of platelet aggregation nor platelet depletion reduced postischemic tissue injury. Instead, postischemic inflammation, as well as liver regeneration and consequently tissue repair, were strikingly impaired. In particular, platelet-derived serotonin mediates hepatocyte proliferation, which is an integral component of postischemic tissue repair. These findings point to a novel role of platelets in hepatic wound healing (Fig. 7.2). The impact of rewarming on the structural integrity of the liver and the mechanism of this type of injury is understood poorly. It probably reflects a combination of cold and warm injury.

Fig.  7.2  Electron micrograph representing warm injury in the mouse liver. Endothelial cell swelling accompanied by hepatocyte necrosis (nH) is observed. Accumulation of polymorphonuclear leukocytes (PMN) and platelets are present in the sinusoids. R red blood cell (Adapted from Selzner et al. [13])

Liver Regeneration Role of Platelets The human body responds to partial hepatectomy by reestablishment of the original volume of the organ, thanks to the unique ability of the liver cells to replicate and increase the remnant segments. The typical scenario of liver volume restoration commences with hyperplasia of various types of intrahepatic cells followed by a phase of cellular hypertrophy [29]. This phenomenon is traditionally known as liver regeneration despite the fact that, in purely biological terminology, neither hyperplasia nor hypertrophy is a synonym for regeneration [30]. Liver regeneration encompasses activation of many intra- and extracellular molecules and pathways [29] (Fig.  7.3). The current concept is that liver regeneration after partial hepatectomy involves a large number of genes organized into three networks: cytokines, growth factor, and metabolic. Marked redundancy exists among them [30]. Many factors pertaining to liver regeneration have been extensively studied, among which platelets and platelet-derived serotonin have recently made exciting advances. Precise integration of growth signals is required for full and

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Fig. 7.3  Pathways of liver regeneration initiated by major hepatectomy. After hepatectomy, nonparenchymal cells, such as stellate cells, Kupffer cells, leukocytes, and platelets, are activated by soluble factors, such as vascular endothelial growth factor and lipopolysaccharide. Interaction between activated vascular components, including platelets, leukocytes, sinusoidal endothelial cells, and Kupffer cells, results in the release of tumor necrosis factor a, interleukin-6, and serotonin. The cytokines cause a priming of the remnant hepatocytes, and concurrently, extracellular proteases such as urokinase-type plasminogen activator convert inactive hepatocyte growth factor to its active form. Inactive hepatocyte growth factor, which is secreted by stellate cells, is a mitogen that induces hepatocyte proliferation. Matrix

metalloproteases convert membrane-bound transforming growth factor a (TGF-a) into the soluble form. In an autocrine loop, TGF-a, along with endothelial growth factor, signals through the endothelial growth factor receptor. The cytokines and the growth factors act in concert to initiate the reentry of quiescent hepatocytes (in the G0 phase) into the cell cycle from the G1 phase to the S phase, resulting in DNA synthesis and hepatocyte proliferation. To signal the end of proliferation, TGF-b blocks further replication. The metabolic load resulting from the loss of hepatocytes is indicated by the accumulation of bile acids in the blood. The bile acids enter the hepatocytes and drive bile acid receptors such as the farnesoid X receptor, resulting in increased protein and DNA synthesis (Adapted from Clavien et al. [29])

synchronized regeneration. Failure to activate these signaling cascades may result in delay of the onset of regeneration, inadequate recovery of liver volume and eventually clinical signs of liver failure. The liver and platelets display a very intimate, albeit complex, interconnection [31]. The liver plays a critical role even during the synthesis of platelets from megakaryocytes through TPO. TPO, the most important growth factor in the regulation of megakaryocyte development and platelet production, is produced mainly in the liver and kidney [31]. Hence platelets are not expected to function properly in diseased liver states [32]. A number of proteins, which induce opposing effects on liver regeneration, are present in platelets. For instance, platelets harbor important growth factors

for execution of liver regeneration, e.g., hepatocyte growth factor HGF [33]. Inversely, platelets contain transforming growth factor-a TGF-a [34], which is required for the termination of liver regeneration. Thus, it is plausible that platelets may participate in orchestrating liver regeneration through harmonized stimulation and inhibition of growth-related signals. Until 2006, it was unclear whether platelets are promoters, inhibitors of, or not even active contributors to, liver regeneration. Many in vitro studies demonstrated that platelets contain several growth factors (Table 7.1) which may theoretically contribute to the process of liver regeneration [35]. However, the only in vivo study on the role of platelets in liver regeneration in rats failed to identify a correlation between platelets and

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liver regeneration [36]. In a previous study on regeneration of liver in rats, it was noted that splenectomy increases platelet counts and accelerates liver regeneration via an unclear mechanism [37]. To investigate more closely the role of platelets in liver regeneration in mice, our team applied inhibitors of platelet function which remarkably reduced liver regeneration. In a second step, we depleted platelets to less than 5% of their normal count by applying a platelet-specific antibody. After 70% liver resection, these mice exhibited significantly impaired liver regeneration, suggesting that a factor contained in platelets may be required to induce or maintain liver regeneration [38]. In another study, thrombocytotic mice exhibited increased liver regeneration while a thrombocytopenic group showed impaired regeneration [39]. Matsuo et al. reported that direct contact between platelets and hepatocytes is necessary for the proliferative effect [40]. These authors concluded that platelet-hepatocyte contact initiates signal transduction involved in growth factor activation. HGF, VEGF and insulin-like growth factor-1 were found to contribute chiefly to hepatocyte proliferation [40] (Fig.  7.4). Murata et  al. have recently demonstrated that platelets promote liver regeneration even under conditions of Kupffer cell depletion, by stimulating HGF and insulin-like growth factor-1 expression [41]. More recently, the same team investigated the effect of thrombocytosis on liver regeneration after

Fig.  7.4  Transmission electron microscopic photograph of residual liver of a thrombocytotic group 5  min after hepatectomy. Magnification ×7,500. Arrowheads indicate a platelet protruding into Disse’s space through the porosity of a flattened process in a sinusoidal endothelial cell (Adapted from Murata et al. [39])

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90% hepatectomy in mice, which was previously considered fatal. They reported that, under the condition of thrombocytosis, platelets accumulated in a large quantity in the liver remnant, liver regeneration occurred and the survival rate was significantly higher compared to mice with normal platelet counts [41].

Role of Platelet-Derived Serotonin Platelets maintain a sort of mutual cooperation with serotonin. Platelets are responsible for picking up serotonin from the gut and lungs and provide the main peripheral storehouse of serotonin [42]. On the other hand, serotonin has a mitogenic influence on megakaryocytes via 5-HT2 receptor [43]. In their dense bodies platelet store and release serotonin, which is able to act as a growth factor [44, 45]. In vitro, serotonin is a potent mitogen and stimulates mitosis of smooth muscle cells [44]. The role of serotonin in liver regeneration in hepatocyte cell cultures has been reported where serotonin caused a dose-dependent increase in (3H)-thymidine incorporation into hepatic DNA in the presence of insulin and epidermal growth factor EGF [46]. The 5-HT2A and 2C receptors appear to mediate mitogenic effects in fibroblasts [47, 48], and the 5-HT2B receptor is involved in the development of the heart [49] and the enteric nervous system [50]. Furthermore, in transfected fibroblasts and renal mesangial cells a mitogenic cross-talk of serotonin receptors 5-HT2A and 5-HT2B has been shown with EGF receptor and platelet-derived growth factor (PDGF) receptor, respectively [51, 52]. Very recently an additional crosstalk of SERT and PDGFb-receptor in smooth muscle cells has been described leading to proliferation and migration [53]. With the exception of a mention in the Russian radiobiology literature about 20 years ago [54, 55], the effect of serotonin on hepatic regeneration was unknown. We recently showed that platelet-derived serotonin is involved in the initiation of liver regeneration [38]. In thrombocytopenic mice, a serotonin agonist restored the deficient hepatic proliferation. The mRNA expression of serotonin receptor subtypes 5-HT2A and 2B increased after partial hepatectomy, and antagonists of these receptors inhibited liver regeneration. Furthermore, knockout mice, lacking peripheral serotonin due to an absence of the rate-limiting

7  Platelets: A New Cell Type in Liver Physiology

synthetic enzyme TPH1, exhibited an impaired liver regeneration after partial hepatectomy. This failure of regeneration was rescued by reloading serotonin-free platelets with the injection of the serotonin precursor 5-HTP. Papadimas et al. [56] investigated the effect of 5-HT2 blockage by ketanserin on liver regeneration after partial hepatectomy in rats. Administration of ketanserin can arrest liver regeneration only when administrated close to G1/S transition point, which suggests that serotonin may be a cofactor for DNA synthesis.

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and prostacyclin, two important endothelial-derived ­platelet inhibitors, may contribute to defective platelet activation in vivo [69]. Although spontaneous bleeding caused by low platelet counts or defective platelet function in cirrhosis is an infrequent event, low platelet counts become clinically relevant when performing liver biopsies, liver transplantation or giving myelosuppressive agents as antiviral treatment (interferon or cytostatics) [70].

Platelet Involvement in Liver Diseases Platelets in Chronic Liver Disease Alterations in Platelets in Patients with Liver Disease Primary hemostasis in patients with acute or chronic liver disease is characterized by two major alterations: thrombocytopenia and functional platelet defects [57, 58]. Thrombocytopenia in patients with liver disease is at least partly caused by an increased sequestration of platelets in the spleen due to portal hypertension, resulting in congestive splenomegaly [59]. In patients with advanced liver cirrhosis, up to 90% of the platelets can become sequestrated in the spleen. However, even in these patients, the peripheral platelet count decreases only moderately. Other mechanisms that explain the reduced platelet count in patients with liver disease are a reduced production of TPO by the diseased liver [60] and a reduced platelet half-life, which is possibly related to autoantibodies [61]. In patients with alcohol-induced cirrhosis, defective platelet production occurs as a result of folic acid deficiency or toxic effects of ethanol on megakaryocytopoiesis [62]. Defective platelet function may be a result of an acquired storage pool defect [63], defective transmembrane signal transduction [64], decreased levels of arachidonic acid required for thromboxane A2 production in the membrane [65], decreased levels of functional platelet receptors as a result of proteolysis by plasmin [66], the presence of abnormal high-density lipoprotein [67], or a reduced hematocrit [68]. In patients with cirrhosis and concomitant renal failure, a  uremic thrombocytopathia may develop on top of this. Moreover, increased production of nitric oxide

Less well studied is the recent observation that platelets can contribute to the progression of liver disease like fibrosis, viral hepatitis, nonalcoholic steatohepatitis (NASH) and cholestatic liver disease.

Platelets and Viral Hepatitis The role of platelets in the development of hepatitis and liver fibrosis has been speculative for a long time. Activation of blood platelets in chronic hepatitis and liver cirrhosis has been shown through measurement of P-selectin expression or platelet factor-4 [71, 72]. Some observations indicate that platelets play key roles in adaptive responses to microbial and antigen challenge, and participate with leukocytes in inflammatory reactions [73]. More recently, important studies showed that platelets are directly involved in infection and virus-induced hepatitis [74, 75]. Hepatic damage resulting from infection by noncytopathic viruses, such as hepatitis B and C virus, is a consequence of the antigen-specific cytotoxic T lymphocytes response (CTL). It has been shown that activated platelets contribute to CTL-mediated liver immunopathology independent of procoagulant function [74]. Indeed, platelet depletion reduces intrahepatic accumulation of virus-specific CTLs and organ damage in mouse models of acute viral hepatitis. More deeply, Lang et  al. investigated the role of platelet-derived serotonin during virus-induced CD8+ T cell-dependent immunopathological hepatitis in mice [75]. During infection with a noncytopathic virus, serotonin release is responsible for sinusoidal microcirculatory failure. Serotonin-dependent alterations in sinusoidal microcirculation account for viral-induced liver cell damage

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by delaying the infiltration of activated virus-specific CD8+ T cells and thereby protracting viral control. They concluded that platelet-derived serotonin aggravates viral control and therefore favors the onset of chronic hepatitis.

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livers from a CCl4-cirrhosis model. Finally, HSCs were shown to express a functional SERT and to participate in both active uptake and release of serotonin. HSCs could respond to serotonin in a profibrogenic manner and 5-HT2 receptor antagonists may also be used for the treatment of liver disease.

Platelets and Liver Fibrosis/Cirrhosis Platelets in Nonalcoholic Steatohepatitis Many studies investigated the impact of liver cirrhosis on platelets, but the role of platelets on liver cirrhosis is less well known. Once again, the increasing knowledge about platelet-derived serotonin has opened new fields and platelets may act on liver fibrosis process through serotonin. Different studies have shown decreased concentrations of intra-platelet serotonin in cirrhotic patients [63, 76, 77]. Beaudry et al. [78] found higher plasma levels of  serotonin in patients with cirrhosis, whereas levels of the serotonin metabolite 5-hydroxyindol acetic acid (5-HIAA) were significantly decreased. Very recently, Watanabe et al. investigated whether platelets reduce liver fibrosis and promote liver regeneration in fibrotic liver in mice [79]. Liver fibrosis was induced by carbon tetrachloride (CCl4) intoxication and thrombocytosis was achieved by giving TPO or splenectomy. Their results suggest that platelets attenuate the progression of liver fibrosis by decreasing expression of transforming growth factor-b (TGF b) and increasing expression of metalloprotinease-9, as well as by promoting liver regeneration. Whether platelets directly interact with hepatic stellate cells (HSC) and whether platelets directly influence some functions of HSCs still remains unknown. HSC is recognized as one of the key mediator in the progression of hepatic fibrosis [80, 81]. In a normal healthy liver, HSCs regulate sinusoidal blood and the traffic of macromolecules across the space of Disse, and also act as a store for vitamin A. Following chronic injury, HSCs are involved in hepatic wound healing and fibrosis. Serotonin was found to attenuate apoptosis and to costimulate proliferation of HSCs in the presence of PDGF in cultured rat HSCs [82]. Proliferation was inhibited and apoptosis rate was increased with 5-HT2B receptor antagonists. The transcription of connective tissue growth factor was increased in rat HSCs after incubation with serotonin. 5-HT2B receptor expression was associated with fibrotic tissue in immunohistological stainings of rat

Very recently, our group suggested that platelet-derived serotonin degradation plays a major role in the pathogenesis of non alcoholic steatohepatitis (NASH) [83]. NASH is thought to result from a two hit process [84]. The first hit is the hepatocellular accumulation of fatty acids, which sensitizes the liver to further injury. Oxidative stress acts as a second hit, leading to lipid peroxidation, mitochondrial damage (megamitochondria), hepatocellular injury (ballooning, Mallory bodies), and finally to chronic inflammation and fibrosis. Degradation of serotonin is catalyzed by the mitochondrial enzyme monoamine oxidase A, generating 5-HIAA as well as reactive oxygen species (ROS) such as hydrogen peroxide. ROS generated by monoamine ­oxidase-mediated catabolism of serotonin were recently reported to play a pivotal role in cardiomyocyte death [85]. In a murine model of diet-induced steatohepatitis, TPH1−/− mice displayed an equal degree of steatosis, yet reduced hepatocellular injury and less severe inflammation compared to wild type mice. The difference in these two NASH-defining features were attributed to an increased uptake and catabolism of serotonin, yielding enhanced levels of ROS and lipid peroxides, which mediated hepatocellular injury by mitochondrial damage and inflammation. This was the first study disclosing SERT expression in the mouse liver, as well as in patients with NASH and provides evidence that platelet-derived serotonin plays a crucial role in the pathogenesis of steatohepatitis. SERT might offer a novel target for the prevention and treatment of NASH.

Platelets in Chronic Cholestasis Cholestasis triggers immediate liver injury and the absence of bile in the intestine facilitates bacterial translocation, which, in turn, may cause sepsis and further liver injury [86]. In a model of bile duct ligationinduced cholestasis in mice, it has been shown that

7  Platelets: A New Cell Type in Liver Physiology

platelets play an important role in cholestasis-induced liver injury by promoting leukocyte recruitment and deteriorating microvascular perfusion [87]. Indeed, depletion of platelets not only reduced hepatic recruitment of leukocytes but also protected against liver injury in cholestatic mice. Moreover, inhibition of P-selectin prevented cholestasis-induced platelet and leukocyte recruitment as well as the associated hepatocellular damage. Thus, targeting platelet accumulation may be a useful strategy against liver damage associated with obstructive jaundice. A recent study has shown the expression of the serotonin receptors 5-HT1A and 5-HT1B in rat cholangiocytes [88]. Their activation markedly inhibits the growth and choleretic activity of the biliary tree. Cholangiocytes from cholestatic rats overexpressed and oversecreted serotonin and treatment with a neutralizing antibody of serotonin enhanced cholangiocyte proliferation. The authors stressed the existence of an autocrine loop based on serotonin that limits the growth and the functional activity of the biliary tree in the course of chronic cholestasis. These findings suggested the involvement of serotonin-mediated pathways in cholestatic disease.

Conclusion Recent advances in bio-medical research have shown that platelets play a crucial role in regulating hepatic function and response to injury. Far from the idea that platelets have mainly haemostatic properties there is now increasing evidence that they also have an important role in liver pathophysiology. Among the molecule contained in platelets, serotonin is one of the molecules with a strong focus of research during the last decade, particularly in the field of liver regeneration. Serotonin, platelets and liver are unique examples of a fruitful cooperation between molecule, corpuscle and organ [89, 90]. Serotonin and platelets are essential, with a close cross-talk with the liver. The role of platelets and serotonin in liver pathophysiology may hopefully contribute to the understanding of pathways involved in liver I/R injury, liver regeneration and chronic liver diseases. Due to the availability of a number of selective agonists and antagonists for the various serotonin receptor subtypes and knockout mice lacking specific serotonin receptors, there is a real possibility for significant

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advances to be made in the field of serotonin and the liver. These studies may ultimately lead to new ­therapies that may decrease the progression of hepatic fibrosis or enhance the regenerative capacity of the liver, helping the many people suffering from chronic liver disease. In the field of liver transplantation, new innovative therapeutic strategies acting on platelets may minimize cold I/R injury and, therefore, increase the pool of liver grafts. However, future laboratory and clinical studies are needed to discover the precise action of serotonin and platelets in these different processes.

Summary

›› Beside their well-known role in primary hae-

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mostasis, there is increasing evidence that platelets are also involved in many pathways of the physiopathology of the liver. In cold hepatic ischemia/reperfusion injury, platelets act in concert with leukocytes and Kupffer cells and induce endothelial cell apoptosis. After partial hepatectomy, platelets promote liver regeneration mainly through plateletderived serotonin release. While acute or chronic liver diseases are characterized by thrombocytopenia and functional platelet defects, platelets could be involved in liver disease like fibrosis, viral hepatitis, NASH and cholestatic liver disease. A better understanding of the role of platelets and serotonin in liver physiopathology may open new strategies to treat patients suffering from chronic liver disease.

Multiple Choice Questions 1.  Regarding platelets, which one of these features is wrong? (a) Platelets are the smallest type of blood cells (b) They come from megakaryocyte maturation (c) They contain a nucleus and two types of secretary granules (d) Their mean duration of life is 9 days (e) In human, normal platelet count is 150– 300 × 109 per liter

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2.  Which is the correct sentence concerning serotonin metabolism? (a) Serotonin is synthesized in platelets (b) Serotonin is synthesized from the amino acid tryptophan (c) Only 5% of serotonin found in blood is store in platelets (d) Serotonin can cross the cell membrane without transporter (e) Products of serotonin degradation are excreted in bile 3.  During cold liver ischemia/reperfusion injury: (a) Platelets interact with hepatic stellate cells (b) Platelets adhere to the sinusoidal endothe­lium (c) Platelets induce necrosis of sinusoidal endothe­ lial cells (d) Platelets are sequestrated in the spleen (e) Kupffer cells activation prevents liver injury 4.  After partial hepatectomy, platelets: (a) Accumulate in the spleen (b) Promotes liver regeneration through insulin release (c) Need Kupffer cells to promote liver regen­eration (d) Bind to 5-HT2A receptors on hepatocyte surface (e) Promote liver regeneration through plateletderived serotonin 5.  In patients with liver cirrhosis, platelets: (a) Usually have a normal blood count (b) Accumulate in the liver (c) Exhibit a decreased concentration of sero­tonin (d) Promote portal hypertension by sequestration in the spleen (e) Exhibit always normal haemostatic function

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8

Immune Cell Communication and Signaling Systems in Liver Disease Ricky H. Bhogal and Simon C. Afford

Introduction The relationship that exists between the different ­components of the immune system within the human liver is a highly complex and dynamic one. In recent years, there has been a rapid expansion of our knowledge of the nature of the liver as an immunologically distinct organ, with numerous unique features that have evolved as a result of its specialized physiological niche. It is now apparent that the immune system plays a crucial role in determining the progression of many liver diseases via chronic inflammatory processes that fail to resolve, through to end stage cirrhosis and on occasions, malignant diseases. The liver receives a substantial blood supply from three sources: from the abdominal aorta via the hepatic artery and two venous plexuses, the portal vein, and inferior vena cava. The liver is therefore continuously exposed to bloodborne pathogens, toxins, tumor cells, and dietary antigens as well as those that can enter the liver from the gut via the ascending biliary tree. The structural organization of the liver has profound implications for its immune function (Fig.  8.1), for review please see Racanelli et al. [1]. The healthy adult human liver contains approximately 1010 lymphocytes and because of vascular blood supply, approximately 108 peripheral blood lymphocytes pass through the liver every 24 h [2]. Lymphocytes are usually the predominant infiltrating leukocyte ­pop­ulation numerically, and are found throughout the ­parenchyma and the portal tracts as part of normal S. C. Afford () The Liver Research Group, School of Infection and Immunity, College of Medicine and Dentistry, University of Birmingham, UK e-mail: [email protected]

immunological surveillance and recirculation. The unique structure of liver architecture permits contact between lymphocytes and antigen presenting cells (APC), which promotes lymphocyte extravasations. This extravasation is further facilitated by fenestrations in the monolayer of hepatic sinusoidal endothelial cells (HSEC) that allow lymphocytes to access the space of Disse, where they can interact with the underlying extracellular matrix, hepatic stellate cells (HSC), and hepatocytes and various nonparenchymal cells. The lymphoid cells within the liver comprise of T-cells, natural killer (NK) cells, and B-cells (Fig. 8.2). T-cells can be divided into conventional and unconventional subgroups on the basis of cell surface markers known as cluster of differentiation antigens (CD). Conventional T-cells comprise CD8+ and CD4+ T-cells. Both populations display diverse T-cell populations with ab-chain T-cell receptors (TCR) that recognize antigens in the context of major histocompatibility complex (MHC) class I and II molecules, respectively. Unconventional T-cells comprise various cell types that are categorized into two major populations: those that express NK cell markers (also called NKT cells) and those that do not. Classical NK cells arise in the thymus, display a very restricted TCR (typically the TCR Va24 and Vb11 chains in humans) and recognize antigens in the context of the MHC class I molecule CD1d. They can be either CD4-positive or CD4/CD8-double negative. By contrast, nonclassical NKT cells encompass TCRab and TCRgd T-cells, do not use TCR Va14 chain and do not express the CD8b chain [3]. Classical and nonclassical NKT cells are more abundant in the liver than in other organs and constitute up to 30% of the intrahepatic lymphocyte population [4]. Importantly, NK cells represent a lymphoid population with a potent cytolytic activity and the capacity to regulate the migration and expansion of NKT cell subsets [5].

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_8, © Springer-Verlag Berlin Heidelberg 2010

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SPACE OF DISSE HSC

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Fig. 8.1  The immune cell composition of the healthy liver. Over 90% of normal liver tissue is composed of hepatocytes which are arranged in adjacent pairs forming tight junctions with canaliculi for bile transport on one side and bordering the vascular sinusoids on the other, through which antigen rich hepatic arterial and portal venous blood flows. Blood in the sinusoids is separated from hepatocytes by a fenestrated monolayer of hepatic sinusoidal endothelial cells (HSEC) and the Space of Disse. Lymphocytes along with other inflammatory cells pass

through sinusoids and are in close contact with HSECs. Resident macrophages (Kupffer cells) are also found within the sinusoids, with professional antigen presenting cells (Dendritic cells) being found largely within the portal tract. The Space of Disse contains stellate cells of myeofibroblast lineage within a loose extracellular matrix. Thus the liver architecture allows for direct and indirect interaction between many cell types. DC dendritic cell; HSC hepatic stellate cell; KC Kupffer cell; NK natural killer cell

Non-parenchymal Cells ≈ 20-40%

Lymphocytes ≈ 25%

Fig. 8.2  The relative proportions of lymphoid cells within a healthy human adult liver. Parenchymal cells or hepatocytes constitute approximately 60–80% of cells within the liver. Nonparenchymal cells constitute approximately 20–40% of cells (endothelial cells 50%, Kupffer cells 20%, lymphocytes 25%, biliary cells 5% and stellate cells <1%). The vast majority of the lymphocytes within the liver are T-cells which can be further divided into conventional and unconventional subgroups (see text for details) (adapted from [1])

T Cells ≈ 63%

NK Cells ≈ 31%

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B Cells ≈ 6%

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8  Immune Cell Communication and Signaling Systems in Liver Disease Fig. 8.3  The basic structure of a toll-like receptor (TLR). Most TLRs appear to function as homodimers, TLR2 forms heterodimers with TLR1 or TLR6, each dimer having different ligand specificity. Coreceptors are sometimes needed for full ligand sensitivity such as MD2, LBP, and CD14 in TLR4’s recognition of LPS. Four adaptor molecules are known to couple to TLRs; MyD88, Tirap, Trif, and TRAM [276–278]. These adaptor molecules activate other protein kinases and initiate intracellular signaling

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LPS LBP CD14

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Unconventional T-cells that do not express NK cell markers include the major group of TCRgd T-cells. This group represents 15–25% of the entire intrahepatic T-cell population, thereby rendering the liver one of the richest sources for these cells within the body. These T-cells possess invariant TCRs that recognize a limited range of antigens such as stress proteins and nonprotein antigens.

MyD88

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referred to as pathogen-associated molecular patterns (PAMPs). They form a receptor super family with the interleukin (IL)-1 receptor as they all share a common Toll-IL-1 receptor (TIR) domain. TLRs function as either homo- or heterodimers and may require coreceptors for full ligand sensitivity. TLRs recruit the cytosolic adaptor molecules MyD88, Tirap, TRIF, and Tram to modulate intracellular signaling (Fig.  8.3). For a comprehensive review of TLR function, please read Akira S [6].

TLR Signaling (See Chap. 9) The Toll like receptor (TLR) superfamily are an important class of at least ten cell surface receptors which provide critical molecular linkage between the innate and adaptive immune system in the liver. TLRs are a class of single membrane-spanning noncatalytic receptors that recognize structurally conserved molecules derived from microbes. They have a key role in innate immunity and are integrally linked to the initiation of the adaptive immune responses. TLRs are pattern recognition receptors (PRR) that recognize molecules that are broadly shared by pathogens but distinguishable from host/self molecules, collectively

Innate Immunity Within the Liver The innate immune system is the first line of defense against infection. Although it is capable of mounting a rapid response on potential pathogens, the innate immune system does not have the capacity of immunological memory. Innate immunity protects against pathogens via a limited repertoire of PRRs that recognize specific PAMPs expressed by potentially harmful invading pathogens [7]. Among the best defined PAMPs is the bacterial antigen Lipopolysaccarhide (LPS). PRRs are divided into three categories: secreted PRR such as

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Fig. 8.4  The potential cross-talk between different immunological cells within the liver. KC and DC are able to release a variety of cytokine-mediators after activation. The leads to the activation of NK/NKT cells which can secrete IFNg and interact with and stimulate HSEC/hepatocyte to secrete chemokines. This results in increased effector cell function and T-cell recruitment

IFN /IL-12/IL-15

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complement, membrane bound PRRs such as TLR [6], NOD-like receptors and RIG-like receptors [8], and phagocytic PRRs expressed on the surface of cells. Once the innate immune system is initiated in the liver, a complex network of interactions between immune cells is triggered. The liver is selectively enriched with cells of the innate immune system including KCs, DCs, NK, and NKT cells. KCs express an array of TLRs that can indirectly or directly recruit and activate other innate immune cells to amplify downstream signaling events. KCs can recruit NK cells into the inflammatory milieu. NK cells can modulate liver signaling pathways by regulating the local production of proinflammatory (Th1) and anti-inflammatory (Th2) cytokines based upon whether it is activated through its activating or inhibitory receptors. As inhibition dominates over activation, the threshold for NK cell activation is lowest in the absence of ligands such as MHC class I that bind to inhibitory receptors [9]. In the absence of inhibitory signals and in the presence of cytokines such as type I interferon (IFN) and type I-IFN-induced CXCL3 [10], ligation of

activating NK cell receptors results in NK cell activation. This results in target cell lysis and rapid production of IFNg which in turn stimulates HSEC and hepatocytes [11] to secrete the chemokine CXCL9, thereby recruiting T-cells to the liver. DCs and KCs also produce IL-12 which can activate NKT cells and results in Fas-mediated cell lysis [12]. Furthermore, NKT cells activation is CD1-restricted [13], which is expressed by hepatocytes and APCs (Fig. 8.4)

Dendritic Cells (DCs) Through their ability to enhance the innate immune response, DC may profoundly influence the development and subsequent shape of the hepatic immune response. DCs are the most potent APCs of the immune system and characterize the pathogencity of the invading antigen [14]. There are distinct subsets of DCs which are likely to undertake overlapping and distinct functions

8  Immune Cell Communication and Signaling Systems in Liver Disease

[15]. The subtypes include myeloid (mDC), lymphoid (lDC), and plasmacytoid (pDC) DCs. mDCs and lDCs are found in the peripheral/central vein [16] and pDCs are found in the liver parenchyma [17]. Interestingly, a variant of DC known as natural killer dendritic cell (NKDC) [18], after stimulation by IL-12 and/or Il-18, secretes IFNg and can lyse target cells and activate naïve T-cells [19]. When activated, DCs show increased phagocytosis and enhanced NK cell and T-cell activation and promote chemokine- and cytokine-dependent immune responses [20]. HSECs and KCs are the other APCs within the liver and are present in an immunologically competent state expressing costimulatory molecules and possessing the ability to secrete IFNg and IL-1 [21]. Indeed IL-1 promotes DC maturation in vitro and the lack of IL-1 leads to aberrant DC maturation with Th2 cytokine production leading to severe hepatocellular damage [22]. Therefore IL-1 secreted by KCs and HSECs may have a role in DCs development. DCs, along with NK cells, provide a critical link between the systemic innate and adaptive immune system but they would seem to be particularly important within the liver. Exposure to certain “danger signals” (typically antigens) cause DCs to cease phagocytosis and upregulate adhesion molecules and chemokine receptors that are necessary for their migration to draining lymph nodes where they can play their part in orchestration of the adaptive immune as appropriate [14]. While there is a controversy over the nature of the repertoire of TLR expression by mDC and pDC, within the human liver both appear to be able to stimulate naïve T-cells via TLR dependent mechanisms. As hepatic DCs have a low expression of costimulatory molecules, it is thought that this renders them poor T-cell activators. However, studies using hepatitis C virus (HCV) show that TLR-dependent maturation of DCs enables them to produce IL-6 and IFNb which can potentially activate NK cells and CD4+ T-cells [23]. Other studies using Francisella tularensis have shown that TLR-dependent IL-12 production by DC is important in inducing T-cell responses [24]. In steady state conditions, DCs are at an immature stage of development and induce tolergenic T-cell responses [25]. However upon TLR recognition in an inflammatory microenvironment, DC can migrate to lymph nodes and induce immune responses [26]. Whilst migrating to the coeliac lymph node, DCs maintain an immature phenotype dependent on CXCR7 [20] and the ligands CXCL19/CXCL21 [27]. Furthermore KC derived CXCL3/MIP-1a and portal tract-associated lymphoid tissue Portal Associated Lymphoid Tissue

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(PALT) derived CXCL21 (see later) also influence TLRdependent DC migration to areas of inflammation [28]. Once active DCs can stimulate the innate immune system through activation of NK cells [29] probably via TLRs subject to regulation by CD45 [30]. For review of the cross-talk between DCs and NK cells please read Brilot et al. [31]. Furthermore, DCs are also known to activate NKT and Tgd cells [32]. In an inflammatory microenvironment the TLR/DC axis may therefore emerge as a potential conductor of the entire hepatic inflammatory orchestra (Fig. 8.5). Interestingly DCs and NK cells share a common developmental pathway implying that these cells may influence each other during differentiation [33]. In an inflammatory microenvironment DCs differentiation can certainly be influenced by interactions with other immune cells particularly NK cells [34]. Granulocyte macrophage colony stimulating factor (GM-CSF) and IL-3, produced by resting NK cells, stimulate colony formation of DCs precursors while TNFa secreted after NK cell activation exert inhibitory effects on DCs [35]. Indeed DCs matured in vitro with TNFa have deficient functional roles [36]. The pivotal role of NK cells in DC maturation may imply that NK cells are vital for co-coordinating the immunological response in the liver. For instance, in vitro, in the presence of cytokines, NK cells can encourage the maturation of immature DCs [37] as shown by CD83, MHC class II and costimulatory molecule upregulation and priming of an efficient antitumor CD8+ T-cell response [38]. Further­more, coculturing CD14+ monocytes precursors with autol­ogous NK cells in the presence of IL-15 induces morphological and phenotypic changes associated with DC activation such as down regulation of CD14 and up regulation of co stimulatory molecules including CD40, CD80, CD86, DC-SIGN, and DEC205 and antigen uptake [34]. In addition, IFNa increases expression of HLA class I and II and CD86 on DC and also increases IL-12 and TNFa production [39]. CD83 expression is required for full DC activation and may require further TLR-dependent signaling [40]. Also, in addition to NK-cell-induced DC activation being dependent upon TNFa, IFNg, cell–cell contact involving NKp30 [41], KIR and NKG2A [42] is also required. Finally for a Th1 polarized response DC maturation is dependent upon NK cells [43]. This evidence supports the hypothesis that NK cells provide a cytokine milieu allowing the differentiation of DC precursors. Immature hepatic DCs express high levels of CXCR2 and moderate levels of CXCR1/CXCR5/

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Fig. 8.5  The functions of hepatic DCs. Through the secretion of various cytokines and chemokines DCs are able to activate NK and NKT cells which can then carry out their effector functions. In addition in an inflammatory hepatic microenvironment DCs can recruit T-cells. Furthermore DCs can migrate to regional lymph node or Portal Associated Lymphoid Tissue (PALT) in a ­chemokine-dependent manner to active the adaptive immune response

NKT & NK CELL ACTIVATION

IL-6/IL-12/IL-15/IL-18

IL-12/CXCL3/CXCL20/MIP-1

CXCL19/CXCL21/CXCL3/MIP-1

MIGRATION TO LYMPH NODE/PALT

T-CELL RECRUITMENT

CXCR7. CXCR1 is activated by CXCL5/RANTES/ CXCL3/MIP-1a and CXCR2 is activated by MCP. CXCR1/CXCR2/CXCR5 is responsible for the migra­ tion of immature DCs into areas of inflammation [44]. Immature DCs when activated by GM-CSF all upregulate expression of mRNA of CXCR7 [44] allowing hepatic DC to migrate towards CXCL19/ CXCL21 secreting lymph nodes. Mice deficient in CXCR7 have deficient migratory ability and furthermore produce less IL-12 [27]. In addition hepatic DCs secrete CXCL3/MIP-Ia, CXCL1, and CXCL2/ MCF-1 and have a strong expression for CXCL5/ RANTES [45]. Activated DCs are the most potent stimulators of NK proliferation and cytotoxicity (see Fig.  8.5). IL-12, IL-18, IL-15, IFN produced by DCs enhance NK cell function [46]. NK cells and DC colocalize in lymphoid organs. IL-12 secreted by DCs is essential for IFNg secretion by NK cells. Furthermore membrane bound IL-15 on DCs is essential for NK cell proliferation and survival [47]. Upon stimulation with different TLR ligands DCs can activate NK cells [48]. In vitro DCs activation of NK cells requires direct cell contact resulting in secretion of IL-12 by DCs towards NK cells and a

synapse between both cells [49]. In addition IL-15Ra expression on DCs is required for NK priming [50]. Hence, IL-15, IL-2, IL-18, and type 1 IFN are critical for this cross-talk. In lymph nodes NK cells play an important role in the initiation of specific T-cell responses by influencing DCs maturation. In vitro if DCs are not stimulated with type 1 IFN, NK cells cannot license DCs to prime T-cell responses [38]. Consequently mature DCs activate NK to produce IFNg which is required for Th1 polarization [47]. In some cases T-cell mediated tumor rejection is dependent upon DCs activation by NK cells [51]. NK cell/DC cross talk may bypass the Th arm in Cytotoxic T-lymphocyte (CTL) induction against tumors expressing NKG2D ligands. A critical role for NKG2D-mediated NK/DC interaction in generating a robust CD8+ T-cell immunity against intracellular parasites is well documented [52]. These studies add further weight to a TLR/DC/NK axis in the early initiation of the immune response of the liver. Similar bi-directional interactions take place between DCs and NKT cells. Activated DCs are able to increase NKT cell secretion of cytokines [53]. Conversely CD40 signaling and release of TNFa and IFNg by NKT cells induce DC maturation [54]. DC maturation induced by

8  Immune Cell Communication and Signaling Systems in Liver Disease

TLR activation is enhanced by NKT cells [55, 56]. DC maturation in vivo with a variety of TLR ligands is greatly enhanced when there are active NKT cells [57]. Plasmacytoid DC stimulated through TLR-9 a recognition receptor for viral ligands, induce NKT cell activation markers and allow these cells to respond to antigen bound on DCs [58]. Therefore NKT cell interaction with DCs subsets contribute to further propagation of the immune response. Further evidence for the importance of NKT cell interaction with DCs is seen in Leishmaniasis where the differentiation stage of DCs, as well as their interaction with NKT cells determines Th1/Th2 polarization [59]. NKT cell/DC interactions can signal downstream IFNg producing CD4+/CD8+ T-cells. DCs activation by NKT cells can activate NK cells to induce antitumor effects [55]. As discussed earlier CD1d are MHC-like molecule involved in the presentation of glycolipids. Hepatic DCs express CD1d. AlphaGalceramide can activate NKT cells, via CD1d, in the liver [17] thereby inducing high levels of IFNg and IL-12 production. This correlates with suppressed tumor activity in a CM54 murine tumor model. Taken together these findings imply that both NK and NKT cells crucially interact with DCs to ensure their maturation and help to propagate the immunological responses including antitumor responses within the liver. The “unconventional” T-gd cells can also effect DC activation as evidenced by CD86 and MHC class I [60] and CD83 upregulation. They also induce IL-12 production by DCs [61]. Tgd cells can also enhance DCs maturation via TLR-2 and TLR-4 ligands [62]. Tgd cells activated by DCs infected in vitro by the bacillus Calmette-Guerin as evidenced by CD69 expression on the cell surface, secrete IFNg and have cytotoxic activity. Consequently DC stimulated Tgd cells help DC to prime a significantly stronger antimyobacterial CD8+ T-cell response. Tgd cells activated by phosphoantigen induce IL-12 by DC, an effect involving IFNg production. Tgd cells activation results in DC maturation and enhanced TCRab T-cell response [61]. In turn DCs induce CD25 and CD69 upregulation on Tgd cells as well as IFNg and TNFa secretion [60]. Activated DCs recruit other immune cells, including other T-cell subtypes, into the liver via CXCL3/ MIP-1a [45] (Fig. 8.5) and mice deficient in CXCL3/ MIP-1a show an attenuation of liver injury induced by ConA [63]. CXCL20/MIP-3a is the most commonly expressed chemokine in the liver and is produced by

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activated DCs. Intrahepatic T-cells express corresponding CXCR6 and can therefore be recruited to the liver via this pathway [45]. Hepatic DC can come into contact with circulating T-cells depending upon the cytokines with the liver microenvironment. For T-cells to be activated by hepatic DC, a combination of specific extracellular matrix protein and antigens are required to overcome the inherent tolergenic nature of hepatic DCs. Furthermore DCs are able to induce specific CTL responses [64]. Most T-cell activation occurs within lymph nodes but there is also evidence of T-cell proliferation in detected PALT. DCs capture antigens and migrate to either draining lymph nodes, which remain at the site and form a granulomatous lesion, or associate with T and B cells to form PALT [65]. Portal associated lymphoid tissue is an area within the portal triad where hepatic DC may interact with immature T or B cells. Hepatic DC can interact with T-cells without going to regional lymph nodes. The center of PALT is comprised of structures closely resembling B-cell follicles and populated by antigen loaded DCs. In between B-cell follicles are CD4+ T-cells. The vascular endothelium of PALT can also secrete CXCL21 and via CXCR7 crucial to the recruitment of mature DC and naïve T-cells to the area. DC maturation by innate lymphocytes is a very early and vitally important event as there is otherwise an absence of inflammation. Furthermore the presence of pathogen-associated molecules alone does not result in DC maturation and subsequent antigen presentation [48]. Interactions between mature DC result in NK/NKT cell activation, lymphocyte proliferation, IFNg production, and cytolytic activity against tumors, virus, or pathogen-infected cells [29]. This function of DC is important prior to engagement of the adaptive immune response. Thus in summary it is the above interactions that provide the bridge between innate and adaptive immunity and affect the subsequent magnitude and polarization of the T-cell mediated response.

Natural Killer (NK) Cell, Natural Killer T (NKT)-Cells, and T-gd Cells The reciprocal interaction of NK cells, NKT cells, and Tgd cells with DCs is critical for initiating the innate immune response within the liver. NK cells, NKT

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cells, and Tgd cells share a common expansion pathway, secrete similar soluble factors and attain cytolytic activity upon activation. NK cells represent a class of lymphocytes distinct from B and T cells and do not express a clonally distributed antigen receptor. The function of NK cells is regulated by a balance of signals from the stimulatory and inhibitory receptors expressed on their cell surface. Stimulatory receptors can be activated by stimulatory ligands expressed on infected, transformed or stressed cells, whereas binding of inhibitory receptors to self class I MHC molecules leads to inhibition of NK cell activation [66]. For a comprehensive review of NK cell receptors and ligands please see Biassoni et al. [67]. Hepatic NK cell numbers are greatly elevated in patients with malignant disease but show reduced cytolytic activity associated with the progression of hepatocellular carcinoma. The antitumor/antiviral action of NK cells in the liver is likely mediated via direct killing of target cells. Like cytolytic T cells, NK cells possess an extremely powerful cytotoxic armory including various TNF ligands such as TRAIL and also the perforin/

granzyme system. In cholestatic liver injury, increased TRAIL expression is thought to be one potential cause of NK cell-mediated hepatocyte injury and malignant liver epithelia is thought to show increased sensitivity to TRAIL mediated apoptosis compared to normal cells [68]. The mechanism through which NK cells mediate antiviral cytotoxicity appears to be organ dependent [69] but is influenced by IL-8 and IFNa [70]. Whilst NK cell cytotoxicity via the perforin/granzyme system is a very potent mechanism it may not have a central role in the liver [71]. Ligand–receptor pairs belonging to the TNF superfamily are also expressed by T cells and CD68+ positive macrophages and appear to be far more likely to play the key role in mediating liver epithelial cell damage [72] via TRAIL [73]. CD40/ CD154 [74–76] as well as the Fas/FasL system which remains a key pathway [74, 77] (see Fig. 8.6). NK cells, in humans, have commonly been classified on the basis of the CD56 expression (NKCD56lownoncytolytic and CD56high-cytolytic respectively). After DC-dependent activation NK cells proliferate and produce IFNg and acquire cytolytic activity probably

FasL

Fig. 8.6  The signaling pathways involved in Fas or TRAIL mediated hepatocyte (or cholangicyte) apoptosis. A potent mechanism of amplifying Fas mediated apoptosis via cooperative interaction between Fas and CD40 has also been described and extensively characterized elsewhere [74–76]. Fas forms a death inducing signaling complex (DISC) upon ligand binding. Membrane bound Fas ligand causes trimerisation of the Fas receptor. The adaptor molecule FADD binds to the death domain of Fas. FADD contains a death effector domain (DED) which can bind to caspase-8. Active caspase-8 is then released from the DISC into the cytosol which can cleave the pro-apoptotic BH3-only protein Bid into t-Bid. Following activation of other caspases this process leads to apoptosis

TRAIL DISC

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in a TLR-dependent fashion [16, 23]. The loss of this critical reciprocal cross-talk between NK cells and DC can lead to altered DC activity, defective NK cell func­ tion and aberrant T-cell responses [78]. Polyinosinic – polytidylic acid (Poly IC) is a double stranded RNA mimic which can bind to TLR3 expressed on APCs such as DCs. The injection of Poly I:C or murine cytomegalovirus (CMV) induces NK cell activation (characterized by CD60 expression) and accumulation in the liver which is followed by IFNg-production and partial clearance of the virus accompanied with massive cellular necrosis [79]. NK cells expressing TRAIL show increased cytotoxicity against primary hepatocytes in the liver injury which is probably enhanced on malignant cells [80] (see earlier). Natural killer T cells are central to the development of the diverse array of inflammatory responses seen within the liver and their presence can be identified by the expression of several NK markers (KIR, Ly49, CD161). NKT cells are also considered a bridge between innate and adaptive immune systems [81]. As previously stated, NKT cells respond to infections or inflammatory challenges prior to conventional adaptive T-cells. NKT cells may therefore potentially regulate a wide variety of autoimmune (see later) and inflammatory diseases [82, 83]. NKT cells are divided into two distinct subgroups: the Type I or iNKT cells (invariant NKT cells) that have a highly conserved invariant abTCR and the Type II NKT cells that (noninvariant NKT cells) express a more diverse TCR repertoire [3, 83]. The highest NKT cell/conventional T-cell ratio is found in the liver. The IL-15 receptor is expressed on NKT cells and IL-15 is vital for proliferation, survival and homoeostasis of NKT cells [84]. HSC secrete IL-15 [85] implying that the HSC/IL-15/NKT cell communication pathway is important for the relative enrichment of NKT cells within the liver. NKT cells recognize antigen presented by the MHC class I-related antigen CD1 [82, 83]. Self and glycolipid molecules presented on nonpolymorphic CD1d molecules can induce NKT cell activation [86, 87]. The reason for the relative enrichment of the liver with NKT cells may be due to the high expression of CD1d on KC, HSEC, and hepatocytes [83]. AlphaGalceramide is the most widely used activator of NKT cells but specific antigens have yet to be identified. Exogenous glycolipid NKT cell ligands include those derived from Gram-negative LPS-negative bacteria and from the spirochete Borrelia burgdorferi [88]. In the liver NKT cells reside mainly in the sinusoids and have

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motility rates calculated at around 20 µm/min [89]. Moreover, upon T-cell antigen activation, hepatic NKT cells abruptly stop moving [89]. The integral role of this cell type in liver immunology was seen after the injection of Concavalin A (ConA), which was seen to induce liver necrosis in mice, apoptosis of hepatocytes and lymphocyte infiltration [90]. This was prevented by steroids leading to the conclusion that the phenomenon was immune mediated and probably involved CD4+ T-cell. However, further work in CD1d knockout mice, which have a deficiency in NKT cells were resistant to ConA-induced liver injury indicating that this process was CD4 NK cell dependent [91]. Chemokines potently regulate NKT cell trafficking [92]. NKT cells have a high surface expression of a number of chemokine receptors CXCR3 and CXCR6 [92, 93] and express a chemokine receptor profile similar to Th1 inflammatory homing cells [92]. Mice deficient in CXCR3 or CXCR6 have significant reductions in hepatic NKT cell numbers, suggesting that these chemokine receptors play important roles in the hepatic recruitment and/or retention of NKT cells [89, 94]. IL-12 derived from activated DCs plays an important role in NKT cell activation [83]. Indeed mature IL-12 producing DCs induce prolonged IFNg producing NKT cells and offer better protection against tumor cells [95]. Moreover, IL-12 immunotherapy, in combination with GM-CSF, for liver tumors induce high levels of IFNg producing NKT cells as well as NK cells and CD8+ T-cells [96]. NKT cells also require CD40/CD154 mediated cell contact for activation [97]. NKT cells are able to rapidly produce large amounts of Th1-type (IFNg and TNFa) and Th2-type (IL-4 and IL-13) cytokines upon activation [98]. Central to NKT cells ability to induce Th1 and Th2 is the interaction of the costimulatory pathways CD28-CD80/ CD86 and CD40-CD154 which induce Th1 and Th2 responses respectively [99]. NKT cells are also capable of producing IL-17 implicating this cell type in inflammatory conditions characterized by a Th17 phenotype [100]. NKT cell-mediated liver injury and apoptosis is known to involve IFNg [101], Fasl(CD178) [102], and TNFa [103]. NKT cells are able to kill other cells through Fasl-mediated and perforin-dependent pathways [104]. IL-6 can abolish and inhibit NKT cellmediated liver cell injury [105] and IL-15 can alleviate ConA-induced hepatitis [106] and these may offer potential avenues for the modulation of NKT cell function. Interestingly IFNg derived from NKT cells can activate NK cells. The synergized function of NK/NKT

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cells appears to be dependent upon the model used. Whilst these cells can take part in cell-mediated antitumor activity [107], IFNg post ConA injection in vivo inhibits NK/NKT cell function, therefore this bidirectional interaction needs further study particularly in the context of specific liver pathologies. T-gd cells differ from the classical ab T-cell by a TCR with a unique structural and antigen binding feature [108], which endows them with independence of any APC for the recognition of foreign epitopes [109]. Each tissue has its own specific subset of T-gd cells according to the variable (V) gene used to generate the TCR [110]. TCRgd T cells represent a minority of T-cells in lymphoid organs and peripheral blood but are relatively abundant within the liver accounting for up to 15–25% of total liver T-cells. The percentage of gd T cells in the liver is significantly increased in the liver of tumor-bearing mice. Elevation of gd T cells was also found in the livers of patients with viral, but not nonviral, hepatitis. Emerging evidence suggest that gd T cells may play a prominent role in innate defenses against viral and bacterial infection and against tumor formation [111]. Thus elevated gd T-cells in the liver may also play an important role in innate defenses against pathogens and transformed cells. Studies have shown that hepatic gd T-cells release IFNg, TNFa, IL-1, and IL-4 and play a role in the response to liver tumors [112]. The signaling pathways activated as a consequence of activating this important T cell subset offers an exciting new avenue for treatment of immune mediated liver disease.

Adaptive Immunity Within the Liver The adaptive immune system is composed of T- and B-lymphocytes and is either initiated or downregulated by the innate immune system in highly complex ways which we are now beginning to understand. In no other organ is this linkage more important and tightly regulated than in the liver, an immunologically highly specialized environment. The adaptive immune system has the capacity of memory. B-cells play an integral role in the humoral immune response whereas T-cells are involved in the cell-mediated immune response. T- and B-cells have a least three stages of differentiation; naïve cells, effector cells and memory cells. Naïve cells are mature but have

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yet to encounter their cognate antigen whereas effector cells have been activated by their cognate antigen. Antigen is displayed on MHC also known as human leukocyte antigen (HLA) in humans. MHC class II molecules activate CD4+ (helper) T-cells and MHC class I molecules display antigen to CD8+ (cytotoxic) T-cells. MHC class I is expressed on all nucleated cells whereas MHC class II are present on APCs. In general CD8+ T-cells are active against intracellular pathogens and CD4+ T-cells against extracellular pathogens.

T-Cells The DC/NK/NKT cell axis is the first line of defence against invading pathogens and these cells drive the initial activation of T-cells. Much of our understanding of the function of T-cells within the liver comes from work performed with hepatotropic viruses. Dendritic cells may be involved in local antigen presentation to T cells trafficking through draining lymph nodes and PALT. As discussed previously, KC and PALT derived cytokines (CXCL3/MIP-1a and CXCL21 respectively) attract DC into the liver in a TLR dependent manner. Antigens can then be presented in situ by liver-resident DCs and activate infiltrating naïve lymphocytes [113]. Interestingly recent evidence shows that DC function locally within the liver and their ability to prime naïve T-cells may be modulated by HSECs [114]. Studies suggest that effector functions of CD8+ T-cells vary as to whether they have been activated in the liver or lymph node [115]. Antigen-specific lymphocytes, activated by DCs in lymph nodes, can then enter the blood stream and home to the liver where they exert their effector functions. When APCs themselves are infected or when their function is inhibited by pathogens, they cannot process endogenous antigen, and direct priming of T-cells becomes absent or inefficient. The only pathway for activating CD8+ T-cells is then the cross-priming pathway [116]. This involves an antigen inside one cell being endocytosed by another cell and cross-­presentation by that cell’s MHC class I to CD8+ T cells. Chemokines play a critical role in leukocyte trafficking. CXCR5 and CXCR3 are important in hepatic T-cell trafficking. Both receptors are associated with type 1 cytokine T-cell responses and the majority of tissue-infiltrating T-cells are found to coexpress these

8  Immune Cell Communication and Signaling Systems in Liver Disease

receptors [117]. During viral infection, CXCR5 and CXCR3 are expressed on NK cells and activated CD4+ and CD8+ T-cells [118]. Furthermore, CXCR5, CXCR3, and CXCR6 are expressed on liver-­infiltrating lymphocytes and there the expression correlates with the severity of the hepatitis. Furthermore, the chemokines IP-10, Mig, and I-TAC through their interaction with CXCR3 and CXCR5 interact with RANTES, and MIP1a and MIP1b attract T-cells to the liver in chronic HCV [119]. Of interest is that, mature hepatic DCs produce RANTES and MIP1a and may themselves be able to recruit T-cells into the liver [120]. The recruitment of lymphocytes to the liver also involves the increased expression of adhesion molecules such as VCAM-1, p-selectin, e-selection, and VAP-1 by hepatocytes [121]. Studies on murine liver show that ICAM-1-mediated trapping of activated CD8+ T-cells depends on the capacity of the vasculature and/or parenchymal cells to present antigen, whereas VCAM-1 trapping is antigen independent [122]. Other studies have suggested that in addition to physical trapping Th1 and Th2 cells use alpha(4)-integrin and VAP1 respectively to adhere within the liver sinusoids [123]. The recruitment of T-cells into the liver involves chemokine-mediated and adhesion molecules-mediated phases. Resting, naïve CD8+ T-cells are preferentially located in secondary lymphoid compartments and require two independent signals to become fully activated. CD8+ T-cells recognize peptides 8–11 amino acids long in the context of MHC class I molecules. This is the first signal and is transduced through the specific TCR. The second signal is independent of the antigen receptor and is critical to allow full activation and differentiation of CD8+ T cells [124]. Thus, whereas primed effector CD8+ T cells can be activated by any target cell that express the cognate antigen in the context of MHC class I, only few, appropriately licensed bone-marrow derived professional APC have the ability to initiate CD8+ T cell responses [125], probably because they express costimulatory molecules and because they carry antigens from the site of infection into lymphoid organs [124]. To date there is no clear evidence whether antigens are presented to T-cells in the liver, or whether it occurs in draining lymph nodes. Several recent studies provide new insights into the kinetics and effector functions of CD8+ T cell responses in the liver. Adoptive transfer of HBV-specific CD8+ T cells into either transgenic mice that bear replication-competent copies of

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HBV in their hepatocytes or into nontransgenic littermates, results in rapid recruitment of the transferred CD8+ T cells into the liver [126]. Consistent with previous reports [127] this finding confirms that activated CD8+ T cells are recruited to and/or are trapped in the liver irrespective of their antigen-specificity. Only upon recognition of their cognate antigen, however, do these CD8+ T cells undergo rapid proliferation [126]. Prolif­ eration presumably occurs directly in the liver in this scenario, as increased numbers of antigen-specific T cells are not detectable in draining lymph nodes during the early days after adoptive transfer. Most importantly, IFNg production of antigen-specific T cells appears early and then declines, whereas granzyme B expression and cytotoxicity are relatively delayed but sustained. This sequential activation and downregulation of CD8+ T cell effector functions is reminiscent of the sequence of early, cytokine-mediated suppression of HBV replication and late appearance of liver injury, which have been described in chimpanzees with acute HBV [128]. Effector functions of intrahepatic CD8+ T cells include the production of cytokines, such as IFNg and TNFa, and cytolytic mechanisms [129]. In particular, CD8+ T cells exert their cytolytic activity by releasing granule contents such as perforin and granzyme and by triggering Fas-mediated apoptosis [130]. The Fasstimulated pathway appears central to the ability of CD8+ T-cells to be cytotoxic. CD8+ T-cells are positive for FasL expression, and interaction of Fas and FasL is a key pathway in triggering apoptosis of HCV-infected hepatocytes. Activated CTL recognize appropriate antigens via the TCR in the context of MHC molecules and express death molecules such as FasL, the activation of which results in caspase 8 activation and ultimately in apoptosis. Alternatively activated CTL release cytotoxic granules containing perfornin and granzyme B. Internalization of granzyme B by hepatocytes is mediated by the formation of perforin-induced membrane pores, Granzyme B cleaves procaspases that also lead to apoptosis. Further acute liver injury continues as a nonspecific, chemokine-mediated amplification of the intrahepatic infiltrate [131]. A key step in this nonspecific amplification of the inflammatory infiltrate is the recruitment of neutrophils. Neutrophils are recruited by adhesion molecules, cytokines and chemokines, many of which are produced by activated KCs. For details of the role of neutrophils in liver injury please see Ramaiah et al. [132].

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B-Cells

Regulation of the Immune Response

Little is known about the function of B-cells as they make up a small proportion of the lymphocytes within the liver (Fig. 8.2) and secondly there is considerably a technical difficulty in isolating and analyzing these cells. Th2 cytokines, which activate B-cells and induce their differentiation into antibody-producing cells, are most commonly seen in autoimmune liver diseases (AILD see later) where much of our understanding of B-cell function comes from. In AILD there is an increase in the number of infiltrating B-cells. Indeed the proportion of CD19+/CD69+ activated functional B-cells is markedly higher in the liver of patients with primary biliary cirrhosis (PBC). Furthermore the antibody-cell complex may cause damage to target cells [133]. Intriguingly, auto reactive B-cells may also act as APCs for T-cell priming. It is not clear how B-cells traffic to the liver and how this process is regulated but CXCL13/CXCR5 may play a crucial role. There is evidence that in murine models of SLE, DCs are able to secrete CXCL13 increasing the number of B-cells seen in the kidney [134]. Stimulation of DC with microbial motifs in the presence of IL-10 may result in CXCL13 production and could lead to the  accumulation of CXCR5-bearing autoantibody-­ producing B-cells in the target organ. Classically, B-cell activation, differentiation and proliferation occur in lymphoid follicles of secondary lymphoid organs but can occur in ectopic germinal centers. The HCV-infected liver can be considered to be an ectopic lymphoid organ in which B-cells proliferate and differentiate antibody-secreting cells within germinal center-like structures of intraportal lymphoid follicles. These findings corroborate the idea that intrahepatic follicle-like structures are functionally similar to those of lymph nodes with respect to B-cell activation, expansion and maturation. Recent studies have indicated that B-cells may play a critical role in the development of hepatic fibrosis. In B-cell-depleted mice, fibrosis did not develop subsequent to liver injury induced by carbon tetrachloride. Similarly, in a-naphthylisothiocyanate (ANIT)-induced liver injury, B-cell depletion inhibited the development of fibrosis [135]. These effects appear to be antibodyindependent and maybe caused by MMPs and TIMPS. There is therefore scope and a need to increase our understanding of B-cell biology and signaling within the liver, however this does remain technically challenging.

Within the liver there is a powerful mechanism for downregulating established immune responses. The (antiinflammatory) cytokine IL-10, which is produced by monocytes, lymphocytes, KCs, hepatocytes, HSEC, and HSC, has a crucial role in immunoregulation. The receptor of IL-10 is highly expressed on immune cells [136]. IL-10 downregulates Th1 cytokines, MHC class II molecules and other costimulatory molecules and can inhibit IFNg synthesis by Th1 cell clones [137]. IL-10 inhibits the presentation of antigen to T-cells and the production of IL-12 [138]. IL-10 also alters the expression of chemokine receptors on DCs and downregulates their migration to the draining lymph node [139]. Furthermore if T-cells are primed in the presence of IL-10 they are prone to lose cytokine production and effector functions [140] and IL-10 also dramatically reduces hepatic necrosis after, in mice challenged with Con A treatment [141]. The immunoregulatory role IL-10 is highlighted by Wahl et al. Hepatic NKT cells produce IL-4 when stimulated by hepatocytes but IFNg when stimulated by DCs. CD8+ T-cells produce no IL-10 when primed by hepatocytes or NKT cells. However IL-10 producing CD8+ T-cells are generated if CD8+ T cells, NKT cells, and hepatocytes are in close contact in the presence of IFN. This leads to downregulation of cytokine production and DC function [142]. HSEC-mediated presentation of antigens to CD8+ T-cells results in tolerance rather than effector functions [143]. In addition CD8+ T-cells cocultured with HSEC exhibit low IL-2 and IFNg production and low cytotoxicity, reduced proliferative capacity, and are prone to undergo apoptosis with the effect being reversed with the addition of IL-2. Antigen-presentation by HSEC can therefore be tolerogenic [144]. Downregulation of IFNg production by T-cells coincides with the upregulation of PD-1 on CD8+ T-cells, a receptor that is known to modulate TCR signaling [145]. This results in inhibition of cytokine production and proliferation in the mouse model of acute HBV. The PD-1 ligand PDL-1 is constitutively expressed on HSEC and KCs which suggests that these liver-resident APC actively downregulate specific effector functions of activated liver-infiltrating T-cells. Thus, the downregulation of the immune response within the liver involves a variety of mechanisms including cytokines and cell-mediated effects.

8  Immune Cell Communication and Signaling Systems in Liver Disease

Immune Cell Function in Liver Disease We have considered the function and interactions of the different immune cells within the liver. The following section focuses on the interplay of the various cells described in different pathological processes affecting the liver.

Liver Transplantation Orthotopic liver transplantation (OLT) is the only option for end-stage liver failure (ESLF) and rejection remains

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an important cause of allograft failure, along with eventual recurrence of original disease in some instances. Destructive immunologic processes can underlie 50–75% of allogenic liver graft rejection. Evidence suggests that liver-derived DCs can downregulate immune responses (Fig. 8.7), thereby inducing and maintaining peripheral T-cell tolerance [146]. For a comprehensive review of the role of DCs in liver transplantation please see Sumpter et al. [147]. T-cells are central to the immune mediated injury after OLT. The context in which antigen is presented to T-cells determines whether the responding T-cell is activated or tolerized. As described earlier, T-cells require 2 distinct but coordinated signals to achieve optimal activation. Two

Activation and/or direct killing of endothelial cells +

CD4 T-cells

Monocytes (self APCs) Recirculation through lymphoid organs

+

CD8 T-cells

HSEC

Macrophages

Direct killing of parenchymal graft cells

Indirect allorecognition

Acute rejection

Chronic rejection

Fig. 8.7  Interactions among HSEC, T-cells, and recipient APCs in allograft rejection. Recipient monocytes recruited by HSECs to the graft tissue are transformed to become highly efficient antigen-presenting DCs that recirculate to peripheral lymphoid organs for maturation. The DCs and intragraft macrophages

Dendritic cells

present donor peptides via the indirect pathway to CD4+ T-cells. CD8+ T cells, on the other hand, are activated by donor HSEC and either can directly kill HSEC or traverse the endothelium and kill parenchymal graft cells (adapted from [279])

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costimulatory molecule–ligand pairs are also important in T-cells, CD28/B7 [148], and CD40/CD154 [148]. TCR stimulation and CD28 ligation leads to increased IL-2, cellular proliferation and induction of antiapoptotic proteins. CD40/CD154, the second receptor/ligand interaction, is critical for alloimmune responses. CD40 is expressed on all APCs whereas CD154 is expressed on CD4+ T-cells following activation. CD40/CD154 ligation augments APC function, it increases MHC class II, CD80, CD86 expression and IL-12 production. This enhances B- and T-cell responses to alloantigen. In liver recipients with acute cellular rejection CD86 and CD154 is seen to be upregulated [149]. CD154 is also detected in livers with chronic rejection and T cell and macrophage derived CD154 may activate CD40 on epithelial cells and amplify FasL mediated death directly via mechanisms involving the NFkB, AP1, and STAT 3 signaling pathways [74–76, 150, 151]. CD40/CD154 blockade appears to result in prolonged rat allograft survival and decreased biliary/ endothelial injury and prolong renal allograft survival in chimpanzees [152]. Many other molecules are likely to be involved in T-cell activation and allograft responses [148]. Cytotoxic T-lymphocyte associated antigen (CTLA4) is a CD28-related protein that is upregulated upon T-cell activation and like CD28 binds to B7-1 and B7-2. CTLA4 delivers a negative signal that attenuates T-cell function and may downregulate T-cell responses and induce tolerance. Indeed antiCTLA-4 also results in prolonged rat allograft survival. The recognition of allograft MHC antigen is the ­primary event leading to graft rejection. Recipient T-lymphocytes recognize donor alloantigens utilizing two methods; the direct and indirect pathways. The direct method involves host T-lymphocytes recognizing native MHC molecules on graft associated APCs. The indirect pathway involves host T-lymphocytes recognizing donor alloantigen-derived peptides in the context of self MHC molecules expressed on recipient APCs. Both these pathways are known to operate in the liver allograft. Early after liver transplantation the direct pathway predominates and is likely to have a role in acute rejection since graft-derived APCs expressing donor alloantigen rapidly egress from the graft and enter secondary lymphoid tissue, where they encounter allospecific T-cells [153]. Host T-cells then have the ability to engage the allograft directly and mediate effector functions. The indirect pathway probably emanates from donor alloantigen that is shed from damaged graft tissue or perhaps in the case of the liver, from soluble MHC molecules,

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that are picked up and presented by self APCs in particular DCs [148]. Therefore, the direct pathway may be important in initiating the classic form of acute rejection and the indirect pathway may predominate later by sustaining a response fueled by epitope spreading as a variety of allopeptides are presented successively by self APC [148]. At the same time, it is important to recognize that the indirect pathway may be involved in immune regulation because T-cells with allopeptide specificity were shown to have regulatory function through inhibition of IFNg production in other organ recipients [154]. The contribution of the indirect pathway to hepatic allograft tolerance recently was confirmed [155]. For both acute and chronic rejection, the targets of mature effector T-cells are donor-derived bile duct epithelial cells (BECs) and vascular endothelium [156]. CTL kill BECs via either perforin-dependent pathways or activation of members of the TNF receptor superfamily, particularly Fas [156] (see Fig. 8.6). The interaction between CTL and target cells is facilitated by the integrins LFA-1 and VLA-4 which enhance adhesion and provide survival signals for effector cells [156]. Bile ducts constitutively express CXCL19 which attracts DCs (see Fig. 8.4) and promotes their localization at the biliary epithelium where they are ideally situated to respond to antigen. Although human BECs lack CD80 and CD86, minimizing their ability to activate naïve T cells [157], the majority of T-cells that come into contact with activated BECs are already primed cells, and therefore, BECs may promote the local proliferation and survival of such cells [156]. There is evidence that the granzyme/perforin pathway is activated in human allograft rejection [156]. The Fas/FasL pathway is also active during liver allograft rejection [158] (Fig.  8.6). CD40 ligation on BECs results in upregulation of FasL; thus, although CD40 does not directly activate caspases, it dramatically enhances the expression of FasL and via NFkB AP1 and STAT-3 pathways, results in autocrine or juxtacrine apoptosis via indirect Fas activation [156].

Autoimmune Liver Disease The triads of the most common autoimmune diseases that can affect the liver are PBC, primary sclerosing cholangitis (PSC) and autoimmune hepatitis (AIH). PBC is characterized by autoimmune destruction of the small and medium sized intrahepatic bile ducts. The

8  Immune Cell Communication and Signaling Systems in Liver Disease

serological marker of PBC is antimitochondrial antibody (AMA) which is directed against the E2 subunit of the mitochondrial pyruvate dehydrogenase complex (PDC). PSC is characterized by fibrosing inflammatory destruction of the intrahepatic and extrahepatic bile ducts. The autoantibody in PSC is unknown. In PBC and PSC, much the same as orthotopic liver transplant rejection, the target cell is BEC or cholangiocytes, although the reason as to why BEC is targeted remains elusive. In PBC, some authors suggest, that transcytosis of the AMA-IgA complex into BEC may cause caspasemediated cell death, while others suggest a bacterial initiator. The third disease entity is AIH where the hepatocyte is the target cell. The role of innate immunity in autoimmunity is only partially understood [159]. NK cell function can be regulated by the receptors such as KIRs. Ligands for inhibitory KIRs on NK cells include HLA class I molecules, whereas the ligands for activating KIRs have yet to be fully characterized. Previous studies have demonstrated that particular combinations of KIR and HLA class I ligands can result in reduced NK cell inhibition increasing susceptibility to autoimmunity. Furthermore, stimulation of TLR-expressing BECs with bacterial motifs, such as LPS, produced high levels of cytokines and chemokines, which would potentially result in recruitment of inflammatory cells. BEC injury or dysfunction in PSC may be the result of an exaggerated activation of the innate and adaptive immune in response to bacterial products in the liver [160]. In PBC there is a marked increase in the NK cells [161] which have increased cytotoxicity, perforin expression, and increased levels of IL-8 and the IL-8R. These alterations in innate immunity might come to influence the initiation and perpetuation of the subsequent adaptive autoimmune response. BECs of patients with PBC overexpress TNFa and its TNFR supporting the idea of paracrine activity leading to eventual apoptosis [162]. In transgenic mice impaired TGFb signaling of T-cells gives PBC like disease pattern [163]. Indeed both TGFb and TNFa have been reported to play a role in PBC progression [164]. NKT cells have potent immunomodulatory properties and have been implicated in the pathogenesis of autoimmunity [165]. Hepatic NKT cell numbers are increased in patients with PBC [166]. IFNg secreted from NKT cells can augment autoimmunity whereas production of Th2-type cytokines can show improvement in experimental models of autoimmity [165]. Injection of ConA in murine models can mimic many

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aspects of human AIH [167]. In this model hepatic NKT cells are central to the severe hepatitis caused by ConA administration as NKT cell −/− mice are completely prevented from hepatitis after ConA treatment [91]. IFNg and TNFa secreted from NKT cells are integral to the development of ConA-induced hepatitis [168]. This work is supported by work which showed that the specific NKT cell ligand a-GalCer [169] also causes AIH similar to ConA. NKT cell secreted IL-4 may also have an important role in ConA-induced hepatitis [168, 169] a finding which parallels observations in children with AIH [170]. Therefore although IL-4 is a Th2-type cytokine in liver disease it has proinflammatory properties [171]. This is supported by findings that IL-4 induces hepatocyte apoptosis in vitro [172] and in vivo [173]. Moreover IL-4 secreted by NKT cells upregulates FasL expression on the NKT cell surface in an autocrine fashion which subsequently interacts with Fas expressed on hepatocytes which lead to hepatocyte apoptosis [174]. In addition IL-4 is linked to the production of IL-5, by NKT cells, during the course of ConA-induced hepatitis [175]. IL-5 can recruit eosinophils to augment hepatitis caused by ConA [176]. This may have implications for OLT as IL-5 recruitment of eosinophils may contribute to allograft rejection [177]. IL-17 is a potent inflammatory cytokine [178] capable of inducing tissue inflammation and autoimmunity [179]. In vivo and in vitro NKT cells can produce IL-17 [100] which may have a role in hepatic inflammation as high serum IL-17 levels are associated with severe hepatitis [180]. This area however needs further investigation. It is known that BEC increase can increase levels of adhesion molecules ICAM-1 [181], VCAM-1, and intergrins including LFA-1 [182] in diseases including PBC allowing interaction with lymphocytes. In addition BECs ability to express T-cell ligands CD80 and CD86 may explain the reason for bile duct loss. Apoptotic BECs in PBC express CD40 and Fas and FasL [150]. Autoreactive CD4+ and CD8+ T-cells are noted to infiltrate into the liver in PBC [183] and both cell types recognize epitopes of PDC-E2. Regulatory T cells (Tregs defined as fox P3+ve, CD4+ CD25high) may have an important role in human autoimmune diseases [184] as a decrease in Tregs is noted in PBC [185]. It is postulated that Tregs are important for the prevention of autoimmunity and maintenance of self-tolerance. For instance transfer of T cells lacking the Treg subset into athymic nude mice results in the development of various T cell-mediated

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autoimmune diseases [186]. PBC patients display significantly lower frequencies of Tregs as percentages of total TCR-ba+/CD4+ T cells may contribute to the failure in tolerance in PBC [185]. Decreased hepatic Treg cells may contribute to the induction of autoreactive B-cells resulting in AMA production. Specific microbial motifs may enhance TLR-dependent IgM production by B-cells [187]. Indeed in PBC, CD86+ CD38+ B-cells produce IgM and AMA via a TLR9-dependent mechanism [188]. Th2 cytokines, which activate and induce differentiation B-cells, dominate in AIH. Indeed CTLA-4 may favor Th2 predominance. IL-4, a marker of B-cell differentiation, and CD23, marker of humoral immunity, are also increased AIH. CTLA-4 polymorphism may also contribute to impaired Treg function in PBC resulting in a decreased B-cell regulation and enhanced B-cell autoimmunity. Similarly in AIH there is significant reduction in peripheral Treg cells with reduced proliferative activity [189]. Also in AIH Tregs do not inhibit CD8+ T-cell proliferation and cytokine production. Interestingly there may be bi-directional interactions between NKT cells and Treg cells. NKT-cell-derived IL-2 increases Treg cell proliferation and enhances CTLA-4 expression [190]. This increase in Treg cells may inhibit autoimmune responses [191]. Therefore defects in NKT cell/Treg cell cross-talk with regards to recruitment or maintenance of Treg cell populations within the liver may underlie autoimmune liver disease. In addition, activated NKT cells upregulate CD154 expression which can interact with CD40 expressed on DCs [97]. CD40 stimulation drives the subsequent semi-maturation of DCs, which, as seen in OLT, plays an important role in the development of tolerance and possibly regulates the ultimate predisposition to autoimmune liver disease [192]. Other molecules may be important in the development of AIH. For instance mice deficient in programmed death-1 (PD-1), which is induced by T-cell activation, develop autoimmune like syndromes. PD-1 binds to PD ligand PDL-(1) and PDL-(2) and regulates Th1 and Th2 cytokines. PDL-1 deficient mice display a spontaneous accumulation of CD8+ T-cells in the liver [48]. Furthermore, PDL-1 absence leads to AIH suggesting that PDL-1 plays a role in CD8+ T-cell deletion in the liver. Our understanding of the role of immune cells in the development of autoimmunity is rapidly expanding. It is likely that a number of known molecules and as yet

R. H. Bhogal and S. C. Afford

unknown molecules play an important role and that the interactions of various immunological cells underlie the development of the disease.

Alcoholic Liver Disease Alcohol abuse has become a major health and social problem with nearly 20% of alcoholics developing fibrosis. Persistent alcohol abuse can cause alcoholic liver disease (ALD) which can eventually lead to liver cirrhosis. Immune mechanisms are partially responsible of the evolution of ALD. Immune dysfunction plays a role in the multifactorial etiology of ALD which include metabolism, genetics, nutrition and environmental factors. Alcohol is metabolized via NADH and NAD in the GI tract [193]. Severe alcohol consumption results in alcohol metabolites that form stable protein adducts which can react together and form adducts such as Malondialdehyde-acetaldehyde (MAA) which are immunogenic. Studies on humans have shown that increased levels of adducts in the serum correlated with the severity of ALD [194]. Chronic ethanol abuse impairs receptors on HSEC and impairs their ability to clear MAA adducts [195] allowing them to interact with the immune system. Indeed in ALD there is an increased infiltration of immune cells [196]. MAA adducts increase TNFa, MCP-1, MIP-2 and increased expression of ICAMs, P-selectins, L-selectin and reactive immune cells. Furthermore alcohol metabolites induce HSEC expression of selectins and increase the adhesion molecules on the surface of SECs [197] to which attach leucocytes [198]. Indeed mice deficient in ICAM-1 had reduced lymphocytic infiltrate in the liver following ethanol treatment [199]. Acetaldehyde adducts (AA) increase cytokines/chemokines from endothelial and HSC [200] and can attract lymphocytes. MAA can stimulate proinflammatory cytokines (TNFa, IL-1, IL-6) [200] and induce a strong antibody response [200]. Furthermore in vitro MAA-adducts can induce T cell expansion. KCs and DCs initiate the hepatic innate immune response (Fig. 8.4). In ALD, similar to the postulated bacterial trigger in PBC and PSC, KCs can be activated by LPS [200] derived from increased intestinal wall permeability [201]. KCs release TNFa and super oxides which supports an inflammatory response

8  Immune Cell Communication and Signaling Systems in Liver Disease

[200]. KCs may serve a key role in ALD as they produce ethanol-derived free radicals that can induce hepatocellular injury. Furthermore, TNFa, derived from KCs and DCs is essential for the induction of hepatocellular injury seen in ALD [200]. The key in the evolution in ALD and progression to fibrosis is thought to be HSC activation. Indeed TNFa, IFNg, and IL-2, produced by DCs, NK, and NKT cells activate HSCs early in ALD. HSCs are capable of producing large amounts of collagen when activated. Growing evidence suggests that HSC may participate in the hepatic innate immune response [202]. Little is known about the role of both NK and NKT cells in ALD [203]. In the murine models of chronic ethanol feeding, increased numbers of hepatic NKT cells have been associated with the development of liver injury, an observation not seen in NKT-cell-deficient mice [204]. Moreover activation of NKT cells by treatment of mice with a-GalCer in the setting of experimental alcoholrelated liver injury, leads to marked hepatocellular injury [204] mediated by hepatic NKT cells in a Fasand TNFa-dependent manner [204]. It is known that there is a significant reduction in T-cell reactivity and proliferation and altered T-cell composition in the livers of chronic alcohol consumers [205], which has a close association with the severity of ALD [206]. T-cells isolated from alcoholics express markers associated with activation and memory (CD45RO, CD57, and CD11b) [207]. These cells are able to rapidly produce IFNg and TNFa again demonstrating the ability of ethanol to prime hepatic T-cells [207, 208]. Coculturing lymphocytes from alcohol cirrhotic patients with ethanol increases adhesion molecule expression and TNFa and IFNg compared to control [206]. Indeed these T-cells show increased reactivity when compared to normal controls [206]. For instance, ethanol is also able to prime liver T-cells to show an exaggerated response to Con A and in rats transferring T-cells from ethanol fed rats control induces liver injury [208]. Increased levels of CD8+ T-cells is seen in the liver with regenerating nodules and fibrosis implicating this cell type in the profibrotic process of ALD [209]. Furthermore CD8+ T-cells can also be activated by AA-modified spleen cells and many other AA-modified proteins [210]. The continuing presence of AA could lead to immune-stimulation and cirrhosis. Moreover ALD is associated with circulating antibodies specific to hepatic antigens [211] that correlate with ALD [212].

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Viral Hepatitis Hepatitis B virus (HBV) and HCV are both hepatotropic viruses that are a significant cause of chronic liver disease worldwide. HBV is a DNA virus consisting of an outer surface coat hepatitis B surface antigen (HBsAg), an inner core antigen (HBcAg) and e antigen (HBeAg). HCV is a RNA virus. Both viruses can cause acute or chronic infections although HCV is associated with a more chronic course. Both can cause progressive liver injury resulting in ESLF unless effectively eradicated [213, 214]. DCs and resident macrophages capture viral antigens. Viral components stimulate DCs through ligation of TLR 7–9 [215]. DCs then migrate to lymphoid tissue and perhaps PALT (Fig. 8.5). Cytokines released by DCs then shape the immune response to viral hepatitis [216]. DCs can activate NKT cells. Moreover increased numbers of activated hepatic NKT cells are seen in HBV [217] and chronic HCV livers [218], which may also be activated in response to viral antigen-expressing hepatocytes. NKT cells through the secretion of IFNg inhibit HCV replication through a noncytolytic mechanism [219] and HBV replication in a murine HBV transgenic model [220]. Virus mediated downregulation of CD1d expression on DCs may allow evasion from NKT cells [221]. NKT cells may therefore be pivotal in the clearance of HCV [222] and HBV-infected hepatocytes [217] which is central to the aim of the immune response. Further evidence for this comes from the role of NKT cells in the development of immunity following HBV vaccination [223]. NKT cells mediate cytokine- and/or FasL-mediated killing of infected hepatocytes. However, if this is overly robust it can lead to severe liver inflammation and even fulminant liver failure. Interestingly activated NKT cells have a negative effect on hepatic regeneration (see below) which can potentially negatively affect hepatic repair following viral hepatitis. Whilst the role of NK cells in human HBV and HCV infections is poorly understood, NK cell-derived IFN inhibits HBV replication in vivo and direct killing of infected hepatocytes in a similar way to NKT cells [224]. In vitro NK cells inhibit HCV replication in human hepatoma cells via an IFNg-dependent mechanism. Indeed individuals with enhanced NK function have improved spontaneous clearance of acute HCV infection [225]. However HCV can escape the antiviral

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response of NK cells through inhibition of its function and may contribute to the chronic refractory HCV infection seen in the majority of patients. Induction of the adaptive immune response is crucial during the course of viral hepatitis with virus-specific T-cell responses [226] being a key influence on the course of the disease. Although HBV does not induce any genes during entry and expansion [227] a large number of T cell-derived IFNg-regulated genes are induced during viral clearance [227]. In acutely infected patients the adaptive T-cell response inhibits viral replication and kills infected hepatocytes terminating viral infection [226, 228]. In chimpanzee models the early phase of HBV clearance was temporally associated with the appearance of CD3, CD8, and IFNg in the liver, which reflects the influx of T-cells into the liver [128]. Although almost all hepatocytes were infected there is only limited liver disease suggesting that noncytopathic mechanisms are active during early viral clearance. Late elimination of the virus takes several weeks occurring in the presence of significant liver disease mediated by virus-specific T-cells exerting cytopathic effector functions [128]. There is a differential role for T-cells in this process. CD4+ T-cells did not have a significant role whereas CD8+ T-cell depletion greatly prolongs viral infection and delays viral clearance [229]. The complete elimination of the virus took several months again associated with increased numbers of intrahepatic CD8+ T-cells and increased intrahepatic IFNg. Therefore intrahepatic HBV-specific CD8+ T-cells are required for rapid viral clearance functioning early in HBV infection through noncytolytic-IFNg-dependent mechanism and later via a cytolytic mechanism that clears the remaining infected hepatocytes. The exact effector function of CD8+ T-cells is probably mediated by its interactions with APCs [126]. CD4+ T-cell are, however, involved in HCV eradication [230]. CD4+ T-cells do not mediate hepatocyte cell injury but facilitate CD8+ T-cells effector functions. In HCV CD8+ T-cells again may act via two mechanisms to clear HCV. The first involves a cytolytic mechanism involving FasL, perforin/granzyme, TRAIL, and TNFa (Fig.  8.6). Indeed, HCVinfected hepatocytes upregulate Fas expression and this correlates with liver inflammation [231]. The second mechanism involves releasing IFNg to suppress HCV replication (noncytolytic) [219]. In contrast to the strong and multispecific CD8+ T-cell immune response in acute self limiting HBC infection the response in chronic HBV is relatively weak [228] and narrowly focused [226]. The same

R. H. Bhogal and S. C. Afford

CD8+ T-cell response is seen in chronic HCV. This suggests that CD8+ T-cells function to control early or acute viral infection [232]. In chronic HCV infection, CD8+ T-cell activity is detected in the liver and the periphery implying an active immune response even in the absence of acute viral infection [233]. One possible explanation is NK/NKT and macrophage derived IL-12/IL-15 may continue to fuel CD8+ T-cell function [234]. However NK cells impairment has also been implicated in the development of chronic HCV infection [235]. As discussed above HBV-specific CD8+ T cell responses are detectable during the acute phase of self-limited infection and then decline [236]. However in chronic infection, HBV-specific CD8+ T-cells remain detectable at high frequencies even after HBsAg seroconversion [237]. Furthermore relatively high levels of HCVspecific CD4+ and CTL cells have been detected in livers from chronic HCV patients [238]. These T-cells are functionally different from HCV-specific T-cells in the peripheral blood, suggesting that T-cells are sequestered within the liver [239]. In the absence of complete virologic recovery, CD8+ T-cells may mediate hepatocellular injury. Studies suggest that CD4+ T-cells and CTL responses differ among patients who recover compared to those who develop chronic HCV infection. There is a strong CD4+ T-cell response in patients with long-lasting chronic HCV [240] which correlates with histological fibrosis. In chronic HCV-infected patients with intrahepatic CD8+ T-cells activity, lower levels of viremia are associated with more active hepatocellular damage [241]. In the HBV transgenic mouse model virus-specific T-cells and NK/NKT cells can abolish HBV expression and replication without hepatocytes injury via IFNg- and TNFa-dependent pathways [129, 219]. Therefore T-cell dysfunction may be central to the persistence of the virus hepatitis. However these mechanisms are poorly understood but may include T cell deletion, anergy, exhaustion and ignorance. Viral infections may also persist via the development of viral escape mutations. Despite a strong Th1 response in HCV the virus persists because of the high mutation rates making it possible to escape immune surveillance [242] and develop resistance to cytokines produced by HCV-specific CD8+ T-cells thus continuing to induce T-cell activity. The viral strains are not cytopathic, but in the presence of persistent, chronic infection, Th1 cells continue to release TNF-a, IFN-g, and IL-2, causing self-inflicted inflammation and necrosis [243]. Furthermore, TNF-a-dependent macrophages activation leads to hepatic fibrosis in a TGF-b-dependent

8  Immune Cell Communication and Signaling Systems in Liver Disease

manner by activating HSC to produce collagen I [244]. Examination of liver specimens from HBV- and HCVinfected patients has revealed increased hepatic NKT cell numbers in chronically infected livers [245]. Moreover, these NKT cells had sig­nificant alterations in their effector functions demonstrated by a skewing in their cytokine-producing profiles to a more Th2-type consisting of IL-4 and IL-13 production, cytokines linked to the development of hepatic fibrosis [245]. Interestingly the increase in hepatic NKT cell numbers in chronic HBV- and HCV-infected livers was associated with a striking upregulation of CD1d expression in APCs in cirrhotic livers [245]. These observations suggest that hepatic NKT cells in patients with chronic viral hepatitis may contribute to the development of progressive liver fibrosis via a mechanism that involves an NKT cell CD1d-associated enhancement of profibrinogenic cytokine secretion. Finally, activation of NK cells has also been implicated in liver injury and

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fibrosis [246]. Taken together, these data show that NK cells not only play an important role in innate response against pathogens in the liver but also contribute to the pathogenesis of liver disease. Little is currently known about the intrahepatic T-cell response during chronic HBV infection although T cell dysfunction and high viral load might also contribute to viral persistence. HBsAg-specific CD8+ T cells display abnormal HLA/ peptide tetramer binding properties in contrast to HBcAg-positive CD8+ T-cells [247]. However antiviral treatment can overcome CD8+ T-cell hypo-responsiveness in subjects with chronic HBV infection, suggesting that the T cells are present but suppressed [248]. A lack of virus-specific CD4+ T-cells and Tregs may contribute to this T cell suppression [249]. Clearly, additional studies are required to better understand the complex host-virus interactions that determine the outcome of chronic HBV and HCV infection. Viral Antigen/Virus

Viral Persistance

Viral mutations Defective NK/NKT cell function Th2 cytokine response Poor CTL response

IFN /TNF Persistent Hepatic Injury

Inhibit Viral Replication Viral Clearance

Fig. 8.8  The immunopathogenesis of viral hepatitis. The interaction between APCs and NK.NKT/T-cells influence whether the virus is cleared or results in fulminant hepatic failure

Adequate

Viral Clearance/Immunity

Fibrosis and/or Cirrhosis

Robust

Acute Liver Failure

ACUTE VIRAL INFECTION

HCC

Death

CHRONIC VIRAL INFECTION

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Hepatic Immune Cell Regeneration The liver has an extraordinary capacity to regenerate following severe injury from toxins and/or invading pathogens. Liver regeneration is divided into three phases. In the primary phase hepatocytes are rendered sensitive to growth factors. In the growth phase the cells enter the cell cycle. Finally liver growth is terminated via an equally impressive coordinated network of inhibitory pathways. During liver regeneration immune cells repopulate the liver and also regulate the process. Partial hepatectomy (PH) animal models provide a useful tool for studying liver regeneration. The mechanism by which nonparenchymal cells, such as immune cells, repopulate the liver after injury is largely unknown. Six hours after PH, immature liver DCs increase significantly reaching prehepatectomy levels 24 h later. DCs may also influence hepatocyte proliferation [250]. Thirty six hours after PH, macrophages and KC repopulate the liver. Interestingly, in this study, NK/NKT cells were not shown to accompany KCs, whereas others have shown that 4–12 h after PH there is a marked hepatic lymphocytosis predominantly made up of NK/NKT cells [251]. One explanation may be that hemopoietic stem cells in the liver can differentiate into several kinds of lymphocyte including NK/NKT cells [252], implying that these cells may develop within the liver after PH. However these cells are thought to be functionally immature [253, 254]. Similarly, CD3+ T-cells can develop in situ within the liver [251]. Importantly, IL-12 can impair liver regeneration and accumulation of NKT [255]. As discussed above, liver regeneration in chronic hepatitis occurs in an inflammatory microenvironment and requires cross-talk between immune and parenchymal cells. After injury, liver regeneration is achieved largely as a result of hepatocyte and BEC proliferation. If hepatocye proliferation is inhibited during liver regeneration it may be undertaken by the putative hepatic progenitor cells (Oval cells), which proliferate, migrate, and differentiate into hepatocytes and BEC. All innate immune cells have an inhibitory role during liver regeneration except KCs. It is known that a lack of KCs can impair liver regeneration [256]. KC-derived IL-6, IL-1,TGFb, and TNFa are pivotal cytokines during regulation of hepatocyte proliferation as well as other aspects of restoration of liver tissue [257] IL-6 stimulates cell cycle entry via STAT3 activation [258], and TNFa may promote IL-6 release early in liver regeneration [259] but later TNFa signaling is

R. H. Bhogal and S. C. Afford

inhibitory via a TGFb-dependent process. NK cells also regulate liver regeneration [260] and exert an inhibitory effect. Immunosuppressants, such as FK506 and Cyclosporin A [261], improve liver regeneration by inhibiting NK cells [262] possibly by ensur­ing that NK-cell derived IFNg cannot activate STAT1 thereby ameliorating the suppression of STAT3 by STAT1 [246]. Anti-NK1.1 antibody inhibits NK/NKT cells again promoting liver regeneration [253]. Of note, PH in severe combined immunodeficient (SCID) mice does not attenuate liver regeneration implying that many immune cells regulate liver regeneration. In general proliferating hepatocytes express, stress induced ligands such as MHC class I chain-related A (MICA), MICB and UL-16-binding proteins [263]. However hepatocytes also express FasL and TRAIL and this may encourage NK cell mediated death. It is important to note however that alymphoid mice die from acute liver failure. This is possibly because they are unable to mount an adequate oval cell response, indicating paradoxically that mice harboring NK or T-cells ­survive as these cells may help oval cell development [264]. The role of NKT cells in liver regeneration is controversial. They are considered negative regulators as evidenced by CD1d−/− NKT cells resisting impaired liver regeneration [265]. Furthermore in muring HBV models, liver regeneration is impaired by IFNg secreting NKT cells with the absence of both improving liver regeneration. However after PH in mice hepatic NKT cell numbers are increased [251] and decreased hepatocyte mitosis is noted in hepatectomized CD1d-knockout mice that lack NKT cells [266]. Moreover a-Galcer activated NKT cells posthepatectomy improve hepatic regeneration via TNFa - and Fas/FasL-dependent pathways [266]. This also suggests that TNFa and Fas play beneficial roles during hepatic regeneration [267, 268]. Again this theory is controversial as some authors suggest that activated hepatic NKT cells can cause severe liver injury post-PH via the same mechanism [255]. NK/NKT cell derived IFNg negatively regulates hepatic regeneration in the murine PH model [246] but NKT cells can alter their response in liver regeneration by decreasing IL-6 production and increasing TGFb ­secretion as detected by Smad2 phosphorylation [269] giving rise to differential effects. NKT cells may also interact with oval cells affecting the ability of these cells to undertake regenerative function. Although IFNg adversely effects hepatocyte proliferation it may help oval cell function [269].

8  Immune Cell Communication and Signaling Systems in Liver Disease

During liver regeneration T-cells within the liver are increased in terms of absolute number and proportion [251, 255]. Indeed IL-1 and TNFa released by KCs and HSEC can induce T-cell activation during liver regeneration but the significance of this remains equivocal [270]. However recent studies do show that CD4+ T-cells localize to oval cells and CD8+ T-cells to bile ducts helping their regeneration although earlier studies had shown that adoptive transfer of T-cells reduced liver cell proliferation after PH [271]. The role of the B-cell in liver regeneration remains to be defined. Isolated studies have shown however that LPS infection after PH increases IgM antibody production and may mediate liver injury but whether there is an effect on liver regeneration is not known [272]. In summary the immune system has a complex role to play in liver regeneration as well as disease. The process is likely to be more intricate than outlined above as other regulators, such as complement, may regulate cytokine responses and prosurvival signals [272]. Further research in this field may help with novel treatments for liver regeneration post liver transplantation and liver injury.

Summary

›› The human liver is an immunologically dis-

tinct organ wherein the innate and adaptive immune responses are uniquely linked to provide protection against a wide range of inflammatory, infectious or toxic insults. The interplay between immunological cells located within the liver influences the phenotype of the eventual immune response. This process is mediated through cell-cell contact between immune cells and by the secretion of soluble mediators such as cytokines. These numerous and often complex interactions modulate the response to liver diseases. We have discussed , in detail, the role of both innate and adaptive immune cells in shaping the hepatic immune response along with their specific functions in liver diseases such as viral hepatitis. The emerging role of Th17 and T-regulatory cells in diseases such as autoimmunity is also highlighted.

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Multiple Choice Questions 1. Under which category of cell surface receptor the Toll-like receptors (TLRs), which are integral in initiating the hepatic immune response, are classi­ fied? (a) Inonotropic receptors (IRs) (b) Guanylyl cyclase receptors (GCRs) (c) Receptor tyrosine kinases (RTKs) (d) Pattern recognition receptors (PRR) (e) G-protein coupled receptors (GPCRs) 2. Which cytokine is required by dendritic cells to promote a Th2 cytokine response, the lack of which may contribute to severe hepatocellular damage? (a) IL-1 (b) IL-6 (c) IL-8 (d) IL-12 (e) IL-17 3. During viral hepatitis which CXCRs are involved in the hepatic recruitment of type1 cytokine T-cells and NK cells? (a) CXCR 7 and CXCR 3 (b) CXCR 5 and CXCR 7 (c) CXCR 2 and CXCR 4 (d) CXCR 3 and CXCR 4 (e) CXCR 3 and CXCR 5 4. In liver allograft rejection (acute and chronic), mature effector T-cells target which cells types? (a) Bile duct epithelial cells (BECs) and posthepatic venules (b) BECs and vascular endothelium (c) BECs, hepatocytes and vascular endothelium (d) Hepatocytes and BECs (e) Vascular endothelium and posthepatic venules 5. At the onset of viral hepatitis, dendritic cells are stimulated through which TLRs to initiate an immunological response? (a) TLR 3–7 (b) TLR 1–4 (c) TLR 7–9 (d) TLR 12 (e) TLR 8 and 9

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Part The Pathways

II

9

Toll-Like Receptors Gyongyi Szabo and Pranoti Mandrekar

Introduction Since the discovery of toll-like receptors (TLRs) in ­Dro­sophila in 1996 [1], scientific knowledge has exploded on the role of pattern recognition receptors in immunity, host defense, and in various noninfectious conditions including liver diseases. Studies demonstrated that TLR expression in immune and nonimmune cells plays an important role in tissue homeostasis, response to infections, and injury in the liver. Links between TLR activation and modulation of functions of TLRs in liver diseases are being discovered. Here, we discuss the characteristics and signaling pathways of TLRs and summarize current knowledge on their role in liver diseases.

Toll-Like Receptors and Their Ligands TLRs are evolutionarily preserved receptors that have the unique capability to sense microbe-specific sequences in pathogens so as to provide danger signals to the host [2]. To date, 11 human and 13 mouse TLRs have been identified, each with distinct pathogen associated molecular pattern (PAMP) recognition capacity from various microorganisms including bacteria, viruses, protozoa, and fungi [3]. TLRs are type I transmembrane glycoproteins that share a common basic structure, including an extracellular ligand recognition domain that contains repeated leucine-rich repeats, a short transmembrane domain, and an intracellular C-terminal, TIR domain [4]. The G. szabo () Department of Medicine, University of Massachusetts Medical School, 364 Plantation Street, Worcester, MA 01605, USA e-mail: [email protected]

TIR domain provides means for interaction of TLRs with intracellular signaling molecules to trigger downstream activation [2–5]. Ligand recognition of TLRs results in their oligomerization that is associated with induction of downstream signaling. TLRs, such as TLR4 or TLR3 form homodimers, while TLR2 can homodimerize or heterodimerize with TLR1 (TLR1/2), TLR6 (TLR2/6) or in humans with TLR10 (TLR2/10) [5]. The localization of TLRs within the cells correlates with their pathogen recognition profile. For example, TLR2, TLR1, and TLR6 are expressed on the cell surface as these TLRs recognize mostly extracellular danger signals (Fig. 9.1). TLR4, the receptor for LPS, and TLR5, a sensor of flagellin, are also expressed on the cellular membrane (Fig.  9.1). In contrast, TLRs that sense nucleic acid sequences, mostly from organisms entering the host cells, include TLR3, TLR7, TLR8, and TLR9. These TLRs are expressed in endosomes and are activated at low pH (Fig. 9.2) [2–5]. TLRs are classified into general groups based on the types of PAMPs they recognize. The ligands for each TLR are summarized in Table 9.1. Briefly, TLR1, 2, 4, and 6 recognize mostly lipids. TLR2 forms heterodimers with TLR1 or TLR6 to discriminate a wide range of PAMPs including peptidoclycans, lipopeptides and lipoproteins from gram-positive bacteria, mycoplasma lipoproteins and fungal zymosan [2–4]. TLR1/2 and TLR2/6 can discriminate triacyl- and ­diacyl-lipopeptide, respectively [2–5]. TLR4, with its extracellular coreceptors CD14 and MD-2, recognizes lipopolysaccharide from gramnegative bacteria. LPS consists of three parts: lipid A, a core oligosaccharide, and an O side chain of which the active portion is lipid A which provokes TLR4 activation and causes septic shock [6, 7]. TLR5 and 11 recognize protein ligands. TLR5 recognizes flagellin, and mouse TLR11 recognizes a yet

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_9, © Springer-Verlag Berlin Heidelberg 2010

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G. szabo and P. Mandrekar TLR1

TLR2

TLR2

TLR6

TLR2

TLR5

TLR4

TLR4

TRAF6

MyD88

MyD88

MyD88

TIRAP

MyD88 IRAK1

TIRAP IRAK4

MyD88

TIRAP

MD2

extracellular space cytoplasm

TRAM TIRAP

TRIF

IKKε

TBK

IRF5

TAK1 p38

CD14

IRF3

JNK

P

NEMO IKKβ

AP-1

IKKα

NF-κB

IKBα p65 p50

Proinflammatory cytokines

P p65 p50

(TNFα, IL-6, IL-1, IL-12)

IRF3

IRF3

P IFNβ

nucleus

Fig. 9.1  Membrane associated toll-like receptors (TLRs) and their signaling adapters

extracellular space cytoplasm

endosome

endosome TLR7/8

TLR3

TRIF

MyD88

RIP TBK1

IRF3

TAK1 NEMO IKKα IKKβ

p

TRAF6 Ub IKKκ

IRAK4 IRAK11

TRAF3

IKKβ

TLR9

p

IRF7 TRAF3

IKBα

p

p IRF3 IRF3

p65 p50 p

p

Fig. 9.2  Endosomal TLR and ­signaling pathways

IRF3 IRF3

IFNß

p

p

IRF7 IRF7

p

p65 p50

p

IRF7 IRF7

IFNβ

inflammatory cytokines nucleus

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TLR Adapters and Signaling

Table 9.1  Toll-like receptor (TLR) and their ligands TLR TLR2

Ligand Peptidoglycan

Extracellular expression Endosomal expression

Pathogen Gram-positive bacteria Hemagglutinin Measles virus Core, NS3 Hepatitis C virus tGPI-mutin Parasite TLR2/1 Triacyl lipopeptide Bacteria/ mycoplasma TLR2/6 Diacyl lipopeptide Mycoplasma Zymosan Saccharomyces. cerevisiae TLR4 LPS Gram-negative bacteria Envelope proteins RSV, MMTV TLR 5 Flagellin Bacteria TLR 11 Uropathogenic bacteria Bacteria components Prolin-like Molecules Toxoplasma gondii TLR3 dsRNA Viruses Poly IC Synthetic compound TLR 7/8 ssRNA Viruses Imidazoquinolines Synthetic compound TLR9 CpG DNA Bacteria DNA Viruses Hemozin Plasmodium

unknown component of uropathogenic bacteria, and a component of Toxoplamsa gondii in mice; however, in humans TLR11 is not functional [5]. Nucleic acids derived from viruses or bacteria are ­recognized by TLR3, TLR7, TLR8, and TLR9 [8]. TLR3 senses double-stranded DNA (dsDNA); TLR7 detects synthetic imidazoquinoline-like molecules, guanosine analogs, single stranded RNA (ssRNA) derived from viruses, and small interfering RNA [2–4, 8]. Human TLR8 that has the highest homology with TLR7 participates in recognition of ssRNA. In mice, the function and ligands of TLR8 remain unknown. TLR9 recognizes CpG DNA motifs present in bacterial and viral genomes as well as non-nucleic acids such as hemozoin from the malaria parasite [3, 8]. The role of TLRs has recently been evaluated in sterile inflammation where microbial infection is not present. TLR4 in particular is at the interface of microbial and sterile inflammation by responding to both bacterial endotoxin and multiple endogenous ligands [9]. The significance of endogenous ligands and their receptor systems in the liver awaits further investigations.

TLRs trigger signaling from the cytoplasmic TIR domain via recruitment of different intracellular adaptors and culminate in the production of proinflammatory cytokines and/or Type I IFN to activate innate immune responses [3–5]. There are five TIR domaincontaining adaptor proteins: MyD88 (myeloid differentiation primary response gene 88), TIRAP (TIR domain containing adaptor protein, also known as Mal) TRIF (TIR domain containing adaptor inducing IFNb) TRAM (TRIF-related adaptor molecule) and SARM (sterile a and HEAT-Armadillo motifs-­containing protein) [4, 10]. The MyD88-dependent signaling is common to all TLRs except for TLR3, and it is essential for the induction of proinflammatory cytokines via NF-kB activation [7, 10, 11]. Interaction of the intracellular TIR domain of TLRs with the TIR domain of TIRAP/Mal results in MyD88 recruitment. As summarized in Figs. 9.1 and 9.2, the MyD88 signaling recruits members of the IL-1 receptor associated kinase (IRAK) family and hyper-phosphorylates IRAK1, leading to dissociation of IRAK1 from MyD88 and interaction with TNF receptor-associated factor 6 (TRAF6). IRAK1/TRAF6 form a complex with transforming growth factor beta (TGF-b)-activating kinase (TAK1) leading to activation of IkB kinase (IKK) a and b, phosphorylation/degradation of IkB proteins to result in NF-kB nuclear translocation and activation of NF-kB dependent genes [7, 10, 11]. These involve proinflammatory cytokine genes such as TNFa, IL-1b, IL-6 and other stress-inducible genes [11]. Ligand stimulation of TLR7/8 and TLR9 also results in recruitment of MyD88 resulting in IRAK1/4 kinase activation and IKKe activation [2, 3, 8]. Through IKKe, IRF7 is phosphorylated and the IRF7 dimer activates IFNa production (Fig. 9.2). Through activation of TAK1, TLR7/8 and 9 also can activate the IKK complex to result in NF-kB and proinflammatory cytokine induction (Fig. 9.2). The MyD88-independent signaling initiated by TLR3 or TLR4 activation results in the activation of IFN-inducible genes and the late activation of NF-kB [12, 13]. In TLR4-induced signaling, the adapter molecules TRAM and TRIF are recruited (Fig. 9.1) while TLR3 activation recruits only TRIF (Fig.  9.2). TRIF activates TBK1/IKKe that phosphorylates IRF3 and IRF3 nuclear translocation induces production of type I IFNs [12–14]. The MyD88-independent pathway also

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activates NF-kB, an important transcription factor in alcoholic liver disease through TRIF and TRAF6 interactions [4, 14]. A late phase NF-kB activation is mediated by TRAF6 engaging TRIF, leading to activation of the IKK kinase ­complex that phosphorylates the inhibitory kB (IkB) [15, 16]. This leads to IkB dissociation from p65/p50 that in turn can translocate into the nucleus and bind the kB-site in the promoter region of proinflammatory genes [11, 15, 16].

Negative Regulation of TLR Signaling While TLR sensing is a critical component of host defense against pathogens, excessive activation of TLR signaling may cause liver damage. Indeed, TLR signaling is strictly regulated at various levels by intracellular negative TLR regulators (Fig. 9.3). The membrane-associated TLRs, TLR2, and TLR4 signaling can be inhibited though RP105 (a homologue of TLR4, essential for LPS recognition associated with MD-1 in B cells), ST2L (also known as the IL-33 receptor), single

RP-105 (TLR2,4)

immunoglobuling IL-1R-related molecule (SIGGR/ TIR8), suppressor of cytokine signaling-1 (SOCS-1), and Toll-interacting protein (Tollip), IRAK-M, A20, MyD88s (MyD88-short, lacks the interdomain), sterile alpha and TRI-motif-containing 1 (SARM1/MyD885) and TRIAD3A [reviewed in 17, 18]. On the other hand, endosomal TLRs such as TLR3 and 9 are negatively regulated by SIGRR, ST2L, SOCS-1, TRIAD3A, IRAK-M, and A20 [reviewed in 17, 18]. Intracellular IRAK activity is inhibited by IRAK-M, a molecule described in monocytes but also expressed in other cell types, SOCS-1, and Toll-interacting protein (Tollip) [18–20]. Loss of IRAK-M upregulation was associated with an increase in proinflammatory cytokine activation and loss of TLR tolerance in monocytes of patients with chronic HCV infection [17, 18]. TIRAP, also known as Mal, is negatively regulated by SOCS-1, ST2L and TRIAD3A [18–20]. The TRIF-dependent pathway is inhibited by sterile alpha and ­TRI-motif-containing 1 (SARM1/MyD885) and TRIAD3A [reviewed in 18–20]. A20 is a protease dequitylating enzyme shown to restrict MyD88-dependent signaling and TRIF-dependent NF-­ kB activation through deubiquitylating TRAF6 [18–20].

CD14

MyD88s MyD88 TIRAP

SARM (TLR3, 4)

TRIF

TLR TLR

SOCS-1

(TLR4, 9)

IRAKs

Tollip

Fig. 9.3  Negative regulators of TLR signaling

ST2L (TLR2, 4, 9)

(TLR4, 9)

(TLR2,4)

A20

SIGRR (TLR4, 9)

SOCS-1

(TLR4, 9)

IRAK-M

(TLR4)

TRAF6

Inflammatory gene expression

TRIAD3A (TLR4, 9)

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The particular role of these different negative regulators in liver diseases awaits further understanding.

TLR Expression in the Liver The function, expression, and clinical implication of TLRs in the liver are being explored. Studies so far have shown that various TLRs are expressed on all cell types of the liver and manifest different actions in various liver disease [20, 21].

Kupffer Cells Kupffer cells, the resident macrophages of the liver, are important in antigen recognition and proinflammatory cytokine production. These cells express TLR2, TLR4, and its coreceptors CD14 and MD-2; respond to their respective ligands resulting in NF-kB activation, production of pro- and anti-inflammatory cytokines; and reactive oxygen species (ROS) [22, 23]. Kupffer cells recognize lipopolysaccharide or endotoxin in circulation via TLR4 resulting in the ­production of pro- and anti-inflammatory cytokines and chemokines. Cross-regulation and cross-talk of TLRs on Kupffer cells is evident from studies that showed that poly I:C pretreatment down-modulated TLR4 expression on Kupffer cells, also suggesting a role for IRF3 [24]. Selective inhibition of Kupffer cells using gadolinium chloride or dechloromethylene diphosphonateliposomes in various models of liver disease alters hepatic cytokine expression in response to TLR 2, 4, and 7/8 ligands, indicating a pivotal role for Kupffer cells in liver disease [25].

Dendritic Cells Besides Kupffer cells, the important innate immune cells of the liver, myeloid (MDC), and lymphoid (plasmacytoid, PDC) dendritic cells also contribute to hepatic immunity [21]. Both hepatic MDC and PDC recognize and present antigens to T cells and have distinct TLR expression and cytokine production profile [26]. However, liver DCs have been shown to exhibit low inherent T cell stimulatory capacity and determine

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the balance between tolerance and immunity [26]. Hepatic PDCs express TLRs 7 and 9 and produce large amounts of IFNa whereas MDCs express TLR2, 3, 4 and TLR9 and produce proinflammatory cytokines and IFNb but not IFNa [26]. Notably, a variety of other immune cells particularly hepatic NK cells are rich in TLR1, 2, 4, 5 and 9, while B cells express high levels of TLR1, 6, 7, and 9 [27].

Hepatocytes Hepatocytes, the predominant nonparenchymal cell compartment in the liver express mRNA for all TLRs [28]. Uptake and clearance of endotoxin from circulation by hepatocytes has suggested functional activity of TLR2 and TLR4 [29]. While TLR2 or TLR4 activation led to NFkB binding, TLR3 stimulation with polyI:C resulted in activation of the type 1 IFNs in ­primary hepatocytes and cell lines [30].

Endothelial Cells Liver endothelial cells lining the hepatic sinusoids express TLR4, upregulate NFkB activity, and produce proinflammatory cytokines such as TNFa as well as ROS in response to TLR4 ligands . Further, LES can become tolerant to LPS [31].

Biliary Epithelial Cells The biliary epithelium expresses mRNA for all TLRs, the TLR4 coreceptor, MD-2 and negative regulators of TLR signaling (IRAK-M, Tollip, SIGGIR and ST-2) [32]. Harada et al. identified that biliary epithelial cells develop tolerance to TLR2 and TLR4 ligands via induction of IRAK-M, a negative regulator of TLR signaling [33].

Stellate Cells Stellate cells express TLR4, TLR2 and CD14, respond to TLR2 and 4 ligands by activation of JNK, ERK and NFkB, and production of proinflammatory cytokines,

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chemokines and expression of adhesion molecules [34]. These cells are located in the Space of Disse and are the prime producers of extracellular matrix in the liver [34, 35]. TLR4 activation has been shown to lead to profibrogenic features such as enhancement of TGF-b signaling and collagen production resulting in stellate cell activation [36].

Clinical Implications of TLR Activation in Liver Disease Viral Hepatitis TLRs are being increasingly recognized in viral hepatitis in various capacities for instance as pathogen sensors, modifiers of immune response, and potential targets for therapeutic modification. Hepatitis B, a DNA virus infection is mostly selflimited but results in chronic infection in about 20% of adults. Decreased expression of TLR1, 2, 4, and 6 transcripts were found in PBMCs of patients with chronic HBV infection and the decreased TLR levels correlated with an impaired cytokine production after TLR4 or TLR2 ligand stimulation [37]. In an in vivo model of chronic HBV infection, administration of ligands specific for TLR3, TLR4, TLR5, TLR7, and TLR9, but not TLR2, inhibited HBV replication in the liver within 24  h in a type I interferon-dependent manner [38, 39]. This was associated with induction of antiviral mediators in dendritic cells but not in hepatocytes during in vivo treatment of HBV infection with TLR ligands. Immune cells are the main producers of type 1 IFNs and their stimulation via TLR 3, 4, 7, 8 and 9, but not TLR2, leads to production of IFNa, suggesting that endogenous IFNs play a critical role in antiviral defense [40, 41]. Indeed, type 1 IFN and IFN-inducing TLR ligands (CpG) could be successfully used as therapeutic agents in viral hepatitis ,or as potent adjuvants for anti-HBV vaccines [42]. In vitro data also support the contention that the innate immune system of the liver controls HBV replication after activation with TLR antagonists [43]. Hepatitis C virus results in chronic hepatitis in about 70–80% of infected individuals. Recent reports using HCV subgenomic replicon-harboring Huh cells showed that HCV has developed strategies to interfere

G. szabo and P. Mandrekar

with IFN pathways through pattern recognition signals. NS3/4A protein interacts directly with TBK1 to decrease TBK1/IRF3 interaction, while NS3 protein alone induces degradation of the TLR adaptor TRIF, both leading to downregulation of IRF3 activity and hamper IRF-3 mediated type 1 IFN induction [44]. These data indicate that HCV subverts TLR signaling pathways to escape from clearance in hepatocytes. In addition, both TLR9- and TLR7/8-induced IFN-a production is diminished in peripheral plasmacytoid ­dendritic cells of HCV infected patients [45]. TLR2 and TLR4 appear to be involved in immune dysregulation in chronic HCV infection. HCV core and NS3 proteins activate monocytes through TLR2 to produce IL-8, IL-6 and TNF-a, and inhibit differentiation and antigen-presenting functions of myeloid dendritic cells [46]. Marked upregulation of TLR2 and TLR4 was reported in patients with chronic HCV infection irrespective of HCV genotype and viral load and was detected in hepatocytes, Kupffer cells, and peripheral blood monocytes [47, 48]. The increased inflammatory cytokine activation in monocytes of HCV patients has been linked to a loss of TLR tolerance due to ongoing TLR4 (through LPS) and TLR2 activation (by HCV core) permitting ongoing proinflammatory cell activation [49].

Alcoholic Hepatitis The role of TLR4 has been defined in alcoholic liver disease where gut-derived endotoxin is implicated in Kupffer cell activation to induce proinflammatory cytokines such as TNF-a, leading to hepatocyte damage. Kupffer cell inactivated, CD14-deficient, TLR4 mutant or TLR4-deficient mice showed significant reduction in alcohol-induced liver injury despite increased endotoxin levels [49, 50]. Investigation of the TLR4 downstream signaling in mice revealed that the lack of the MyD88 TLR adapter did not prevent alcohol-induced steatohepatitis suggesting that the MyD88-independent TLR4 pathway is affected by alcohol. Indeed, the IRF3-inducible gene, IRF7 was increased in mouse Kupffer cells after alcohol feeding [51]. It has also been shown that chronic alcohol upregulates the expression of several TLRs and CD14 at the mRNA level and sensitizes to the corresponding TLR ligands to increased TNF-a production [52] (Table 9.2).

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Table 9.2  Role of TLRs in pathogenesis of liver diseases Liver Diseases Hepatitis B virus

Abnormalities in expression and function of TLRs TLR 1, 2, 3, 4, 6 TLR 7, 9

Hepatitis C virus

TLR 1, 2, 3, 4, 6 TLR 7, 8, 9

Alcoholic hepatitis

TLR 2, 3, 4, 6 TLR 7, 8, 9 CD14

Nonalcoholic fatty liver Liver fibrosis Primary biliary cirrhosis Acute liver failure

TLR 4

Ischemia-reperfusion injury

TLR 2, 4 TLR 2, 3, 4, 5 TLR 9 TLR 4 TLR 9 TLR 2, 4

Mechanisms of action and consequences

References

TLR 1, 2, 4, 6, 7, 9 – downregulation of mRNA and cytokine production TLR 3, 9 – decreased HBV replication in Kupffer cells and liver sinusoidal endothelial cells TLR 3 – recognition of virus; Hijacked by NS3/4A TLR 2 – dendritic cells, monocytes recognition of NS3 and HCV core TLR4 – hyperresponsive monocytes to LPS; Loss of TLR tolerance TLR 7, 9 – decreased IFNa by plasmacytoid dendritic cells TLR 2, 3, 6, 7, 8, 9 – upregulation of mRNA in monocytes and T cells TLR 4 – pathogenesis through gut-derived LPS TLR 2, 3, 4, 5, 6, 9, CD14 – upregulation in the liver TLR 2, 3, 4, 5, 6, 9, CD14 – increased sensitization to ligands and pro-inflammatory cytokine production TLR 4 – pro-inflammatory cytokine production

[37] [38, 39] [43, 44] [46] [48] [46] [47, 48] [49, 51] [50–52] [52] [53, 54]

TLR 2, 4 – stellate cell activation TLR 3 – upregulation of mRNA TLR 2, 3, 4, 5, 9 – sensitization to ligands in monocytes TLR 4, 9 – sensitization to LPS injury

[34, 35, 56] [57] [57, 59] [62]

TLR 2, 4 – activation of inflammatory responses leading to injury

[63]

Nonalcoholic Fatty Liver Disease Nonalcoholic steatohepatitis characterized by lipid accumulation and inflammation in the liver can lead to fibrosis. In the methionine-choline deficient diet model of NASH, enhanced susceptibility to TLR4 and not to TLR2 ligands was observed. The role of endogenous TLR4 stimulation was suggested by a recent study where the loss of TLR4 attenuated hepatic lipid accumulation and mRNA levels of liver fibrogenic markers in a MCD diet-induced steatohepatitis [53]. Unlike TLR4, TLR2 deficiency did not prevent NASH in the MCD model [54].

coreceptor, CD14 and MyD88 were found to be critical in hepatic fibrogenesis induced by bile duct ligation [34, 35]. TLR4 expressed on hepatic stellate cells and KCs plays a central role in induction and perpetuation of liver fibrosis. Hepatic stellate cells are directly activated by LPS through TLR4 [34, 35]. TLR4activated HSC produce chemokines that recruit KCs to the site of injury [55]. TLR4 signaling also activates TGF-b production and inhibits the TGFbRdecoy receptor towards a net result of profibrogenic ­activation [36, 56].

Other Liver Conditions Liver Fibrosis Hepatic fibrosis develops in livers affected by chronic hepatitis caused by infectious (HCV, HBV, HIV) or noninfectious causes (alcohol, metabolic disease, biliary obstruction, autoimmune process, etc). TLR4, its

Emerging data indicates that TLR4 and TLR9 may also be involved in biliary diseases such as PBC [57,  58]. In liver regeneration, there was no requirement for TLR2, TLR4 or TLR9 after partial hepatectomy. However, the  TLR adapter, MyD88 played a crucial role in the activation of NF-kB and induction of

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TNF-a, IL-6 and early immediate genes such as c-myc, c-fos and c-jun during liver regeneration in a partial hepatectomy model [59, 60]. Excessive TLR activation by administration of TLR4 or TLR3 ligands suppressed liver regeneration after partial hepatectomy [61]. Experimental models in mice indicate that exposure to heat-killed P. acnes or TLR9 plus TLR2 ligands results in sensitization to TLR4-induced liver injury [62]. Although a role for ischemia-reperfusion injury in the liver can be independent of LPS recognition due to oxidative stress, evidence suggests a role for TLRs in ischemic-reperfusion injury [63]. LPS levels were elevated early after reperfusion and multiple components of the LPS signaling pathway, including CD14, LBP, as well as TLR2, are activated during ischemia/ reperfusion injury after liver transplantation [64]. Interestingly, TLR4 activation mediates inflammatory responses via both IRF3 and MyD88-dependent pathways after ischemia/reperfusion [60, 63, 64]. TLR4deficient, but not wild-type or TLR2-deficient, mice were protected against ischemia/reperfusion induced liver damage, suggesting an important role for TLR4.

G. szabo and P. Mandrekar

TLRs could also be targets to boost immune responses. Basic and clinical investigations are ongoing with TLR7 and TLR9 ligands that may prove to be effective in boosting antiviral immunity or as adjuvants for vaccines in liver diseases. With the promise of recent discoveries, careful investigation of TLR in therapy may provide novel approaches for the amelioration of different liver diseases.

Summary

›› TLRs ›› ›› ››

›› Future Perspectives Increasing evidence suggests that activation or modulation of various TLRs has a role in the pathogenesis of several liver diseases. However, further studies are needed to specifically define the involvement of individual TLRs particularly in human liver diseases. Knowledge so far indicates that modulation of TLR signaling could be considered a therapeutic target in different liver diseases. Based on the involvement of LPS in various types of liver injury (acute liver failure, alcoholic hepatitis) it is tempting to speculate that pharmacological inhibition of endotoxin-induced, TLR4mediated responses could be beneficial. Strategies explored to date include blocking of TLR4 signaling by targeting the TLR4 coreceptor, MD-2 [65], development of a synthetic nontoxic lipid A derivative with TLR4 blocking capacity [66] and low molecular weight mimic of Toll-IL-1 receptor/resistance domain that inhibits IL-1R-mediated responses [67]. TLR signaling could be disrupted at a downstream level (MyD88, TRIF, IRAK or TRAF proteins), by using short ­interfering RNAs [68], however, the safety of in vivo application of such an approach is yet to be tested.

›› ›› ›› ››

are evolutionarily preserved receptors that have the unique capability to sense PAMP. TLRs are transmembrane glycoproteins which upon ligand binding oligomerize to induce downstream signaling. TLR4 recognizes LPS, TLR5 flagellin, TLR3, TLR7, TLR8, TLR9 nucleic acids derived from viruses and bacteria. MyD88 is a common adaptor to all TLRs except for TLR3 and it is essential for the induction of proinflammatory cytokines via NF-kB activation. Kupffer cells express TLR2, TLR4 and its coreceptors MD-2 and CD14. In hepatocytes TLR2 and TLR4 stimulation activates NF-kB, TLR3 type I interferon. Hepatitis C virus subverts TLR signaling pathways to escape from clearance in hepatocytes. MyD88 is important during liver regener­ ation. TLR4 has been implicated in the pathogenesis of NASH, in fibrosis and in ischemia/reperfusion injury.

Multiple Choice Questions 1. Which one of the following TLRs does not recognize nucleic acid sequences, but extracellular signal? (a) TLR3 (b) TLR5 (c) TLR7 (d) TLR8 (e) TLR9

9  Toll-Like Receptors

2. What is true regarding TLR4? (a) It recognizes LPS (b) Its coreceptor is CD14 (c) Its coreceptor is MD-2 (d) It dimerizes (e) All the above statements are true 3. What is true regarding the cell expression of TLR4? (a) It is expressed on hepatocytes (b) It is expressed on cholangiocytes (c) It is expressed on Kupffer cells (d) It is expressed on stellate cells (e) It is expressed on all the above cited cell types 4. HCV subverts TLR signaling by (a) Stimulating the TBK1/IRF3 interaction (b) Upregulating IRF3 activity (c) Preventing degradation of TRIF (d) NS3 and NS3/4A proteins (e) Downregulation of TLR2 and TLR4 5. Reduction in alcohol-induced liver injury were observed in following mice except (a) CD-14 deficient (b) TLR4 deficient (c) With inactivated Kupffer cells (d) MyD88 deficient (e) TLR4 mutant

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158 29. Matsumura T, Degawa T, Takii T, Hayashi H, Okamoto T, Inoue J et al (2003) TRAF6-NF-kB pathway is essential for interleukin-1-induces TLR2 expression and its functional response to TLR2 ligand in murine hepatocytes. Immunology 109:127–136 30. Li K, Chen Z, Kato N, Gale M Jr, Lemon SM (2005) Distinct poly(I-C) and virus-activated signaling pathways leading to interferon-beta production in hepatocytes. J Biol Chem 280:16739–16747 31. Urig A, Banafsche R, Kremer M, Hegenbarth S, Hamann A, Neurath M, Gerken G et al (2005) Development and functional consequences of LPS tolerance in sinusoidal endothelial cells of the liver. J Leukoc Biol 77:626–633 32. Harada K, Isse K, Nakanuma Y (2006) Interferon (gamma) accelerates NF-(kappa) B activation of biliary epithelial cells induced by Toll-like receptor and ligand interactin. J Clin Pathol 59:184–190 33. Harada K, Isse K, Sato Y, Ozaki S, Nakanuma Y (2006) Endotoxin tolerance in human intraheptatic biliary epithelial cells is induced by upregulation of IRAK-M. Liver Int 26: 935–942 34. Paik YH, Schwabe RF, Bataller R, Russo MP, Jobin C, Brenner DA (2003) Toll-like receptor 4 mediates inflammatory signaling by bacterial lipopolysaccharide in human hepatic stellate cells. Hepatology 37:1043–1055 35. Brun P, Castagliuolo I, Pinzani M, Palu G, Martines D (2005) Exposure to bacterial cell wall products triggers an inflammatory phenotype in hepatic stellate cells. Am J Physiol Gastrointest Liver Physiol 289:G571–G578 36. Seki E, DeMinicis S, Osterreicher CH, Klue J, Osawa Y, Brenner DA et al (2007) TLR4 enhances TGF-beta signaling and hepatic fibrosis. Nat Med 13:1324–1332 37. Chen Z, Cheng Y, Xu Y, Liao J, Zhang X, Hu Y, Zhang Q et  al (2008) Expression profiles and function of Toll-like receptors 2 and 4 in peripheral blood mononuclear cells of chronic hepatitis B patients. Clin Immunol. 128:400–408 38. Isogawa M, Robek MD, Furuichi Y, Chisari FV (2005) Tolllike receptor receptor signaling inhibits hepatitis B virus replication in vivo. J Virol 79:7269–7722 39. McClary H, Chisari KR, FV GLG (2000) Relative sensitivity of hepatitis B virus and other hepatotropic viruses to the antiviral effects of cytokines. J Virol 74:2255–2264 40. Ito T, Amakawa R, Fukuhara S (2002) Roles of Toll-like receptors in natural interferon-producing cells as sensors in immune surveillance. Hum Immunol 63:1120–1125 41. Szabo G, Dolganiuc A (2007) The role of plasmacytoid dendritic cell-derived IFNa in antiviral immunity. Crit Rev Immunol 28:61–94 42. Verthelyi D, Wang VW, Lifson JD, Klinman DM (2004) CpG oligodeoxynucleotides improve the response to hepatitis B immunization in healthy and SIV-infected rhesus macaues. AIDS 18:1003–1008 43. Otsuka M, Kato N, Moriyama M, Taniguchi H, Wang Y, Dharel N, Kawabe T et al (2005) Interaction between the HCV NS3 protein and the host TBK1 protein leads to inhibition of cellular antiviral responses. Hepatology 41:1004–1012 44. Li K, Foy E, Ferron JC, Nakamura M, Ferreon AC, Ikeda M, Ray Sc et al (2005) Immune evasion by hepatitis C virus NS3/4A protease-mediated cleavage of the Toll-like receptor 3 adaptor protein TRIF. Proc Natl Acad Sci USA 102: 2992–2997

G. szabo and P. Mandrekar 45. Dolganiuc A, Chang S, Kodys K, Mandrekar P, Bakis G, Cormier M, Szabo G (2006) HCV Core protein-induced, monocyte-mediated mechanisms of reduced IFN-alpha and plasmacytoid dendritic cell loss in chronic HCV infection. J Immunol 177:6758–6768 46. Dolganiuc A, Oak S, Kodys K, Golenbock DT, Finberg RW, Kurt-Jones E, Szabo G (2004) Hepatitis C core and nonstructural 3 protein trigger toll-like receptor 2-mediated path­ways and inflammatory activation. Gastroenterology 127: 1513–1524 47. Visvanathan K, Skinner M, Chang J, Lewin S, Thompson A, Locarnini S, Riordan SM (2005) Up-regulation of Toll-like receptors expression in chronic hepatitis C: correlation with circulating pro-inflammatory cytokine levels and hepatic necro-inflammatory activity. Hepatology 42:547A 48. Dolganiuc A, Norkina O, Kodys K, Catalano D, Mandrekar P, Bakis G, Marshall C, Szabo G (2007) Viral and host factors induce macrophage activation and loss of toll-like receptor tolerance in chronic HCV infection. Gastroenterolgy 133:1627–1636 49. Uesgi T, Froh M, Arteel GE, Bradford BU, Thurman RG (2001) Toll-like receptor 4 is involved in the mechanism of early alcohol-induced liver injury in mice. Hepatology 34:101–108 50. Yin M, Bradford BU, Wheeler MD, Uesugi T, Froh M, Goyert SM, Thurman RG (2001) Reduced early alcoholinduced liver injury in CD14-deficient mice. J Immunol 166:4737–4742 51. Hritz I, Mandrekar P, Velayudham A, Catalano D, Dolganiuc A, Kodys K, Kurt-Jones E, Szabo G (2008) The critical role of Toll-like receptor 4 in alcoholic liver disease is independent of the common TLR adaptor, MyD88. Hepatology 48:1224–1231 52. Gustot T, Lemmers A, Moreno C, Nagy N, Quertinmont E, Nicaise C, Franchimont D et al (2006) Differential liver sensitization to toll-like receptor pathways in mice with alcoholic fatty liver. Hepatology 43:989–1000 53. Rivera CA, Adegboyega P, van Rooijen N, Tagalicud A, Allman M, Wallace M (2007) Toll-like receptor-4 signaling and Kupffer cells play pivotal roles in the pathogenesis of non-alcoholic steatohepatitis. J Hepatol 47:571–579 54. Szabo G, Velayudham A, Romics L Jr, Mandrekar P (2005) Modulation of non-alcoholic steatohepatitis by pattern ­recognition receptors in mice: the role of Toll-like receptors 2 and 4. Alcohol Clin Exp Res 29:140s–145s 55. Zhang X, Yu WP, Gao L, Wei KB, Ju JL, Xu JZ (2004) Effects of lipopolysaccharides stimulated Kupffer cells on activation of rat hepatic stellate cells. World J Gastroenterol 10:610613 56. Kisseleva T, Brenner DA (2007) Role of hepatic stellate cells in fibrogenesis and the reversal of fibrosis. J Gastroenterol Hepatol 22:S73–S78 57. Wang AP, Migita K, Ito M, Takii Y, Daikoku M, Yokoyama T, Komori A, Nakamura M, Yatsuhashi H, Ishibashi H (2005) Hepatic expression of toll-like receptor 4 in primary biliary cirrhosis. J Autoimmun 25:85–91 58. Kikuchi K, Lian ZX, Yang GX, Ansari AA, Ikehara S, Kaplan M, Miyakawa H, Coppel RL, Gershwin ME (2005) Bacterial CpG induces hyper-IgM production in CD27(+) memory B-treated cells in primary biliary cirrhosis. Gastroenterology 128:304–312

9  Toll-Like Receptors 59. Seki E, Tsutsui H, Iimuro Y, Naka T, Son G, Akira S, Kishimoto T et al (2005) Contribution of Toll-like receptor/ myeloid differentiation factor 88 signaling to murine liver regeneration. Hepatology 41:443–450 60. Campbell JS, Riehle KJ, Brooling JT, Bauer RL, Mitchell C, Fausto N (2006) Proinflammatory cytokine production in liver regeneration is MyD88-dependent, but independent of CD14, TLR2, and TLR4. J Immunol 176:2522–2528 61. Sun R, Gao B (2004) Negative regulation of liver regeneration by innate immunity (natural killer cells/interferongamma). Gastroenterology 127:1525–1539 62. Velayudham A, Dolganiuc A, Kurt-Jones E, Szabo G (2006) Critical role for Toll-like receptors and the common adaptor, MyD88, in granulomas and sensitization to injury. J Hepatology 45:813–824 63. Zhai Y, Shen XD, O’Connell R, Gao F, Lassman C, Busuttil RW, Cheng G et al (2004) Cutting edge: TLR4 activation mediates liver ischemia/reperfusion inflammatory response via IFN regulatory factor 3-dependent MyD88independent pathway. J Immunol 173:7115–7119

159 64. Tsoulfas G, Takahashi Y, Ganster RW, Yagnik G, Guo Z, Fung JJ, Murase N et al (2002) Activation of the lipopolysaccharide signaling pathway in hepatic transplantation preservation injury. Transplantation 74:7–13 65. Visintin A, Halmen KA, Latz E, Monks BG, Golenbock DT (2005) Pharmacological inhibition of endotoxin responses is achieved by targeting the TLR4 co-receptor, MD-2. J Immunol 175:6465–6472 66. Kawata T, Bristol JR, Rossignol DP, Rose JR, Kobayashi S, Yokohama H, Ishibashi A et al (1999) E5531, a synthetic nontoxic lipid A derivative blocks the immunobiological activities of lipopolysaccharide. Br J Pharmacol 127: 853–862 67. Bartfai T, Behrens MM, Gaidarova S, Pemberton J, Shivanyuk A, Rebek J Jr (2003) A low molecular weight mimic of the Toll/IL-1 receptor/resistance domain inhibits IL-1 receptor-mediated responses. Proc Natl Acad Sci U S A 100:7971–7976 68. Xie FY, Woodle MC, Lu PY (2006) Harnessing in vivo siRNA delivery for drug discovery and therapeutic development. Drug Discov Today 11:67–73

TNF/TNF Receptors

10

Jörn M. Schattenberg and Mark J. Czaja

Introduction Tumor necrosis factor-a (TNF) is a pleiotropic cyto­ kine whose biological functions regulate the cellular responses of injury and repair, inflammation and immunity, and proliferation. In the liver, TNF exerts autocrine and paracrine effects that mediate a variety of pathophysiological states that involve liver injury and cell death and/or hepatocellular proliferation. Thus, TNF is a central regulator of hepatic physiology and delineation of the complex signaling pathways that mediate the disparate effects of this cytokine has contributed to our understanding of its function. In particular, investigations have attempted to determine how this factor could promote either cell proliferation or death in hepatocytes under different physiologic circumstances. With these studies has come an increased understanding of the complex events that determine whether a hepatocyte undergoes apoptosis or proliferation following TNF stimulation. This chapter will focus initially on signaling events that follow TNF ligand– receptor interaction, and subsequently on the precise functions of TNF signaling in specific pathophysiologic states. Although considerable progress has been made in defining TNF signaling pathways in hepatocytes, the challenge remains to determine how these signal cascades regulate disease states in order to mani­ pulate these pathways for the treatment of human liver diseases.

M. J. Czaja (*) Marion Bessin Liver Research Center, Albert Einstein College of Medicine, 1300 Morris Park Avenue, Bronx, NY 10461, USA e-mail: [email protected]

TNF and TNF Receptors: Molecules and Structure In response to infection or inflammation, TNF is produced and secreted primarily by activated macrophages, although epithelial cells, adipocytes, and endothelial cells can also be sources of this protein [1, 2]. TNF is produced as a 26-kDa type II transmembrane protein with an extracellular C-terminal domain for receptor interaction, a single transmembrane domain, and an intracellular N-terminal domain essential for cell signaling [3]. The membranous form of TNF is a homo­ trimer [4]. A soluble, 17-kDa form (sTNF) is produced through proteolytic cleavage of the transmembrane form by the metalloprotease TNF converting enzyme (TACE), a member of the mammalian adamalysin family [5, 6]. The cleavage of TNF by TACE can be inhibited by the tissue inhibitors of metalloproteases (TIMP), including TIMP-3 [7]. The bioactivity of membranebound and soluble TNF differs with regard to receptor activation (see below), and sTNF homotrimers dissociate below nanomolar concentrations and lose their bioactivity [8]. TNF exerts its biological effects by binding to either TNF receptor type 1 (TNF-R1, p55/65, CD120a) or TNF receptor type 2 (TNF-R2, p75/80, CD120b). These receptors belong to the TNF receptor superfamily that shares unique protein–protein interaction domains which determine their cell signaling functions. These pro­teins are type I transmembrane molecules with an extracellular N-terminus and an intracellular C-terminus [3]. They are characterized by the presence of extracellular cysteine rich domains (CRD) that are critical for ligand binding and receptor interactions [9]. The intracellu­lar C-terminus of TNF-R1 contains the death domain, the middle acidic sphingomyelinase activating domain (ASD) and neutral

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sphingomyelinase activating domain (NSD) that are responsible for the formation and regulation of the intracellular signaling complex (see Sect.  10.3.1) [10]. Although the membranous and soluble forms of TNF induce equivalent activation of TNF-R1, full activation of TNF-R2 requires membrane-bound TNF [11]. Crystallographic studies have suggested that a trimeric TNF-R1 interacts with the homotrimeric ligand [4]. Recent studies have demonstrated that rather than receptor trimerization occurring after ligand binding, the TNF-R exists as a preformed multimer to which the ligand binds in its homotrimeric form [3, 12]. The extracellular region responsible for receptor interaction and aggregation has been termed as the preligand binding assembly domain (PLAD) [9]. This region is physically distinct from the ligand binding domains, but crucial for TNF signaling. The widespread effects of TNF stimulation are directly related to the ubiquitous presence of TNF-R1 on all cells, while TNF-R2 expression is generally restricted to immune derived and endothelial cells [8]. Both receptors are expressed in the liver on nonparenchymal cells such as Kupffer cells and hepatocytes [13]. The expression of TNF-R1 and TNF-R2 is regulated differentially and in a cell type dependent manner. While TNF-R1 is controlled by a constitutive promoter, TNFR2 expression is highly inducible and varies widely among cell types [14]. The relative contribution of the two TNF receptors to TNF signaling is tissue dependent. In most cell types including hepatocytes, TNF-R1 is required for the induction of apoptosis, while the role of TNF-R2 is less defined. Interestingly, TNF-R2 possesses a lower binding affinity and a higher dissociation rate for TNF than TNF-R1. This suggests that TNF-R2 may transiently bind and then release TNF, serving to increase local concentrations of TNF that then act on TNF-R1 [15]. This phenomenon is known as ligand-passing and may allow TNF-R1 activation at much lower TNF concentrations [16, 17]. Alternatively, overexpression of TNF-R2 may inhibit TNF signaling by competing with TNF-R1 for ligand [17]. Thus the overall biological effect of TNF may depend in part on the relative ratio of the two receptors [18]. However, TNF-R1 is capable of transducing all of the biological effects attributed to TNF at a much lower receptor density than TNF-R2. Therefore TNF-R2 is currently thought to play an accessory role, enhancing, modulating or synergizing with TNF-R1 [3, 19]. As will be discussed subsequently, the biological effects of TNF in the liver have been largely attributed to signaling through TNF-R1 [20]. However models of liver injury

J. M. Schattenberg and M. J. Czaja

have shown involvement of both TNF-R1 and TNF-R2 in cell death signaling events [13, 21, 22]. TNF-R cleavage normally occurs and yields soluble receptor fragments that have been implicated as decoy receptors capable of neutralizing TNF activity [23]. However, the binding affinity of the soluble receptors is low in comparison to the membrane form, making their function unclear. Increases in circulating levels of these receptors have been reported in human liver diseases [24, 25]. These circulating receptors may serve to bind and neutralize the activity of TNF, or they may temporarily bind and later release TNF, prolonging its biological effects. Soluble receptors have been used experimentally to block the effects of TNF in the liver. The administration of an engineered dimeric soluble TNF-R has been successfully employed in the prevention of toxic liver injury in rats [26], although the effectiveness of such a treatment strategy in human liver disease has been unsuccessful to date as discussed subsequently.

TNF Signaling Pathways Intracellular Death Signaling Complex Binding between TNF and TNF-R occurs at the plasma membrane through interactions between the CRD of the receptor and the trimeric ligand. Binding results in a conformational change in the receptor and translocation of the receptor–ligand complex to lipid-enriched membrane microdomains known as lipid rafts [27]. Follow­ ing ligand–receptor interaction, an early intracellular signaling complex is formed to which signaling molecules are recruited [28]. While the outcome of TNF binding to its receptor can result in divergent cellular effects, these early events are common to all of the biological effects of TNF signaling [29]. The intracellular domains of TNF-R1 and TNF-R2 are devoid of intrinsic kinase activity and therefore depend on homophilic protein–protein interactions between motifs of approximately 80 amino acids for the initiation of cell signaling [30]. Based on the existence of one of the two distinct domains, the TNF-R superfamily members are divided into two subgroups, the death domain (DD)-containing receptors and the TNF-R-associated factor (TRAF) interacting receptors [31]. TNF-R1 belongs to the first group, while TNF-R2 is devoid of an intracellular DD.

10  TNF/TNF Receptors

The earliest molecule recruited to the intracellular DD of the TNF-R1 is the TNF-R-associated death domain protein (TRADD) which acts as an adaptor for other DD-containing proteins [32]. Association with the DD is normally inhibited by the binding of the silencer of the death domain protein (SODD) which masks this site. Dissociation of SODD from the receptor occurs following ligand binding and conformational changes [33]. At this level of the TNF signaling ­pathway, the survival and death signaling pathways bifurcate and recruit different downstream effector molecules [29]. The apoptotic cell death pathway is activated following the recruitment of the Fas-associated death domain (FADD) protein to TRADD through interactions between the DD in each protein [34]. While critical for TNF death signaling, FADD was first described as an adaptor molecule mediating Fas-dependent apoptosis [35]. In contrast, TRADD is not required for death by the Fas pathway, but is unique to TNF signaling and deletion of TRADD prevents TNF-R1 induced cell death [36]. Besides the C-terminal DD, FADD carries a second domain, the death effector domain (DED) in its N-terminal region [37]. This domain recruits proteins from the caspase (cysteine aspartate protease) family of enzymes. These proteases are capable of cleaving substrates after a loosely specific series of amino acids that contain aspartate in the first position. Crucial to their catalytic activity is the presence of a cysteine residue in the active center of the molecule [38]. The family can be divided into upstream initiator caspases, such as caspase 8 and 10, and the downstream effector caspases, caspase 3, 6, and  7. The effector caspases are responsible for the cleavage of proteins whose functional loss induces apoptosis [39]. Caspases are constitutively expressed as inactive zymogens or procaspases that require cleavage into smaller active subunits. Procaspases contain DEDs, and the DED of caspase 8 allows its recruitment to the DED of FADD. Upon colocalization with FADD, high, localized concentrations of procaspase 8 undergo autoproteolytic cleavage, releasing activated caspase 8. This complex has been termed the death-inducing signaling complex (DISC), and the mode of activation is referred to as the induced proximity model of activation [40, 41]. Upon TNF binding to TNF-R1 initially only a weak and transient formation of the DISC occurs, and signal transduction is regulated through the recruitment of antagonistic proteins such as the TNF-R-associated factor-2 (TRAF-2) mediated recruitment of inhibitor of apoptosis

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proteins (IAP) which interferes with the activation of caspase 8. The specific involvement of FADD in TNF signaling in the liver has been demonstrated by studies in which adenoviral inhibition of FADD function in vivo blocked caspase 3 activation and liver damage from TNF-induced liver injury [24]. The apparent contradiction between the rapid recruitment of signaling molecules to TNF-R1, and the long delay before TNF-induced death occurs, has been resolved by studies in nonhepatic cells. The complex colocalizing with procaspase 8 in cells following TNF stim­ ulation is undetectable before 30–60 min and not fully formed before 4–8 h [42–44]. This complex is composed of TRADD, FADD, the serine-threonine kinase receptor interacting protein (RIP), and TRAF-2, but is devoid of TNF-R. In accordance with this lack of receptor, the complex is detectable only in the cytosol, and not in membrane-enriched fractions [43, 45]. Thus, TNFinduced caspase 8 activation appears to occur following the dissociation of TRADD from the TNF-R, although the events triggering TNF-R-TRADD dissociation are unknown. The delay in formation of this intracellular DISC complex presumably explains why TNF-induced apoptosis in hepatocytes occurs at a much slower rate than that induced by the Fas death receptor.

Mitochondrial Amplification of the Death Signal After DISC formation, TNF-induced hepatocyte death results from the mitochondrial death pathway in which caspase 8 activation leads to functional changes in mito­chondria such as the mitochondrial permeability tran­sition (MPT). As a result, mitochondrial proteins such as cytochrome c are released into the cytosol and activate downstream caspases. Thus, the inhibition of cyto­chrome c release from mitochondria by the MPT inhibitor cyclosporine A prevents hepatocyte apoptosis at a point downstream of FADD binding to the TNF-R, but upstream of caspase 3 activation [46]. The mechanism of cytochrome c release involves cleavage of the Bcl-2 family member Bid by caspase 8 [47]. Truncated Bid (tBid) migrates to the mitochondria and triggers oligomerization of the proapoptotic Bcl-2 family members Bax and Bak. These molecules then insert into the mitochondrial membrane, resulting in release of mitochondrial proteins including cytochrome c [48]. Bid

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mediates hepatocyte death from TNF as hepatocytes deficient for either Bid alone or both Bak and Bax have increased resistance to TNF-induced cell death in association with the prevention of mitochondrial depolarization and cytochrome c release [48, 49]. Bid-deficient mice are partially protected from TNF-dependent toxic liver injury [50]. However cytochrome c release still occurs in a delayed fashion in Bid-deficient mice, indicating that Bid-independent mechanisms of mitochondrial activation exist [51]. Following release into the cytoplasm, cytochrome c triggers formation of the apoptosome, a complex with apoptosis protease activating factor-1 (APAF-1) and procaspase 9. Caspase 9 becomes activated and in turn activates caspase 3, resulting in apoptosis [52]. Hepatocytes and other cell types that are dependent on this mitochondrial death pathway have been termed as type II cells. In contrast, type I cells generate high levels of caspase 8 that directly activate caspase 3. Accordingly, expression of the antiapoptotic factors Bcl-2 and Bcl-XL that inhibit Bid and Bax activation prevent apoptosis in type II, but not in type I cells [53]. In support of the concept of the hepatocyte as a type II cell is that in vivo Bcl-2 or Bcl-XL overexpression is partially effective in preventing liver injury from TNF [54, 55]. The mechanisms by which mitochondria promote hepatocyte death from TNF are likely to be even more complex than those that are outlined. For example, the release of mitochondrial proteins other than cytochrome c may be involved. The mitochondrial protein SMAC/ DIABLO has been implicated in TNF-induced apoptosis [56], but the involvement of this protein in hepatocyte death is unknown. Another mechanism of mitochondrial death pathway activation in hepatocytes is through the lysosomal cysteine protease cathepsin B. Hepatocyte death from TNF in vitro and in vivo is dependent on the release of cathepsin B from acidic vesicles [57–59]. The proapoptotic effect of cathepsin B occurs above the level of mitochondrial cytochrome c release as this process is blocked in TNF-treated cathepsin B null hepatocytes and this protease induces mitochondrial cytochrome c release in a cell-free system [57]. How the action of cathepsin B integrates with the other components of the hepatocyte TNF-mitochondrial death pathway is not yet known. Although apoptosis via the mitochondrial death pathway (summarized in Fig. 10.1) is an important mechanism of hepatocyte death from TNF, it is clearly not the only form of TNF death signaling. The partial effects of  Bid ablation [50], or Bcl-2/Bcl-XL overexpression

J. M. Schattenberg and M. J. Czaja

[54, 55], on TNF-dependent liver injury in vivo suggest that the mitochondrial death pathway may be less important in vivo than in cultured hepatocytes. Recent investigations suggest that the Bid-dependent mitochondrial pathway may be a rapid form of TNF-induced death but a more delayed Bid-independent activation of the mitochondrial death pathway occurs when Bid function is inhibited [51]. This finding is in contrast to Fas-induced hepatocyte death which is almost completely Biddependent [60]. Alternatively, forms of death other than caspase-dependent apoptosis may be involved in hepatocyte death from TNF. TNF-induced cell death independent of caspase activation occurs in a differentiated hepatocyte cell line and in an in vivo model of TNFdependent hepatitis [61, 62]. TNF can also cause necrosis as well as apoptosis as demonstrated by the ability of cytochrome P450 2E1 overexpression to sensitize rat hepatocytes to TNF-induced necrosis [63]. If these pathways of caspase-independent cell death occur in vivo, then their existence would affect the design of anti-TNF treatments for liver disease. It is possible that therapeutic agents aimed at blocking the mitochondrial apoptotic pathway or caspase activation will not be effective because the hepatocyte is then shunted into an alternative, caspase-independent death signaling pathway.

NF-kB The ability of TNF to induce either cellular proliferation or death has led to the concept that TNF signaling acts as “a double-edged sword” [12]. The mechanism by which hepatocytes block the TNF death signaling pathway has been delineated. Investigations in cultured hepatocytes and animals have identified nuclear factor-kB (NF-kB) signaling as critical for the maintenance of hepatocyte resistance to TNF killing. The NF-kB family of transcription factors consists of NF-kB1/p50, NF-kB2/p52, c-Rel, RelA/p65 and RelB. In addition, precursor proteins of NF-kB1 (p105) and NF-kB2 (p100) exist and have been implicated in the regulation of NF-kB activation [64, 65]. The common feature of these NF-kB family members is the presence of the Rel homology domain (RHD) which is important for protein dimerization, DNA binding, nuclear localization and interactions with inhibitory proteins [66]. NF-kB activation occurs in response to a number of inflammatory mediators including TNF. The initial

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Fig. 10.1  TNF induced proliferative and cell death signaling pathways. The binding of membrane bound or soluble TNF to the TNF-R1 induces the recruitment of intracellular proteins that then activate the NF-kB and JNK cellular signaling pathways. These signaling molecules lead to proliferative or cell death responses in the hepatocyte

steps of TNF-dependent NF-kB activation are identical to the TNF-initiated apoptotic pathway described previously (Fig. 10.1). Following receptor–ligand inter­ action, recruited TRADD binds to TRAF family proteins [67]. This family consists of six distinct members, TRAF 1-6, all of which are capable of binding to the TRADD death domain by means of their highly conserved C-terminal TRAF-C domain [68]. TRAF-2, TRAF-5, and TRAF-6 have been implicated in the activation of NF-kB, and deletion of Traf-2 and Traf-5 in mice blocks TNF-induced NF-kB activation [69]. These proteins trigger NF-kB activation through their recruitment of the IkB kinases (IKK) to the TNF-R signaling complex. In resting cells, NF-kB is sequestered in the cytoplasm as an inactive heterodimeric complex bound to an inhibitory counterpart IkB. Phosphorylation of IkB by IKK results in IkB polyubiquitination, proteasome-dependent degradation and dissociation from NF-kB [70]. As a result, the nuclear

localization signal of NF-kB is unmasked, leading to its translocation to the nucleus and activation of gene expression. The IKK complex consists of two catalytically active kinases IKK1 (IKKa) and IKK2 (IKKb), and the regulatory kinase IKKg (NEMO). TRAF proteins recruit the IKK complex to the membrane-bound TNF-R complex, but a second receptor-bound protein, RIP, is also required for full IKK activation [71]. Ripdeficient cells are able to recruit IKK1 and IKK2 to the TNF-R complex following TNF binding, but recruitment of the regulatory subunit IKKg is significantly reduced [72]. Additional regulation of the transcriptional activity of NF-kB can result from direct phosphorylation of its subunits [73]. Numerous kinases have been implicated in NF-kB phosphorylation, including mitogen-activated protein kinases (MAPK), protein kinase C isoforms, casein kinase II and the nuclear DNA repair enzyme poly(ADP-ribose) polymerase-1 (PARP-1) [74–76].

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The initial suggestion that NF-kB signaling was important in hepatic homeostasis was the finding that genetic ablation of the NF-kB p65 subunit resulted in embryonic lethality from hepatocellular apoptosis [77]. Subsequent studies demonstrated that the hepatic apoptosis in p65-NF-kB null mice was induced by TNF, because the mice were rescued from their lethal phenotype through simultaneous inactivation of TNF-R1 [78]. Knockouts of Ikk2 or Ikkg are also embryonic lethals, stressing the physiological importance of IKK-depen­ dent IkB phosphorylation in hepatic NF-kB activation [66]. The initial confirmation that NF-kB acti­vation mediated hepatocyte resistance to TNF toxicity came from studies of NF-kB inactivation in cultured hepatocytes. In both primary rat hepatocytes and a nontransformed rat hepatocyte line, inhibition of NF-kB activity by adenoviral delivery of a phosphorylation defective mutant IkB sensitized these cells to death from TNF [46, 79]. Despite virtual total inhibition of NF-kB activity with this adenovirus, only partial cell death occurred, suggesting that other signaling pathways may compensate for the loss of NF-kB. Investigations in human hepatocytes have demonstrated the additional involvement of NF-kB-independent survival pathways mediated by sphingosine kinase and phosphatidylinositol 3-kinase/Akt in hepatocyte resistance to TNF toxicity [80]. However, in mouse hepatocyte studies Akt signaling promoted resistance to TNF through induction of NF-kB activation [81]. Additional studies in IKK-deficient mice have confirmed the function of NF-kB signaling in hepatocyte resistance to TNF toxicity and clarified the specific roles of the IKK subunits in this process. IKK2 and IKKg null mice die from massive hepatic apoptosis indicating a critical hepatic function for both kinases early in life [82, 83]. However, a hepatocyte-specific knockout of IKK2 failed to affect NF-kB activation or sensitize to death from TNF, suggesting that in the adult hepatocyte IKK1 could compensate for the loss of IKK2 [84]. In contrast, the specific knockout of IKKg led to a complete block in TNF-induced NF-kB activation and apoptosis [85], confirming the role for the IKK complex and NF-kB signaling in hepatocyte resistance to TNF toxicity. NF-kB inactivation sensitizes hepatocytes to TNFinduced death through the classical mitochondrial death pathway as evidenced by mitochondrial changes, cytochrome c release, and resultant caspase-dependent apoptosis [46, 79]. The fact that hepatocyte resistance

J. M. Schattenberg and M. J. Czaja

to TNF toxicity requires transcription and translation [86], and activation of the transcription factor NF-kB, suggests that NF-kB up regulates a gene(s) that protects against TNF cytotoxicity. In TNF-treated hepatocytes, the TNF-responsive, NF-kB-dependent gene inducible nitric oxide synthase (iNOS) has been demonstrated to be protective against TNF toxicity [87]. However, the protective effects of endogenous iNOS were only partial, suggesting that its induction cannot completely explain NF-kB-mediated hepatocyte resistance to TNF toxicity. Subsequent studies demonstrated that the protective effect of NF-kB activation was mediated through crosstalk with the MAPK member c-Jun N-terminal kinase (JNK) as discussed in the next section.

c-Jun N-Terminal Kinase A second signaling event following recruitment of TRAF proteins to TRADD is the activation of the MAPK JNK which requires TRAF-2 [88]. JNK is a key ­regulator of the pathways of cell death, ­proliferation, inflammation, and insulin signaling. Activated JNK ­phos­phorylates substrates include the activator protein-1 (AP-1) transcri­ ption factor subunit c-Jun, leading to increased AP-1 transcriptional activity [89]. However, JNK also has effects independent of transcriptional regulation such as through its ability to alter rates of proteasomal protein degradation [90]. In all cells including hepatocytes TNF treatment in the setting of an inhibition of NF-kB signaling leads to prolonged JNK and AP-1 activation that induces cell death [88, 91]. Cell death from TNF and NF-kB inactivation was blocked in a rat nontransformed cell line by the inhibition of c-Jun function through adenoviral expression of the c-Jun dominant negative TAM67. Blocking JNK/c-Jun function with TAM67 prevented mitochondrial cytochrome c release and caspase activation, suggest­ing that JNK overactivation mediated the TNF mitochondrial death pathway [88]. Pharmacological JNK inhibition also blocked death in primary mouse hepatocytes sensitized to TNF killing by either NF-kB inhibition or transcriptional blockage from actinomycin D, suggesting that JNK promoted cell death independently of effects on transcription [92]. The in vivo function of JNK in hepatic TNF signaling has been examined using mice null for the JNK1 and JNK2 isoforms. In hepatocytes two JNK genes,

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jnk1 and jnk2, are expressed and both generate the p54 and p46 isoforms of the JNK protein [93]. Initially the different JNK isoforms were thought to have redundant functions based on findings that mice null for either gene lacked a phenotype whereas deletion of  both genes was an embryonic lethal [94]. Studies in the galactosamine/lipopolysaccharide (LPS) model of TNF-dependent hepatic injury demonstrated that cotreatment with the toxin galactosamine and LPS resulted in sustained JNK activation [95], similar to what occurred in hepatocytes with inhibited NF-kB signaling. Hepatic injury was dependent on jnk2 signaling as mice null for jnk2 but not jnk1 were protected from liver injury and had reduced mortality [95]. In the absence of JNK2, activation of caspase 8 and the mitochondrial death pathway failed to occur, indicating that that the classical apoptotic TNF death pathway was mediated by JNK2. These findings contrasted with those by Chang et al. [90] that indicated that TNFdependent liver injury from concanavalin A (ConA) or galactosamine/LPS was mediated by JNK1 and not JNK2. In this work the prodeath effect of JNK1 was linked to its activation of E3 ligase and enhanced degradation of the antiapoptotic factor c-FLIPL [90]. A recent study examining injury from ConA alone or in combination with galactosamine indicated that JNK2 mediated apoptosis via activation of caspase 8 and the mitochondrial death pathway whereas necrotic death was JNK1 dependent [96]. All of these in vivo studies clearly link JNK activation to the induction of the TNF death pathway. However, to elucidate the specific effects of the two jnk gene products more studies are required. Current evidence suggests that JNK2 is critical for caspase-dependent apoptosis and that JNK1 may also have a role depending on the injurious stimulus and type of death. The activation and involvement of MAPK family members other than JNK in hepatocyte TNF signaling have not been described. Although extracellular signalregulated kinase 1/2 (ERK1/2) has been implicated as a cytoprotective pathway in several forms of cell death, ERK1/2 has not been shown to be regulated by TNF or to protect against TNF toxicity in hepatocytes. TNF is known to activate p38 MAPK in a RIP-dependent manner in fibroblasts [97]. However, p38 MAPK activation is not seen in a rat hepatocyte cell line after TNF stimulation, and inhibition of this MAPK does not affect death from TNF in these cells (Czaja, unpublished observation).

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Function of TNF Signaling in Hepatic Pathophysiology Liver Regeneration The liver has the unique capability to switch from a quiescent to a proliferative state in response to a loss of mass secondary to surgical reduction or cell death. A critical function for TNF signaling in hepatic regeneration has been demonstrated by investigations in both models of partial hepatectomy and liver injury. Within 1  h, TNF is produced in the liver in response to the regenerative stimulus of a partial hepatectomy [2]. In contrast to macrophage-dependent TNF generation during liver injury, there is evidence that the sources of TNF production after partial hepatectomy are biliary and endothelial cells [2]. Neutralizing anti-TNF, antibodies inhibit hepatocyte proliferation following partial hepatectomy, clearly implicating TNF as a direct or indirect hepatocyte mitogen in this model [98]. Sub­ sequent studies of partial hepatectomy in Tnf-r1 knockout mice also demonstrated reduced DNA synthesis and liver mass in these mice as compared to wild-type animals [99]. Inhibition of the biological activities of TNF led to reduced activation of the downstream transcriptional regulators NF-kB, signal transducer and activator of transcription 3 (STAT3), and AP-1, but not of CCAAT/enhancer binding protein (C/EBP), as measured by levels of DNA binding [99]. In contrast, posthepatectomy liver regeneration was normal in Tnf-r2 null mice [100]. Absence of the type 2 receptor had no effect on NF-kB and STAT3 binding or IL-6 production, but caused a delay in AP-1 and C/EBP binding [100]. These data suggest that TNF-induced proliferation occurs exclusively through signaling by the type 1 receptor, again demonstrating the predominant role for the TNF-R1 in hepatic TNF responses. Studies of carbon tetrachloride-induced liver regeneration also demonstrated that a lack of TNF-R1-mediated signaling decreased the regenerative response [101]. However, in all of these studies loss of TNF signaling merely delayed liver regeneration, but did not prevent the eventual return of the liver to a normal mass. Thus, TNF seems to directly or indirectly promote the initiation of hepatocyte proliferation, but TNF signaling is not obligatory for liver regeneration to occur. Subsequent investigations in Il-6 null mice suggested that this cytokine is also critical for liver regeneration

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and survival after partial hepatectomy [102], in part through the activation of STAT3 [103]. In another study, similar effects were seen although some mice survived and had a normal regenerative response [104]. In carbon tetrachloride-treated mice lacking Tnf-r1, the reduction in liver regeneration resulting from the absence of TNF signaling was prevented by the injection of IL-6 [101]. These investigations all suggested that the effect of TNF on liver regeneration was not direct, but rather mediated through IL-6 signaling. A possible signaling cascade for liver regeneration is that TNF-induced activation of NF-kB leads to production by Kupffer cells or other nonparenchymal cells of IL-6 that activates STAT3 and other downstream signals required for hepatocyte proliferation. However, other studies in Il-6 null mice and mice with ablation of the IL-6 downstream signaling molecule glycoprotein 130 (gp130), have revealed no significant effect of the loss of IL-6 signaling on DNA synthesis after partial hepatectomy [105, 106]. Treat­ ment with LPS after partial hepatectomy did result in a reduction in DNA synthesis and survival in Il-6 null mice associated with decreased Bcl-XL levels and increased hepatocyte apoptosis [106]. Studies in hepatocyte specific knockouts confirmed that this effect was dependent on hepatocyte gp130 signaling and STAT3-mediated [107]. This work suggests that the function of IL-6 after partial hepatectomy is not to regulate DNA synthesis, but to up regulate protective acute phase proteins. The direct effects of TNF on hepatocyte proliferation after partial hepatectomy therefore need to be re-evaluated. The importance of TNF-induced NF-kB activation after partial hepatectomy was suggested by studies in which inhibition of NF-kB signaling by adenoviral expression of a phosphorylation defective mutant IkB blocked liver regeneration and caused massive hepatic apoptosis [108]. DNA synthesis was unaffected by NF-kB inhibition, but cell cycle block in late S or G2 occurred along with apoptosis. The p65 NF-kB subunit seems most critical for this NF-kB function as liver regeneration after partial hepatectomy was unaffected by the absence of its heterodimeric partner p50 [109]. A potential NF-kB-dependent gene responsible for protection against the cytotoxic effects of TNF after partial hepatectomy was identified in studies that revealed marked postpartial hepatectomy apoptosis in iNos knockout mice [110]. However, it is unclear whether NF-kB activation in hepatocytes mediated these effects as hepatocyte specific ablation of NF-kB or IKKb failed to affect postpartial hepatectomy-induced liver regen­

J. M. Schattenberg and M. J. Czaja

eration [111, 112]. One study also noted an absence of apoptosis, but the 45% inhibition of NF-kB activation achieved in this study may have been insufficient to fully block the protective effects of NF-kB [111]. The discordant results of global versus hepatocyte specific NF-kB inhibition may reflect species differences, nonspecific effects of adenoviral vectors, or the fact that additional effects on nonparenchymal cell NF-kB activity are required to trigger apoptosis after partial hepatectomy. The complexity of crosstalk between hepatocytes and nonparenchymal cells was underscored by recent findings that a hepatocyte specific knockout of IKK2 that blocked NF-kB activation in these cells augmented the postpartial hepatectomy proliferative response apparently secondary to increased NF-kB activation and TNF production in the nonparenchymal cells [113]. Less is known about the role of TNF signaling in chronic states of liver injury and regeneration associated with elevated TNF levels such as alcoholic liver disease. Rats chronically fed with alcohol have a decreased regen­erative response after partial hepatectomy despite serum TNF and IL-6 levels similar to normal rats [114]. ­Ethanol-fed rats had decreased NF-kB and c-Jun activation following partial hepatectomy suggesting an ­alcohol-induced impairment of these proliferative signaling pathways [115]. However, ethanol-fed rats were clearly ­responsive to TNF as its inhibition reduced liver regeneration to a greater extent in ethanol-fed animals than in control rats [114]. Thus, there is contradictory evidence that ethanol-fed animals are both more dependent on the regenerative stimulus of TNF, as well refractory to its effects on signaling pathways thought to be involved in proliferative TNF signaling.

Toxin-Induced Liver Injury Formerly toxin-induced liver injury was thought to result simply from the direct and passive biochemical effects of the toxin or its metabolite on the hepatocyte. However, it is now apparent that liver injury results in large part from the effects of inflammatory cell products on the hepatocyte and is actively regulated by cell signaling pathways. Prominent among the products of inflammation that mediate toxin-induced liver injury is TNF. TNF is produced as part of the liver’s response to hepatotoxins such as carbon tetrachloride and galactosamine in rodents [116], and alcohol in humans

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[117]. It was first demonstrated in the rat carbon tetrachloride model that TNF induced most of the liver injury from this hepatotoxin. Neutralization of TNF by a dimeric soluble receptor markedly reduced carbon tetrachloride-induced liver injury as measured by transaminases, histology and mortality [26]. Decreased injury from carbon tetrachloride was also subsequently demonstrated in both Tnf and Tnf-r1 null mice [118]. The involvement of TNF in hepatotoxic liver injury has now been demonstrated for a number of toxins ­including alcohol [119]. The similar ability of agents that neutra­ lize LPS to prevent liver injury from carbon tetrachloride and ethanol suggests that a common mechanism of toxic injury is through LPS-induced macrophage activation and the resultant production of TNF that triggers hepatocyte injury and death. The ability of TNF to act as a hepatocyte cytotoxin had to be reconciled with its known proliferative effects after partial hepatectomy and the fact that although TNF was cytotoxic to many transformed cells, normal cells were resistant to TNF toxicity. It had been known for a long time that cells normally resistant to TNF, including hepatocytes, become sensitized to death from TNF by transcriptional or translational arrest [86]. This fact suggested that: (1) resistance to TNF requires transcriptional up regulation of a protective gene(s); (2) following partial hepatectomy, the remaining normal cells are able to up regulate this protective gene, resulting in a nontoxic, proliferative TNF effect; and (3) because hepatotoxins interfere with macromolecular synthesis, they might block the protective response and thereby sensitize hepatocytes to death from TNF. Alternatively, toxins might induce TNF injury by triggering a massive outpouring of TNF that overwhelms the cellular protective mechanisms. Although some toxins do augment the induction of TNF by LPS [120], most toxins are thought to act by interfering with the ability of hepatocytes to up regulate protective genes [121]. Attempts to identify the protective factor(s) that mediates hepatocyte resistance to TNF toxicity ­initially focused on antioxidant factors because of the ability of TNF to induce cellular oxidative stress. Although the antioxidant enzyme manganese superoxide dismutase (MnSOD) acts as a protective factor, it was demonstrated that MnSOD was not the inducible protective factor in hepatocytes because although TNF up regulated MnSOD gene expression, no increase in protein occurred [121]. Levels of the principle hepatic non­­ enzymatic antioxidant glutathione (GSH) are also not

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regulated at a transcriptional level by TNF [122]. How­ ever, depressed levels of GSH do worsen TNF-induced toxic liver injury in vivo [122]. The effects of reduced GSH levels may be mediated through changes in cell signaling as in vitro studies have demonstrated that GSH depletion can down regulate the activation of NF-kB in response to TNF [123]. Alcohol in particular may sensitize hepatocytes to TNF injury by selectively depleting mitochondrial GSH [124]. GSH depletion in macrophages may also increase their production of TNF [125]. Contradictory reports of GSH depletion preventing TNF-induced hepatocyte death are an artifact of the ability of sudden, profound GSH decreases to block DISC formation [126]. Oxidative stress even in the absence of antioxidant depletion may also act to promote death from TNF. Chronic oxidative stress gener­ated by cytochrome P450 2E1 overexpression sensitizes cultured hepatocytes to death from TNF [63], and may play a role in promoting TNF injury in steatohepatitis (see Sect. 10.4.3). Thus, although antioxidant genes are not the transcriptionally regulated protective factor against TNF cytotoxicity, impaired antioxidant defenses or increased oxidative stress may promote TNF-induced liver injury. The previously discussed in vitro findings that NF-kB inactivation and resultant JNK/AP-1 overactivation sensitize hepatocytes to death from TNF suggest that these pathways may be involved in TNF-mediated hepatotoxic injury. However, evidence of in vivo im­­pairment of NF-kB signaling in response to TNF in the setting of toxin exposure is currently lacking. Although increased liver injury from LPS administration occurred in mice chronically fed with ethanol as compared to control-fed mice [127], NF-kB or JNK activation was not altered in the ethanol-fed mice. Despite caspase 3 activation in LPS-treated control mice, caspase activation, cytochrome c release and DNA fragmentation were all ab­­sent in the ethanol-fed mice after LPS ­treatment [127]. As previously discussed, there is in vivo evidence for JNK overactivation in response to toxic injury from galactosamine [95]. In this model NF-kB activation may be unaffected by toxin but the inability of the toxin-damaged hepatocyte to up regulate NF-kB-dependent genes probably leads to JNK overactivation. Although it is possible that NF-kB acts solely to down regulate JNK activation, other NF-kB-dependent genes are likely to be involved in the resistance against TNF toxicity. Although a number of potential, TNFinducible, NF-kB-dependent protective genes have

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been identified in nonhepatic cells, their function in hepatocytes is largely unknown. One gene specifically studied in hepatocytes is iNOS, which was previously discussed for its protective effects after partial hepatectomy. TNF induces hepatic iNOS in mice, and this up regulation is blocked with NF-kB inhibition [87]. TNF by itself caused liver damage in iNos null mice, but the degree of injury suggested that loss of iNOS alone was not sufficient to fully sensitize hepatocytes to death from TNF [87]. Nitric oxide generated by iNOS may also protect hepatocytes from TNF-induced toxic injury by the inactivation of caspases and inhibition of the mitochondrial death pathway [128]. A second, TNFinducible protective factor, A20, has also been shown to block the lethality of galactosamine/LPS when overexpressed in mice [129]. However, it is not known if toxins prevent the normal TNF-induced up regulation of this factor. The protective effect of A20 overexpression occurred in the absence of any reduction in TNF production and was associated with increased liver regeneration, suggesting that factors that selectively block TNF death signaling may offer an advantage over TNF-neutralizing agents by preserving the proliferative effects of TNF [129, 130]. Strong animal and human data demonstrating TNF induction and its mechanistic involvement in alcoholinduced liver injury have suggested that this disease would be an ideal target for anti-TNF therapy. Pharmacologically soluble TNF receptor fragments are used as TNF antagonists in the treatment of chronic inflammatory diseases like rheumatoid arthritis, inflammatory bowel disease and psoriasis [131]. However, anti-TNF therapy in human alcoholic liver disease has given conflicting results [132, 133]. These findings probably result from the fact that global TNF inhibition blocks the beneficial effects of TNF (proproliferative, protective against infections) as well as its cytotoxicity. Thus, rather than proving that TNF is not involved in human liver disease, these findings indicate the importance of developing therapeutic agents that specifically target the components of the TNF death signaling pathway such as JNK2.

Nonalcoholic Steatohepatitis The most common liver disease in industrialized countries results from the accumulation of excess fat in hepatocytes or hepatic steatosis and is termed nonalcoholic

J. M. Schattenberg and M. J. Czaja

fatty liver disease (NAFLD). NAFLD encompasses a spectrum from simple steatosis to steatosis combined with varying degrees of necro-inflammation and fibrosis [134]. The initial hepatic lipid accumulation, and subsequent progression to cellular injury and inflammation termed nonalcoholic steatohepatitis (NASH), are thought to result from distinct mechanisms. Risk factors that contribute to the development of NASH are obesity, dys­ lipidemia, and insulin resistance, a group of disorders that constitute the metabolic syndrome [135]. However the mechanisms which contribute to the progression of steatosis to steatohepatitis and hepatocellular injury and the relevance of the above mentioned risk factors in this progression are not well understood. There is in vivo evidence in animal models of a function for TNF signaling in NASH. In leptin-­deficient ob/ob mice, hepatic and serum TNF levels are markedly increased in parallel with the presence of insulin resistance and increased levels of oxidative stress [136]. Diet-induced insulin resistance, a risk factor for the development of NASH, is blunted in mice lacking TNF [137]. In contrast, studies in TNF and TNF-R knockout mice using a dietary model of steatohepatitis suggested that TNF had no causal role in disease development as animals developed steatohepatitis irrespective of their TNF or TNF-R expression [138]. This study did demonstrate a TNF-independent role of NF-kB in the development of NASH. Animals with steatohepatitis had increased NF-kB binding activity, and blockage of NF-kB activation resulted in decreased hepatic injury and inflammation without influencing steatosis [138]. TNF may function to promote NASH not only through the cytokine’s cytotoxic effects but also through its ability to impair insulin signaling, thereby promoting steatosis. TNF produced primarily by adipose tissue with obesity can inhibit insulin signaling pathways as TNF deficient mice are protected from the development of insulin resistance [139, 140]. Finally, TNF may directly promote lipid accumulation. Mice administered TNF had increased accumulation of fat in hepatocytes that occurred in parallel to increased levels of sterol regulatory element binding protein-1c (SREBP-1c) [141]. In studies of human NAFLD, patients with biopsyproven steatohepatitis had increased TNF and TNF receptor levels in the liver and serum [142, 143], and increased serum levels of lipopolysaccharide-binding protein (LBP) [144]. LBP augments the ability of LPS  to trigger the release of TNF from target cells. In accordance with this observation, increased levels of

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TNF secretion in response to LPS were found from peripheral blood cells in patients with steatosis or steatohepatitis [145]. Additional indirect evidence for the importance of TNF in the induction and progression of steatohepatitis comes from genetic studies evaluating TNF promoter polymorphisms in defined ethnic groups. These studies found a predisposition for NAFLD progression in patients with polymorphisms of the TNF promoter gene region [146, 147]. Also, the presence of fibrosis characteristic of end-stage liver disease was found to correlate with levels of soluble TNF receptor in NAFLD patients [148]. Thus, despite the contradictory findings in animal models of steatohepatitis, indirect evidence does exist for a potential role for TNF in human NAFLD.

Viral Hepatitis The pathogenesis of liver cell injury in acute and chronic viral hepatitis B and C is poorly understood, but proinflammatory cytokines are thought to play a central role in modulating the cellular immune response, virus replication and liver injury. The presence of virus-infected hepatocytes triggers an immune response that is characterized by the infiltration of cytotoxic T-lymphocytes that mediates the majority of hepatocellular injury observed in chronic hepatitis. This immune response is determined by viral and host factors such as TNF promoter polymorphisms [149, 150]. In chronic HBV infection, TNF was shown to inhibit viral replication through a strong activation of NFk-B that impaired viral capsid formation [151]. In transgenic hepatoma cell lines TNF suppresses HBV production and secretion and induces cellular apoptosis synergistically with INFg [152, 153]. Central to the proapoptotic effects of TNF in infected hepatocytes is the HBV protein X (HBx). Expression of HBx increases the susceptibility of primary hepatocytes to cell death from TNF through a caspase-dependent mechanism [154]. These signaling events rely on the activation of the MAPK p38, since inhibition of p38 protected HBxtrangenic hepatoma cells from TNF-induced apoptosis [155]. HBx protein also interferes directly with TNF signaling pathways through the inhibition of the caspase 8 homologue c-FLIP that results in increased caspase activation in response to TNF

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stimulation [156]. These results point towards multiple effects of HBV on TNF signaling. Although strong evidence for the proapoptotic effects of HBx protein exists, an antiapoptotic effect through activation of the NF-kB subunits RelA/p65 and c-Rel has also been reported [157]. In chronic hepatitis C infection, the role of TNF signaling is not as well established as for HBV. Viral factors may promote a proinflammatory and proliferative response through increased secretion of TNF from inflammatory cells. Transient expression of the nonstructural HCV protein NS-3 in hepatoma cell lines resulted in increased NF-kB binding activity and JNK activation, suggesting that this viral protein may augment hepatocyte signaling in response to TNF [158, 159]. Inhibition of TNF-mediated cell death occurred in HCV core transgenic hepatocytes secondary to increased NF-kB activation [160]. However, another study found increased sensitivity towards TNF-induced cell death in HCV-core transgenic hepatoma cell lines resulting from the direct interaction of core antigen with the cytoplasmic domain of TNF-R1 [161]. These apparently conflicting reports on the roles of TNF and NF-kB signaling in chronic viral hepatitis must be resolved by further studies.

Conclusion The involvement of TNF in a number of pathophysiologic conditions is evidence for the importance of TNF signaling in the liver. Although considerable progress in understanding hepatic TNF signaling pathways has been made, significant gaps in our knowledge still exist particularly in the area of the final gene products that mediate the biological effects of TNF. Evidence of alterations in the levels of TNF ligand and receptors in human studies suggests that TNF signaling may affect the outcome of human liver disease. While therapeutic efforts to block TNF function in these diseases may prove beneficial, neutralization of all TNF function may prove ineffective in treating these diseases as has been proven in the case of anti-TNF therapy for alcoholic hepatitis. An additional understanding of hepatic TNF signaling pathways may lead to agents that selectively target the beneficial or cytotoxic effects of TNF, and thus have added effectiveness in treating human liver diseases.

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Summary

›› TNF is produced and secreted primarily by acti››

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›› ›› ›› ››

vated macrophages, i.e., in the liver by Kupffer cells. TNF is produced as a transmembrane protein; a soluble form is produced through proteolytic cleavage of the transmembrane form by the metalloprotease TACE. Although the membranous and soluble forms of TNF induce equivalent activation of TNF-R1, full activation of TNF-R2 requires ­membranebound TNF. TNF-R2 expression is generally restricted to immune derived and endothelial cells. In contrast to TNF-R1, its expression is highly inducible, but TNF-R1 is capable of transducing all the biological effects attributed to TNF at a much lower receptor density than TNF-R2. TNF-R1 recruits the adaptor TRADD, which can bind TRAF-2 and RIP to activate NF-kB and JNK or FADD to activate caspase 8 and apoptosis. TNF treatment in the setting of an inhibition of NF-kB signaling leads to a prolonged JNK and AP-1 activation with sensitization to cell death. Examples of TNF-inducible protective factors are A20 and iNOS. After partial hepatectomy, TNF seems to promote the initiation of hepatocyte proliferation. The involvement of TNF in hepatotoxic liver injury has been demonstrated for a number of toxins including alcohol.

Multiple Choice Questions 1. Important differences exist between TNF and Fas pathways. What is true? (a) FADD is not required in Fas pathway (b) TNF-induced apoptosis in hepatocytes occurs at much faster rates than by Fas (c) Fas-induced apoptosis is almost completely bid-dependent (d) TRADD is required in Fas pathway (e) SODD belongs to both pathways

2. Which of the following statements is false? Lipopoly­ saccharide (LPS) (a) Is part of the external membrane of gram negative bacteria (b) Induces an inflammatory response (c) Triggers release of TNF from macrophages (d) Binds to a receptor at the cell membrane following interaction with lipopolysaccharide binding protein (LBP) (e) Is a hepatocyte-specific transcription factor 3. Choose the appropriate answer. The mitogen-­activated protein kinase c-Jun N-terminal kinase (JNK) (a) Antagonizes TNF through inhibitory phosphorylation of TNF-R1 (b) Promotes cell death in hepatocytes in response to TNF when NF-kB is inhibited (c) Causes polyubiquitination of TNF (d) Integrates into mitochondria to release cytochrome c (e) Migrates into the nucleus to act as a specific transcription factor 4. Which of the following statements is false? TNF (a) Is a proinflammatory cytokine (b) Binds to a cell membrane bound receptor (c) Exerts its physiological function following endocytosis through clathrin coated pits (d) Binds as homotrimeric complex to the corresponding receptor (e) Can result in either apoptosis or proliferation of hepatocytes 5. Which of the following statements is false?: (a) The receptor to which TNF binds can be either the CD120a (TNF-R1) or CD120b (TNF-R2) receptor (b) Only membrane bound TNF binds to a TNF receptor (c) Shedding of membrane bound TNF-R occurs through the actions of the metalloprotease TNF converting enzyme (TACE) (d) Following TNF binding to its receptor, an intracellular signaling complex associates (e) TNF receptors are expressed on hepatocytes, macrophages and endothelial cells Acknowledgments  Supported in part by National Institutes of Health grants DK044234 and DK061498 to MJC and a Deutsche Forschungsgemeinschaft (DFG) grant to JMS.

10  TNF/TNF Receptors

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11

Fas/FasL Maria Eugenia Guicciardi and Gregory J. Gores

Introduction Fas (CD95, APO-1) belongs to the death receptor family, a subgroup of the tumor necrosis factor/nerve growth factor (TNF/NGF) receptor superfamily. These cell surface cytokine receptors are able to initiate an apoptotic signaling cascade after binding a group of structurally related ligands or specific antibodies [1]. The members of this family are type-I transmembrane proteins with a C-terminal intracellular tail, a membrane spanning region, and an extracellular N-terminal domain. Through interaction with the N-terminal domain, the receptors bind their cognate ligands (called death ligands), the majority of which are type-II transmembrane proteins belonging to the TNF family of proteins and comprised of an intracellular N-terminal domain, a transmembrane region, and a C-terminal extracellular tail. The signature features of the death receptors are represented by a highly homologous region in their extracellular domains containing one to five cysteine-rich domains (CRD) and a ~80-amino acid cytoplasmic sequence known as death domain (DD), which is required to initiate the death signal. Engagement of death receptors results in the initiation of the so-called extrinsic pathway of apoptosis, one of the two main signaling pathways leading to apoptotic cell death. The second one is generated by a mitochondrial dysfunction and is referred to as the intrinsic pathway [2] (Fig. 11.1). Although both signaling pathways are sufficient to trigger apoptosis, they

G. J. Gores (*) Mayo Clinic College of Medicine, 200 First Street SW, Rochester, MN 55905, USA e-mail: [email protected]

are not mutually exclusive and can be simultaneously activated in the same cell through cross-talk between pathways, especially in hepatocytes. Apoptosis is essential to preserve liver function and health, as it ensures the efficient removal of unwanted cells (i.e., aged or virus-infected cells) in a highly controlled manner. Apoptotic cells are ultimately fragmented into membrane-bound, organelle-containing corpses (apoptotic bodies) which are readily engulfed by neighboring phagocytes, mainly Kuppfer cells; this engulfment process may, under pathologic conditions, promote liver inflammation and damage by amplifying Fas-mediated hepatocyte apoptosis through FasL production by the Kuppfer cells themselves (Fig.  11.2) [3]. Although apoptosis in the liver can occur through activation of both the extrinsic and the intrinsic pathways, the extrinsic pathway seems to be by far the most relevant, probably due to the high level of expression of death receptors in hepatic cells. In particular, Fas is constitutively expressed by every cell type in the liver [4], rendering all liver cells sensitive to Fas-mediated apoptosis in vivo. Fas-induced apoptosis plays a fundamental role in liver physiology by contributing to the elimination of senescent cells and maintaining liver homeostasis [5], as well as, in pathologic conditions, by ensuring the removal of virus-infected or mutated cells via the interaction between FasL-positive cytotoxic T lymphocytes and Fas-expressing target cells [6, 7]. However, excessive or defective Fas-mediated apoptosis leads to disease pathogenesis such as liver failure, fibrosis, and carcinogenesis. In this chapter, we review the molecular mechanisms and regulation of Fas signaling and the role of the Fas/FasL system in the pathophysiology of the liver.

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_11, © Springer-Verlag Berlin Heidelberg 2010

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180 Fig. 11.1  Apoptotic pathways. Apoptosis is induced via activation of the extrinsic pathway or the intrinsic pathway. Death receptor engagement mediates the extrinsic pathway, whereas different stimuli can trigger the intrinsic pathway, which is regulated by the Bcl-2 family proteins (see text for details)

M. E. Guicciardi and G. J. Gores EXTRINSIC PATHWAY

INTRINSIC PATHWAY Growth factor deprivation

Death Receptor engagement

DNA damage

UV light

Activation of proapoptotic Bcl-2 family members

Other stimuli

Antiapoptotic Bcl-2 family members

DISC formation

Mitochondrial dysfunction Activation of Initiator caspases (casp-8, casp-10)

Activation of initiator caspase-9

Activation of effector caspases (casp-3, -6, -7)

Apoptosis

FasL/CD95L

Fas/CD95

Hepatocyte

N

Apoptosis

Fig. 11.2  A vicious cycle of apoptosis. Schematic representation of the amplification of Fas-mediated hepatocyte apoptosis by Kupffer cell-generated FasL. Apoptotic hepatocytes fragment into apoptotic bodies, whose engulfment by Kupffer cells induces their activation and production of FasL, which, in turn, exacerbates the apoptotic damage

Apoptotic Bodies

Engulfment of Apoptotic Bodies

N Kupffer cell

Production of FasL FasL

11  Fas/FasL

Fas (Cd95/Apo-1) and Fas Ligand (Fasl/CD95L 4) Fas (CD95/APO-1) Fas is a glycosylated cell-surface protein, ubiquitously expressed in various tissues, in particular in thymus, liver, heart, kidney, pancreas, and activated mature lymphocytes and virus-infected lymphocytes. Although soluble forms of the receptor also exist, whose functions are still largely unknown, the membrane-bound form is largely predominant and highly biologically active [8]. In order to avoid unnecessary activation of the apoptotic pathway, Fas expression and localization are tightly regulated through a variety of mechanisms. First of all, only minimal amount of Fas is expressed on the plasma membrane in unstimulated cells, whereas the majority of the receptor localizes in the cytosol, in particular, in the Golgi complex and the trans-Golgi network [9, 10]. After a proapoptotic stimulus, Fascontaining vesicles translocate to the cell surface, increasing Fas expression on the plasma membrane and initiating the apoptotic signal. This mechanism provides an effective tool to regulate the plasma membrane density of the death receptor, and avoid its spontaneous activation [10, 11]. Fas can also be modulated at a posttranslational level, by glycosylation and palmitoylation of the receptor [12, 13], as well as at the transcriptional level, by direct regulation of Fas expression via activation of the transcription factors NF-kB and p53 [14–16].

Fas Ligand (Fasl/CD95l) FasL (CD95L) is a type II transmembrane protein, mainly expressed in preassociated homotrimeric structures on the cell surface of activated T cells [17]. FasL also exists in a soluble form generated after the cleavage by a metalloprotease between Ser126 and Leu127 in its extracellular domain, but its activity is likely not biologically significant due to a much lower apoptoticinducing capacity of the soluble form compared to the membrane-bound FasL [18].

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Fas/Fasl Signaling On the cell surface, Fas is found in the form of preassembled receptor oligomers [19]. Engagement of Fas by either agonistic antibodies or FasL induces posttranslational modification of the receptor, (i.e., palmitoylation), and the formation of progressively higher order receptor aggregates [13, 20, 21], which recruit the adaptor molecule FADD (Fas-associated protein with death domain), procaspase-8 and -10, and the short and long forms of the cellular FLICE/caspase 8-like inhibitory protein (FLIP), to form the so-called death inducing signaling complex (DISC) [22, 23]. These complexes move into lipid rafts [24, 25], and they are eventually internalized through clathrinmediated endocytosis and delivered to the early endosomal compartment, where larger amounts of DISC are formed [22] (Fig. 11.3). Recruitment and accumulation of procaspase-8 and/or -10 at the DISC result in their spontaneous activation and initiation of a proteolytic cascade leading to apoptosis. While c-FLIPS competitively inhibits procaspase-8 recruitment to the DISC, the function of c-FLIPL, which structurally resembles caspase-8, except that it has no cysteine protease activity, remains controversial [26]. Indeed, cFLIPL has been described to both interfere with and promote caspase 8 activation at the DISC [27–29]. The role of FLIPL is likely determined by a variety of factors, including its expression level and cellular levels relative to caspase 8. Downstream the DISC formation, activation of effector caspases, such as caspase-3, -6 and -7, which are ultimately responsible for the degradation of key cellular components, can occur via two different signaling pathways. Based on the signaling pathway preferentially activated after Fas stimulation, cells have been classified into type I and type II [30] (Fig. 11.3). In type I cells, large amounts of DISC are rapidly assembled and internalized, and caspase-8, which is mainly activated at the DISC, directly cleave and activate caspase-3. In these cells, prevention of mitochondrial dysfunction by over-expression of the anti- apoptotic proteins Bcl-2 or Bcl-XL does not block the activation of caspase-8 or caspase-3, nor does it inhibit apoptosis, suggesting a mitochondria-independent activation of a caspase cascade. In contrast, in type II cells, DISC formation is

182 Fig. 11.3  Fas signaling pathways. Schematic representation of Fasmediated apoptotic and nonapoptotic signaling pathways (see text for details)

M. E. Guicciardi and G. J. Gores FasL Fas aggregates

Pre-associated Fas

Type I Internalization

Plasma membrane

FADD Procaspase 8/10

Fa

s

he

te ro zy go

ci ty

cFLIP

NF-?B MAPK Activation

Type II Bid

tBid

Bax/Bak

Abundant DISC formation cFLIP

AIF

SMAC/Diablo Cytochrome c

PROLIFERATION MOTILITY INVASION

strongly reduced, and activation of caspases, including caspase-8, occurs mainly downstream of mitochondria, as both caspase activation and apoptosis can be prevented by over-expression of Bcl-2 or Bcl-XL. Notably, Fas induces mitochondrial dysfunction in both type I and type II cells, but only in type II cells mitochondria are essential for the execution of the apoptotic program, whereas in type I cells they probably function solely as amplifiers of the apoptotic signal [30]. Mitochondrial dysfunction in type II is initiated by caspase-8-mediated cleavage of Bid, a proapoptotic, BH3-only member of the Bcl-2 family of proteins [31, 32]. Truncated Bid (tBid) translocates to the mitochondria and contributes to the outer mitochondrial membrane permeabilization, resulting in the release of apoptogenic factors such as cytochrome c [31, 32], AIF (apoptosis-inducing factor)

Caspase 3

APOPTOSIS

[33], and SMAC (second mitochondria-derived activator of caspases)/Diablo (direct IAP-binding protein with low pI) [34, 35]. In the cytosol, cytochrome c associates with the adaptor Apaf-1 (apoptosis-activating factor 1) and procaspase-9 to form a complex named apoptosome. Through an energy-requiring reaction, procaspase-9 in the apoptosome is processed into the mature enzyme and, in turn, activates downstream caspases.

Fas/FasL in Liver Diseases Dysregulation of hepatocyte apoptosis often associates with liver diseases. Defective hepatocyte apoptosis leads to diseases associated with excessive cell growth,

11  Fas/FasL

such as hepatocellular carcinoma. On the contrary, excessive hepatocyte apoptosis is a feature of viral and autoimmune hepatitis, acute hepatic failure, cholestatic diseases, alcoholic and nonalcoholic hepatitis, chemotherapeutic-induced liver damage, as well as transplantation-associated liver damage, such as ischemia/ reperfusion injury and graft rejection. The role of the Fas/FasL system in several human liver diseases associated with disruption of apoptosis is described in greater details in this section.

Pathologic Conditions Associated with Reduced Fas-Mediated Apoptosis Hepatocellular Carcinoma Hepatocellular carcinoma, the most common primary malignancy of the liver, has multiple etiologies, including environmental, nutritional, and metabolic factors, as well as chronic viral infections. Downregulation of Fas is frequently observed in tumors, including hepatocellular carcinoma, and correlates with advanced stages of the tumor and poor prognosis [36–38]. The loss of Fas represents an advantageous adaptation of the cancer cell, because it allows the cell to survive the attack by FasL-expressing cytotoxic T-lymphocytes and NK cells [39]. However, tumors showing complete loss of Fas expression are extremely rare, while mutations in the Fas gene, especially in the DD region, are very common. Interestingly, these tumors almost never display loss of heterozygocity, suggesting that maintaining one wild-type allele may confer an oncogenic advantage [40]. Recent studies showed that the signaling threshold to activate NF-kB after Fas engagement is significantly lower than the one required for the internalization of the receptor and assembly of the DISC, and it can be achieved even in the presence of only one functional Fas allele [41]. Consistently, in cells carrying heterozygous mutations in the Fas gene or cells expressing reduced levels of Fas, Fas stimulation does not induce apoptosis, but results in activation of MAP kinases- and NF-kB-mediated prosurvival and tumorigenic pathways involved in invasion and metastasis [22] (Fig. 11.3). Therefore, therapeutic approaches aimed to restore Fas expression and sensitivity to Fasmediated apoptosis in tumor cells may be effective in the therapy of hepatocellular carcinomas. Several

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chemotherapeutic drugs induce tumor cell apoptosis by causing DNA damage and activation of p53, which, in turn, have been found to up-regulate Fas expression and increase sensitivity to Fas-mediated apoptosis [15, 16]. Unfortunately, p53 is frequently mutated in tumor cells, rendering this approach ineffective. In addition to mutated Fas, tumor cells often express elevated FasL, which promotes tumor growth by actively killing the tumor-infiltrating lymphocytes [39, 42, 43] and by recruiting neutrophils to the tumor, which causes a sustained inflammatory response [44]. Persistent inflammation leads to massive cell loss and liver regeneration that, in turn, may significantly increase the chance of mutagenic events. Thus, early in the disease process, inhibiting FasL and its inflammatory signaling would prevent the milieu necessary for carcinogenesis to occur. These new findings have to be considered in order to design a better therapeutic approach for hepatocellular carcinomas.

Pathologic Conditions Associated with Excessive Fas-Mediated Apoptosis Viral Hepatitis Viral hepatitis is mainly caused by infection with Hepatitis B (HBV) or C virus (HCV). However, the virus itself has very mild cytopathic effects on the infected host cells, and the extensive tissue damage associated with viral hepatitis is generally the result of host immune response to viral antigen. During viral hepatitis, specific classes of cytotoxic T lymphocytes (CTL) recognize and kill viral antigen-expressing, HBV-or HCV-infected hepatocytes to clear the virus from the liver. This causes the initial liver damage, which is subsequently exacerbated by the influx of antigen-non-specific inflammatory cells. The killing of viral antigen-positive hepatocytes by CTL occurs via apoptosis, as demonstrated by the presence of apoptotic bodies, once referred to as Councilman bodies, in the liver of patients with viral hepatitis. In particular, Fas, although not the only apoptotic pathway involved, seems to play a key role in this process. Indeed, Fas expression is increased in the liver of patients with chronic hepatitis B and C, and directly correlates with disease activity such as periportal and intralobular inflammation [45–49]. It is not clear whether Fas

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expression is mainly regulated by virus-specific protein or by inflammatory cytokines, such as interleukin-1, generated after the first immune response. Areas of FasL-positive infiltrating mononuclear cells are also common in the liver of HBV- and HCVinfected patients, confirming the importance of the Fas/FasL system in the removal of infected cells by CTL during viral hepatitis [45, 47–49]. Nevertheless, the role of the HBV and HCV proteins in Fas-mediated apoptosis remains controversial. The HBV X-gene product (HBx) has been shown to stimulate the apoptotic turnover of hepatocytes [50], as well as to activate NF-kB and JNK pathways and therefore protect liver cells from apoptosis [51, 52]. Similarly, HCV proteins inhibit Fas-mediated apoptosis and death in transgenic mice by preventing the release of cytochrome c from the mitochondria [53]. Therefore, hepatitis virus proteins may either sensitize hepatocyte to Fas-induced apoptosis or inhibit apoptosis to maintain persistent infection.

Alcoholic Hepatitis Although the pathogenesis of alcoholic hepatitis and cirrhosis is still poorly understood, apoptosis certainly plays an important role both in vitro and in vivo. Apoptosis is a characteristic feature of experimental ethanol-induced liver injury [54, 55]. Moreover, hepatocyte apoptosis in liver biopsies of patients with alcoholic hepatitis correlates with the disease severity, being most abundant in patients with high bilirubin and AST levels, and grade 4 steatohepatitis [56, 57]. Among the several mechanisms proposed to explain alcohol-induced hepatocyte apoptosis there is the activation of death receptor pathways, in particular, the Fas/FasL and TNF-a/TNF-R1 signaling. Patients with alcoholic hepatitis express higher levels of Fas and FasL in the hepatocytes compared to healthy subjects, which renders the cells more susceptible both to cytotoxic T-lymphocyte-mediated apoptosis and to cell death by autocrine and/or paracrine mechanisms [56]. The increased expression of Fas and FasL may result from TNF-a-induced activation of NF-kB, a transcription factor which can up-regulate both these genes [14]. Indeed, TNF-a serum levels are elevated during alcoholic hepatitis, and are directly involved in hepatocyte apoptosis [58]. In addition to a direct cytotoxic effect on the hepatocyte, the TNF-a/TNF-R1 system

M. E. Guicciardi and G. J. Gores

is also required for Fas-mediated cell death, as demonstrated by the increased resistance of TNF-R1/TNFR2 double knock-out mice to Fas-induced fulminant liver injury [59]. Thus, it appears that both Fas and TNF-R1 ­contribute to ethanol-mediated liver injury through a synergistic action in inducing hepatocyte apoptosis.

Cholestatic Liver Disease Cholestasis is defined as an impairment of bile flow through the liver. As a consequence, high concentrations of bile acids accumulate within the hepatocytes, causing tissue damage and liver failure. Several studies demonstrated that hydrophobic bile acids, such as deoxycholic and glycodeoxycholic acids, are able to cause hepatocyte apoptosis in vitro [10, 60–63]. More remarkably, massive hepatocyte apoptosis is clearly detectable in the liver of bile duct-ligated mice, an animal model of extrahepatic cholestasis [64]. Although bile acids have detergent properties and could potentially exert their toxic effect by damaging the cell membranes, they actually need to be transported into the cell to trigger apoptosis, as cells lacking a functional bile acid transporter are resistant to bile acid-induced apoptosis [65]. It has been shown that elevated concentrations of bile acids within the hepatocyte can induce Fas translocation from its intracellular locations to the plasma membrane, where the increased surface density triggers its oligomerization and initiates the apoptotic signal [10]. Indeed, bile acids-induced apoptosis largely occurs via a Fas-dependent, FasL-indepen­dent mechanism, both in vitro [61], and in vivo [64]. Hydrophobic bile acids-induced, Fas-dependent hepatocyte apoptosis involves activation of epidermal growth factor receptor (EGFR) and EGFR-catalyzed Fas tyrosine phosphorylation, which is required for its oligomerization [66, 67]. In addition, in a model of chronic cholestasis, Fas-mediated cytoxicity promotes the development of liver fibrosis, the result of excessive deposition of extracellular matrix during wound healing response which follows a prolonged injury to the liver [68]. In the absence of Fas, long-term, bile duct-ligated mice showed reduced markers of fibrosis as compared to Fas-expressing ­animals, suggesting that inhibition of Fas-mediated ­hepatocyte apoptosis may prevent liver fibrogenesis. Although Fas plays a major role in executing bile acid-mediated apoptosis,

11  Fas/FasL

other death receptors have also been involved, including TRAIL-R2/DR5 [62]. Both Fas and TRAIL-R2 signal apoptosis through activation of caspase-8/-10 and Bid, and therefore targeted inhibition of caspases or Bid could have therapeutic relevance in the treatment of cholestatic liver diseases.

Wilson’s Disease Wilson’s disease is a genetic disorder caused by excessive copper storage in different organs and tissues, including the liver. Liver sections from patients with Wilson’s disease show significant hepatocyte apoptosis associated with upregulation of Fas and FasL on the hepatocyte cell membrane [69]. Similarly, hepatocyte apoptosis and Fas expression have been found increased in a model of copper overload in vitro [69]. As already suggested in alcoholic hepatitis, the simultaneous expression of Fas and FasL on the same cell membrane may promote fratricide killing of neighboring cells. Copper accumulation within the hepatocyte causes oxidative stress which, in turn, may promote Fas activation and apoptosis [70]. The upregulation of Fas likely occurs via the activation of the tumor suppressor gene p53, which follows the oxidative stress-induced DNA damage. Indeed, treatment of hepatoma cells with copper results in a transient increase in p53 and Fas expression, the latter being a consequence of p53 transcriptional activity [69, 71]. Inhibition of either FasL or caspases effectively reduces apoptosis with similar results, suggesting that Fas might be the only apoptotic signal involved in copper-induced apoptosis. Therefore therapies aimed to inhibit either Fas or FasL, or caspases might be useful in the treatment of Wilson’s disease and could reduce the need for transplantation in the acute form of this disease.

Nonalcoholic Steatohepatitis (NASH) NASH is the most severe form of nonalcoholic fatty liver disease (NAFLD), characterized by the presence of macrovesicular steatosis along with inflammatory activity, and sometimes associated with fibrosis. The molecular mechanisms involved in tissue damage ­during NASH are poorly understood. However, Fas expression, activation of caspase-3 and -7 and hepatocyte apoptosis are enhanced in the liver of NASH

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patients, and positively correlated with the biochemical and histopathologic markers of liver injury [72, 73]. Moreover, recent studies showed increased sensitivity to Fas-mediated liver damage in obese ob/ob mice, an animal model of NAFLD, possibly due to decreased ­hepatocyte mitochondrial membrane potential [74]. Consis­tently, mitochondrial function is often impaired in the liver of subjects with NASH [75]. Activation of Fas results in mitochondrial dysfunction as a consequence of the activation of Bid and its translocation to the mitochondria. Moreover, mitochondrial dysfunction is associated with generation of reactive oxygen species, which are also able to induce apoptosis, further exacerbating tissue injury and inflammation. Thus, Fas inhibition may be an effective therapy to reduce liver damage and prevent development of cirrhosis in NASH.

Summary

›› The Fas/FasL system plays a key role in main-

taining liver homeostasis and function through regulation of cell death and survival. Several liver diseases are associated with either Fas over-expression or down-regulation. Tumor cells often acquire resistance to Fas-mediated apoptosis – in these cells stimulation of Fas does not result in apoptosis, but triggers activation of prosurvival and pro-oncogenic signaling pathways, which promote cancer growth. Hepatocyte apoptosis often represents the early stage of many liver diseases, independent of their etiology, and it occurs mainly through engagement of death receptors on the plasma membrane, especially Fas. Uncontrolled hepatocyte apoptosis can progress into liver injury if the number of cells dying is significantly higher than the number of cells replaced by cell division. The presence of a large number of apoptotic bodies that overwhelms the clearance capacity by phagocytes can exacerbate the tissue damage by eliciting a sustained inflammatory response, and can generate a profibrogenic response by the hepatic stellate cells. Therapeutic strategies aimed to modulate Fas-mediated apoptosis may ultimately be effective in reducing liver damage in several human liver diseases.

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Multiple Choice Questions 1. Which region of the death receptors is required to initiate an apoptotic response? (a) The cysteine-rich domain (CRD) (b) The death domain (DD) (c) The death effector domain (DED) (d) All of the above 2. Fas-mediated apoptosis in the liver is beneficial and essential in: (a) Physiologic conditions, to maintain tissue homeostasis by eliminating senescent cells and counterbalancing cell proliferation (b) Pathologic conditions, to eliminate virus-infected and/or mutated cells (c) Both the above (d) Neither of the above – apoptosis in the liver is always associated with tissue damage 3. What is the difference between type I and type II cells? (a) Type I cells do not depend on mitochondrial permeabilization for Fas-mediated apoptosis (b) Fas-mediated apoptosis can be inhibited by Bcl-2 or Bcl-XL in type I cells, but not in type II cells (c) Only type I cells are sensitive to Fas-mediated apoptosis (d) Type II cells generate larger amounts of caspase-8 at the DISC 4. In several tumors, including hepatocellular carcinoma, Fas is frequently: (a) Over-expressed (b) Down-regulated (c) Unchanged 5. In which of these diseases does Fas-mediated apoptosis not require binding to FasL to initiate the death cascade? (a) Viral hepatitis (b) Hepatocellular carcinoma (c) Nonalcoholic steatohepatitis (NASH) (d) Cholestatic liver disease

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187 40. Peter ME, Legembre P, Barnhart BC (2005) Does CD95 have tumor promoting activities? Biochim Biophys Acta 1755:25–36 41. Legembre P, Barnhart BC, Zheng L et al (2004) Induction of apoptosis and activation of NF-kappaB by CD95 require different signalling thresholds. EMBO Rep 5:1084–1089 42. Hahne M, Rimoldi D, Schroter M et al (1996) Melanoma cell expression of Fas(Apo-1/CD95) ligand: implications for tumor immune escape. Science 274:1363–1366 43. Griffith TS, Brunner T, Fletcher SM et al (1995) Fas ligandinduced apoptosis as a mechanism of immune privilege. Science 270:1189–1192 44. Philip M, Rowley DA, Schreiber H (2004) Inflammation as a tumor promoter in cancer induction. Semin Cancer Biol 14:433–439 45. Mochizuki K, Hayashi N, Hiramatsu N et al (1996) Fas antigen expression in liver tissues of patients with chronic hepatitis B. J Hepatol 24:1–7 46. Luo KX, Zhu YF, Zhang LX et al (1997) In situ investigation of Fas/FasL expression in chronic hepatitis B infection and related liver diseases. J Viral Hepat 4:303–307 47. Galle PR, Hofmann WJ, Walczak H et al (1995) Involvement of the CD95 (APO-1/Fas) receptor and ligand in liver damage. J Exp Med 182:1223–1230 48. Hiramatsu N, Hayashi N, Katayama K et al (1994) Immunohistochemical detection of Fas antigen in liver tissue of patients with chronic hepatitis C. Hepatology 19:1354–1359 49. Yoneyama K, Goto T, Miura K et al (2002) The expression of Fas and Fas ligand, and the effects of interferon in chronic liver diseases with hepatitis C virus. Hepatol Res 24:327–337 50. Terradillos O, de La Coste A, Pollicino T et al (2002) The hepatitis B virus X protein abrogates Bcl-2-mediated protection against Fas apoptosis in the liver. Oncogene 21: 377–386 51. Pan J, Duan LX, Sun BS et al (2001) Hepatitis B virus X protein protects against anti-Fas-mediated apoptosis in human liver cells by inducing NF-kappa B. J Gen Virol 82: 171–182 52. Diao J, Khine AA, Sarangi F et al (2001) X protein of hepatitis B virus inhibits Fas-mediated apoptosis and is associated with up-regulation of the SAPK/JNK pathway. J Biol Chem 276:8328–8340 53. Machida K, Tsukiyama-Kohara K, Seike E et al (2001) Inhibition of cytochrome c release in Fas-mediated signaling pathway in transgenic mice induced to express hepatitis C viral proteins. J Biol Chem 276:12140–12146 54. Goldin RD, Hunt NC, Clark J et al (1993) Apoptotic bodies in a murine model of alcoholic liver disease: reversibility of ethanol-induced changes. J Pathol 171:73–76 55. Benedetti A, Brunelli E, Risicato R et al (1988) Subcellular changes and apoptosis induced by ethanol in rat liver. J Hepatol 6:137–143 56. Natori S, Rust C, Stadheim LM et al (2001) Hepatocyte apoptosis is a pathologic feature of human alcoholic hepatitis. J Hepatol 34:248–253 57. Kawahara H, Matsuda Y, Takase S (1994) Is apoptosis involved in alcoholic hepatitis? Alcohol Alcohol Suppl 29:113–118 58. McClain C, Hill D, Schmidt J et al (1993) Cytokines and alcoholic liver disease. Semin Liver Dis 13:170–182 59. Costelli P, Aoki P, Zingaro B et al (2003) Mice lacking TNFa receptors 1 and 2 are resistant to death and fulminant liver

188 injury induced by agonistic anti-Fas antibody. Cell Death Differ 10:997–1004 60. Patel T, Bronk SF, Gores GJ (1994) Increases of intracellular magnesium promote glycodeoxycholate-induced apoptosis in rat hepatocytes. J Clin Invest 94:2183–2192 61. Faubion WA, Guicciardi ME, Miyoshi H et al (1999) Toxic bile salts induce rodent hepatocyte apoptosis via direct activation of Fas. J Clin Invest 103:137–145 62. Higuchi H, Bronk SF, Takikawa Y et al (2001) The bile acid glycochenodeoxycholate induces trail-receptor 2/DR5 expression and apoptosis. J Biol Chem 276:38610–38618 63. Guicciardi ME, Gores GJ (2002) Bile acid-mediated hepatocyte apoptosis and cholestatic liver disease. Dig Liver Dis 34:387–392 64. Miyoshi H, Rust C, Roberts PJ et al (1999) Hepatocyte apoptosis after bile duct ligation in the mouse involves Fas. Gastroenterology 117:669–677 65. Guicciardi ME, Faubion WA, Bronk SF et al (2000) Mechanisms of bile acid-induced cell death. In: Andus T, Rogler G, Schlottmann K (eds) Cytokines and cell homeostasis in the gastrointestinal tract. Kluwer, Dordrecht, pp 284–289 66. Reinehr R, Becker S, Wettstein M et al (2004) Involvement of the Src family kinase yes in bile salt-induced apoptosis. Gastroenterology 127:1540–1557 67. Eberle A, Reinehr R, Becker S et al (2007) CD95 tyrosine phosphorylation is required for CD95 oligomerization. Apoptosis 12:719–729

M. E. Guicciardi and G. J. Gores 68. Canbay A, Higuchi H, Bronk SF et al (2002) Fas enhances fibrogenesis in the bile duct ligated mouse: a link between apoptosis and fibrosis. Gastroenterology 123:1323–1330 69. Strand S, Hofmann WJ, Grambihler A et al (1998) Hepatic failure and liver cell damage in acute Wilson’s disease involve CD95 (APO-1/Fas) mediated apoptosis. Nat Med 4: 588–593 70. Aust SD, Morehouse LA, Thomas CE (1985) Role of metals in oxygen radical reactions. J Free Radic Biol Med 1:3–25 71. Narayanan VS, Fitch CA, Levenson CW (2001) Tumor suppressor protein p53 mRNA and subcellular localization are altered by changes in cellular copper in human Hep G2 cells. J Nutr 131:1427–1432 72. Feldstein AE, Canbay A, Angulo P et al (2003) Hepatocyte apoptosis and fas expression are prominent features of human nonalcoholic steatohepatitis. Gastroenterology 125: 437–443 73. Feldstein AE, Canbay A, Guicciardi ME et al (2003) Diet associated hepatic steatosis sensitizes to Fas mediated liver injury in mice. J Hepatol 39:978–983 74. Siebler J, Schuchmann M, Strand S et al (2007) Enhanced sensitivity to CD95-induced apoptosis in ob/ob mice. Dig Dis Sci 52:2396–2402 75. Perez-Carreras M, Del Hoyo P, Martin MA et al (2003) Defective hepatic mitochondrial respiratory chain in patients with nonalcoholic steatohepatitis. Hepatology 38: 999–1007

Interferon Signaling

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Markus H. Heim

Interferons Interferon (IFN) was identified more than 50 years ago by Isaacs and Lindenmann during their studies of the phenomenon of viral interference, the ability of an active or inactivated virus to interfere with the growth of an unrelated virus [1]. Today, more than ten mammalian IFN species and numerous subspecies have been discovered, each with individual properties, but all having antiviral activity [2]. They are currently classified into three groups: type I, type II and type III IFNs. The type I IFNs include all IFN-as, IFN-b, IFN-e, IFN-k, IFN-w and IFN-n [3]. Humans have 12 different IFN-as and a single IFN-b. Type I IFN genes are clustered on the human chromosome 9. Each subtype is encoded by its own gene and regulated by its own promoter, and none of them contain introns [3]. The different IFN-as and IFNbs have substantial differences in their specific antiviral activities and in the ratios of antiviral to antiproliferative activities. However, the molecular basis of these differences is not yet known. All type I IFNs bind to the same interferon alpha/beta receptor (IFNAR) that consist of two major subunits: IFNAR1 (a subunit in the older literature) [4] and IFNAR2c (the bL subunit) [5, 6]. There is only one class II IFN, IFN-g. IFNg is produced by T lymphocytes when stimulated with antigens or mitogens. IFNg binds to a distinct receptor, the IFN gamma receptor (IFNGR), which consists of the two subunits IFNGR1 (previously a chain) [7] and IFNGR2 (previously b chain or accessory factor) [8, 9].

M. H. Heim Division of Gasteroenterology and Hepatology, University Hospital, Basel, 4031 Basel, Switzerland e-mail: [email protected]

The recently described type III IFNs, IFN-l2, IFN-l3 and IFN-l1, are also known as IL-28A, IL-28B and IL29, respectively. Same as type I IFNs, they are also induced by viral infections [10]. They signal through the IFN-l receptor consisting of the IL-10R2 chain, which is shared with the IL-10 receptor, and a unique IFN-l chain [11, 12].

Induction of Type I Interferons Cells produce IFN-as and IFN-bs in response to infection by a variety of viruses. Unlike bacteria and fungi, which have microbe-specific structures distinguishable from host cell structures, viruses are made predominantly of host-derived components. Given the lack of virus-specific proteins or lipids, the cellular receptors that detect viruses have instead evolved to recognize the presence of the viral genome composed of nucleic acids. Two important pathways that detect viral genomes and induce type I IFNs have been discovered and characterised during recent years: the toll-like receptor (TLR) dependent pathway [13, 14] and the cytosolic pathway triggered by binding of viral RNA to the RNA helicases retinoic acid inducible gene-I (RIG-I) and melanoma differentiation antigen 5 (MDA5) [15, 16]. TLRs are a family of transmembrane pattern recognition receptors (PRRs) that recognize microbial pathogen associated molecular patterns (PAMPs) and activate the expression of genes involved in inflammatory and immune responses [14]. There are at least 10 human TLRs, and 3 of them are involved in the recognition of viral infections: TLR3, TLR7 and TLR9. TLRs are expressed not only on various immune cells such as macrophages, dendritic cells (DCs), B cells, but also on fibroblasts and epithelial cells. While TLRs involved in the recognition of bacterial components are expressed

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_12, © Springer-Verlag Berlin Heidelberg 2010

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on the cell surface, TLR3, TLR7 and TLR9 are localised in intracellular compartments such as endosomes. TLR3 recognizes dsRNA (e.g., HCV-RNA) [17], TLR7 detects ssRNA [18, 19] and TLR9 interacts with unmethylated DNA with CpG motifs [20]. TLR activation induces signaling cascades that mainly involve the key transcription factors NF-kB and various interferon regulatory factors (IRFs) (Fig.  12.1). Specifically, IRF3 and IRF7 have both distinct and essential roles for virus-induced transcriptional activation of IFN-b [21]. IRF3 is constitutively expressed in most cells, whereas IRF7 is expressed at low amounts and is strongly expressed only after stimulation of cells with type I IFNs [22]. TLR3 uses the adapter protein Trif and the kinase TBK1 to activate mainly IRF3 in conventional DCs and macrophages, whereas TLR7 and TLR9 induce the expression and secretion of large amounts of type I IFNs in plasmacytoid DCs through the adaptor molecule MyD88 that directly interacts with IRF7 (not IRF3) [23, 24]. The MyD88 pathway requires the IRAK4-IRAK1-IKKa kinase cascade to activate both IRF7 and the NF-kB pathway [25].

M. H. Heim

The cytosolic pathway of type I IFN induction is initiated by the recognition of viral 5¢ triphosphate RNA and dsRNA by RIG-I and MDA5. Binding of viral RNA induces a conformational change of these sensors that results in the binding to Cardif (IPS-1, MAVS, VISA), an essential downstream adaptor in the cytosolic pathway [26–29]. Through as yet unidentified mediators, Cardif then propagates the signal to the TBK1 and IKKi kinases that finally activate IRF3 and NF-kB (Fig. 12.1).

Interferon Signaling Through the Jak–Stat Pathway The Receptor–Kinase Complex All type I IFNs bind to the same IFNAR that consists of two major subunits: IFNAR1 (a subunit in the older literature) [4] and IFNAR2c (the bL subunit) [5, 6]. Each receptor subunit constitutively binds to a single specific member of the Janus kinase (Jak) family: IFNAR1 to tyrosine kinase 2 (TYK2) and IFNAR2c to JAK1. Upon binding of the two chains by type I IFNs, TYK2 and JAK1 transactivate each other by mutual tyrosine phosphorylation, and then initiate a cascade of tyrosine phosphorylation events on the intracellular domains of the receptors and on signal transducer and activator of transcription (STAT) 1, STAT2, and STAT3.

Signal Transducers and Activators of Transcription (STATs)

Fig.  12.1  Viral infections are sensed by two important pathways: the toll-like receptor (TLR) dependent pathway and the cytosolic pathway triggered by binding of viral RNA to the RNA helicases retinoic acid inducible gene-I (RIG-I) and melanoma differentiation antigen 5 (MDA5). RIG-I and MDA5 signal through Cardif (MAVS, IPS-1, VISA) and TBK1 to activate the transcription factors IRF3 and NFkB. TLR3 signaling depends on the adaptor TRIF to activate TBK, IRF and NFkB, whereas TLR7 and TLR9 use the MyD88 – IKKa pathway to activate IFN-b gene transcription

In most cells, type I IFNs activate STAT1, STAT2, and STAT3. STAT1 and STAT2 combine with a third transcription factor, IRF9, to form interferon stimulated gene factor 3 (ISGF3). ISGF3 binds to interferon stimulated response elements (ISREs) in the promoters of IFN stimulated genes (ISGs). Alternatively, IFN activated STAT1 and STAT3 can form homodimers or STAT1–STAT3 heterodimers. These STAT dimers bind a different class of response elements, the gamma activated sequence (GAS) elements. Once bound to the promoters of ISGs, STATs induce the transcription of genes involved in the generation of an antiviral state (Fig. 12.2) [30, 31]. STAT proteins are between 750 and 850 amino acids long. They share well-defined, structurally and

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Fig.  12.2  IFNa/b signaling through the Jak–STAT pathway. Binding of IFNa/b to the receptor brings together the two receptor subunits IFNAR1 and IFNAR2c, and leads to the mutual tyrosine phosphorylation and activation of Tyk2 and Janus kinase 1 (Jak1). The activated kinases phosphorylate tyrosine residues on the receptors which then become docking sites for STATs that bind with their SH2 domains to the phosphotyrosines. STATs are then phosphorylated on a single tyrosine

c-terminal of their SH2 domain, and form dimers through mutual phosphotyrosine-SH2 domain interactions. STAT1–STAT2 heterodimers combine with IRF9 to form the transcription factor ISGF3 that binds to ISRE elements in the promoters of ISGs. STAT1 and STAT3 can form homo- or heterodimers that bind to a different class of promoter elements, the GAS elements, to stimulate an overlapping but distinct set of ISGs. Negative regulators are shown in green

Fig. 12.3  Domain structure of human STAT proteins. Differential mRNA splicing and/or post-translational proteolytic processing generates multiple isoforms. The long isoforms are designated a, the shorter isoforms b. Domains are labelled as follows: N N-terminal domain, C–C coiled-coil domain, DNA DNA binding domain, LD linker domain, SH2 SH2 domain, P phosphorylated tail segment,

T transactivation domain. Three important sites of post-translational modifications are indicated: the arginine residue in the N-terminus (Rxx), the tyrosine that is phosphorylated upon activation of the STATs (Yxxx), and the serine phosphorylation site in the transactivation domain (Sxxx). The numbers of the amino acid residues that constitute the boundaries of the domains are shown

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functionally conserved domains including the aminoterminal (NH2), coiled-coil, DNA-binding, linker, SH2, tyrosine activation, and transcriptional activation domains (Fig.  12.3) [32]. The N-terminal domain is important for homotypic dimerization of inactive STATs and for cooperative DNA binding to tandem GAS elements [33, 34]. The coiled-coil domain is a protein interaction domain. Binding to GAS elements is provided by the adjacent DNA binding domain. The SH2 domain has a central role for the recruitment of STATs to tyrosine phosphorylated receptors and for dimerization of activated STATs, and importantly, provides specificity of signaling through the Jak–STAT pathway [35]. The carboxy-terminal residues constitute the transactivation domain. Alternative splicing at the 3¢ end of the gene transcripts generates shorter isoforms of STAT1, 3, 4, 5A, and 5B. The shorter isoforms lack a functional transcriptional activation domain, but retain the capacity to occupy specific binding sites in the promoters of target genes. By competing with full length STATs for DNA binding sites, they can inhibit transcriptional activation of target genes, and when overexpressed can be dominant negative regulators of transcription. However, in multimeric complexes with other transcription factors, these short isoforms need not be negative regulators of transcription. STAT1b can combine with STAT2 and IRF9 to form the transcription factor ISGF3, and STAT3b and c-Jun cooperatively bind to an IL-6 responsive promoter element in the a2-macroglobulin gene and activate its transcription [36, 37]. In both cases, the transcriptional activation domain is provided by the partner proteins of the short STAT isoforms.

Negative Regulators of Interferon Signaling

M. H. Heim

feedback loop. Type I IFNs induce SOCS1 and SOCS3 [39], and overexpression experiments have demonstrated that both inhibit IFN signaling through the Jak–STAT pathway [39, 40]. SOCS1-deficient mice develop severe inflammatory disease [41], but are very resistant to viral infections, most likely because of enhanced type I IFN signaling [42].

USP18 Ubiquitin specific peptidase 18 (USP18/UBP43) is another important negative regulator in type I IFN signaling. USP18/UBP43 was originally identified as a protease cleaving ubiquitin-like modifier ISG15 from target proteins, but was recently found to play a negative regulatory role independently of its ISGdeconjugating ability [43, 44]. UBP43 was reported to inhibit the activation of Jak1 by interfering with the binding of Jak1 to IFNAR2c [45]. UBP43 deficient mice show a severe phenotype characterised by brain cell injury, poly-I:C hypersensitivity, and premature death [46, 47]. Interest­ingly, they are resistant to otherwise fatal cerebral infections with LCMV and VSV [48].

Protein Inhibitor of Activated STAT1 (PIAS1) and PIAS3 PIAS1 and PIAS3 specifically bind to tyrosine phosphorylated STAT1 and STAT3, respectively, and inhibit the DNA-binding of STAT dimers [49]. PIAS1 selectively inhibits IFN-inducible genes and is important in innate immunity. As a consequence, PIAS1 deficient mice show increased protection against pathogenic infection [50].

TcPTP Suppressor of Cytokine Signaling (SOCS) SOCS proteins are important negative regulators of Jak–STAT signaling [38]. The family consists of eight members, CIS and SOCS1 to SOCS7. CIS, SOCS1, SOCS2, and SOCS3 are induced by a large number of cytokines and inhibit cytokine receptors in a negative

STAT1 is deactivated in the nucleus by dephosphorylation of the tyrosine 701 by T cell protein tyosine phosphatase (TcPTP) [51]. TcPTP deficient mice develop progressive systemic inflammatory disease as shown by chronic myocarditis, gastritis, nephritis, and sialadenitis as well as elevated serum IFN-g [52].

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Refractoriness of Interferon Signaling

Interferon Regulated Genes

It has been known for many years that cultured cells become refractory to IFN within hours and remain unresponsive for up to 3 days [53]. Maximal activation of the IFN signaling pathways is observed within the first 2 h of IFN treatment. Continuous exposure to IFN results in a “desensitization” characterised by a return to pretreatment levels of interferon stimulated gene (ISG) transcription. Moreover, during the 48–72 h following the initial IFNa stimulation of the cells, any further IFN treatment fails to reinduce the transcription of ISGs [53]. To investigate the sensitivity of the liver during prolonged exposure to therapeutic concentrations of IFNa, we treated mice repeatedly with subcutaneous injections of IFNa and prepared extracts from their livers at various time points. IFNa signaling was investigated by phospho-STAT Western blots, gel shifts and quantification of IFNa target gene induction. In as yet unpublished work we showed that liver cells in vivo become refractory within hours after the first injection of IFNa and remain so for at least 2 days. A systematic analysis of the negative regulators of IFNa signaling surprisingly revealed that SOCS are responsible for the early inhibition of STAT phosphorylation within the first 2–4 h, but not for the observed long-term refractoriness. Rather, a long lasting upregulation of USP18/UBP43 was found to be responsible for the observed unresponsiveness of liver cells to prolonged IFNa exposure.

Stimulation of cells with type I IFNs usually leads to the induction of several hundred genes (IFN stimulated genes, ISGs), but there are also some genes that are negatively regulated by IFNs [58–60]. There is considerable variation between different cell types with regard to the number and also the identity of the regulated genes [59]. Gene expression analysis in human and chimpanzee has shown that systemic administration of (pegylated) IFNa induces overlapping but clearly distinct sets of genes in liver and peripheral blood mononuclear cells (PBMCs) [60, 61]. The mRNA levels of most of the genes are increased two to tenfold through IFN stimulation, but some genes are induced even stronger [60]. In the liver, most of the ISGs are upregulated within hours after administration of pegylated IFNa and rapidly downregulated again within the first 8–24 h [61].

Effects of Type I Interferons Interferons exhibit a wide spectrum of biological activities in target cells, including antiviral, immunomodulatory, antiangiogenic, and growth inhibitory effects. They exert their effects mainly through Jak– STAT mediated regulation of gene transcription. However, there are also Jak–STAT independent effects, notably the activation of the p38 Map kinase signaling cascade [54, 55], and the activation of the phosphatidylinositol 3 (PI3) kinase – Akt kinase – mTOR/p70 S6 kinase pathway that regulates mRNA translation [56, 57].

Antiviral Effects Type I IFN induced regulation of hundreds of genes establishes an “antiviral state” in the cell [62, 63]. The term “antiviral state” implies protection of the cell against viral infection, but it is a generic term, and the lack of precise criteria for its definition reflects the fact that we still have only an elementary understanding of what exactly it is. Indeed, a large number of these regulated genes have as yet unknown functions. Some ISGs have broad antiviral effects. For example, protein kinase R (PKR), a member of the eukaryotic initiation factor 2a (eIF2a) kinase family, phosphorylates eIF2a with a consequent blockade of translation of most cellular and viral mRNAs [64]. Members of the IFNinduced protein with tetratricopeptide repeats (IFIT1 (ISG56) and IFIT2 (ISG54)) also inhibit translation by binding to eIF3 [65]. Another well-studied antiviral effector is 2¢–5¢ oligoadenylate synthetase (OAS). Both the gene transcription and the enzymatic activity are regulated: the enzymatic activity is stimulated by viral dsRNA, and OAS expression is upregulated several-fold by IFNa. The 2¢–5¢oligoadenylates produced by activated OAS in turn activate the latent RNA

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nuclease RNase L, resulting in the degradation of viral and host RNAs [64]. Recently, the ISG15 system has been found to be another broadly active non-specific antiviral effector. ISG15 is one of the most prominent ISGs. It is an ubiquitin-like protein that conjugates to more than 150 cellular target proteins [48, 66–68]. The conjugation is executed by an enzymatic cascade that includes an E1 activating enzyme (UBE1L) [69], an E2 conjugating enzyme (UbcH8) [70, 71], and an E3 ligase (HERC5 and TRIM25) [72, 73]. The conjugation can be reversed by ubiquitin protease 43 (UBP43, also known as USP18) [44]. All these enzymes are induced by type I IFNs. Many of the ISG15 target proteins have important roles in the IFN response, for example Jak1, STAT1, RIG-I, MxA, PKR and RNaseL [67]. Consistent with its role in the IFN system, mice deficient in ISG15 have increased susceptibility to infection with several viruses [74]. In addition to these relatively non-specific effector systems there are a number of ISGs with activities against distinct classes of viruses. For example, the MX proteins have protective effects against influenza and VSV by binding to viral nucleocapsids and the viral polymerase [75], and the members of the APOBEC3 family of cytidine deaminases have activity against HIV [76]. Several ISGs have been implicated in the host defense against hepatitis C virus (HCV). Viperin, a member of the radical S-adenosyl methionine domain containing enzymes, inhibits replication in HCV replicon system [77, 78]. PKR and ISG20, a 3¢–5¢ exonuclease with a strong preference for single-stranded RNA, also strongly inhibit HCV replicons [78].

Antiproliferative Effects In addition to their well-known antiviral effects, type I IFNs inhibit cell growth and control apoptosis, activities that affect the suppression of cancer and infection [79]. Different cells in culture exhibit varying degrees of sensitivity to the antiproliferative activity of IFNs. Lymphoblastoid Daudi cells are exquisitely sensitive to the antiproliferative effects of IFNa, which lead to a rapid shutdown of c-myc transcription, possibly through a decrease in the activity of the transcription factor E2F [80]. The antiproliferative effects of IFNa are the rational basis for their use in the treatment of metastatic malignant melanoma and renal cell carcinoma [81].

M. H. Heim

Interferon Signaling in Viral Hepatitis Since more than 20 years, IFN-a2a and IFN-a2b have been used for the treatment of patients with chronic hepatitis B (CHB) and C (CHC). Unmodified IFN-as have now been largely replaced in clinical use by pegylated IFN-a2a and pegylated IFN-a2b, in which a large molecule of poly(ethylene glycol) (PEG) is covalently attached to recombinant IFN-as, resulting in an active molecule with a longer half-life, better pharmacokinetic profile and better rate of virological response [82–85]. Therapies with pegylated IFN-a can induce long-lasting remission in CHB and sustained virological response in CHC in 30–60% of patients.

Interference of Hepatitis C Virus with Interferon Signaling In order to escape from the powerful antiviral effects of the IFN system, many viruses have evolved strategies to block IFN signal transduction [86, 87]. Interference of HCV with IFN signaling has been suggested in several studies, sometimes with controversial results [88, 89]. One proposed mechanism is the inhibition of STAT1 activation through an upregulation of SOCS3 by HCV core protein, which has been found after transient transfection of HepG2 and Huh7 cells [90, 91]. Another group reported that the expression of HCV proteins in Huh7 cells leads to a proteasomedependent degradation of STAT1 [92]. A third group, also using HCV protein expression in Huh7 cells, reported normal STAT1 expression and phosphorylation, but an inhibition of nuclear translocation of phosphorylated STAT1 [93]. We have found an inhibition of DNA binding of activated STATs not only in cells transfected with the HCV genome, but also in the liver of HCV transgenic mice, and in liver biopsies of patients with CHC [94, 95]. In all cases, STAT1 protein expression and tyrosine phosphorylation were not impaired. Further investigations of the molecular mechanisms of HCV interference with IFN signaling identified protein phosphatase 2A (PP2A) as an important mediator in the inhibitory pathway [96]. The catalytic subunit of PP2A, PP2Ac, was found to be overexpressed as a result of an endoplasmatic reticulum (ER) stress response induced by HCV protein

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195

Fig. 12.4  HCV interference with IFN-a signaling. HCV induced upregulation of PP2Ac results in an inhibition of PRMT1, the enzyme responsible for STAT1 methylation. Unmethylated

STAT1 can be bound by the inhibitor PIAS1. STAT1-PIAS1 association lowers the affinity of STAT1 homodimers to a subset of ISGs and thereby decreases their transcription

expression [97]. PP2Ac was overexpressed in cells after HCV protein expression, in liver extracts of HCV transgenic mice, and in liver biopsies of patients with CHC [96]. Furthermore, expression of a constitutive active form of PP2Ac in Huh7 cells resulted in an inhibition of STAT1 DNA binding [96]. PP2A can directly bind to protein arginine methyltransferase 1 (PRMT1) and inhibit its enzymatic activity [98]. This inhibition of PRMT1 results in a decreased methylation of a number of proteins, amongst them STAT1 [96]. It has been reported that the arginine methylation of STAT1 regulates the association of STAT1 with the inhibitor PIAS1 [99], a finding that is still controversial [100]. Nonetheless, we have found that inhibition of PRMT1 by increased expression of PP2Ac leads to an increased association of STAT1 with PIAS1, a finding that could

well explain the impaired DNA binding of activated STATs in HCV infected cells [96]. Interestingly, treatment of cells with the methyl group donor S-adenosylmethionine restored normal IFN signaling in cells with HCV protein expression and increased the potency of IFNa in the replicon system [101]. Our current working model of HCV interference with IFN signaling is shown in Fig. 12.4.

Hepatitis C Virus Induced Activation of the Interferon System The infection of cells with HCV leads to the induction of IFN-b through the activation of the RIG-I and of

196

the TLR3 pathways [16, 102, 103]. Interestingly, the HCV NS3/4A protease has been shown to cleave and inactivate Cardif (MAVS, IPS-1, VISA) and TRIF, two important adaptor proteins in the RIG-I and the TLR3 pathway, respectively [27, 102]. Hepatic gene expression studies with chimpanzees and humans, however, provide compelling evidence that the inhibition of IFN-b induction is in many cases incomplete and cannot prevent the activation of the endogenous IFN system. It has been shown, for example, that the acute infection of chimpanzees with HCV leads to the rapid activation of the endogenous IFN system [104]. Moreover, chronically infected chimpanzees have an ongoing induction of a large number of ISGs, suggesting a continuous stimulation of the endogenous IFN system [105]. Interestingly, a single chimpanzee infected with a genotype 3 HCV showed less induction of ISGs compared with genotype 1 infected animals [105]. Many patients with CHC also have a permanent induction of ISGs in the liver [60, 106, 107]. There is a strong association between such a preactivation of the endogenous IFN system and the failure to respond to pegIFNa/ribavirin therapy [60, 106, 107]. Interestingly, patients with a preactivated IFN system have ISG expression levels comparable to those achieved in responders by the treatment with pegIFNa, and it is presently not known, why such a high expression level of ISGs in preactivated patients does not induce a spontaneous clearance of HCV [60]. Patients with genotype 1 infections significantly more often had a preactivated IFN system than those infected with genotypes 2 and 3, providing a possible explanation of why the treatment is more often successful in the latter group [60].

Interferon Signaling in Chronic Hepatitis B Much less is known about HBV interference with the IFN system. The stable expression of HBV polymerase in 2fTGH cells inhibits DNA binding of ISGF3 [108], a finding that was confirmed in Huh7 cells harbouring a replication competent HBV genome [109]. Interestingly, HBV protein expression in these cells induced an upregulation of PP2Ac, and PP2Ac expression was increased in liver biopsies of patients with CHB compared to

M. H. Heim

controls [109]. Therefore, similar molecular mechanisms might be responsible for interference with IFN signaling both in CHB and CHC.

Interferon Signaling in Chronic Hepatitis D Using an in vitro system that allows HDV replication in Huh7 cells [110] it was recently shown that HDV inhibits phosphorylation of STAT1 and STAT2 and thereby inhibits the induction of ISGs [111]. There are presently no biopsy studies to show if HDV inhibits IFN signaling in the liver of patients with CHD.

Summary

›› IFN

are a family of cytokines with antiviral and antiproliferative properties. They are classified into three groups: type I, type II and type III IFNs. In response to viral infections, most cells can induce the transcription of type I IFNs such as IFN-b and several IFN-as. The induction of IFNs is mediated by two main sensory pathways that converge on the transcription factors NF-kB and IRF3/7: the TLR pathway and the RIG-I/MDA5 – Cardif pathway. Secreted IFN-b and all IFN-as bind to a same receptor on the cell surface, IFNAR. IFN-a/b signal transduction from IFNAR to the nucleus requires the activation of two tyrosine kinases, Jak1 and Tyk2, and STAT proteins. Binding of activated STAT1 and STAT2 to specific promoter elements of IFN target genes predominantly results in the transcriptional induction of these genes, with only few target genes being repressed. The induced genes collectively establish an antiviral state in the cell, who’s exact nature is still elusive. Important negative regulators of the IFN signal transduction pathway are the SOCS proteins, PIAS1 and USP18/UBP43. There is increasing evidence that IFN signaling is impaired in CHC, and that despite an activation of the endogenous IFN system, HCV persists in most infected individuals.

12  Interferon Signaling

Multiple Choice Questions 1. Which statement is correct concerning IFN-a induced signaling? (a) IFN-a signals through the receptor-associated kinases JAK1, JAK2, and TYK3 (b) IFN-a activates transcription factors that are able to bind interferon stimulated response elements (ISREs) and gamma activated sequence (GAS) elements (c) JAKs phosphorylate the adaptor molecule Cardif (IPS-1, MAVS, VISA) (d) STAT proteins contain tandem kinase or pseudokinase domains (e) JAK proteins consist of amino-terminal (NH2), coiled-coil, DNA-binding, linker, SH2, tyrosine activation, and transcriptional activation domains 2. Toll-like receptors (TLRs) (a) Activate the type II IFN signaling pathway (b) Detect viral genomes what ultimately leads to the induction of type I IFNs (c) Facilitate viral replication (d) Recognize cytosolic bacterial components bound to retinoic acid inducible gene-I (RIG-I) and melanoma differentiation antigen 5 (MDA5) (e) Are never located on endosomal membranes 3. Find the wrong answer concerning negative regulation of the IFN signaling pathway (a) SOCS1-deficient mice are highly susceptible to viral infections (b) UBP43 deficient mice are resistant to otherwise fatal cerebral infections (c) PIAS proteins bind to activated STATs and inhibit IFN-a induced signaling (d) The phosphatase TcPTP is crucial to prevent excessive inflammatory disease in mice due to an excessive activation of IFN-induced signaling (e) Continuous exposure of cells to IFN results in a “desensitization” 4. The type I IFN-induced regulation of hundreds of genes in a cell is leading to an “antiviral state.” Which of the following statements is correct? (a) Protein kinase R (PKR) mRNA is downregulated by type I IFNs (b) Type I IFNs stimulate cell proliferation and can therefore not be used as anticancer drugs

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(c) None of the known ISG15 target proteins are interferon stimulated genes (ISGs) (d) Activation of 2¢–5¢ oligoadenylate synthetase (OAS) and the RNA nuclease RNase L result in degradation of viral RNA (e) Viperin enhances viral replication in a HCV replicon cell culture system 5. Many patients with chronic hepatitis C have an induction of ISGs in the liver before treatment initiation, (a) And this is especially common in patients with HCV genotypes 2 and 3 (b) And this leads to spontaneous viral clearance in all of these patients (c) And this preactivation of the endogenous IFN system is strongly associated with failure to respond to pegIFNa/ribavirin therapy (d) And therefore respond well to pegIFNa/ribavirin therapy (e) And this indicates a coinfection with hepatitis B virus

Selected Readings 1. Darnell JE Jr, Kerr IM, Stark GR (1994) Jak-STAT pathways and transcriptional activation in response to IFNs and other extracellular signaling proteins. Science 264:1415–1421 (this classical paper describes the fundamentals of Jak-STAT signaling) 2. Akira S, Uematsu S, Takeuchi O (2006) Pathogen recognition and innate immunity. Cell 124:783–801 (an up-to-date review of IFN induction through the activation of pathogen recognition pathways) 3. Stetson DB, Medzhitov R (2006) Type I interferons in host defense. Immunity 25:373–381 (a comprehensive review of the induction of type I IFNs and the IFN effector systems) 4. Feld JJ, Hoofnagle JH (2005) Mechanism of action of interferon and ribavirin in treatment of hepatitis C. Nature 436:967–972 (concise overview of IFNa in the treatment of chronic hepatitis C)

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NF-kB Tom Luedde and Christian Trautwein

Introduction Nuclear factor (NF)-kB was first described in 1986 by the group of Nobel Prize winner David Baltimore as a nuclear factor necessary for immunoglobulin k light chain transcription [1, 2]. It exists in virtually all known cell types and mitochondria [3, 4] and regulates the transcription of an exceptionally large number of genes, including those involved in immune and inflammatory response, cell death, and proliferation [5]. In the last few years, tremendous progress has been made in advancing our understanding of the complex functions of this pathway in vivo. In particular, studies using conditional knockout technology in mice to specifically inactivate this pathway in certain tissues have highlighted the crucial function of NF-kB in linking innate immune responses and cytokine signaling to all kinds of pathological conditions that had not initially been associated with inflammation [6]. As the liver is a preferred source of and target for cytokines, it is not surprising that the NF-kB pathway influences nearly all physiological processes in the liver and hepatic diseases such as acute liver failure, hepatocarcinogenesis, and hepatic fibrogenesis [7]. On the basis of its fundamental importance, it is more than likely that novel molecular therapies targeting members of the NF-kB pathway will be developed in the near future and will enter daily clinical practice. Therefore, to understand the basic principles of this pathway, we will give a short introduction to the structure and basic activation pathways of NF-kB, describe its role in modulating

T. Luedde () Medical Department III, University Hospital RWTH Aachen, Pauwelsstrabe 30, 52074 Aachen, Germany e-mail: [email protected]

hepatocyte cell death, and then focus on its role in hepatic disease conditions, namely hepatocarcinogenesis, liver fibrosis, and hepatitis.

The NF-kB Transcription Factor Family The NF-kB signaling pathway developed early during the course of evolution and is also found in Drosophila and mollusks [8]. In NF-kB-like transcription factors are activated in order to combat infections [9], and the functions of NF-kB in the immune response and in the components of the pathway have been evolutionarily conserved in mammals. NF-kB is a dimer of members of the Rel family of NDA-binding proteins. Nearly all vertebrate NF-kB proteins have been crystallized and their structures determined. The human NF-kB family includes five cellular DNA-binding subunit proteins: p50 (encoded by NFKB1), p52 (NFKB2), c-Rel (REL), p65/RelA (RELA), and RelB (RELB) [10]. The domain architectures of these subunits and the IkB proteins are schematically shown in Fig. 13.1. The NF-kB DNAbinding subunits share an N-terminal Rel homology domain (RHD). They form a unique butterfly-shaped structure composed of b strands arranged in a pattern similar to immuno­globulin domains. This region is responsible for ­DNA binding, dimerization, nuclear translocation, and interaction with the inhibitory IkB proteins [11]. The proteins p65 (RelA), RelB, and c-Rel contain C-terminal transactivation domains that trigger target gene transcription. Of these proteins, p65, which contains two potent transactivation domains (TADs) within its C-terminus, mediates the strongest gene activation [12]. The other two members, p52 and p50, are derived from larger precursors (p105–p50, p100–p52) by either constitutive (p105) or regulated (p100) processing

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_13, © Springer-Verlag Berlin Heidelberg 2010

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steps [13]. They are generally not activators of transcription unless they form heterodimers with p65, RelB, or c-Rel. NF-kB commonly refers to a p50/p65 heterodimer. It is one of the most avidly forming dimers and is the major Rel complex in most cells [14]. Although most NF-kB proteins are transcriptionally active, some combinations such as p50/p50 homodimers can be transcriptionally repressive [15]. In the nucleus, NF-kB recognizes kB sites bearing a consensus sequence of 5′ GGG.Pu.N.W.Py.Py.CC 3′ (Pu is purine, Py is pyrimidine, W is adenine or thymine, and N is any base) [16]. The activity of NF-kB is controlled by IkBs:IkBa (NFKBIA), IkBb (NFKBIB), IkBe (NFKBIE), IkBg (alternative transcript of NFKB1), IkBz (NFKBZ), and Bcl-3 (BCL3) – a family of cytoplasmic inhibitory proteins that share a number of protein/protein interaction domains called ankyrin repeats. The precursor forms p105 (NFKB1) and p100 (NFKB2) are also included in this family, as they contain IkB-like

repeats and therefore inhibit NF-kB activation [11, 17] (Fig.  13.1). NF-kB is effectively sequestered in the cytoplasm by IkB in an inactive state via complex formation and the ability of IkB to mask the nuclear localization site (NLS) of NF-kB [16, 18]. As IkBa is an NF-kB target gene, it also terminates NF-kB activation at the transcriptional level: increased synthesis of IkBa shuts down NF-kB-induced gene expression by IkBa-mediated nuclear export of the DNA-binding subunits, thereby acting within a negative feedback loop [19].

Regulation of NF-kB Activation Different pathways have evolved to activate NF-kB, all of which lead to the generation of ­DNA-binding dimers. At present, the so-called canonical and noncanonical pathways and the DNA damage induced NF-kB

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Fig. 13.1  Structures of the NF-kB and IkB proteins and the components of the IKK complex. Numbers indicate the numbers of amino acids in the human proteins. RHD, Rel homology domain; TAD, transactivation domain; LZ, leucine zipper domain; HLH, helix-loop-helix domain; Z, zinc finger domain; NBD, NEMO-binding domain; CC1/2, coiled-coil domains

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pathway have been identified. More­over, further mechanisms such as p65 posttranslational modifications regulate the activity of this transcription factor. The canonical pathway is the best described and probably most important mediator of NF-kB activation in response to cytokines such as tumor necrosis factor (TNF) or interleukin 1. An essential step in this pathway is the disruption of cytoplasmic NF-kB/IkB complexes, initiated by the phosphorylation of the most important IkB family member, IkBa, at serines 32 and 36 through a high-molecular IkB kinase (IKK) complex. It consists of three tightly associated IkB kinase (IKK) polypeptides. IKK1 (also called IKKa) and IKK2 (IKKb) are the catalytic subunits of the kinase complex and have very similar primary structures with 52% overall similarity [20–22] (Fig. 13.1). (The terms IKK1 and IKKa as well as IKK2 and IKKb are used synonymously. As the names IKK1 and IKK2 distinguish these proteins more clearly from IkBa and IkBb, we will use this nomenclature in this chapter.) In vitro, IKK1 and IKK2 can form homo- and heterodimers [23]. Both IKK1 and IKK2 are able to phosphorylate IkB, but IKK2 has a higher kinase activity in vitro compared with IKK1 [21, 24–26]. Moreover, the IKK complex contains a regulatory subunit called NEMO (NF-kB essential modulator), IKKg, or IKKAP-1 [27–29]. While no direct catalytic domain responsible for IkB phosphorylation has been identified in the NEMO protein, it is essential for IKK activation in a complex process involving adapter proteins such as “TNF-receptor-associated factors” (TRAF) and receptor-interacting protein (RIP), as well as auto phosphorylation and probably regulatory ubiquitination steps [4, 30] (Fig. 13.2). Phosphorylation is a prerequisite for the subsequent polyubiquitination of IkBa by  a ­specific, constitutively active ubiquitin ligase belonging to the SCF family [31]. The ubiquitin-marked IkB proteins are then rapidly degraded by the 26S proteasome, leading to the unmasking of the nuclear localization site (NLS) of NF-kB, and thus allowing nuclear entry, DNA binding, and transcriptional activity of NF-kB. Whereas the two IkB kinases IKK1 and IKK2 fulfill similar functions in TNF-induced NF-kB activation [32], IKK1 also withholds distinct functions independent of IKK2 and canonical NF-kB signaling.  As  such, a noncanonical NF-kB pathway has been described, particularly in B cells. The activation of this IKK1-dependent pathway involves the

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kinase NIK and results in the release of p52/RelB and p50/RelB dimers [33, 34]. It is induced by lymphotoxin b (LTb) for example, and leads to NIK- and IKK1-dependent processing of the p100 precursor protein, which results in the release of p52 [33] (Fig. 13.2). In addition to this noncanonical NF-kB pathway, IKK1 has been demonstrated to exert a direct nucleosomal function by phosphorylating histone H3 [35]. In keratinocytes, IKK1 is directly recruited to retinoic-acid-dependent genes and regulates their transcription [36], underlining that this IKK subunit might withhold specific functions that are independent of NF-kB. The functions of IKK1dependent pathways in the liver have not been as extensively studied as those for canonical NF-kB signaling, but they seem to be important for the prevention of cholangitis, and they also seem to modulate hepatocarcinogenesis by a mechanism that is yet to be defined [32]. DNA-damage-induced NF-kB activation has been observed mainly in vitro, for instance after doxorubicin stimulation or UV radiation, and involves mitogenactivated protein kinase (MAPK)-dependent alternative IkBa phosphorylation [37–40]. Moreover, the activation of NF-kB by different forms of DNA damage has been suggested to involve nuclear-to-cytoplasmatic shuttling and SUMOylation of NEMO [41, 42]. However, the biological role of this pathway in liver physiology and pathology is not yet known. A growing number of studies have suggested that, besides the formation and nuclear translocation of NF-kB dimers, posttranslational modifications of NF-kB subunits might also influence NF-kB activation. In particular, the phosphorylation of p65 and its acetylation appear to significantly modify its transcriptional activity. Numerous phosphorylation sites have been described on the p65 protein, which are targeted either by a single or by several kinases. Serine 276 is the best-characterized phosphorylation site of p65. Fibroblasts containing a mutant p65 with serine 276 replaced with alanine instead of the wildtype form showed an impaired tumor necrosis factor (TNF)-induced expression of the NF-kB target gene interleukin (IL)-6, and these cells lost their NF-kBdependent protection against TNF-induced apoptosis [43]. Whereas both of the catalytic subunits of PKA, PKAc, and MSK-1 have been identified as serine 276 kinases [43, 44], phosphorylation at serines 468 and 536 appears to involve IKK1 and IKK2 but

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Fig.  13.2  Canonical NF-kB activation (left part): upon TNF binding to its receptor, the IKK complex (IKK1, IKK2, and NEMO) gets activated and mediates phosphorylation and ubiquitination of the inhibitory protein IkBa, thereby releasing the p50/p65 NF-kB dimer and unmasking its nuclear localization site. NF-kB translocates to the nucleus and induces transcription of antiapoptotic and inflammatory genes. Noncanonical NF-kB activation (right part): binding of lymphotoxin-b to its

receptor leads to NIK-dependent activation of IKK1 homodimers. IKK1 cleaves the inactive precursor protein p100 to the active NF-kB protein p52, which associates with RelB (and other factors) and translocates to the nucleus. TNF, tumor necrosis factor; TNF-R1, TNF receptor 1; FADD, Fas-associated death domain; TRADD, TNF-receptor-associated death domain; TRAF2, TNF-receptor-associated factor; JNK, c-Jun-(N)-terminal kinase; LT-b lymphotoxin-b

not NEMO [32, 45, 46], demonstrating an ­additional function of these IKK subunits in NF-kB ­activation besides IkB phosphorylation. Another path to the posttranslational modification of NF-kB is the acetylation of p65, which might enhance its transactivation potential [11]. However, the significance of these posttranscriptional p65 modifications in vivo and especially in the liver are presently not well understood, and studies with genetically modified animals bearing point mutations in the Rela gene are  needed for a better understanding of their biological roles.

NF-kB in the Regulation of Hepatocyte Cell Death NF-kB Protects Hepatocytes from TNF-Induced Apoptosis Tumor necrosis factor (TNF) family ligands such as TNF, Fas, and “tumor necrosis factor related apoptosis inducing ligand” (TRAIL), as well as their corresponding receptors, are associated with several hepatic disease conditions and influence different concepts of hepatocyte

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cell death [47] (see Chaps. 13 and 14). Necrotic cell death is the result of acute metabolic disruption with ATP depletion, ion dysregulation, mitochondrial and cellular swelling, and activation of degradative enzymes. On the other hand, apoptosis, synonymously used for programmed cell death, is an important mode of cell death that occurs when specific cells need to be removed during development or normal tissue turnover. Classic apoptosis leads to the orderly resorption of target cells without severe impairment of cellular metabolism, and its onset is triggered by specific and active signaling through the activation of a cascade of cysteine–aspartate proteases called caspases [48, 49]. TNF signals through two distinct cell surface receptors, TNF-R1 and TNF-R2, among which TNFR1 initiates the majority of TNF’s biological activities [50] (see Chap. 10). Aside from the activation of caspases, the ligand binding of TNF to its receptor also leads to the activation of the NF-kB pathway (see Sect.  13.3 and Fig.  13.2). Numerous studies have shown that NF-kB provides survival signals in the context of death-receptor-induced apoptosis in hepatocytes. Evidence that NF-kB governs critical antiapoptotic genes comes from well-described animal models. Injection of TNF into mice and addition of TNF to hepatocyte cell cultures resulted in the activation of NF-kB binding [51]. In contrast to Fasmediated apoptosis, hepatocytes are resistant to apoptosis induced by TNF or bacterial lipopolysaccharides (LPS) – potent inducers of endogenous TNF secreted by immune cells [52] – unless they are treated with inhibitors of transcription such as cycloheximide or actinomycin D [53–55]. Further evidence that NF-kB mediates this protective transcriptional program comes from genetic experiments. Knockout mice lacking the Rela gene die between days E15 and E16 post coitum as a result of fetal hepatocyte apoptosis [56]. This is caused by increased sensitivity towards TNF, as Tnf/Rela doubledeficient mice are rescued from embryonic lethality [57]. c-Rel may partially compensate for p65 in blocking liver apoptosis in that Rel/Rela double-mutant mice show liver degeneration approximately 1.5 days earlier than Rela single-knockout mice [58]. Genetic experiments have also highlighted the crucial functions of the IKK subunits in TNF-mediated liver apoptosis. Ikk2−/− mice die in utero at embryonic

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day (E) 12.5 as a result of massive apoptosis in the liver, and fibroblasts from these mice show no activation of NF-kB in response to TNF and interleukin (IL)-1 [59–61]. A similar phenotype was noted in mice lacking Nemo, which also show massive hepatocyte apoptosis and a defect in NF-kB activation upon TNF stimulation in primary murine embryonic fibroblast (MEF) cells [62]. In contrast, mice lacking Ikk1 die shortly after birth and display a phenotype marked by thickening of skin and limb as well as skeletal defects but normal liver development [63, 64]. Therefore, during embryogenesis, IKK2 and NEMO appear to be the critical subunits for NF-kB activation and the protection of hepatocytes from proinflammatory cytokines such as TNF, whereas the distinct phenotype of Ikk1deficient animals underlined the functional diversity of the catalytic IKK subunits. Recent studies using conditional cre/loxP-based knockout technology to target the different IKK subunits specifically in hepatocytes demonstrated the essential functions of NF-kB and the IKK complex in the prevention of TNF-induced liver failure in the adult mouse liver. As such, deletion of Nemo in hepatocytes resulted in complete blockage of TNF-induced NF-kB activation, along with massive hepatocyte apoptosis in isolated hepatocytes and acute liver failure in vivo in response to low-dose LPS injections [65, 66]. Surprisingly, hepatocyte-specific deletion of Ikk2 resulted in significant remnant NF-kB activation in response to TNF in hepatocytes and did not sensitize the livers of adult mice to TNF- or LPS-induced liver failure [65–67], suggesting that in the adult liver IKK1 and IKK2 might demonstrate more redundancy in the canonical NF-kB pathway than during embryonic development. Indeed, mice with combined hepatocyte-specific deletions of Ikk1 and Ikk2 exhibited acute liver failure and complete blockage of the NF-kB pathway in response to LPS injection, comparable with Nemo-mutant mice [32]. Thus, in contrast to the embryonic situation, IKK1 can substitute for the function of IKK2 in canonical NF-kB activation in the adult mouse liver. However, the fact that mice with combined ablation of IKK1 and IKK2 in parenchymal liver cells, but not Nemo-mutant mice, develop a lethal cholangitis [32] underlines the notion that IKK1 withholds additional, noncanonical functions in these cells that need to be determined in the future.

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Mediators of the Antiapoptotic NF-kB Function: Interactions with Stress-Related Signaling Pathways NF-kB induces a variety of target genes with potential antiapoptotic functions. Among these are the cellular inhibitors of apoptosis (c-IAPs) such as c-IAP1, c-IAP2, XIAP, antiapoptotic Bcl-2 family members (A1 and Bcl-xL), c-FLIP, and TRAF1/TRAF2, which are induced upon TNF stimulation in an NF-kBdependent manner [68, 69]. However, the mechanism, by which NF-kB exerts its antiapoptotic function in TNF-induced apoptosis remained enigmatic until recent genetic studies unraveled a complex regulatory network regulated by NF-kB that involves interactions with other stress-related pathways, namely the c-Jun-(N)-terminal kinase (JNK) and the p38 mitogen-­activated protein kinase (MAPK) signaling cascades (Fig. 13.3). The JNK cascade is strongly activated in TNFmediated apoptosis via TRAF2 and RIP [70] (see Chap. 14). Functional interconnection between NF-kB and JNK was revealed by two studies which showed that TNF-induced activation of JNK is prolonged in cells that are deficient in NF-kB activation (p65 and Ikk2 knockout MEFs and cells stably expressing degradation­ -resistant IkBa) and that prolonged JNK activation in these cells promoted apoptosis [71, 72]. Moreover, it was shown that prolonged JNK activation induced by TNF depends on the production of reactive oxygen species (ROS) and is negatively regulated by NF-kB, which in turn blocks ROS accumulation [67, 73]. Accumulation of ROS in NF-kB defective cells is caused by reduced expression of an NF-kB-dependent gene coding for an important ROS-metabolizing enzyme called superoxide dismutase 2 (Sod2) [66, 74]. This in turn leads to oxidation and inactivation of JNKinactivating phosphatases through the conversion of their catalytic cysteine to sulfenic acid, and subsequently prolonged JNK activity upon TNF stimulation, which can be prevented by antioxidant treatment [74]. Finally, a further study revealed the connection between prolonged JNK activation and TNF-induced apoptosis in NF-kB-defective hepatocytes by demonstrating that TNF-mediated JNK activation accelerates turnover of the NF-kB-induced antiapoptotic protein c-FLIP, an inhibitor of caspase-8. This is mediated by JNKmediated phosphorylation and activation of the E3

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Fig.  13.3  Mediators of the protective function of NF-kB in TNF-induced apoptosis. Activation of NF-kB induces transcription of the c-FLIP gene, which in turn inhibits TNF-dependent activation of caspase-8 and subsequent caspases. Moreover, it prevents ROS-dependent prolonged JNK activation and activation of the E3 ligase Itch, and thereby prevents c-FLIP ubiquitination and degradation. p38a inhibits MKK4 hyperactivation of JNK and thus collaborates with NF-kB to prevent TNFinduced cell death. SOD2, superoxide dismustase 2; MKK4, MAPK kinase 4; c-FLIP, FLICE-inhibitory protein

ubiquitin ligase Itch, which specifically ubiquitinates c-FLIP and induces its proteasomal degradation [75] (Fig. 13.3). It was recently demonstrated that the p38MAPK signaling cascade also collaborates with the NF-kB pathway to protect hepatocytes from TNF/LPS-induced liver injury. The p38MAPK pathway regulates cellular responses to stress and is implicated in cell proliferation, differentiation, and apoptosis [76, 77]. Conditional deletion of p38a, the main p38 isoform, in ­parenchymalliver cells results in excessive JNK activation in response to LPS injection, demonstrating a physiological function of p38a in suppressing JNK activation upon TNF stimulation [78]. However, the mechanism for this ­suppression is different from that used by NF-kB to control JNK activity, and involves the MAP kinase

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kinase MKK4. Furthermore, combined p38a/Ikk2 liver-mutant mice, but not the respective single mutants, developed liver failure upon LPS injection [78]. Therefore, p38a deficiency only sensitizes the liver to cytokine-induced damage when combined with partial NF-kB inhibition. These collective findings underline the idea that the network of NF-kB, JNK, and p38MAPK is a crucial regulator of hepatocyte cell death. The close interactions of its components in hepatocytes will have to be taken into consideration when molecular drugs are developed in the future to target specific axes of this network.

NF-kB Promotes Inflammatory Necrosis of Hepatocytes upon Ischemia/Reperfusion In contrast to models of death-receptor-ligand stimulation, NF-kB has been demonstrated to promote cell death in a model of ischemia/reperfusion injury. Ischemia/reperfusion (I/R) injury in rodents represents a complex model that reflects liver damage after organ  transplantation, tissue resections, or hemorrhagic shock, whose molecular mechanisms are poorly understood. In most studies, the injury detected after temporary clamping of the hepatic blood flow and subsequent reperfusion leads to an excessive inflammatory response followed by necrotic cell death. It has been proposed that, instead of linear apoptotic signaling, I/R injury consists of two different phases, one displaying acute cellular injury and a secondary, subacute phase resulting from inflammatory responses [79]. Genetic experiments identified TNF but not Fas as being a critical component in the mediation of I/R injury to the liver [80], suggesting that NF-kB might also withhold a specific function in this context. NF-kB DNA binding occurs quickly upon hepatic I/R [81]. It has been suggested that this process might occur as an alternative to that which occurs during death-receptor-dependent signaling. Whereas TNF stimulation leads to IKK-dependent phosphorylation of both IkBa and IkBb on serine residues, it was suggested that, during I/R injury, NF-kB gets activated by c-Src-dependent tyrosine phosphorylation of IkBa but not IkBb on Tyr42, and that this process might occur in the absence of IkBa ubiquitin-dependent degradation

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[82, 83]. However, data from our own laboratory provided evidence that this process is still dependent on IKK2, as conditional knockout mice for IKK2 in hepatocytes show a defect in NF-kB activation upon I/R [65]. This partial NF-kB inhibition surprisingly did not increase cell death, but instead resulted in a dramatic attenuation of cell death and necrosis, along with significantly lower aminotransferases [65]. At present, it is not clear how this cell-death promoting action of NF-kB in the I/R model is mediated. Mice with hepatocyte-specific Ikk2 deletion showed reduced infiltration of immune cells, pointing towards a proinflammatory role of NF-kB in this model. However, the protective effect of Ikk2 deletion was demonstrated at very early time points after reperfusion, even before immune cell invasion could be detected in WT animals, suggesting that NF-kB directly promotes hepatocyte cell death on an intracellular level by an unknown mechanism. These data demonstrate that whether NF-kB withholds a preventive or promoting function in the mediation of hepatocyte cell death depends on the experimental model.

The Role of the NF-kB Pathway in Liver Disease NF-kB in Liver Cancer Many chronic liver diseases are not sufficiently treatable at present and often progress to liver cirrhosis and hepatocellular carcinoma (HCC), a tumor that is rapidly increasing in incidence worldwide [84]. Whereas eradicating the primary cause of liver tumors, such as chronic viral hepatitis, is currently unattainable in a significant number of cases, the identification and characterization of central signaling pathways that play a role in the mediation of hepatocarcinogenesis and the subsequent possibility of a targeted pharmacological modification of the central molecules involved in these pathways may be a very promising and probably more realistic objective in the future. In contrast to many other tumors, an HCC usually arises on the basis of chronic inflammation caused for instance by viral hepatitis [84]. NF-kB is constitutively activated in many hepatocellular carcinomas [85], and it is not surprising that a pathway like NF-kB, which links inflammation and cytokines with basic

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cellular processes like apoptosis or proliferation, plays a crucial role in hepatocarcinogenesis. Again, recent genetic studies in mice have provided a novel insight into the role of this pathway in the development of HCC [66, 86, 87]. However, the specific experimental procedures of these studies with regard to genetic targeting and tumorigenesis protocols have highlighted that the function of NF-kB might differ depending on the stage of tumorigenesis (initiation, promotion, and metastasizing), on the fine tuning of NF-kB activation, and especially on the hepatic cell type (parenchymal vs. nonparenchymal cells) in which NF-kB is active or inhibited, respectively. In one study, the role of the NF-kB pathway was examined in mdr2-KO mice which spontaneously develop a cholestatic hepatitis due to defective biliary transport, and subsequently HCCs [88]. In this model, inhibition of NF-kB by a hepatocyte-specific inducible IkB super-repressor transgene results in a  dramatic reduction in tumor development [86]. Moreover, hepatocyte transformation and initiation of HCCs in this model is rather unaffected by the inhibition of NF-kB, whereas an inhibition of NF-kB during the later stages of tumor promotion contributes mainly to the observed phenotype. From their study, the authors convincingly concluded that NF-kB is a hepatic tumor promoter that is needed during later stages of tumor promotion to protect the transformed liver cells from TNF-induced cell death. Two other studies suggested an opposite, tumorsuppressive role of NF-kB in the liver during the early steps of tumor initiation. Mice with hepatocyte-­specific deletion of Ikk2 (and thus partial NF-kB inhibition, see Sect.  13.4.1) injected at the age of 2 weeks with the DNA-damaging agent diethyl nitrosamine (DEN) develop more and larger liver tumors than their respective wild-type litter mates [87]. In line with this finding, mice with the hepatocyte-specific deletion of Nemo and thus, complete NF-kB inhibition develop spontaneous hepatitis at a young age, and subsequently multiple liver tumors [66]. Carcinogenesis in these models is based on increased hepatocyte cell death upon DEN treatment [87], or even spontaneous hepatocyte apoptosis in Nemo-mutant mice triggered by small intrinsic doses of LPS and cytokines released by commensal bacteria in the guts of these animals. Hepatocyte apoptosis induced by increased oxidative stress (see Sect. 13.4.2) then results in the release of interleukin 1a [89] and the activation of nonparenchymal liver

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cells such as hepatic Kupffer cells (see Chap. 4). These cells in turn produce mitogenic cytokines such as IL-6 in an NF-kB-dependent manner and induce compensatory liver regeneration, completing a vicious cycle of hepatocyte apoptosis and ­compensatory proliferation that culminates in hepatocarcinogenesis [90]. Interestingly, deletion of Ikk2 in hepatic Kupffer cells in addition to hepatocytes prevents NF-kB-dependent cytokine release from these immune cells and thus inhibits liver regeneration and hepatocarcinogenesis [87], demonstrating that, in contrast to hepatocytes, NF-kB has a procarcinogenic function in Kupffer cells. Thus, the function of NF-kB varies not only between the early and late phases of carcinogenesis but also between parenchymal and nonparenchymal liver cells, highlighting the need for a deeper understanding of this pathway before the application of systemic drugs  targeting NF-kB in the prevention or treatment of HCC.

NF-kB in Liver Fibrosis Liver fibrosis and cirrhosis represent the common final pathways of virtually all chronic liver diseases and are a massive health care burden worldwide [91]. At present, effective pharmacological treatment options to limit liver fibrosis in response to chronic injury are very limited, underlining the need to study the molecular mechanisms that underlie this process. In the chronically injured liver, repeated and overlapping phases of inflammation and wound healing overwhelm the normal regenerative process and instead promote the net deposition of fibrillar collagen [92]. The principal cells responsible for promoting the hepatic deposition of cross-linked fibrillar collagen are activated hepatic stellate cells (HSC) hepatic myofibroblasts (HM), which are scarce in the normal liver but increase dramatically in number at sites of injury [91]. NF-kB may play an important role in not only the initial inflammatory response of hepatocytes to injury but also HSC and HM function. NF-kB is found in an unusual persistent activated form in HM, where its key role might be to promote the inflammatory phenotype [93]. Moreover, with regard to its function in hepatocytes, it may be crucial to prevent the apoptosis of HM, because the proapoptotic activities of insulin-like growth factor (IGF-)1 on rodent HM correlate with decreased NF-kB activity [94]. Conversely, the

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antiapoptotic effects of transforming growth factor (TGF) b and TNF are associated with increased NF-kB activity [95]. It is believed that the resolution of liver fibrosis and the regeneration of normal tissue require the regulated removal of HM by apoptosis [96]. Thus, it is possible that inhibition of NF-kB in HM might diminish the process of liver fibrosis or ameliorate the reversion of the fibrotic liver. Indeed, the treatment of cultured human and rat HMs with the anti-inflammatory drug sulfasalazine induced HM apoptosis in a dose-dependent manner [97]. Single-dose administration of sulfasalazine inhibited hepatic fibrogenesis in experimental liver fibrosis in rats with a corresponding reduction in activated HM, hepatic expression of procollagen, and TIMP-1, and increased MMP-2 activity [97]. However, sulfasalazine is a potent but rather unspecific inhibitor of IKKdependent NF-kB activation and influences the activities of many other intracellular inflammatory pathways [98], meaning that studies with the conditional targeting of NF-kB in HSC/HM are needed to evaluate the specific role of NF-kB in these cells. However, further evidence for a specific role of NF-kB in HSC comes from a recent study which, demonstrated that Toll-like receptor (TLR)-4 activation in hepatic stellate cells promotes liver fibrogenesis by promoting macrophage chemotaxis and enhancing TGF-b signaling via the NF-kB target gene Bambi [99].

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activating NF-kB, which may lead to viral persistence and possibly hepatocarcinogenesis [107]. Moreover, the HCV core protein can promote the proliferation of human hepatoma cell lines by activating NF-kB, thereby leading to TGF-a induction and activation of the ERK/MAP kinase pathway [108]. In summary, the NF-kB pathway might be involved in perpetuating liver inflammation and blocking hepatocyte apoptosis in HBV and HBC infections, showing that these viruses acquired the ability to modify NF-kB function to their own advantage [112]. Recent genetic studies have demonstrated an additional role of hepatic NF-kB in modulating systemic metabolic processes and the development of non­ alcoholic steatohepatitis (NASH). Hepatic NF-kB over-activation leads to a type 2 diabetes phenotype characterized by hyperglycemia, profound hepatic insulin resistance and moderate systemic insulin resistance, including effects in muscle [109]. On the other hand, hepatocyte-­specific inhibition of NF-kB prevents the development of a type 2-diabetic phenotype in mice fed on a high-fat diet [110, 111]. In contrast to a beneficial effect of NF-kB inhibition on systemic glucose tolerance, hepatocyte-specific deletion of NEMO results in the spontaneous development of NASH in mice and synergizes with a high-fat diet in the development of liver steatosis as a consequence of increased oxidative stress, decreased peroxisome proliferatoractivated receptor (PPAR-alpha), and increased PPARgamma expression [66, 111].

NF-kB in Viral Hepatitis and Fatty Liver Disease

NF-kB as a Therapeutic Target

Worldwide, the hepatitis B virus (HBV) remains the leading cause of liver disease and hepatocarcinogenesis, followed by hepatitis C virus (HCV) infection [100]. These viruses interact with the NF-kB pathway on multiple levels. Both HBV and HCV modulate the NF-kB pathway. Activation of NF-kB by HBV has been shown to be mediated by an interaction of the viral HBx protein with IkBa and the precursor/inhibitor subunit p105 [101, 102]. In this context, NF-kB suppresses HBx-mediated apoptosis [103]. Moreover, both HBs and HBc antigen can activate NF-kB [104, 105]. Regarding HCV, both nonstructural and HCV core proteins influence NF-kB [106]. HCV infection may cause antiapoptosis by

Numerous chemical compounds and viral vectors that modulate NF-kB have been developed or are under development, including antioxidants, protea­some inhibitors, and molecular IKK inhibitors [112]. As outlined above, genetic studies using conditional knockout technology have resulted in an enormous gain in knowledge on the role of NF-kB in the liver. However, these data have highlighted the ­differential functions and partially opposing effects of NF-kB activation in different hepatic cell types (­hepatocytes/Kupffer cells), different disease models (e.g.,  TNF-induced liver failure/ischemia/reperfusion injury), and even different stages of the same disease (early vs. late stages of hepatocarcinogenesis). On the basis of the data presented previously,

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an NF-kB inhibition in hepatocytes might prove beneficial in, say, the prevention of ischemic liver injury, the prevention of hepatic fibrogenesis, the treatment of type 2 diabetes, or the treatment of advanced hepatocellular carcinoma, whereas specific targeting of this pathway in Kupffer cells might prevent early stages of hepatocarcinogenesis. However, a cell-specific application of drugs is not possible at present. In terms of systemic drugs, antioxidants appear to be a safe way to try to prevent hepatocarcinogenesis on the basis of chronic hepatitis [66]. Molecular inhibitors of IKK2 that are under development allow a more specific modulation of this pathway than proteasome inhibitors and have shown similarly beneficial effects to genetic targeting in certain hepatic disease models [65, 113]. Specific inhibition of IKK2 would also retain the function of IKK1 in preventing spontaneous hepatocyte apoptosis, as seen in Nemo-conditional knockouts. However, systemic IKK2 inhibition withholds the risk of excessive IL-1b processing and secretion, and thereby a generalized inflammatory response [114]. Therefore, further genetic and preclinical studies are needed to better understand the role of NF-kB, to further dissect the pathway, and possibly to find new mediators of NF-kB-dependent signals that would allow more specific modulation of this pathway in certain hepatic diseases without hepatic or systemic toxicity.

Summary

›› The NF-kB family of transcription factors reg-

››

››

ulates the transcription of an exceptionally large number of genes, including those involved in immune and inflammatory response, cell death, and proliferation. It consists of five members–p50, p52, p65, c-Rel, and RelB– that build homo- and heterodimers, among which p50/p65 is the most important dimer in the canonical NF-kB pathway. NF-kB activation in response to cytokines such as TNF relies on phosphorylation and degradation of the inhibitory protein IkBa by the IKK and complex, consisting of the catalytic subunits IKK1and IKK2, and the regulatory subunit NEMO.

›› Genetic studies have demonstrated that canon-

››

›› ››

ical, IKK-dependent NF-kB activation prevents TNF- and LPS- induced liver failure and hepatocyte apoptosis in mice. This protective action is mediated by NF-kB-dependent transcription of the caspase-inhibitory protein c-FLIP and the inhibition of JNK-dependent degradation of c-FLIP by NF-kB. The role of NF-kB in hepatocarcinogenesis differs between hepatic cell types and the early and late stages of tumor development. Whereas NF-kB promotes tumor development in hepatocytes in the later stages of inflammatory hepatocarcinogenesis as well as tumor initiation in hepatic Kupffer cells by inducing IL-6, it acts as a tumor suppressor in hepatocytes during the early stages of HCC development. NF-kB influences other hepatic disease processes such as fibrogenesis, viral hepatitis, ischemia/reperfusion injury, and NASH. The prominent role of NF-kB in the liver makes it an attractive candidate for the development of novel therapies based on molecular drugs targeting this pathway, but further studies are needed to better understand its cell-specific functions in order to avoid unwanted side effects.

Multiple Choice Questions 1. Which answer reflects best the current view of the function of the NF-kB pathway in the liver? (a) p52/RelB heterodimers are the most important activators of canonical NF-kB target genes (b) IKK1 masks the nuclear localization site of RelA (c) IKK2 mediates NIK-dependent cleavage of the precursor protein p100 (d) The NF-kB pathway controls IkBa transcription (e) IKK2 and NEMO show a high structural homology. 2. Which answer best reflects the current view of the function of the NF-kB pathway in the liver? (a) IKK2 constitutive knockout mice do not show a significant phenotype

13  NF-kB

(b) I KK1 constitutive knockout mice show a skin phenotype, but develop normal livers (c) The noncanonical NF-kB pathway is only active in lymphocytes and does not play a role in parenchymal organs such as the liver (d) NEMO conditional knockout mice are protected from TNF-induced hepatocyte apoptosis (e) IkBa induces NF-kB degradation. 3. Which answer best reflects the current view of the function of the NF-kB pathway in the liver? (a) NF-kB activation promotes the accumulation of reactive oxygen species in hepatocytes (b) Prolonged JNK activation inhibits apoptosis in hepatocytes (c) NF-kB and p38a modulate JNK activity by the same molecular mechanism (d) NF-kB prevents the inflammatory response upon hepatic ischemia/reperfusion (e) Accumulation of reactive oxygen species leads to the inactivation of JNK-directed phosphatases after TNF stimulation of hepatocytes. 4. Which answer best reflects the current view of the function of the NF-kB pathway in the liver? (a) IL-6 release from hepatic Kupffer cells promotes hepatocyte proliferation and hepatocar­ cinogenesis (b) Hepatocyte-specific deletion of IKK2 in mice leads to spontaneous tumor development in the liver (c) Conditional deletion of IKK2 in hepatic Kupffer cells promotes tumor development in a model of chemical hepatocarcinogenesis (d) NF-kB and p38a have opposing functions in TNF-induced apoptosis in hepatocytes (e) Transformed hepatocytes are resistant to TNFinduced apoptosis independent of NF-kB. 5. Which answer best reflects the current view of the function of the NF-kB pathway in the liver? (a) Present data indicate that NF-kB activation in hepatic stellate cells inhibits fibrogenesis (b) Hyper-activation of NF-kB in hepatocytes protects from insulin resistance (c) Deletion of NEMO in hepatocytes protects from fatty liver disease (d) Systemic IKK2 inhibition might induce an inflammatory response through a mechanism involving IL-1b (e) Hepatitis B virus but not hepatitis C virus interacts with the NF-kB pathway.

211 Acknowledgments  Work in the laboratory of T.L. is supported by starting grants from the European Research Council (No. 208237), the German Research Foundation (SFB/TRR 57), and the Interdisciplinary Centre for Clinical Research “BIOMAT.” within the Faculty of Medicine at RWTH Aachen University (to T.L. and C.T.). Work in the laboratory of C.T. is supported by the German Research Foundation (SFB/TRR 57)/ (SFB 542) and the German Cancer Foundation.

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JNKs in liver diseases

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R. Schwabe

Introduction c-Jun NH2-terminal kinases (JNKs) are members of the mitogen-activated protein kinase (MAPK) family. Like other MAPKs, JNKs are conserved throughout eukaryotic evolution, activated via three-tiered phosphorylation cascades, and involved in a wide range of cellular responses to stress. JNKs have been shown to play important roles in proliferation, cell death, inflammation and cell metabolism. These seemingly unrelated responses are part of an overall stress response program that ensures proper repair of cells sustaining minor damage, elimination of cells sustaining irreversible structural or genetic damage, as well as their proper replacement.

Functions and Targets of JNKs in Cell Biology

addition to the archetypical JNK target c-Jun, JNKs phosphorylate a wide range of other targets which is likely to be the basis for their involvement in a wide range of biological processes. While JNK1 and JNK2 are ubiquitously expressed, JNK3 is largely restricted to neuronal cells [3]. Different affinities to specific substrates are thought to be the reason for the distinct functions of JNK1 and JNK2 (see paragraph below). However, there is still considerable debate on specific functions of JNK1 and JNK2 in the regulation of important biological functions such as proliferation and cell death. It is likely that there is at least some functional redundancy between the JNKs, as JNK1 and JNK2 have common targets such as c-Jun. Moreover, the fact that JNK1- and JNK2-deficient mice are viable and phenotypically normal and that JNK1-/JNK2-double deficient mice are embryologically lethal suggests that JNK1 can fulfill some functions of JNK2 and vice versa [4, 5].

JNK Isoforms

JNK Activation and Inactivation

The JNK proteins are encoded by three genes, jnk1, jnk2, and jnk3, which are alternatively spliced to create at least 10 isoforms [1]. Each JNK is expressed as a short 46-kDa form and long 54-kDa form [1]. JNKs display greater than 85% identity at the protein level, and have been known for their ability to phosphorylate the target c-Jun at serine 63 and 73 [2]. In

JNKs are activated by a wide range of stimuli including “stress” such as UV- and g-irradiation, cytokines including TNFa, IL-1b, PDGF, and TGFb, bacterial products such as LPS, peptidoglycan, and nonmethylated CpG-DNA, and endoplasmatic reticulum (ER) “stress.” Activation of JNKs is achieved through a three-tier phosphorylation cascade involving several MAPK kinase kinases (MKKKs or MAP3Ks) and two mitogen-­activated protein kinase kinases (MKKs or MAP2Ks) that directly phosphorylate and activate JNKs (see Fig.  14.1). Among the MKKKs, TAK1 plays an important role in initiating JNK activation in response to inflammatory cytokines, TLR3, TLR4,

R. Schwabe Department of Medicine, columbia University, Russ Berrie Pavilion, Room 415, 1150 St. Nicholas Avenue, New York, NY 10032, USA e-mail: [email protected]

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_14, © Springer-Verlag Berlin Heidelberg 2010

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Activators ROS

Cytokines, PAMPs, UC, γ-Irradiation Trx

MKKKs

ASK1 TAK1, MLKs, MEKKs

MKKs JNKs

MKPs

MKK4

MKK7

JNK1

JNK2

JNK targets

Transcription factors, nuclear receptors, Bcl-2 family members, ubiquitin ligases

Responses

Proliferation, Cell death, inflammation, Migration, Wound healing

Fig. 14.1  JNK activation. Cytokines, PAMPs, UV- and g-irradiation activate the JNK pathway through a three-tier kinase cascade. Activated MKKKs (TAK1, ASK1, MEKKs, and mixed lineages kinases) phosphorylate MKK4 and MKK7. Cytokines predominantly activate MKK7, whereas UV- and g-irradiation predominantly activate MKK4. MKK4 and MKK7 then phosphorylate JNKs which in turn phosphorylate various targets to achieve a wide range of biological responses. Reactive oxygen species (ROS) achieve activation of JNKs by blocking the inhibitory effects of thioredoxin (Trx) on ASK1, and the inhibitory effects of MAP kinase phosphatases (MKPs) on JNKs

amplify this signal. The amplification of JNK activation by MKK4 does not require MKK4 activation but is mediated by basal MKK4 activity [12]. ROS can at least partially bypass this three-tier cascade to activate JNKs by inactivating MAP kinase phosphatases (MKPs). A number of different scaffolding proteins such as JIP proteins not only recruit JNKs, MKKs, and MKKKs to promote their proper interaction and initiate signaling, but also recruit negative regulators of signaling such as MKPs. Recent studies support an important role of MKPs in the negative regulation of JNK signal transduction [13]. At least 2 MKPs terminate JNK responses as demonstrated by increased JNK activity in MKP1- and MKP5-deficient mice [13]. Notably, specific targeting and inactivation of MKPs result in sustained JNK activation in some signaling pathways such as the TNF pathway (see below) [14]. Inhibition of MKPs may also be the main mechanism for JNK activation following stimuli such as UV irradiation [15].

Regulation of JNK Activity by Other and TLR9 as well as B- and T-cell receptor activation Kinase Pathways [6–8]. The MKKK ASK1 appears to play a role in mediating JNK activation in response to reactive oxygen species (ROS) and is also involved in cytokineand ER stress-induced JNK activation [9, 10]. ROS-induced activation of ASK1 is achieved through the inactivation of thioredoxin, a redox-sensitive binding partner and inhibitor of ASK1 activity [11]. The contribution of the MKKKs mixed lineage kinases and MEKKs to the activation of the JNK signaling cascade is not well characterized. MKK4 and MKK7 are responsible for JNK phosphorylation in kinase domain VIII which confers JNK activation. Maximal activation of JNKs requires dual phosphorylation of threonine and tyrosine residues within kinase domain VIII [12]. MKK7 preferentially phosphorylates threonine 183, and MKK4 preferentially phosphorylates tyrosine 185. However, only MKK7 is efficiently activated in response to TNFa and IL-1b [12]. Accordingly, while there is no JNK activation in response to TNFa and IL-1b in MKK7-deficient MEFs, the JNK activation in MKK4-deficient MEFs [12] is reduced by 50%. These results have led to the hypothesis that activated MKK7 plays the key role in initiating JNK activation in response to inflammatory stimuli whereas the function of MKK4 is merely to

In addition to activating JNKs, inflammatory stimuli and stress concomitantly trigger the activation of several other kinase pathways such as p38 and IKK. These pathways not only collaborate with the c-Jun/AP-1 pathway at the level of target gene transcription, but also have direct effect on the activation of JNKs. The IKK/NF-kB cascade upregulates molecules such as manganese superoxide dismutase (MnSOD) to eliminate ROS [14]. If the IKK/NF-kB cascade is inactivated, increased ROS levels lead to an inactivation of MKPs and prolonged JNK activation [14]. p38a antagonizes the JNK/c-Jun pathway by inhibiting MKK7, the kinase that is largely responsible for JNK activation following TNFa stimulation. Accordingly, p38sdeficient MEFs and livers display a prolonged JNK activation [16–18].

JNK Substrates and Target Genes c-Jun represent the archetypical target of JNKs and is  phosphorylated at serine residues 63 and 73 in its  transactivation domain [2]. c-Jun dimerizes with

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proteins of the jun, fos, ATF, and Maf families to form activating protein-1 (AP-1) transcription factor complexes [19]. c-Jun is the most potent transcriptional activator of the AP-1 complex, and its transcriptional activity is strongly enhanced by JNK-mediated ­phosphorylation at Ser63 and Ser73 [20]. Moreover, c-jun is an immediate early gene that contains an AP-1 site and is strongly and very rapidly upregulated following JNK activation in a positive amplification loop [21, 22]. Interestingly, JNKs phosphorylate other members of the AP-1 complex such as JunD and ATF-2. Although JunD does not contain an effective JNK docking site, it can be phosphorylated after heterodimerization of either c-Jun or JunB, both of which have efficient docking sites. ATF-2 is another unit of the AP-1 complex that can be phosphorylated by JNKs and dimerizes with c-Jun to bind to a nonconsensus AP-1 site in the c-jun promoter and stimulate c-Jun expression [22]. Phosphorylation of Elk-1 by JNK can upregulate c-Fos levels which has been shown to be important for AP-1 activity as Jun/Fos heterodimers have a higher stability than Jun/Jun homodimers [22]. Besides phosphorylating these AP-1 transcription factors, JNK additionally phosphorylates a number of other targets including nonAP-1 transcription factors, nuclear hormone receptors, scaffold proteins, ubiquitin ligases, kinases, cytoskeletal proteins, and Bcl-2 family members (see Table 14.1) [23]. While it is likely that JNK isoforms exert distinct functions by phosphorylating a different set of targets, a systematic comparison of the ability of JNK1 and JNK2 to phosphorylate specific substrates has not been done. It has been suggested that

Table 14.1  JNK substrates Protein family/function Transcription factors

Nuclear hormone receptors

Scaffold proteins Kinases Ubiquitin ligases Cytoskeletal proteins and associated proteins Histones Bcl-2 family members

Member c-Jun, JunB, JunD, JDP2, ATF-2, Elk-1, c-Myc, p53, NFAT, Foxo4, Stat3, Stat1, Pax2, TCFb1 PPAR-g1, Glucocorticoid receptor, RXR, RARa, Nur77 JIP1, IRS-1, 14-3-3, ShcA Akt, RSK2 Itch Keratin 8, tau, SCG10, DCX H2AX Bcl-2, Bcl-xl, Mcl-1, Bim, Bid, Bad, Bmf

JNK1 and JNK2 exert different roles in the regulation of c-Jun phosphorylation and stability. JNK2 is ­preferentially bound to c-Jun under unstimulated conditions and promotes c-Jun degradation whereas JNK1 is the major c-Jun interacting kinase after stimulation [24]. However, a more recent study came to the conclusion that JNK2 is a positive regulator of c-Jun expression, and that some results in JNK1deficient fibroblasts were due to a compensatory upregulation of JNK1 expression and activity [25]. JNK1 is a more efficient kinase of the E3 ubiquitin ligase Itch than JNK2 and thus efficiently promotes the degradation of c-Jun and the antiapoptotic protein FLIPL [26, 27]. Microarray analysis in JNK1- and JNK2-deficient fibroblasts suggest that JNK1 and JNK2 share some target genes but that many target genes of JNK1 and JNK2 differ [28].

Regulation of Cell Death and Survival by JNKs JNKs exert both cytoprotective and cell-death promoting effects in response to different stimuli [29]. The seemingly opposite effects of JNK in the regulation of cell death can be attributed to temporal differences in JNK activation. Under most circumstances, JNK activation is transient and exerts cytoprotective effects [30]. The prosurvival function of JNK appears to be mediated by several mechanisms that involve the transcriptional upregulation of antiapoptotic genes by phosphorylated JunD [31] and most likely phosphorylated c-Jun [32], as well as transcriptionindependent effects such as phosphorylation of Bcl-2 that enhances its antiapoptotic function [33]. Under specific conditions such as the inactivation of NF-kB, JNK activation is sustained and contributes to the induction of cell death [30, 34, 35]. As described above, sustained activation of JNK occurs when ROS inactivate MKPs, e.g., when NF-kB is activated and ROS-scavenging enzymes such as MnSOD are ­downregulated. The crucial role of ROS in mediating ­prolonged JNK activation and cell death has been highlighted by the finding that the antioxidant butylated hydroxyanisole (BHA) prevents prolonged JNK activation and TNF-mediated cell death [14]. Sustained, but not transient JNK1 activation ­promotes TNFa-induced cell death by phosphorylating the E3 ubiquitin ligase Itch which targets the ­caspase-8

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inhibitor cFLIPL for proteasome-mediated degradation [27]. In addition to the E3 ubitiquitin ligase Itch, several other JNK targets have been suggested to promote apoptosis, predominantly members of the Bcl-2 family. Mcl-1 and Bcl-xl are antiapoptotic Bcl-2 family members that sequester proapoptotic Bcl-2 family members. Mcl-1 and Bcl-xl are inactivated or degraded following JNK-mediated phosphorylation thus contributing to increased activity of proapoptotic Bcl-2 family members and cell death [36, 37]. JNK-mediated phosphorylation of 14-3-3 promotes apoptosis in a similar fashion as 14-3-3 usually sequesters the proapoptotic Bcl-2 family member Bax and dissocates from Bax following JNKmediated phosphorylation [38]. JNK phosphorylation has been suggested to trigger the cleavage+ of the proapoptotic Bcl-2 family member Bid to a specific fragment termed “jBid,” which then acts on the mitochondria to release Smac, a cofactor that is required for caspase-8 activation [39]. JNK-dependent phosphorylation of the histone H2AX, a histone H2A variant, is essential for DNA fragmentation after UV treatment and thus contributes to an essential feature of apoptosis [40]. JNK1, and to a lesser extent JNK2 and JNK3 phosphorylate the transcription factor c-Myc at Ser-62 and Ser-71 to increase its proapoptotic activity [41].

Regulation of Proliferation by JNKs JNKs are activated under various circumstances such as development, carcinogenesis, and exposure to cytotoxic factors, genotoxic stress, or growth factors. All of these stimuli in common trigger a proliferative response that is, at least in part, mediated by the JNK signaling cascade. Fibroblasts lacking both JNK1 and JNK2, or c-Jun display decreased cellular proliferation [24, 42]. In vitro and in vivo studies in knockout and transgenic mice demonstrate an essential role of c-Jun in the transition of the G1-S phase of the cell cycle [42]. This function seems to be largely dependent on JNKdependent c-Jun phosphorylation at S63 and S73 as cells with mutant phosphorylation sites display severe defects in proliferation [43]. The JNK/c-Jun signaling cascade promotes proliferation by two different mechanisms: (1) Activation of cyclin D1- and cyclin E-dependent kinases (CDKs) and transcription factor

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E2F. (2) Repression of the tumor suppressor gene p53 and its target gene, the CDK inhibitor p21 [42]. JNK1 appears to mediate the majority of proliferation as JNK1-deficient fibroblast display reduced c-Jun phosphorylation and decreased proliferation rates [24]. In contrast, JNK2-deficient fibroblasts display increased c-Jun phosphorylation levels and increased proliferation [24]. The proliferation defect in c-Jun-deficient fibroblasts is similar in extent as in JNK1-/JNK2double deficient fibroblasts suggesting that c-Jun is the major target that promotes proliferation in response to JNK activation [24].

Regulation of Inflammation by JNKs The AP-1 transcription factor is known to regulate several inflammatory genes including TNFa [44]. Inhibition or activation of the JNK pathway blocks or increases TNFa release in macrophages [45]. Pharmacological inhibition of JNK by the small molecule inhibitor SP600125 has unveiled a number of inflammatory target genes including TNFa, MMP13, RANTES and MCP-1 [46–49]. Thus, JNK appears to contribute to the induction of proinflammatory gene transcription by AP-1. In addition, JNKs are involved in T-helper cell differentiation [50].

JNKs in Liver Disease TNF-Mediated Liver Injury Tumor necrosis factor is a well-characterized inducer of hepatocyte cell death in vitro and in vivo, and has been implied in the pathophysiology of various hepatic diseases (see Chap. 10). Under normal circumstances, TNFa does not induce death in hepatocytes but requires inhibition of NF-kB, or inhibition of transcription or translation by actinomycin D, cycloheximide, or galactosamine. Interestingly, these treatments lead to a prolonged activation of JNK in hepatocytes, and prolonged JNK activation is required for its cell death promoting effects [14, 51, 52]. In contrast to TNFa, the TNF receptor family member Fas, does not strongly activate JNK in hepatocytes and does not require JNK activity for the induction of hepatocyte death in vitro or in vivo

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[52, 53]. Although JNK induces AP-1-dependent transcription by phosphorylating and transactivating its classical targets c-Jun, ATF2 and JunD, the mechanism by which JNK enhances TNF-induced apoptosis in hepatocytes is independent of c-Jun and transcription, and involves prolonged JNK activation and JNK targets upstream of the mitochondria [51, 52]. Prolonged activation of JNK in TNF-stimulated hepatocytes requires the absence of NF-kB or transcription, and is achieved by the increased levels of ROS in the absence of NF-kB which directly inactivate MKP phosphatases (see Fig.  14.2) [14]. The important role of ROS in mediating prolonged JNK activation and cell death in the liver is emphasized by the finding that the antioxidant BHA prevents prolonged JNK activation and liver injury after administration of concanavalin A [14], a model of T-cell-mediated hepatitis in which liver injury depends on TNFa [27]. A recent study demonstrated that JNK phosphorylates and activates the E3 ubiquitin ligase Itch, which then ubiquitinates c-FLIPL, a wellcharacterized inhibitor of caspase 8 activation, to

induce its proteasomal degradation [27]. Accordingly, livers from JNK1- or Itch-deficient mice showed no c-FLIPL ubiquination after ConA injection. Thus, sustained JNK activation leads to an Itch-dependent reduction in the levels of c-FLIPL allowing for the activation of upstream caspases by the activated TNF receptor complex and subsequent cell death. It is not entirely clear why prolonged activation of JNK is required for its proapoptotic effects. However, it is conceivable that c-FLIPL degradation needs to occur either at a later point of time or over an extended period to allow sufficient activation of caspases. The role of the JNK1-Itch pathway in hepatocyte cell death was confirmed in vivo by the resistance of JNK1 and Itchdeficient mice to liver injury in two models of TNFmediated liver injury, ConA injection and TNFa plus galactosamine injection [27]. Studies in mice with c-Jun deficient hepatocytes further confirm that c-Jun is not the JNK target that mediates hepatocyte death [32]. In fact, mice lacking c-Jun in hepatocytes display increased liver cell death and mortality upon ConA

TNF TNFR Casp 8

FADD TRADD TRAF2 RIP IKKKs

ASK1

ROS IKK

MKK4/7

MKPs

JNK P

P

Itch

P

P

c-Jun

Transcription of NFkB-dependent genes

c-FLIPT

P

IkB p50 p65 P P

P

U

Apoptosis

P

MnSOD

Activated NFkB p50 p65

P P

Proliferation Cytoprotection

Fig. 14.2  TNF-induced JNK activation. TNFa simultaneously activates the IKK/NF-kB and the JNK/c-Jun signaling cascades. Under normal circumstances, NF-kB upregulates ROSscavenging enzymes such as MnSOD and keeps MKPs in an activated state. Functional MKPs in turn allow only short activation of JNK which induces c-Jun mediated proliferation and

cytoprotection. However, when NF-kB is inhibited, the blockade of ROS by MnSOD is lifted. Subsequently, MKPs are inactivated and prolonged JNK activation occurs. Prolonged JNK activation is associated with phosphorylation of the E3 ubiquitin ligase Itch and degradation of FLIPL to promote apoptosis

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injection due to a reduced expression of inducible nitric oxide synthase [32]. Thus, the c-Jun/AP-1 pathway is hepatoprotective in TNF-dependent liver injury whereas JNKs exert proapoptotic roles. Moreover, the nonselective JNK inhibitor D-JNKi efficiently reduced liver injury after ConA, or LPS plus galactosamine injection [27]. Both JNK1- and JNK2-deficient mice displayed strong reductions in Con-A induced liver injury in one study suggesting that both isoforms may contribute to TNF-mediated liver injury [54]. In contrast, ablation of JNK2 but not JNK1 profoundly reduced liver injury after treatment with LPS plus galactosamine [55] and ConA plus galactosamine [56] in two other studies. Notably, the decreased liver injury in JNK2-deficient mice was associated with a significant reduction of mortality following treatment with LPS/GalN [55]. The somewhat conflicting results with regard to the relative role of JNK isoforms in TNFmediated liver injury are not entirely clear and may be attributed to different JNK-deficient mouse strains or slightly different models of liver injury. A recent study even suggests that JNK is largely responsible for TNFa secretion from hematopoetic cells and its subsequent hepatotoxic effects but that hepatocellular JNK is not required for TNF-dependent liver injury [57]. However, this study adds additional uncertainty as the authors compare JNK2-deficient mice that carry a hepatocytespecific JNK1 deletion to wild-type mice (and not JNK2-deficient mice) in most experiments, and do not present data on mice that only carry hepatocyte-­specific deletions of JNK1, JNK2, or both. Nonetheless, all of the above studies suggest a crucial role of JNKs in TNF-mediated liver injury and support the notion that pharmacological pan-JNK inhibitors may represent a powerful approach to block TNF-induced cell death in vivo. Moreover, antioxidants may be an additional therapeutic approach to reduce prolonged JNK activation and TNF-mediated liver injury.

Acetaminophen-Induced Liver Injury Overdoses of acetaminophen (AAP), an effective analgesic and antipyretic drug, are the most common cause of drug-induced liver failure in the United States [58]. While AAP is a very safe drug at therapeutic doses, overdoses of AAP lead to the generation of high amounts of the toxic metabolite N-acetyl-quinoneimine

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(NAPQI) by CYP2E1. NAPQI binds to and depletes GSH and, after GSH depletion, form adducts with thiol-containing proteins [59]. GSH depletion and formation of protein adducts are key mechanisms of AAP-induced cell death leading to increased generation of mitochondrial ROS, mitochondrial permeability transition (MPT), pore formation in the outer mitochondrial membrane by proapoptotic Bcl-2 family members and mitochondrial release of the proapototic factors cytochrome c and Smac [59]. Accumulating evidence is linking JNK activation to AAP-induced hepatocyte death and liver failure. An increase in JNK activity has been demonstrated in AAP-induced liver failure in mice and humans [53, 60]. Importantly, pharmacologic or genetic ablation of JNK activity protects mice from AAP-induced liver injury and has a striking effect on mortality [53, 60, 61]. One key upstream component of the signaling cascade that ultimately leads to JNK activation is the MAPKKK ASK1 [61]. ASK1 is usually bound to thioredoxin which inhibits its activity [11]. Following AAP, thioredoxin dissociates from ASK1 thus allowing ASK1 activation and activation of JNKs (see Fig. 14.3) [61]. As ASK1deficient mice show normal JNK activation at early time points but a strong suppression of JNK activation at late time points, ASK1 seems to be predominantly responsible for a prolonged activation of JNK [61]. Thus, the duration of JNK activation is critical for its ability to promote cell death after AAP, similar to JNK-dependent cell death triggered by other stimuli. The signaling cascade by which ASK1 activates JNK involves MKK4 and both JNK1 and JNK2. Although JNK2 appears to be the main mediator of hepatotoxicity as suggested by two studies that find significant reduction of AAP-induced liver injury in JNK2deficient mice [60, 61], there is evidence that JNK1 also makes important contributions and that inactivation of both kinases is required to inhibit AAP toxicity: (1) JNK1-deficient have display a small but significant reduction of liver injury at early time points [61]. (2) Pharmacological inhibition of JNK using pan JNK inhibitors is much more efficient than deleting one of the two JNK isoforms expressed in the liver [53, 60, 61]. The requirement to inhibit both JNK1 and JNK2 may also explain studies that did not find a reduction of liver injury in JNK1- and JNK2-deficient mice at late time points. [53]. As ASK1 deletion was also less efficient than pharmacological JNK blockade, and did not abolish JNK activation at early time

14  JNKs in liver diseases

221 AAP

NAPQI GSH ROS ASK1 - Trx dissocation

?????

ASK 1 activation

Notably, in AAP-induced liver failure in mice, delayed administration of JNK inhibitor SP600125 was more efficient than the administration of N-acetylcysteine [53], the standard treatment for AAP-induced liver failure in patients. Moreover, liver injury was reduced by SP600125 6  h after AAP, a time point at which GSH depletion already occurred [60]. Thus, JNK inhibition may represent a suitable approach in patients with AAP overdose either at late time points, i.e., when NAC is likely to be inefficient, and may also have positive effects when used in combination with NAC.

MKK4 Late prolonged activation of JNK1/JNK2

Early activation of JNK1/JNK2

?

JNK translocation to Mitochondria MPT Cell death

Fig. 14.3  JNK mediates acetaminophen-induced hepatocyte cell death. The acetaminophen (AAP) metabolite NAPQI reduces intracellular GSH levels and subsequently increases ROS which promote JNK activation through two different pathways: (1) One yet uncharacterized pathway leads to a temporary increase of JNK activation which has only little effect on hepatocyte cell death. (2) In a second pathway, ROS promote the dissociation of ASK1 from its inhibitor thioredoxin (Trx). Activated ASK1 then phosphorylates MKK4 leading to prolonged activation of JNK, and activation of a mitochondrial death pathway in hepatocytes

points, it is likely that JNKs are activated by a second ASK1-independent mechanism to contribute to  liver injury (see Fig. 14.3) [61]. The targets of JNK in AAPinduced liver injury remain to be identified. A recent study has shown that JNK translocates to mitochondria following AAP to promote the onset of the MPT [62]. The MPT is a key mechanism for ­AAP-mediated cell death as demonstrated by the ability of MPT inhibitors to prevent AAP-mediated cell death in vitro and in vivo [63, 64]. Based on the above described mouse studies, pan-JNK inhibitors will most likely be more efficient than specific JNK1 or JNK2 inhibitors  for the treatment of AAP-induced liver disease.

Ischemia-Reperfusion Injury Hepatic ischemia followed by reperfusion (I/R) is a major clinical problem during transplantation, liver resection, and circulatory shock, producing apoptosis and necrosis. TNFa, but not Fas, is a crucial mediator in hepatic reperfusion injury [65]. Inhibition of TNF signaling by TNF-antiserum or genetic inactivation of TNF-R1 ameliorates hepatic reperfusion injury and prolongs survival [65, 66]. JNK is strongly activated following ischemia-reperfusion [67, 68]. Blocking hepatic ROS production by overexpression of SOD1 almost completely prevents hepatic JNK activation and injury suggesting that ROS are a major con­ tributor of JNK activation and injury in I/R [69]. Pharmacological inhibition of JNK improves a 7-day survival after hepatic I/R from 20–40% to 60–100% [70]. JNK inhibition strongly reduces Bid degradation, mitochondrial cytochrome c release, caspase 3 activation and lipid peroxidation suggesting that JNK acts upstream of the mitochondria in I/R. In a model of orthotopic liver transplatation, JNK inhibitors show a similar efficacy in decreasing pericentral hepatocyte necrosis and nonparenchymal cell death [71]. Thus, the ROS-JNK pathway represents a promising new target for the treatment of hepatic I/R injury. JNK2 appears to be the main mediator of hepatic I/R injury as JNK2-deficient mice display decreased transaminase levels and necrosis, and a greatly increased 14-day survival [72]. Moreover, JNK2-deficient mice display a significant decrease in depolarized mitochondria confirming the hypothesis that JNK acts upstream of the mitochondria to promote injury after hepatic ischemia-reperfusion [72],

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Liver Fibrosis JNK has been shown to be an important component of several profibrogenic signaling cascades including PDGF, TGFb and angiotensin [73, 74]. Moreover, JNK plays an important role in the activation of hepatic stellate cells, a cell population that is crucially involved in the development of liver fibrosis (see Chap. 3). Inhibition of JNK signaling in hepatic stellate cells by SP600125 reduces TGFb1- and PDGF-induced migration [73]. Moreover, SP600125 and a dominant negative TAK1 decrease aSMA expression during the culture-activation of hepatic stellate cells [75]. JNK inhibitor SP600125 also abolishes proinflammatory gene expression in hepatic stellate cells [48, 49]. Thus, the JNK pathway plays an essential role in several profibrogenic pathways in hepatic stellate cells. However, the contribution of JNKs to hepatic fibrogenesis in vivo is not yet fully understood. The best evidence that JNK plays a role in fibrogenesis in vivo comes from a study that demonstrated an increase in phospho-JNK levels in hepatic stellate cells isolated from fibrotic livers [73]. Preliminary results demonstrate that pharmacological inhibition of JNK reduces TGFb-, PDGF-, and AngII-mediated hepatic stellate cell activation and proliferation in vitro and well as experimental fibrogenesis in vivo, and that JNK1 but not JNK2 is responsible for promoting fibrogenesis after bile duct ligation (RFS, unpublished data).

Liver Regeneration The liver possesses a unique ability to regenerate. Following massive hepatic injury or two-thirds partial hepatectomy, a number of growth-promoting pathways are activated to initiate several rounds of hepatocyte proliferation and restore liver mass [76]. JNK is strongly activated already 20 min after partial hepatectomy [77]. Blocking JNK activity by SP600125 inhibits hepatocyte proliferation in vitro and impairs liver regeneration after two-thirds partial hepatectomy [78]. Moreover, pharmacological JNK blockade strongly decreases the expression of cyclin D1, a crucial regulator of hepatocyte proliferation. A defect in

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hepatocyte proliferation and liver regeneration is also observed in mice lacking c-Jun in hepatocytes [79]. However, mice that express c-Jun lacking the JNK phosphorylation sites S63 and S73 ­display normal liver regeneration [79]. Thus, it appears that JNK does not mediate its effect on liver regeneration exclusively through c-Jun but through other targets. However, results on liver regeneration in mice expressing mutant c-Jun require cautious interpretation as mutant c-Jun may accumulate upon stress challenges [17]. Thus, the effect of JNK-mediated c-Jun phosphorylation sites might be masked by increased levels of c-Jun and be underestimated in this mouse model. JNK1 but not JNK2 is the isoform that promotes proliferation after partial hepatectomy [80, 81]. Gadd45b is a repressor of JNK activity and strongly induced after partial hepatectomy. Deficiency of Gadd45b leads to a hyperactivation of JNK which suppresses liver regeneration, probably due to the induction of cell death after prolonged JNK activation as suggested by the increased ALT levels in Gadd45b-deficient mice [82].

Hepatocellular Carcinoma c-Jun was originally characterized as a proto-oncogene [83], and the JNK/c-Jun pathway is known to be involved in oncogenic transformation by HA-Ras [20] suggesting a potential involvement in carcinogenesis. Moreover, JNK is known to drive cyclin D1 expression and proliferation in hepatocytes [78]. Both c-Jun and JNK indeed contribute to diethylnitrosamine (DEN)-induced hepatocellular cancer [84, 85]. Notably, an activation of JNK1 but not JNK2 is observed in a large percentage of human HCCs [80]. The mechanisms by which c-Jun and JNK promote HCC seem to be entirely different. c-Jun protects tumor cells from undergoing apoptosis as demonstrated by a 5-fold increase in apoptosis in c-Jun-deficient tumors in comparison to wild-type tumors [84]. This protective effect of c-Jun is mediated in early stages of tumor development following tumor initiation, and depends on the ability of c-Jun to antagonize p53 activity [84]. Remarkably, c-Jun-deficient livers do not show a lower proliferation rate. In contrast,

14  JNKs in liver diseases Fig. 14.4  JNK and c-Jun promote hepatocarcinogenesis through different mechanisms. JNKs and c-Jun promote hepatocarcinogenesis at differ­ent stages. During tumor initiation, JNK is activated in hepatocytes to promote cell death which triggers signals in Kupffer cells that promote compensatory proliferation and tumorigenesis. In the promotion phase, JNK additionally induces tumor cell proliferation. c-Jun on the other hand, prevents cell death in tumor cells independently of JNK to promote tum­origenesis

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I. Initiation Compensalory Proliferation

TNF, IL-6, HGF

II. Promotion

DEN

BHA D-JNKi

ROS JNK1

JNK1

c-Jun

Hepatocyte

IKKβ Proliferation Kupffer cell

JNK1-deficient mice display a strong reduction in tumor proliferation as well as a defect in tumor initiation after DEN treatment [85]. JNK1-deficient mice display a decrease in transaminase ­levels, necrosis, hepatomitogens, and compensatory proliferation [85]. Thus, JNK must act through a target  distinct from c-Jun to promote hepatocyte cell death and compensatory proliferation during tumor initiation, and HCC proliferation at later stages (see Fig. 14.4). Interestingly, knock-in mice that express a c-Jun lacking the JNK phosphorylation sites Ser63 and Ser73 do not have a defect in DEN-induced tumor formation [84]. Based on the results in these knock-in mice and strikingly different results in c-Jun- and JNK1-deficient mice, one has to conclude that JNK does not mediate its tumor-promoting effects in the liver through c-Jun. When JNK activation is increased, e.g., after deletion of IKKb or p38a in hepatocytes, DEN tumor formation is increased [17, 18, 86] further underlining the crucial role of the JNK pathway in liver carcinogenesis. ROS appear to be a main stimulus for prolonged JNK activation during DEN-induced hepatocarcinogenesis. The prolonged JNK activation of mice with a hepatocyte-specific deletion of IKKb was prevented by a diet containing the antioxidant BHA [86]. Moreover, this BHA containing diet also strongly reduced the number of hepatic tumors after DEN [86]. A recent study suggested that JNK phosphorylates c-Myc to repress the cell-cycle inhibitor p21 and increase proliferation during hepatocarcinogenesis [80]. In summary, JNK and c-Jun represent promising

Apoptosis

Protection from Apoptosis

targets in hepatocarcinogenesis. Decreased JNK activation in hepatocarcinogenesis may not only be achieved by specific pharmacologic inhibitors but also by antioxidants. Moreover, it should be established whether simultaneous inhibition of JNK and c-Jun results in additive anti-tumor effects.

Fatty Liver and Steatohepatitis The JNK pathway is activated in genetically obese ob/ ob mice as well as in mice fed a methionine-choline deficient (MCD) or a high fat diet [87–89]. Probiotics decrease hepatic JNK activity in ob/ob mice suggesting that JNK activation is mediated by products from the intestinal microbiota. Deletion of JNK1 but not JNK2 decreases JNK activity and the development of steatohepatitis in the MCD diet model [88]. In this model, JNK1-deficient mice display significantly reduced levels of hepatic triglycerides, inflammation, lipid peroxidation, liver injury, and apoptosis compared to the wild-type and JNK2-deficient mice [88]. In a high fat model of steatohepatitis, JNK1-deficient mice display a strong reduction of phospho-JNK levels and failed to develop excessive weight gain, insulin resistance, or steatohepatitis [89]. In contrast, JNK2deficient mice show only a reduction of the p54 but not the p46 isoform of JNK. Moreover, JNK2-deficient mice remain obese and insulin resistant and display increased liver injury [89]. Notably, established

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steatohepatitis and the associated insulin resistance are decreased by JNK1 knockdown whereas knockdown of JNK2 markedly increases liver injury [89]. The crucial role of JNK1 in steatohepatitis is further confirmed by a preliminary study that observed decreased progression from steatosis to steatohepatitis and liver fibrosis in JNK1-deficient mice [90]. In conclusion, JNK1 promotes the development of steatohepatitis whereas JNK2 exerts cytoprotective effects in fatty liver disease. Targeting JNK1 may be a useful strategy to prevent or reverse the development of steatohepatitis and hepatic fibrosis.

Conclusion and Outlook JNKs have emerged as crucial regulators of proliferation, cell death, inflammation and metabolism in the liver. Genetic and pharmacological approaches have clearly demonstrated that targeting the JNK pathway is beneficial in diseases such as AAP poisoning, hepatocellular cancer, fatty liver, and TNF-mediated liver disease. However, many questions regarding the possible use of pharmacological JNK inhibitors remain to be answered as most studies have been undertaken in animal models. Given the vast array of biological functions of JNK in many different organs including the brain, one has to be concerned about potential side effects of JNK inhibition, especially in diseases that would require long-term treatment such as hepatocellular cancer and fatty liver. Thus, the first step toward employing JNK inhibitors as pharmacological tools in clinical hepatology would probably be to establish their efficacy in diseases that would require only shortterm treatment such as AAP intoxi­cation and acute hepatitis. If the long-term safety of JNK inhibitors can be established, or drugs with a high ­first-pass effect that selectively target the liver can be ­developed, one could also envision JNK inhibition as a preventative approach for chronic hepatitis, hepatic fibrosis, and hepatocellular carcinoma. Antioxidants may represent a promising therapeutic alternative to specific JNK inhibitors, and exert beneficial effects in TNF-mediated liver disease or hepatocellular carcinoma through their ability to block prolonged JNK activation.

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Summary

›› JNK1

and JNK2 are ubiquitously expressed, JNK3 is largely restricted to neuronal cells Activation of JNKs is achieved through a phosphorylation cascade involving MAPK kinases and scaffold proteins. In addition to c-Jun, JNKs phosphorylate a wide range of other targets comprising nuclear hormone receptors, scaffold proteins, kinases, ubiquitin ligases, Bcl-2 family members. Under most circumstances, JNK activation is transient and cytoprotective. Under specific conditions such as inactivation of NF-kB and unchecked ROS production, JNK activation is sustained and contributes to cell death. The JNK/c-Jun signaling cascade promotes proliferation by two different mechanisms (1) activation of cyclin D1 and cyclin E-dependent kinases (2) repression of the tumor suppressor gene p53. c-Jun contributes to the expression of several inflammatory AP-1 regulated cytokines such s TNFa, MCP-1. Pan-JNK inhibition protects mice from acetaminopheninduced liver injury. JNK1 promotes the development of steatohepatitis, and hepatocellular cancer. JNK2 is an important mediator of TNFand hepatic ischemia/reperfusion-induced injury. JNKs are important components of several profibrogenic signaling cascades.

Multiple Choice Questions 1. Which of the following stimuli activates JNKs: (a) UV irradiation (b) ER stress (c) TNFa (d) Nonmethylated CpG-DNA (e) All of the above 2. Which one of the following statements is correct: (a) Maximal activation of JNK requires dual phos­ phorylation (b) MKK7 deficiency prevents JNK activation in response to TNFa

14  JNKs in liver diseases

(c) MKK4 deficiency impairs JNK activation by only 50% (d) Reactive oxygen species may activate JNK via inhibition of MAP kinase phosphatases (e) All the above statements are true 3. Regarding regulation of JNK activity by other pathways, which statement is true: (a) Inactivation of the IKK/NF-kB cascade prevents JNK activation (b) P38 inactivation prevents JNK activation (c) ROS decrease activation of JNK (d) Dissociation of ASK1 from thioredoxin promotes JNKs activation (e) Probiotics stimulate JNK activity in the liver 4. Which one of the following statements is wrong? (a) JNKs phosphorylate transcriptions factors including c-Jun and JunD (b) JNKs phosphorylate Scaffold proteins such as JIP1 and IRS-1 (c) JNKs phosphorylate caspases including caspase 3, 8 and 9 (d) JNKs phosphorylate Bcl-2 family members such as Bcl-2 and Bcl-xl (e) JNKs phosphorylate nuclear hormone receptors such as RXR and PPAR-g1 5. The mechanism of JNK-induced apoptosis in hepatocytes involves (a) Phosphorylation of Itch (b) Activation of an E3 ubiquitin ligase (c) Activation of caspase 8 (d) Degradation of c-FLIPL (e) All of the above Acknowledgements  This work was supported by NIH grants R01DK076920 and U54CA126513.

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R. Schwabe 41. Noguchi K, Kitanaka C, Yamana H et al (1999) Regulation of c-Myc through phosphorylation at Ser-62 and Ser-71 by c-Jun N-terminal kinase. J Biol Chem 274:32580–32587 42. Schreiber M, Kolbus A, Piu F et al (1999) Control of cell cycle progression by c-Jun is p53 dependent. Genes Dev 13:607–619 43. Behrens A, Sibilia M, Wagner EF (1999) Amino-terminal phosphorylation of c-Jun regulates stress-induced apoptosis and cellular proliferation. Nat Genet 21:326–329 44. Wagner EF, Eferl R (2005) Fos/AP-1 proteins in bone and the immune system. Immunol Rev 208:126–140 45. Ishizuka T, Terada N, Gerwins P et al (1997) Mast cell tumor necrosis factor alpha production is regulated by MEK kinases. Proc Natl Acad Sci U S A 94:6358–6363 46. Bennett BL, Sasaki DT, Murray BW et al (2001) SP600125, an anthrapyrazolone inhibitor of Jun N-terminal kinase. Proc Natl Acad Sci U S A 98:13681–13686 47. Han Z, Boyle DL, Chang L et al (2001) c-Jun N-terminal kinase is required for metalloproteinase expression and joint destruction in inflammatory arthritis. J Clin Invest 108:73–81 48. Schwabe RF, Bataller R, Brenner DA (2003) Human hepatic stellate cells express CCR5 and RANTES to induce proliferation and migration. Am J Physiol Gastrointest Liver Physiol 285:G949–G958 49. Marra F, Delogu W, Petrai I et al (2004) Differential requirement of members of the MAPK family for CCL2 expression by hepatic stellate cells. Am J Physiol Gastrointest Liver Physiol 287:G18–G26 50. Dong C, Yang DD, Tournier C et al (2000) JNK is required for effector T-cell function but not for T-cell activation. Nature 405:91–94 51. Liu H, Lo CR, Czaja MJ (2002) NF-kappaB inhibition sensitizes hepatocytes to TNF-induced apoptosis through a sustained activation of JNK and c-Jun. Hepatology 35: 772–778 52. Schwabe RF, Uchinami H, Qian T et al (2004) Differential requirement for c-Jun NH2-terminal kinase in TNFalphaand Fas-mediated apoptosis in hepatocytes. FASEB J 18: 720–722 53. Henderson NC, Pollock KJ, Frew J et al (2007) Critical role of c-jun (NH2) terminal kinase in paracetamol-induced acute liver failure. Gut 56:982–990 54. Maeda S, Chang L, Li ZW et al (2003) IKKbeta is required for prevention of apoptosis mediated by cell-bound but not by circulating TNFalpha. Immunity 19:725–737 55. Wang Y, Singh R, Lefkowitch JH et al (2006) Tumor necrosis factor-induced toxic liver injury results from JNK2dependent activation of caspase-8 and the mitochondrial death pathway. J Biol Chem 281:15258–15267 56. Ni HM, Chen X, Ding WX et al (2008) Differential roles of JNK in ConA/GalN and ConA-induced liver injury in mice. Am J Pathol 173:962–972 57. Das M, Sabio G, Jiang F et al (2009) Induction of hepatitis by JNK-mediated expression of TNF-alpha. Cell 136:249–260 58. Lee WM (2003) Acute liver failure in the United States. Semin Liver Dis 23:217–226 59. Jaeschke H, Bajt ML (2006) Intracellular signaling mechanisms of acetaminophen-induced liver cell death. Toxicol Sci 89:31–41 60. Gunawan BK, Liu ZX, Han D et al (2006) c-Jun N-terminal kinase plays a major role in murine acetaminophen hepatotoxicity. Gastroenterology 131:165–178

14  JNKs in liver diseases 61. Nakagawa H, Maeda S, Hikiba Y et al (2008) Deletion of apoptosis signal-regulating kinase 1 attenuates acetaminophen­induced liver injury by inhibiting c-Jun N-terminal kinase activation. Gastroenterology 135:1311–1321 62. Hanawa N, Shinohara M, Saberi B et al (2008) Role of JNK translocation to mitochondria leading to inhibition of mitochondria bioenergetics in acetaminophen-induced liver injury. J Biol Chem 283:13565–13577 63. Kon K, Kim JS, Jaeschke H et al (2004) Mitochondrial permeability transition in acetaminophen-induced necrosis and apoptosis of cultured mouse hepatocytes. Hepatology 40: 1170–1179 64. Beales D, McLean AE (1996) Protection in the late stages of paracetamol-induced liver cell injury with fructose, cyslosporin A and trifluoperazine. Toxicology 107:201–208 65. Rudiger HA, Clavien PA (2002) Tumor necrosis factor alpha, but not Fas, mediates hepatocellular apoptosis in the murine ischemic liver. Gastroenterology 122:202–210 66. Colletti LM, Remick DG, Burtch GD et al (1990) Role of tumor necrosis factor-alpha in the pathophysiologic alterations after hepatic ischemia/reperfusion injury in the rat. J Clin Invest 85:1936–1943 67. Bradham CA, Stachlewitz RF, Gao W et al (1997) Reper­ fusion after liver transplantation in rats differentially activates the mitogen-activated protein kinases. Hepatology 25: 1128–1135 68. Zwacka RM, Zhang Y, Zhou W et al (1998) Ischemia/reperfusion injury in the liver of BALB/c mice activates AP-1 and nuclear factor kappaB independently of IkappaB degradation. Hepatology 28:1022–1030 69. Lehmann TG, Wheeler MD, Schwabe RF et al (2000) Gene delivery of Cu/Zn-superoxide dismutase improves graft function after transplantation of fatty livers in the rat. Hepatology 32:1255–1264 70. Uehara T, Bennett B, Sakata ST et al (2005) JNK mediates hepatic ischemia reperfusion injury. J Hepatol 42:850–859 71. Uehara T, Xi Peng X, Bennett B et al (2004) c-Jun N-terminal kinase mediates hepatic injury after rat liver transplantation. Transplantation 78:324–332 72. Theruvath TP, Snoddy MC, Zhong Z et al (2008) Mito­ chondrial permeability transition in liver ischemia and reperfusion: role of c-Jun N-terminal kinase 2. Transplantation 85:1500–1504 73. Yoshida K, Matsuzaki K, Mori S et al (2005) Transforming growth factor-beta and platelet-derived growth factor signal via c-Jun N-terminal kinase-dependent Smad2/3 phosphorylation in rat hepatic stellate cells after acute liver injury. Am J Pathol 166:1029–1039 74. Bataller R, Schwabe RF, Choi YH et al (2003) NADPH oxidase signal transduces angiotensin II in hepatic stellate cells and is critical in hepatic fibrosis. J Clin Invest 112: 1383–1394

227 75. Schnabl B, Bradham CA, Bennett BL et al (2001) TAK1/ JNK and p38 have opposite effects on rat hepatic stellate cells. Hepatology 34:953–963 76. Taub R (2004) Liver regeneration: from myth to mechanism. Nat Rev Mol Cell Biol 5:836–847 77. Westwick JK, Weitzel C, Leffert HL et al (1995) Activation of Jun kinase is an early event in hepatic regeneration. J Clin Invest 95:803–810 78. Schwabe RF, Bradham CA, Uehara T et al (2003) c-Jun-Nterminal kinase drives cyclin D1 expression and proliferation during liver regeneration. Hepatology 37:824–832 79. Behrens A, Sibilia M, David JP et al (2002) Impaired postnatal hepatocyte proliferation and liver regeneration in mice lacking c-jun in the liver. EMBO J 21:1782–1790 80. Hui L, Zatloukal K, Scheuch H et al (2008) Proliferation of human HCC cells and chemically induced mouse liver cancers requires JNK1-dependent p21 downregulation. J Clin Invest 118:3943–3953 81. Sabapathy K, Wagner EF (2004) JNK2: a negative regulator of cellular proliferation. Cell Cycle 3:1520–1523 82. Papa S, Zazzeroni F, Fu YX et al (2008) Gadd45beta promotes hepatocyte survival during liver regeneration in mice  by modulating JNK signaling. J Clin Invest 118: 1911–1923 83. Bohmann D, Bos TJ, Admon A et al (1987) Human protooncogene c-jun encodes a DNA binding protein with structural and functional properties of transcription factor AP-1. Science 238:1386–1392 84. Eferl R, Ricci R, Kenner L et al (2003) Liver tumor development. c-Jun antagonizes the proapoptotic activity of p53. Cell 112:181–192 85. Sakurai T, Maeda S, Chang L et al (2006) Loss of hepatic NF-kappa B activity enhances chemical hepatocarcinogenesis through sustained c-Jun N-terminal kinase 1 activation. Proc Natl Acad Sci U S A 103:10544–10551 86. Maeda S, Kamata H, Luo JL et al (2005) IKKbeta couples hepatocyte death to cytokine-driven compensatory proliferation that promotes chemical hepatocarcinogenesis. Cell 121:977–990 87. Li Z, Yang S, Lin H et al (2003) Probiotics and antibodies to TNF inhibit inflammatory activity and improve nonalcoholic fatty liver disease. Hepatology 37:343–350 88. Schattenberg JM, Singh R, Wang Y et al (2006) JNK1 but not JNK2 promotes the development of steatohepatitis in mice. Hepatology 43:163–172 89. Singh R, Wang Y, Xiang Y et al (2009) Differential effects of JNK1 and JNK2 inhibition on murine steatohepatitis and insulin resistance. Hepatology 49:87–96 90. Kodama Y, Kisseleva T, Miura K et al (2008) JNK1 in hematopoietic cells mediates progression from diet-induced hepatic steatosis to steatohepatits and liver fibrosis. Hepa­ tology 48:366A

15

Insulin Pathway Miran Kim and Jack R. Wands

Introduction Insulin is the principal hormone controlling blood glucose levels. Insulin stimulates the uptake of glucose, and amino and fatty acids into cells, and increases the expression and/or activity of enzymes that enhance glycogen, lipid and protein synthesis, while inhibiting the activity or expression of those enzymes that catalyze the degradation of glycogen [1]. The increase in circulating insulin levels stimulates glucose transport into peripheral tissues and inhibits hepatic gluconeogenesis. Decreased secretion of insulin coupled with tissue resistance results in type 2 diabetes and is also associated with central obesity, hypertension, polycystic ovarian syndrome, dyslipidemia, and atherosclerosis. In addition, insulin has a role as a hepatotrophic factor and promotes hepatocyte proliferation, although the mechanisms by which it stimulates liver growth are not completely understood. At the cellular level, insulin action is characterized by diverse effects, including changes in vesicle trafficking, stimulation of protein kinases and phosphatases, promotion of cellular growth and differentiation, as well as activation or repression of gene transcription [2, 3]. The stimulation of the insulin/insulin receptor substrate-1 (IRS-1) system activates a number of intracellular signaling cascades that ultimately lead to important downstream biologic effects critical for cell function (Fig. 15.1). This complexity of cellular actions implies that insulin stimulation must involve multiple signaling pathways that

J. R. Wands () The Liver Research Center, Alpert Medical School of Brown University, 55 Claverick Street, 4th Floor, Providence RI 02903, USA e-mail: [email protected]

diverge at or near the activation of receptor tyrosine kinase. Indeed, it is likely that even individual effects of the hormone require the activities of multiple signaling cascades. Although understanding of the signal transduction pathways that underlie insulin’s major physiologic effects is still incomplete, remarkable advances have occurred in the last decade. It is now clear that activation of insulin receptor tyrosine kinase, acting through the insulin receptor substrate (IRS) proteins as multisite docking molecules, creates binding sites that enable the IRSs to recruit and activate multiple, independent intracellular signal generators [4]. In this chapter, we discuss some of the known structural and functional features of the insulin receptor and IRS proteins and focus on recent advances in the understanding of the role of IRS proteins in insulin signaling

Insulin receptor

Activates downstream signaling, leading to diverse effects on:

Glucose transport

Glycoden synthesis

Lipid metabolism

Protein synthesis

Gene expression Cell growth

Mitogenesis Cell survival

Fig.  15.1  Insulin produces diverse biological effects on cells through the insulin receptor and downstream signal transduction cascades

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_15, © Springer-Verlag Berlin Heidelberg 2010

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effects. We will summarize the evidence regarding the potential role of IRS-1 in the pathogenesis of hepatocellular carcinoma (HCC) and explore insulin action on hepatocyte proliferation and liver development in the setting of chronic ethanol abuse.

either IRA or IRB was associated with an intermediate apoptotic phenotype. These observations suggest that the balance between IRA and IRB receptors may trigger ligand-independent apoptosis in hepatocytes and regulate liver development [12].

Insulin Receptor

IRS Proteins

The insulin receptor (IR), a tetrameric glycoprotein composed of two a- and b-subunits, is highly expressed in adipocytes and hepatocytes. The a-subunit comprises the extracellular domain and contains the ligand binding site(s), whereas the intracellular portion of the b-subunit has tyrosine kinase activity. The unoccupied a-subunit inhibits the tyrosine kinase activity of the b-subunit and the removal of the a-subunits by deletional mutagenesis reverses this inhibition. Tyrosyl autophosphorylation after receptor–ligand interaction is a key mechanism that activates insulin signaling pathways. These cascades transmit the insulin signal by promoting binding to Src homology 2 (SH2) or phosphotyrosine-binding (PTB) domains of downstream signaling molecules such as IRS-1, IRS-2, and SH2 domain-containing (Shc) proteins, and growth factor receptor-bound protein 10 (Grb10) and Grb14 [5–8]. Thus, it is clear that the intrinsic tyrosine kinase activity of the IR is essential for insulin action. Naturally occurring mutations of the IR in humans may cause partial inhibition of tyrosine kinase activity and are associated with severe insulin resistance. Without IRs, mice die shortly after birth, while humans survive for a short time with severe growth retardation and diabetes [9]. Liver-specific IR knockout mice developed secondary hyperinsulinemia from a combination of increased b-cell mass and decreased insulin clearance by the liver; these events promoted progressive liver dysfunction including focal dysplasia and hyperplastic nodules. In addition, albumin, triglycerides, and free fatty acid levels in serum were reduced as well [10, 11]. Recently, the role of IR isoforms A and B were demonstrated using SV40-immortalized neonatal hepatocytes. In this study, an IR deficiency in hepatocytes produced acceleration of caspase-3 activation, DNA laddering, and cell death. Coexpression of IRA and IRB subunits in IR-deficient hepatocytes rescued such cells from apoptosis. The expression of

Overview The IRS proteins, as IR-specific docking proteins, carry out various functions downstream of insulin (and IGF) receptors by (1) providing a juxtamembrane localization signal for PIP3 generation, (2) amplifying the signal engendered by receptor autophosphorylation, and (3) engage a panoply of substrates that account for the diverse actions of insulin [13]. These proteins contain several common structural features: (1) an N-terminal pleckstrin homology (PH) and/or PTB domains that mediates protein–lipid or protein–protein interactions; (2) multiple C-terminal tyrosine residues that create SH2-protein binding sites; (3) proline-rich regions to engage SH3 or WW domains; and (4) serine/threonine-rich regions, which may regulate overall IRS function through other protein­–protein interactions [14]. Such functional domains amplify receptor signals by directly recruiting SH2 proteins to their phosphorylation sites. These adaptor proteins also dissociate the intracellular signaling complex from endocytic pathways that are involved in the recycling of the IR. This property may be ­especially important for insulin-stimulated biological effects such as glucose uptake.

Members of the IRS Protein Family Six IRS proteins have been identified that differ with respect to tissue distribution, subcellular localization, developmental expression, binding to the IR, and interaction with SH2 domain-containing proteins. Although the IRS proteins are highly homologous, they serve complementary roles in insulin signaling rather than representing redundant molecules, as

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231

Table 15.1  Summary of IRS function as determined by knockout mouse models Gene Phenotype IRS-1

IRS-2

IRS-3 IRS-4 IRS-1/IRS-3

Significant growth inhibition Mild insulin resistance, glucose intolerance does not develop because of compensatory hyperinsulinemia Female: increased longevity Reduction of mucosal mass in colon Promote mammary tumor metastasis No insulin resistance during fasting, but after refeeding (liver-specific KO) Insulin resistance in muscle and liver, abnormal b-cell development, type 2 diabetes Males develop dehydration, hyperosmolar coma and death Upregulation of IRS-1 expression, increase IGF-1R signaling Reduced life span Insulin resistance during fasting, but not after refeeding (liver-specific KO) Normal growth, and glucose tolerance Mild defects in growth in male mice Mild defects in reproduction, slight impairment in glucose homeostasis Lipoatrophic diabetes

shown by the studies in knockout mice (Table 15.1). For example, IRS-1 and IRS-2 are the best ­characterized members of this family and are widely expressed in muscle, liver, fat, and pancreatic islet cells [8, 15]. In  this regard, IRS-1 null (-/-) mice are stunted in growth but do not develop diabetes because an alternate substrate such as IRS-2 (pp190) compensates for the lack of IRS-1 in the liver [16–19]. IRS-1 transgenic (Tg) mice showed glucose tolerance and significantly enlarged epididymal fat mass, as well as elevated serum TNF-a concentrations [20]. In contrast, IRS-2 -/- mice develop peripheral insulin resistance in the liver and skeletal muscle and b-cell failure and lose their ability to regulate glucose homeostasis [21]. Selman et al. performed longevity studies in both IRS-1 and IRS-2 knockout (KO) mice. Female IRS-1 KO mice displayed statistically increased longevity (~30%) relative to controls, whereas male and female IRS-2 KO mice had reduced life spans [22]. In addition, conditional knockouts of liver IRS-1 and IRS-2 genes were developed and demonstrated that the two proteins play overlapping roles in insulin action, with IRS-1 acting in the postprandial state and IRS-2 in the fasted state; thus, the two substrates were able to compensate for each another [23, 24], indicating that the specificity of insulin signaling arises from downstream interactions of IRS proteins. IRS-3 gene expression is restricted to adipose tissue and b-cells in rodents and has not yet been identified in the human genome [25]. IRS-2/IRS-3 double KO mice revealed a similar degree of glucose intolerance and insulin resistance to

Reference [9, 16, 17, 24, 47, 121–123]

[21, 24, 122, 124–126]

[28, 127] [28, 29] [128]

IRS-2 KO mice [26]. The IRS-4 gene is expressed predominantly in brain, thymus, and kidney, where it may bind to and transmit signals via phosphoinositide 3-kinase (PI3K) and growth factor receptor-bound protein 2 (Grb2)-mediated cascades. In rat liver membranes, insulin and angiotensin II modulated IRS-4 tyrosine phosphorylation in a PI3K-dependent manner [27]. Knockout of IRS-3 or IRS-4 genes display mild metabolic and growth phenotypes [28]. The IRS-4 KO mice appear normal with the exception of reduced fertility [29]. Finally, IRS-5 and IRS-6 are most abundantly expressed in kidney, liver, and skeletal muscle, respectively [30].

The IRS-1 and Hepatocellular Carcinoma Human (h) IRS-1 was cloned from an overexpressing HCC cell line and serves as the prototype docking protein for the IR. It was initially detected in insulin-­ stimulated Fao hepatoma cells by immunoprecipitation with anti-phosphotyrosine antibody [31]. The IRS-1 protein has a calculated molecular mass of 132 kDa, but owing to extensive phosphorylation it migrates at 185 kDa on sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) [32]. More important, IRS-1 is tyrosyl phosphorylated by insulin receptor tyrosine kinase activity [33]. Tyrosyl-phosphorylated IRS-1 transduces various growth and metabolic signals through interaction with downstream SH2-containing

232

molecules that bind to specific IRS-1 motifs, namely the p85 subunit of PI3K [34, 35], Grb2 [36], SH2 domaincontaining protein tyrosine phosphatase-2 (SHP2 or Syp) [37], and phospholipase Cg (PLCg) [38]. There is evidence to suggest that hIRS-1 may have transforming properties as well as play a prominent role in normal hepatic growth. The hIRS-1 protein found in hepatocytes is highly overexpressed in multiple HCC cell lines and clinical tumor samples. This observation suggests that hIRS-1 may function as a signal transduction molecule during the molecular pathogenesis of HCC [32, 33, 39–41]. Thus, highly expressed and phosphorylated hIRS-1 may enhance intracellular growth signals and contribute to the multistep process of hepatic oncogenesis. Direct evidence for this concept was provided by the construction of a dominant-negative mutant that interfered with endogenous hIRS-1 tyrosyl phosphorylation. The C-terminal truncated hIRS-1 molecule (dominant-negative mutant) inhibited tyrosyl phosphorylation of endogenous hIRS-1 and Shc proteins. Subsequently, the activity of downstream signaling molecules such as PI3K and mitogen-activated protein kinase (MAPK) were inhibited. More important, stable transfection of this dominant-negative mutant into HCC cells reversed the malignant phenotype as characterized by inhibition of transformed foci formation, loss of anchorage-­ independent growth in soft agar, inability to form tumors in nude mice, and strikingly reduced cell proliferate activity [42]. The Grb2 and SHP2 proteins also contributed to the cellular transforming activity of hIRS-1. Stable transfection and overexpression of the hIRS-1 gene in NIH 3T3 cells lead to increased hIRS-1 tyrosyl phosphorylation, enhanced binding of Grb2 and SHP2 but not PI3K, and persistent or constitutive activation of the downstream MAPK cascade. Such transfected 3T3 cells develop a phenotype characterized by transformed foci formation, induction of anchorage-­ independent cell growth and increased cell proliferation. When such cells were injected into nude mice, they were highly tumorigenic [43]. Further studies have revealed that hIRS-1-mediated mitogenic signals are directly regulated by interaction with Grb2 and Syp, and these interactions were followed by the activation of MAPK cascade [35, 44–47]. Mutant (Y897F and Y1180F) hIRS-1 constructs reduced the intracellular interaction of IRS-1 with Grb2 and Syp proteins, respectively. Single IRS-1 mutant molecules did not completely reduce the insulin-dependent transforming

M. Kim and J. R. Wands

activity. However, a double mutant (Y897F/Y1180F) construct strikingly attenuated the transforming ­activity of hIRS-1. Therefore, hIRS-1-induced cellular transformation required an interaction with both Grb2 and Syp signal transduction molecules [42]. It is of interest that over 80% of human HCC tumors had enhanced hIRS-1 gene expression compared with adjacent noninvolved liver tissue [32, 48]. Moreover, there was a significant relationship between the level of hIRS-1 overexpression and the tumor size. The overexpressed hIRS-1 protein was found to be tyrosyl phosphorylated and interacted with PI3K, Grb2, and SHP2 proteins and there was constitutive activation of both the PI3K and MAPK signal transduction cascades. Moreover, overexpression of hIRS-1 in the liver of a transgenic mouse model led to increased hepatocyte DNA synthesis [41]. In addition, hIRS-1 ­transgenic mice revealed high levels of aspartyl-asparaginyl-bhydroxylase (AAH), while a dominant-negative (hIRS1DC) mutant reduced AAH expression, and motility and invasiveness in FOCUS HCC cells suggesting that IRS-1 regulated AAH expression, and HCC cell motility and invasion [49]. In view of the observations to support a possible role for hIRS-1 in tumor growth, there has been no evidence of HCC development in hIRS-1 transgenic mice [50]. Interestingly, chronic ethanol exposure in hIRS-1 transgenic mice demonstrated hepatic micro- and macrosteatosis, focal chronic inflammation, apoptosis and disordered lobular architecture [51]. As HCCs exhibit high degrees of genetic heterogeneity and involvement of multiple signaling pathways, other cofactors in addition to activation of the IRS-1 signal transduction cascade may be required to transform hepatocytes.

IRS and Hepatocyte Proliferation Insulin is one of the regulators of normal hepatocyte proliferation and subsequent liver growth [52, 53]. The cellular pathways responsible for transmitting the insulin-mediated signal from the cell surface to the nucleus in the context of fetal liver growth are under active investigation. Most recently, the role of IRSmediated growth cascades has been studied in rapidgrowing fetal rat liver; expression and tyrosyl phosphorylation of IRS-1 was reduced compared with the adult liver. These developmental changes resulted in a lack of sensitivity to insulin stimulation and

15  Insulin Pathway

subsequent downstream activation of the PI3K and MAPK cascades until they become functional in the postneonatal period. In contrast, a high level of IRS-2 expression and tyrosyl phosphorylation was present as early as embryonic day 15 with robust PI3K binding and activation, which may enhance hepatocyte survival during the rapid growth phase of the liver. In addition, IRS-2 was found to propagate the insulin signal via PI3K in the late-gestation fetal liver. Therefore, IRS-2 is the dominant substrate for insulin receptor kinase activity with respect to tyrosyl phosphorylation and downstream PI3K pathway activation during fetal life and may enhance hepatocyte survival signals. These investigations lead us to believe that IRS-1 may have a major role in the adult liver with respect to mediating hepatic growth via the MAPK pathway [54]. However, the IRS-1 signal transduction pathway does not play a major role in fetal liver growth because IRS-2 functions as the major insulin-responsive molecule. During liver regeneration induced by partial hepatectomy, there was tyrosyl phosphorylation of the insulin receptor b-subunit and IRS-1 followed by an association with PI3K; these events occurred prior to the onset of DNA synthesis in the late G0 phase of the cell cycle [55]. In another setting, IRS-1 protein was significantly increased in cirrhosis compared to normal liver, which may favor enhanced hepatic growth [56]. In the early stage of rat liver regeneration, IRS-1 expression was increased; a finding consistent with a stimulatory role in the regenerative process, whereas it returned to baseline levels 7 days later when the hepatic growth process was complete. The reduced IRS-1 level occurred in the setting of increased IRS-2 and IRS-4 expression. Given that 1 and 7 days after partial hepatectomy, isolated hepatocytes responded similarly to insulin in terms of cell proliferation, a compensatory role was proposed for the induction of IRS-2/4. Since IRS-4 is activated by insulin stimulation of rat hepatocytes, it seems likely that expression and tyrosyl phosphorylation of IRS-4 was a compensatory mechanism to augment liver regeneration. In support of this argument was the association of IRS-4 with PI3K, SHP2, and protein kinase Cz (PKCz), subsequently to transmit the insulin signal [57]. Moreover, IRS-4 was also involved in HepG2 HCC cell proliferation/differentiation through ERK and p70S6K activation [58]. In the liver-specific IGF-1 receptor KO murine model, hepatocye proliferation after partial hepatectomy was significantly reduced in males, but not in females. These

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animals exhibited decreased IRS-1, cyclin D1 and A induction, but not IRS-2 expression, suggesting differential regulation of IRS proteins during liver regeneration in a gender-specific manner [59].

Insulin Signaling Pathways Through IRS Proteins While more detailed information regarding the insulin signaling cascade is provided elsewhere (see Chaps. 19 and 20), we will present a synopsis of the key steps that result from insulin action (Fig. 15.2). PI3K, one of the SH2 domain-containing molecules, interacts with tyrosyl phosphorylated IRS proteins, thereby activating this enzyme [38, 60] to generate phosphatidylinositol3,4,5-trisphosphate (PIP3). Increasing PIP3 concen­ trations bring protein kinase B (PKB)/Akt into proximity  with another pleckstrin homology (PH)domain-containing protein kinase, namely phosphoinositide dependent kinase 1 (PDK1), resulting in Akt phosphorylation at residues Thr 308 and Ser 473 [21, 61]. The identification of Akt substrates has been of great interest to understand the mechanisms by which this kinase impacts cell growth and programmed cell death pathways. The glycogen synthase kinase 3b (GSK3b) protein is a ubiquitously expressed serine/ threonine protein kinase and is one of the principal Akt substrates [62, 63]. The Akt-induced phosphorylation of GSK3b results in GSK3b inactivation and leads to decreased phosphorylation and increased glycogen synthase activity [61, 64]. In addition, GSK3b overexpression elicits apoptosis that can be blocked by Aktmediated GSK3b phosphorylation [65]. Akt has also been implicated in the regulation of Ras protein-­specific guanine nucleotide-releasing factor 2 (Raf) and provides possible crosstalk between PI3K and MAPK signal transduction cascades. Other recently identified targets of Akt include Bcl2-antagonist of cell death (BAD), forkhead box protein 01A (FoxO1), forkhead box protein 03A (FoxO3A), forkhead box protein 04 (FoxO4), endothelial nitric oxide synthase (eNOS) and mammalian target of rapamycin (mTOR) [61]. Activation of the MAPK/endothelial signal-­regulated kinase (ERK) pathway is another major effector mechanism for insulin action [38]. This pathway involves the tyrosine phosphorylation of IRS proteins and Shc  [5], which interact with Grb2 thereby recruiting

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Fig. 15.2  Cartoon of the insulin-signaling pathways through the IRS proteins. Note the importance of signaling through the PI3K and MAPK/ERK cascades

IN IR Gab1 PI3K

SOCS1/3 Shc

IRS

PIP3

PTP1B

Grb2

PTEN

IRS

SHP2

Fyn

SHIP Akt PDK

PKC α/ε/δ

Sos Ras

Grb10

Spred

Akt

Raf

Bad

RKIP

PKA GSK3

Apoptosis

mTOR Glycogen synthesis

Lipolysis

p70S6K

MEK

ERK

Protein synthesis NUCLEUS

Akt

FoxO1,3A,4

ERK Growth

Apoptosis

Son-of-Sevenless (SOS) exchange protein to the plasma membrane for activation of Ras. The activation of Ras also requires the stimulation of SHP2 through its interaction with Grb2-associated binding protein 2 (Gab-1) or IRS1/2. Once activated, Ras operates as a molecular switch stimulating a serine kinase cascade through the stepwise activation of Raf, MAPK/ERK kinase 1 (MEK) and ERK. Activated ERK can translocate into the nucleus where it catalyzes the phosphorylation of transcription factors such as p62TCF important for initiating gene expression required for cellular proliferation. Block of this pathway with dominant-negative mutants or pharmacological inhibitors prevents cell growth induced by insulin signaling but has no effect on the metabolic actions of this hormone [1].

Suppressors of cytokine signaling (SOCS)1 is induced by insulin in vitro and inhibits autophosphorylation of the IR and interacts directly with IRS. There is evidence that SOCS1 targets IRS-1 and -2 for proteasomal degradation [66]. SOCS1 also binds to residues within the kinase domain of the IR that are essential for IRS-2 recognition [67]. SOCS1 knockout mice revealed increased IRS-2 expression and hepatic insulin sensitivity [68]. Protein kinase Ce (PKCe) and PKCd, are involved in  the down-regulation of insulin signaling through IRS-1. In HepG2 human HCC cells, treatment with high glucose concentrations resulted in phosphorylation of serine residues on IRS-1. The high glucose treatment attenuated the insulin-induced association of IRS-1

15  Insulin Pathway

with PI3K and downstream phosphorylation of Akt. This phenomenon was associated with the translocation of PKCe and PKCd from the cytosol to the plasma membrane in association with IRS-1. In contrast, ­insulin-induced association of Shc and Grb2 to the IR was not inhibited. Therefore, PKCe and PKCd may function as inhibitors of the insulin signaling pathway via regulating the phosphorylation of IRS-1 [69]. It has been reported that constitutive association of PKCa with IRS-1 was reversed by insulin in murine liver [70]. PKCa serves as an active inhibitor and regulates the insulin cascade through its association with IRS-1. In this context, stimulation with insulin results in PKCa disassociation from IRS-1 and opens the “gate” for transmission of the insulin signal. Then, PKCa reassociates with IRS-1 to inhibit function. Effects of insulin on PKCa are independent of PI3K. The ultimate activity of PKCa will depend on the sites of tyrosine phosphorylation under various physiologic conditions [71].

Inhibition of Insulin Signaling Insulin signaling can be inhibited by several mechanisms including phosphotyrosine dephosphorylation, serine/threonine phosphorylation, and degradation of IRS proteins. The protein-tyrosine phosphatase-1B (PTP1B) has been implicated as a negative regulator of insulin signaling. For example, PTP1B overexpression in L6 myocytes and Fao HCC cells blocked tyrosine phosphorylation of the IR and IRS-1 by more than 70% and resulted in a significant inhibition of the association between IRS-1 and PI3K. Thus, there was inhibition of downstream Akt and MAPK phosphorylation as well [72]. Reduction of PTP1B protein expression by specific antisense oligodeoxynucleotides in Fao cells also increased insulin-stimulated phosphorylation of Akt and GSK3b without any noticeable change in protein expression levels. These results demonstrate that reduction of PTP1B can modulate key insulin signaling events downstream of the IR [73]. In insulinresistant rats, the increase of PTP1B expression and interaction with the IR contributed to impaired insulin signaling in the liver [74]. In addition, serine/threonine phosphorylation of IRS proteins decreases tyrosyl phosphorylation and thereby attenuates insulin signaling as a key negative regulatory mechanism [75–77]. Thus, serine phosphorylation of IRS proteins is a

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potent factor that inhibits insulin signaling. Liver from insulin-resistant rodents showed elevated serine kinase activity for IRS-1 as demonstrated by a specific Ser789-phosphorylation [78, 79]. Other investigations revealed enhanced serine phosphorylation of IRS-1 in diabetic tissues in vivo [80–82] and GSK3b was responsible for this phosphorylation event [83, 84]. Interleukin-6 (IL-6) is one of the several proinflammatory cytokines that have been associated with insulin resistance and type 2 diabetes. IL-6 exposure reduces tyrosine phosphorylation of IRS-1 and the association with PI3K in both primary mouse hepatocytes and HepG2 cells [85]. SOCS proteins are induced by inflammation. Among them, SOCS1 or SOCS3 targets IRS-1 and IRS-2 for ubiquitin-mediated degradation and therefore blocks insulin signaling. Indeed, SOCS1 and SOCS3 were found to bind both recombinant and endogenous IRS-1 and IRS-2 protein and promote their ubiquitination and subsequent degradation in multiple cell types [66]. Interestingly, the IL-6dependent induction of insulin resistance is mediated by SOCS proteins. In mice exposed to IL-6 hepatic SOCS3, expression was increased and it was ­associated with inhibition of insulin-dependent insulin receptor autophosphorylation and IRS-1 tyrosyl phosphorylation as well. Induction of SOCS3 in liver may be an important mechanism to explain IL-6-mediated insulin resistance [86]. Moreover, ubiquitin/proteasome-mediated degradation of IRS-2, but not IRS-1 in L1 and Fao HCC cells, occurs via a PI3K/Akt-dependent pathway and is closely associated with inhibition of insulin signaling [81]. The Raf kinase inhibitor protein (RKIP), is a metastasis suppressor gene and inhibits insulin signaling in the liver. RKIP directly interacts with both Raf-1 and MEK and disrupts Raf-1/MEK binding thereby preventing the activation of MEK and downstream components of this signaling cascade. Downregu­ lation of RKIP expression was found in over 80% of human HCC, as compared to adjacent peritumoral tissues, indicating that activation of the IGF-I/ERK/MAPK pathway plays a major role during human hepatocarcinogenesis [87]. Ethanol also affects insulin signaling through inhibition of IRS-1 tyrosyl phosphorylation. Therefore, ethanol reduces the interaction between Syp and tyrosyl phosphorylated IRS-1 [88–90]. High ethanol intake is considered a major factor for impaired insulin sensitivity. Acute and chronic ethanol-exposed rats resulted in reduced tyrosyl phosphorylation of IRs, IRS-1 and IRS-2

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proteins. Chronic ethanol exposure impairs survival mechanisms in the liver because of its inhibitory effect on insulin signaling through PI3K/Akt and increases the hepatocyte levels of phosphatase and tensin homolog deleted on chromosome 10 (PTEN), a major negative regulator of the PI3K/Akt signal transduction cascade [91–94]. Thus, chronic ethanol consumption produced liver injury with increased hepatocellular steatosis, inflammation and apoptosis, which was associated with hepatic insulin resistance [95].

Insulin and Growth Hormone The insulin signaling pathway is also linked to growth hormone (GH). Excess GH is associated with secondary hyperinsulinemia through alterations of the early steps of insulin action in the liver. IRs were reduced in a transgenic mouse model that overexpressed GH whereas IR and IRS-1 phosphorylation, the IRS-1/ PI3K interaction and PI3K activity appeared to be maximally activated. Under these conditions, it was not possible to further stimulate this signal transduction cascade in vivo because of a complete insensitivity to insulin action [96]. On the other hand, GH deficiency was associated with increased tissue sensitivity to insulin. In the liver of growth hormone receptor (GHR)-KO mice, the lack of GH effects was associated with increased IR abundance and enhanced autophosphorylation following insulin binding. These alterations may represent an adaptation to the low insulin concentrations leading to or contributing to increased insulin sensitivity [97]. The antagonistic action of GH on insulin signaling is not a consequence of a direct interaction with the IR. Instead, long-term exposure to GH leads to a reduction of IR levels and an impairment of tyrosine kinase activity. The signals induced by GH and insulin may converge on downstream postreceptor proteins. Activation of PI3K appears to be an important site of convergence between the signals generated by these two hormones. Rodent models of chronic GH excess have been useful to investigate the mechanisms by which GH induces insulin resistance. Decreased IR, IRS-1, and IRS-2 tyrosyl phosphorylation in response to insulin stimulation was found in skeletal muscle, whereas a chronic activation of the IRS/PI3K pathway was found in liver. The induction of proteins that inhibit IR signaling such

M. Kim and J. R. Wands

as SOCS-1 and -6 may also be involved in GH-mediated effects. The mechanisms of insulin resistance induced by GH involve uncoupling between PI3K and its downstream signaling mediators. GH also reduces insulin sensitivity by enhancing events that negatively modulate insulin signaling such as stimulation of serine phosphorylation of IRS-1, which prevents its recruitment to the IR as well as induction of SOCS-1 and SOCS-3, which alter IRS protein function [98]. Finally, GH may modulate the life span of cells by altering insulin sensitivity [99].

Transcriptional Regulation by Insulin Insulin, after binding to its receptor, regulates many cellular processes through modulation of gene expression. In a subset of genes, insulin exerts a negative action on transcription while in others it has a positive effect. Insulin controls gene transcription by modifying the binding of transcription factors to insulin-response elements or by regulating their transcriptional activity. Several transcription factors, including FoxO, sterolregulatory element-binding protein family (SREBP) and Sp1, are influenced by insulin action [100]. Insulin changes transcription by influencing the level, localization, and activity of transcription factors. For example, FoxO proteins are major negative regulators of transcription induced by insulin [101]. Recently, Kamagate et al. reported that FoxO1 regulated excessive production of triglyceride-rich VLDL. These investigators demonstrated that microsomal triglyceride transfer protein (MTP) is a target of FoxO1 and that excessive VLDL production is caused by the inability of insulin to modulate FoxO1 transcriptional activation of MTP [102]. It has been observed that the expression and activity of FoxO1 were increased in nonalcoholic steatohepatitis [103]. The SREBP transcription factor family consists of three isoforms (SREBP-1a, -1c, and -2), which control biosynthesis of lipids and play a pivotal role in cellular sterol homeostasis. In the liver, SREBP-1c is the major isoform that regulates biosynthesis of fatty acids and triglycerides. Insulin increases transcription of the SREBP-1 gene via the activation of PI3-kinase and Akt [104–106]. Within the nucleus, SREBP activity appears to be augmented after insulin-induced phosphorylation of specific serine and threonine residues [107–109]. Overproduction of hepatic nuclear SREBP-1c in

15  Insulin Pathway

transgenic mice was associated with activation of lipogenic genes leading to fatty liver, whereas the absence of SREBP-1 (by targeted gene disruption) abolished nutritional regulation of lipogenic enzymes [110, 111]. Insulin and hyperglycemia elevate SREBP-1c expression [112–114]. FoxO1 and SREBP-1c reciprocally regulate IRS-2 expression and insulin sensitivity in the liver [115]. Sp1 is a ubiquitous transactivator that binds to GC-rich motifs in the promoters of several insulinresponsive genes [116]. Hepatic phosphoenolpyruvate carboxykinase (PEPCK) and IGFBP-1 gene transcription was inhibited by insulin through SREBP-1c and Sp1 [117, 118], while fatty acid synthase (FAS) and malic enzyme (ME) were upregulated [119, 120].

Perspectives The molecular mechanisms of insulin action on cells have been under intense investigation. In this context, efforts to understand insulin effects on tissues have led to the discovery of the IR, its primary role as a tyrosine kinase, and more importantly how this tyrosine kinase phosphorylates IRS proteins, especially IRS-1. It is now appreciated that tyrosyl phosphorylated IRS-1 acts as a docking protein. It binds to and activates several cytosolic signaling molecules important in mediating downstream growth and metabolic effects. These major accomplishments aid our understanding of the molecular mechanisms involved in the insulin-­signaling network, as exemplified in Fig. 15.2. Future efforts will need to focus on determining how the various IRS-1associated proteins mediate growth signals related to the multistep process of hepatocarcinogenesis. Other significant areas of research include defining the role of chronic ethanol consumption on phosphorylation of IRS proteins in an attempt to understand better the inhibitory effect of ethanol on liver growth. Therefore, understanding insulin action may have direct relevance to the pathogenesis of acute and chronic liver diseases as well as the development of HCC.

Multiple Choice Questions 1. Insulin receptor (IR) is a family of receptor tyrosine kinases (RTK). Tyrosyl autophosphorylation of IR

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triggered by receptor–ligand interaction is a key mechanism to activate insulin-signaling pathway. Many of these receptors for growth factors belong to RTK. Which receptor(s) is not an RTK? (a) MET (hepatocyte growth factor receptor) (b) IGF-IIR (insulin-like growth factor II receptor) (c) Insulin receptor (d) EGFR (epidermal growth factor receptor) (e) TbR (transforming growth factor beta receptor) 2. Six IRS proteins have been identified so far. Which statement is correct? (a) IRS-3 can compensate for the lack of IRS-1 in the liver (b) IRS-2 gene is expressed predominantly in brain, thymus, and kidney (c) Female IRS-1 knockout mice showed longer life span (d) IRS-4 expression is restricted to adipose tissue and b-cells (e) IRS-3 knockout mice suffer from glucose intolerance 3. The roles of IRS proteins have been defined using murine knockout models. Which IRS knockout mice exhibited normal growth and normal glucose tolerance? (a) IRS-4 (b) IRS-1/IRS-3 (c) IRS-3 (d) IRS-2 (e) IRS4 4. There are several known molecules that inhibit insulin signaling. Which one does not? (a) RKIP (raf kinase inhibitor protein) (b) PTEN (tensin homolog deleted on chromosome 10) (c) IGFBP (insulin-like growth factor binding protein) (d) SOCS1/3 (suppressors of cytokine signaling 1/3) (e) Protein-tyrosine-phosphatase-1B 5. Insulin regulates many cellular processes and the expression of several genes by modifying the binding of transcription factors. Which transcription factor is not involved in insulin signaling? (a) SREBP (sterol-regulatory element-binding protein) (b) FoxO (c) TCF (T-cell factor) (d) Sp1

238 Acknowledgment  This work was supported in part by NIH grants CA-35711 AA-02666 (JRW) and COBRE  RR-­ P20RR017695 (MK).

Selected Reading   1. Nandi A, Kitamura Y, Kahn CR et al. (2004) Mouse models of insulin resistance. Physiol Rev 84:623–647 (this review paper describes animal models including IRSs and IR knockout/transgenic mice in detail)   2. Mounier C, Poster BI (2006) Transcriptional regulation by insulin: from the receptor to the gene. Can J Physiol Pharmacol 84:713–724 (this review addresses the regulation by insulin of gene transcription in the liver)   3. Kubota N, Kubota T, Itoh S et al (2008) Dynamic functional relay between insulin receptor substrate 1 and 2 in hepatic insulin signaling during fasting and feeding. Cell Metab 8:49–64 (this article shows hepatic IRS-1 and IRS-2 function in a distinct manner in the regulation of glucose homeostasis using KO and Tg mice)   4. Longato L, de al Monte S, Califano S et al. (2008) Synergistic permalignant effects of chronic ethanol exposure and insulin receptor substrate-1 overexpression in liver. Hepatol Res 38:940–953 (this recent paper shows the effects of ethanol in IRS-1 transgenic mice)

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M. Kim and J. R. Wands 83. Liberman Z, Eldar-Finkelman H (2005) Serine 332 phosphorylation of insulin receptor substrate-1 by glycogen synthase kinase-3 attenuates insulin signaling. J Biol Chem 280: 4422–4428 84. Sharfi H, Eldar-Finkelman H (2008) Sequential phosphorylation of insulin receptor substrate-2 by glycogen synthase kinase-3 and c-Jun NH2-terminal kinase plays a role in hepatic insulin signaling. Am J Physiol Endocrinol Metab 294:E307–E315 85. Senn JJ, Klover PJ, Nowak IA et  al (2002) Interleukin-6 induces cellular insulin resistance in hepatocytes. Diabetes 51:3391–3399 86. Senn JJ, Klover PJ, Nowak IA et  al (2003) Suppressor of cytokine signaling-3 (SOCS-3), a potential mediator of interleukin-6-dependent insulin resistance in hepatocytes. J Biol Chem 278:13740–13746 87. Lee HC, Tian B, Sedivy JM et al (2006) Loss of Raf kinase inhibitor protein promotes cell proliferation and migration of human hepatoma cells. Gastroenterology 131:1208–1217 88. Bhavani K, de la Monte S, Brown NV et al (1995) Effect of ethanol on p36 protein kinase substrate and insulin receptor substrate 1 expression and tyrosyl phosphorylation in human hepatocellular carcinoma cells. Alcohol Clin Exp Res 19: 441–446 89. de la Monte SM, Ganju N, Tanaka S et al (1999) Differential effects of ethanol on insulin-signaling through the insulin receptor substrate-1. Alcohol Clin Exp Res 23:770–777 90. Xu YY, Bhavani K, Wands JR et al (1995) Ethanol inhibits insulin receptor substrate-1 tyrosine phosphorylation and insulin stimulated neuronal thread protein gene expression. Biochem J 310:125–132 91. He L, Simmen FA, Mehendale HM et  al (2006) Chronic ethanol intake impairs insulin signaling in rats by disrupting Akt association with the cell membrane. Role of TRB3 in inhibition of Akt/protein kinase B activation. J Biol Chem 281:11126–11134 92. Onishi Y, Honda M, Ogihara T et al (2003) Ethanol feeding induces insulin resistance with enhanced PI 3-kinase activation. Biochem Biophys Res Comm 303:788–794 93. Yao XH, Nyomba BL (2008) Hepatic insulin resistance induced by prenatal alcohol exposure is associated with reduced PTEN and TRB3 acetylation in adult rat offspring. Am J Physiol Regul Integr Comp Physiol 294:R1797–R1806 94. Yeon JE, Califano S, Xu J et  al (2003) Potential role of PTEN phosphatase in ethanol-impaired survival signaling in the liver. Hepatology 38:703–714 95. de la Monte SM, Yeon JE, Tong M et al (2008) Insulin resistance in experimental alcohol-induced liver disease. J Gastroenterol Hepatol 23:e477–e486 96. Dominici FP, Cifone D, Bartke A et al (1999) Loss of sensitivity to insulin at early events of the insulin signaling pathway in the liver of growth hormone-transgenic mice. J Endocrinol 161:383–392 97. Dominici FP, Diaz GA, Bartke A et al (2000) Compensatory alterations of insulin signal transduction in liver of growth hormone receptor knockout mice. J Endocrinol 166: 579–590 98. Dominici FP, Argentino DP, Munoz MC et  al (2005) Influence of the crosstalk between growth hormone and insulin signalling on the modulation of insulin sensitivity. Growth Horm IGF Res 15:324–336

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241 and hypertriglyceridemia with sterol regulatory element binding protein involvement. Diabetes 52:2905–2913 114. Shimomura I, Bashmakov Y, Ikemoto S et al (1999) Insulin selectively increases SREBP-1c mRNA in the livers of rats with streptozotocin-induced diabetes. Proc Natl Acad Sci U S A 96:13656–13661 115. Shimano H, Amemiya-Kudo M, Takahashi A et  al (2007) Sterol regulatory element-binding protein-1c and pancreatic beta-cell dysfunction. Diabetes Obes Metab 9(Suppl 2):133–139 116. Samson SL, Wong NC (2002) Role of Sp1 in insulin regulation of gene expression. J Mol Endocrinol 29:265–279 117. Chakravarty K, Wu SY, Chiang CM et al (2004) SREBP-1c and Sp1 interact to regulate transcription of the gene for phosphoenolpyruvate carboxykinase (GTP) in the liver. J Biol Chem 279:15385–15395 118. Finlay D, Patel S, Dickson LM et al (2004) Glycogen synthase kinase-3 regulates IGFBP-1 gene transcription through the thymine-rich insulin response element. BMC Mol Biol 5:15 119. Barroso I, Santisteban P (1999) Insulin-induced early growth response gene (Egr-1) mediates a short term repression of rat malic enzyme gene transcription. J Biol Chem 274:17997–18004 120. Porstmann T, Griffiths B, Chung YL et al (2005) PKB/Akt induces transcription of enzymes involved in cholesterol and fatty acid biosynthesis via activation of SREBP. Oncogene 24:6465–6481 121. Ma Z, Gibson SL, Byrne MA et al (2006) Suppression of insulin receptor substrate 1 (IRS-1) promotes mammary tumor metastasis. Mol Cell Biol 26:9338–9351 122. Selman C, Lingard S, Choudhury AI et al (2008) Evidence for lifespan extension and delayed age-related biomarkers in insulin receptor substrate 1 null mice. Faseb J 22: 807–818 123. Simmons JG, Ling Y, Wilkins H et al (2007) Cell-specific effects of insulin receptor substrate-1 deficiency on normal and IGF-I-mediated colon growth. Am J Physiol Gastro­ intest Liver Physiol 293:G995–G1003 124. Freude S, Leeser U, Muller M et al (2008) IRS-2 branch of IGF-1 receptor signaling is essential for appropriate timing of myelination. J Neurochem 107:907–917 125. Kido Y, Burks DJ, Withers D et al (2000) Tissue-specific insulin resistance in mice with mutations in the insulin receptor, IRS-1, and IRS-2. J Clin Invest 105:199–205 126. Kubota N, Tobe K, Terauchi Y et al (2000) Distribution of insulin receptor substrate 2 causes type 2 diabetes because of liver insulin resistance and lack of compensatory betacell hyperplasia. Diabetes 49:1880–1889 127. Liu SCH, Wang Q, Lienhard GE et al (1999) Insulin receptor substrate 3 is not essential for growth or glucose homeostasis. J Biol Chem 274:18093–18099

Role of PKB/Akt in Liver Diseases

16

Elena Zhuravleva, Oliver Tschopp, and Brian A. Hemmings

Introduction

Description of the PKB Family

PKB/Akt is a ubiquitous and evolutionarily conserved serine/threonine kinase that is recognized as a major coordinator of various intracellular signals. It controls cell responses to extrinsic stimuli and regulates cell metabolism, proliferation, and survival. Proper tuning of PKB activity via direct or indirect mechanisms is of utmost importance for stringent regulation of PKB-dependent cellular activities. Many diseases, such as cancer or metabolic disorders, are the result of, or are associated with, aberrant activity of the PI3K/PTEN/PKB pathway. In many tumors, the PI3K/PTEN/PKB pathway is activated by upstream mutations in PI3K or PTEN or by the amplification/overexpression/mutation of PKB iso­ forms themselves. Liver tumors are not the only pathological condition associated with disorders of this pathway. PKB has also been implicated in the development of hepatic insulin resistance, type 2 diabetes mellitus and, as has become evident over the past few years, in ischemia/reperfusion processes. In this chapter, the role of PKB in major physiological processes of cells is summarized and different liver disease conditions are considered by analyzing their  pathophysiology from the perspective of PKB involvement.

The first publication identifying PKB as a serine/threonine kinase came from Jones et al. in 1991 [1]. The authors initially termed the kinase Rac, for related to the A and C kinases, but was then renamed PKB/Akt [2, 3]. Since the publication of these landmark studies more than 17 years ago, it became clear that PKB/Akt is one of the major targets of phosphatidylinositol 3-kinase (PI3K)-generated signals and is involved in the regulation of cell growth, proliferation, apoptosis, glucose metabolism, angiogenesis, and migration. Readers are referred to other excellent reviews covering the history of PKB discovery and in-depth analysis of its involvement in all these diverse cellular processes [4–7]. The mammalian genome encodes three isoforms of Akt/PKB: Akt1/PKBa, Akt2/PKBb, and Akt3/PKBg, which are highly conserved despite being the products of three different genes located on different chromosomes. Interestingly, worms and flies have a single Akt/PKB protein. PKB family members share a similar domain structure containing an N-terminal pleckstrin homology (PH) domain [8], a catalytic domain and a C-terminal regulatory domain with a regulatory phosphorylation hydrophobic motif (HM) [4]. PKB that fully activated and exerts its biological functions is phosphorylated at two sites, one located within the activation loop of the kinase domain (Thr308 in PKBa) and the other within the HM (Ser473). Although PKB isoforms are highly conserved (approximately 80% amino acid identity), they apparently perform distinct biological functions, which could reflect tissue specific expression. For instance, PKBa mRNA is present in the majority of organs with low levels in pancreas and skeletal muscle [9–11]; PKBb mRNA is highly abundant in insulin-responsive tissues such as skeletal

B. A. Hemmings () Friedrich Miescher Institute for Biomedical Research, Maulbeerstrasse 66, 4058 Basel, Switzerland e-mail: [email protected]

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_16, © Springer-Verlag Berlin Heidelberg 2010

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muscle, liver, and adipose tissue, and PKBg mRNA levels are high in brain, testis, lung, mammary gland, and adipose. Interestingly, in tissues and organs with the highest levels of PKBb, PKBg levels are the lowest [11]. Taking into account the complex cellular composition of the liver, it should be noted that hepatocytes are not the only cell types expressing PKB. Kupffer cells, hepatic stellate cells (HSC), and sinusoidal endothelial cells (SEC) also express this kinase and PKB is involved in their functions in normal physiological conditions as well as in pathological states. For example, Kupffer cells stimulated by lipoteichoic acid and lipopolysaccharide reduce the inflammatory response  via activation of the PI3K/PKB

pathway [12]. It  has  also been suggested that PKB plays a key role in early liver regeneration and in the induction of cytokines secretion and the proliferation of SECs [13, 14].

Activation and Regulation of PKB/Akt Activating Stimuli and Upstream Kinases Growth factor (Table 16.1) binding promotes the recruitment and activation of class I PI3K after autophos­ phorylation of the receptor on tyrosine residues. At the

Table 16.1  List of selected PKB activators (top row) and PKB substrates assigned to two major groups: growth/survival/cell cyclerelated and metabolism-related (middle and bottom rows)

Activators

Growth factors and cytokines

Interactors and other molecules

Insulin Insulin like growth factor (IGF) Epidermal growth factor (EGF) Hepatic growth factor (HGF) Basic fibroblast growth factor (FGFb) Nerve growth factor (NGF) Platelet derived growth factor (PDGF) Vascular endothelial growth factor (VEGF)

Heat shock protein 90 kDa (Hsp90) BTB (POZ) domain containing 10 (BTBD10) Fused toes protein 1 Ft1 T-cell leukemia antigen 1 (Tcl1) Adaptor protein containing PH domain, PTB domain, and Leucine zipper motif (APPL) Akt phosphorylation enhancer (APE)

Chemical compounds and other stimuli Orthovanadate (phosphatase inhibitor) Okadaic acid (phosphatase inhibitor) DNA damage Hydrogen peroxide/reactive oxygen species Hypoxia Heat shock Zinc, cadmium Exercise

Leukemia inhibitory factor (LIF) Interleukins 2,3,4,5,8 (IL2,3,4,5,8) RANTES Stem cell factor Tumor necrosis factor alpha (TNFα)

Substrates

Growth, survival, and cell cycle related BAD Caspase 9 FoxOs YAP Mdm2 IKKα p21 p27 TSC2 PRAS40 Hexokinases GSK3α/β

Becomes a target for 14-3-3-proteins, is released from anti-apoptotic Bcl-2 proteins Inactivation, prevents formation of apoptosome Nuclear exclusion, prevents transcription of pro-apoptotic genes Inactivation, translocates to the cytosol from the nucleus Stabilization, leads to increase of p53 levels Inhibition, induces NF-κB transcriptional activity Cytoplasmic retention, blocks cell-cycle progression Cytoplasmic retention, attenuates cell-cycle inhibition Inhibition, is released from Rheb, which leads to mTOR phosphorylation/activation Inhibition (when non-phosphorylated, binds to mTORC1 and inhibits its activity) Activation, promotes association of HK with mitochondrial membrane Inhibition, leads to stabilization of proteins involved in G1/S phase transition

Metabolism related AS160 ACL PGC-1α GSK3α/β FoxOs Hexokinases PFK2

Inhibition, leads to release of GLUT4 and increase of glucose transport Activation, increases fatty acid synthesis Inactivation of co-activating transcriptional activity for FoxOs, HNF4α, and other genes Inactivation, increase in glycogen synthesis Inactivation (when active, decreases glycolysis and fatty acids synthesis, increases gluconeogenesis) Activation, important for glucose utilization Activation, catalyzes production of fructose 2,6-bisphosphate, leads to increased glycolysis

16  Role of PKB/Akt in Liver Diseases

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membrane the PI3K phosphorylates PtdIns(4,5)P2 to form PtdIns(3,4,5)P3, which then serves as docking site for a subgroup of proteins with PH domains. The PI3K family comprises of three classes, I, II, and III. Class I PI3Ks are heterodimers of distinct regulatory (p50-55/p85) and catalytic (p110a, p110b or p110d) subunits; their activity is directed mainly to the phosphorylation of PIP2 to PIP3. Class II and class III differ from class I in structure and function (for more comprehensive reviews see [15, 16]). PI3K is involved in the regulation of a wide  range of cellular processes, such as cell growth, proliferation, differentiation, motility, survival, and intracellular trafficking; many of these PI3K effects are mediated by downstream PKB. The constitutive activation of PI3K class I due to a gain-of-function mutation (in the p110a subunit, for example) or the downregulation of its

negative regulator PTEN (lipid phosphatase and tensin homolog deleted on chromosome ten) are striking features of many human cancers [17]. Thus, inactive PKB/Akt that is translocated to the plasma membrane (PM), undergoes a conformational change, attaches to phospholipid through a PH domain and becomes phosphorylated (schematic representation of PKB activation is shown on the Fig.  16.1). Once recruited to the PM, PKB is activated in a two-step process that requires phosphorylation on both Thr308 in the activation loop of the kinase domain and Ser473 within the HM of the regulatory domain. Thr308 is subjected to phosphorylation by PDK1 kinase, which is recruited to the PM through its PH domain; precomplexed PDK1 in cytoplasm [18, 19]. Notably, mutation of the PH domain of PDK1 in mice diminishes PKB

GFR GF

PT SH EN IP

P

Ras

PIP2

P

PI3K

P

PIP3

P

P

P

PDK1 KINASE

PH PH

TR

P

PH

P

T308

TORC2

T308

DNA-PKc

HM

HM P

S473

S473

N

TIO

CA

LO

S AN

PIP3

P

PH

Activation of MAPK Pathway

PDK1

PH

ATM

N

IO

AT

C LO

S

AN

TR

HM

Inactive PKB

INACTIVATION / DEPHOSPHORYLATION

PH

PP2A PHLPP P

POSITIVE REGULATORS P

Hsp90, BTBD10 Ft1, Tcl1

HM Grb10, NEGATIVE REGULATORS Trb3, CKIP-1 Keratin10 CTMP

Active PKB

DOWNSTREAM TARGETS

Fig. 16.1  Schematic representation of PKB activation and regulation (adapted from [6])

UPSTREAM S473 KINASES

P

KINASE

P

SOS

PLASMA MEMBRANE

KINASE

P

P

P85

Shc P Grb2

LY294002

P110

P

KINASE

Wortmannin

P

246

a­ ctivation, leading to small size and insulin resistance [20]. PKB, monophosphorylated on Thr308 has ca. 10% of the activity of the fully phosphorylated enzyme. Additional Ser473 phosphorylation stabilizes the active conformation, allowing most PKB molecules to adopt a fully active state [21]. The identity of the kinase phosphorylating PKB on Ser473 remained controversial for many years; today it is accepted that the PI3K-related protein kinase family the TORC2 complex, DNA-PK, and ATM are responsible for this phosphorylation. The fact that mTORC2 is a Ser473 kinase for PKB is widely recognized [22, 23]. Work done on knock-out mice and Drosophila cells has provided genetic evidence favoring the hypothesis that components of the rapamycininsensitive Rictor-mTOR complex have a shared posi­tive role in the phosphorylation of the HM site of PKB [22]. DNA-PK has also been identified as an upstream Ser473 kinase of PKB [24, 25]. However, DNA-PK does not play a role in insulin-promoted activation of PKB in serum-sufficient cells. It has been shown that DNAPKcs specifically phosphorylates PKB on Ser473 after DNA damage, thus promoting survival in response to genotoxic stress in vivo [25]. Activation of growth factor receptors (GFR) by a ligand (insulin, growth factors (GF)) induces their autophosphorylation and recruits the p85 regulatory subunit of phosphatidylinositol 3-kinase (PI3K). Subse­ quent activation of the p110 catalytic subunit leads to phosphorylation of phosphoinositol-(4,5)-bis phosphate (PIP2) and formation of the phosphoinositol(3,4,5)-tris phosphate (PIP3). PIP3 is a substrate for lipid phosphatase, the tensin homolog PTEN, and the SH2-domain-containing inositol polyphosphate 5-phosphatase SHIP, which act as endogenous inhibitors of the PI3K-dependent pathway, indirectly inhibiting PKB activity. Wortmannin and LY294002 also inhibit PI3K. Once formed, PIP3 serve as docking sites for the PH domains of PDK1 and PKB, which translocate to the plasma membrane from the cytoplasm. As a result of this translocation, inactive PKB is phosphorylated by PDK1 on Thr308 in a regulatory kinase domain. The second phosphorylation event on Ser473 in the HM by upstream kinases such as TORC2, DNA-PKc, and ATM are cell-type and stimuli specific. It leads to conformational changes in the PKB molecule and full activation of the kinase. Activated PKB then trans­locates to different subcellular compartments, such as the nucleus, ER, Golgi, and

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mitochondria, where it exerts its biological activity. Protein phosphatase 2A (PP2A) and a PH domain leucine-rich repeat protein phosphatase (PHLPP) dephosphorylate and inactivate PKB. Other negative regulators of PKB are Grb10, carboxyl-terminal modulator protein (CTMP), tribbles homolog 3 (Trb3), casein kinase 2-interacting protein-1 (CKIP1), and keratin 10. Positive regulation of PKB activity may be achieved through interaction with BTBD10 and the heat shock proteins Hsp90 and Hsp27, which protect the PKB molecule from dephosphorylation. T-cell leukemia antigen-1 (Tcl-1) and fused toes protein-1 (Ft1) may also function as positive regulators of PKB. For more details on the activation and regulation of PKB please refer to the corresponding section of this chapter.

Positive Regulation of PKB via Interaction with Other Proteins Positive regulation of PKB activity may be mediated through interaction with Hsp90, Tcl1, and Ft1 (see Table  16.1). Protein chaperone Hsp90, a prominent target for anticancer therapy, protects many kinases bearing activated mutations [26, 27]. It is known that activated PKB requires Hsp90 to maintain the phosphorylation state, which allows cancer cells to proliferate and circumvent apoptosis [28, 29]. T-cell leukemia antigen (Tcl1) interacts with PKB and enhances its kinase activity [30, 31]. It is highly activated in multiple neoplastic conditions, mainly T- and B-cell malignancies. Fused toes protein 1 (Ft1) was identified as a direct PKB interactor, leading to the enhanced phosphorylation of both Thr308 and Ser473 by promoting its interaction with the PDK1 [32].

Negative Regulation of PKB by Phosphatases Certain cellular mechanisms counteract the activation of PKB. Negative regulation of PKB could be mediated either by a direct mechanism, such as intramolecular interactions, or indirectly by modulation of factors impor­ tant for PKB activation.

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The best-studied negative regulator of the PI3K/PKB pathway is PTEN, a tumor suppressor protein that is often inactivated in many disorders characterized by PKB hyperactivation, such as cancers and some metabolic diseases (described in detail below). This molecule acts as a lipid phosphatase by dephosphorylating PIP3 at the D3 position converting it to PIP2. This leads to inhibition of the PI3K pathway and reduces recruitment of PDK1 and PKB to the PM and thus, subsequently decreases PKB activity. At the transcription level, PTEN is positively regulated by p53, Myc, Egr-1, and PPARg, whereas Ras, JNK, Notch, and miR-21 are negative regulators of PTEN transcription (reviewed in [33]). Downregulation due to the loss of promoter activity or loss-of-function mutations of PTEN are distinct characteristics of many neoplastic diseases, including hepatocellular carcinoma (HCC) [34, 35]. Given that PKB activation is achieved by increased phosphorylation, protein phosphatases are direct negative regulators acting on phosphorylated PKB. Protein phosphatase 2A (PP2A) acts as a negative regulator by dephosphorylation of PKB at both sites [36, 37], but particularly at Thr308 [38]. Heat shock protein 90 (Hsp90), a general chaperone to numerous targets may inhibit PP2A-mediated dephosphorylation, offering PKB protection from inactivation. BTB (POZ) domain containing 10 (BTBD10) has also been reported to interact with PKB and protect it from PP2A-mediated dephosphorylation [39]. There is no structural similarity between BTBD10 and Hsp90 and, unlike Hsp90, BTBD10 does not affect PDK1 activity [40]. BTBD10 probably acts by binding to PP2A and its target. It remains to be established whether BTBD10 is a PKB-specific activator or whether it protects other PP2A targets. A further phosphatase directly dephosphorylating Ser473 is the PHLPP, which has a PP2C-like catalytic core, is not sensitive to okadaic acid, and binds directly to PKB via a C-terminal PDZ motif [38]. This phosphatase is markedly reduced in several colon cancer and glioblastoma cell lines with elevated PKB phosphorylation. Recently, a second PHLPP isoform, PHLPP2, has been cloned [41]. In parallel to the different tissue expression patterns and substrate specificities of the different PKB isoforms, the PHLPP isoforms have been reported to show specificity for distinct PKB isoforms. PHLPP1 was shown to affect PKBb/g while PHLPP2 influences the activity of PKBa/g, with marked differences in the affected PKB substrates. These data led to the speculation that PHLPP1 may regulate PKBb and affect glucose

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metabolism, while PHLPP2 is involved mostly in the regulation of PKBa and cell survival [41].

Further PKB/Akt Interactors Molecules other than phosphatases also negatively regulate PKB activity. Grb10, a growth factor receptorbound protein 10, interacts with numerous receptor tyrosine kinases and has been implicated in the regulation of the PI3K pathway downstream of the insulin receptor. Whether Grb10 has positive or negative effects on insulin- or IGF-receptor signaling remains controversial [42–44]. In 2005, it was shown that Grb10 and PKB form a constitutive complex that promotes kinase translocation to the PM; thus, it has been proposed that Grb10 functions as a PKB coactivator [45]. However, these studies were performed mainly in vitro and the in vivo role of Grb10 was unknown until recently. First, it was shown that overexpression of Grb10 in mice leads to insulin resistance [46] and the Liu group reported later that disruption of this gene enhances insulin signaling and sensitivity [43]. CTMP, shown to interact with PKB, delays Ser473 phosphorylation and thus inhibits kinase activation [47]. Keratin K10 is also a negative regulator of PKB, to which it binds and inhibits its intracellular translocation, leads to downregulation of PKB activity [48]. Casein kinase 2-interacting protein-1 (CKIP-1) has been shown recently to interact with the PH domain of PKB and inhibit kinase active, thus suppressing tumor growth [49]. Tribbles homolog 3 (Trb3) is a pseudokinase that binds preferentially to nonphosphorylated PKB, thereby blocking Thr308 phosphorylation induced by insulin and other GF [50]. Trb3, the abundance of which increases during fasting, may be especially relevant for liver tissue. It was suggested that Trb3 activation is beneficial under fasting conditions, but that pathological overexpression after food intake may contribute to insulin resistance and hyperglycemia. However, genetic deletion of Trb3 in mice had a minimal effect on insulin-induced PKB activation in hepatic tissues, arguing that Trb3 has a minor effect on glucose and energy homeostasis [51]. Further studies are clearly needed to evaluate the importance of Trb3 in other contexts such as hypoxia and other stresses, and after tissue-specific deletions of Trb3. Overall, it may be concluded that fine tuning of PKB signaling occurs at different levels, beginning

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with activation by a variety of exogenous signals, which is then contra-regulated via either direct dephosphorylation or interaction of numerous proteins with different domains of the kinase.

PKB Substrates and Their Functions Regulation of PKB has been studied extensively ever since its discovery and all the results obtained have made clear that PKB, being so stringently regulated, is one of the most versatile proteins in the cell. The long list of PKB substrates can be divided into several subgroups corresponding to the processes in which PKB is involved. These include regulation of cell size, transcription, survival, antiapoptotic activity, and metabolism (for a recent review see [7]).

Regulation of Cell Size and Survival Once activated following GF/insulin treatment (Table 16.1), PKB translocates from the cell membrane to its targets and phosphorylates them at the consensus site RXRXXS/T. Constitutive activation of PKB contributes greatly to aberrant cell cycle regulation, resulting in uncontrolled cell proliferation and suppressed apoptotic pathways. This prosurvival and antiapoptotic activity is exerted through phosphorylation of target proteins, leading to either activation or inhibition of their activities. The predominant mechanism by which PKB regulates cell size (cell mass) is activation of the mTOR complex 1 (mTORC1). This is exerted via phosphorylation of TSC2 within the complex TSC1/2 and abrogation of TSC2 activity. When nonphosphorylated, TSC2 prevents formation of Rheb-GTP. Upon TSC2 phosphorylation, Rheb-GTP accumulates and activates mTORC1, which in turn activates S6K1 and 4E-BP1, stimulating translation initiation and ribosome biogenesis [52–56]. Like TSC2, PKB also phosphorylates PRAS40 (prolinerich Akt substrate of 40 kDa) [57–59]. Overexpression of a mutant, which cannot be phosphorylated by PKB, blocks PKB-mediated activation of S6K1. Recently, PRAS40 has been found to associate with and be phosphorylated by mTORC1 and is thought to negatively regulate its signaling [59–62]. DNA double strand breaks and promotes PKB activation. Recently published work by Bozulic et al. has

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implemented the role of PKBa isoform in promotion of cell survival and antiapoptotic processes upon genotoxic stress [25]. DNA-PK recruitment to doublestranded, damaged DNA results in activation of PKBa; DNA-PK serves as an upstream kinase for PKBa and Ser473 PKBa phosphorylation occurs in the nucleus, presumably leading to the activation of transcription factors with prosurvival activity. The importance of PKB for cell proliferation and survival is further illustrated by overexpression experiments in which myristoylated or constitutively active variants of PKB were introduced into either cells or mice in a tissue/organ-specific manner. For example, overactive PKB alone contributes to tumorigenesis; the overexpression of PKB in the prostate gives rise to neoplasia. Increase in PKB activity enhances the number and size of thymocytes, cardiomyocytes [63], pancreatic b-cells [64, 65] prostate epithelial cells [66], and hepatocytes [67]. In contrast, PKBa knock-out mice are small, suffer increased neonatal lethality and increased spontaneous apoptosis in the thymus [11, 68, 69]. This isoform was shown to be important for trophoblast differentiation and it was concluded that PKBa regulates placental development and differentiation. These multiple in vivo studies emphasize a role for PKB at different stages of development and point to an indispensable role of the PKBa isoform in guiding cell size and survival processes.

Regulation of Apoptosis The first identified PKB target involved in apoptosis was the BH3-only protein Bad, which is a negative regulator of prosurvival Bcl-2 family members [70]. Phosphorylated on Ser136 by PKB, Bad becomes a ­target for 14-3-3 proteins, which triggers release of Bad from Bcl-2 proteins. Caspase 9, which is another PKB  substrate with proapoptotic activity, participates in the  formation of the apoptosome. Phosphorylation on Ser 196 inhibits protein activity and blocks caspase 9 induced cell death [71]. PKB substrates from the FoxO family (FoxO1-4) are inactivated upon phosphorylation via their exclusion from the nucleus, where they normally act as transcription factors. FoxOs target genes promote apoptosis, cell-cycle arrest, and various metabolic processes (see below); among these are Bim, Bad, and the proapoptotic protein FasL [72, 73]. Similar

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to FoxO proteins, the proapoptotic protein YAP translocates to the cytosol after being phosphorylated, whereas when active in the nucleus, it activates transcription of proapoptotic genes such as p73 [73]. Overall, activation of the PKB pathway provides cells with survival signals that allow them to withstand apoptotic stimuli.

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i­nsulin-mediated glucose uptake in muscle and in ­adipocytes and an inability to lower hepatic glucose production [76–78]. As a result, these mice are insulin resistant and glucose intolerant and exhibit subsequent b cell failure. All these phenomena are typical of type 2 diabetes mellitus (T2D), which is characterized by an increased insulin requirement compounded by decreased insulin secretion.

Cell Cycle Control Role of PKB in Glucose Metabolism PKB-mediated control of cell-cycle progression has been shown to be crucial for the G1/S transition. This activity is mediated mainly by regulation of FoxO localization and p53 intracellular concentration (via Mdm2 stabilization). PKB also phosphorylates p21, which act as cellcycle inhibitor. When phosphorylated it accu­mulates in the cytoplasm and cannot bind and inhibit CDK2, which allows cell-cycle transition. p21 can be regulated also at the transcriptional level by FoxO proteins, promoting antiproliferative effect. The Mdm2/p53 signaling pathway plays an extremely important role in the regulation of cell cycle progression [74]. Mdm2, which is an E3 ubiquitin-ligase targeting p53 to its degradation, is phosphorylated and consequently stabilized by active PKB [75]. Mdm2 stabilization consequently leads to p53 ubiquitination and degradation. Notably, the p53 pathway is the most mutated in cancer cells, including HCC. Its deregulation, i.e., loss-of-function, promotes cell cycle transition and blocks apoptosis, leading in many cases to cancer progression. Cell cycle transition, mediated by PKB, is closely associated with proliferation and apoptosis, greatly contributing to the events of transformation and regeneration.

Metabolism The role of PKB in the regulation of metabolic pathways is tightly interconnected with its positive effects on cell growth and proliferation. The main PKB isoform important for executing these effects in insulinsensitive tissues is PKBb (and to some extent PKBa). As already discussed, the principal sites of expression of the b isoform are liver, muscle, and adipose tissue. Analysis of PKBb KO mice revealed the importance of this particular isoform in mediating metabolic effects of PKB. Notably, PKBb KO mice exhibit impaired

PKB exerts its biological activity by actively stimulating glucose uptake. Here, the important substrate is AS160 (Akt substrate of 160 kDa, or TBC1D4), a molecule with Rab-GAP activity that has up to nine potential PKB phosphorylation sites. Upon phosphorylation, AS160 loses its GAP activity, which leads to the accumulation of Rab-GTP and, consequently, to the translocation of the glucose transporter molecule Glut4 to the PM and glucose uptake. In line with this, insulin-­ stimulated AS160 phosphorylation is substantially reduced in muscles of T2D patients. Thus, after uptake, glucose is exploited either in the pentose-phosphate pathway (PPP), is oxidized via glycolysis, or is stored in the form of glycogen. PKB can be viewed as an important signaling node that coordinates signals of metabolic and antiapoptotic pathways. This effect is mediated by hexokinases (HKs), which phosphorylate glucose to glucose-6-phosphate; it is used later in PPP or glycolysis. HKs are bound to the outer membrane of mitochondria and PKB activation maintains or enhances this association. This leads, in turn, to antiapoptotic effects of PKB/HKs on the cell via mitochondrial membrane stabilization [79]. Such effects require glucose and may be considered as a system linking cell survival and metabolism. To a large extent PKB signaling affects both glycolysis and glycogenesis. Glycogen synthase, the enzyme catalyzing the last step in glycogen synthesis, is phosphorylated by glycogen synthase kinase 3 (GSK3) [80]; this phosphorylation leads to inactivation of glycogen synthase. Both the a and b isoforms of glycogen synthase kinase are PKB substrates. Phosphorylation on Ser21 and Ser9 of GSK3a and b, respectively, leads to inactivation of the kinase, which in turn stimulates glycogenesis. The subsequent activity is of utmost importance in liver and muscle. Most of the research has been focused on the GSK3b isoform, implicating it in

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muscle glucose metabolism [81]; Woodgett’s group has reported recently that the GSK3a isoform is a major regulator of glucose turnover in the liver [82]. PKB activation also increases rates of glycolysis, which has been shown in numerous cancer lines [83]. Its overexpression correlates with increased glucose consumption and the accumulation of NADH and lactate without increase in oxygen usage, which suggests that it does not affect mitochondrial respiration. The fact that many tumors are characterized by increased PKBa activity raises the interesting possibility that PKBa regulates Glut1 membrane localization. Tumors often display overexpression of Glut1 on the cell surface as well as elevated glucose transport, which would identify PKBa as a cause of increased Glut1 expression and glycolysis.

Regulation of Lipid Metabolism Insulin is a potent anabolic hormone and, in addition to glucose metabolism, it also regulates lipid homeostasis by increasing lipogenesis in the case of nutrient excess. PKB was shown to mediate insulin/PI3K effects through GSK3 inactivation, which when actively promotes degradation of sterol regulatory element binding proteins (SREBPs). It should be mentioned that a further PI3Kdependent kinase, atypical PKC, mediates the insulin effect on increased lipogenesis via increasing SREBP expression [84]. Three isoforms of SREBPs, -1a, -1c, and -1b, trigger expression of more than 30 genes involved in cholesterol metabolism, fatty acid, triglyceride, and phospholipid biosynthesis. In liver, as well as in adipose tissue, the predominant isoform is SREBP-1c, which upregulates transcription and expression of lipogenic enzymes such as fatty acid synthase (FAS) and ATP citrate lyase (ACL). ACL is also a PKB substrate important for fatty acid synthesis. ACL links glucose metabolism to lipid synthesis by cleaving citrate to acetyl-CoA and oxaloacetate in the cytosol, thus supporting the synthesis of fatty acids, cholesterol, etc. [85]. ACL has been proposed as a target for anticancer therapy, since it is positively regulated in PKB-overexpressing tumors it mediates metabolic switches characteristic of cancer cells [86]. HCC, a neoplastic disease of liver, is associated with a marked increase in lipogenic enzymes and it was suggested that SREBP-1c is involved in this process [87].

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FoxOs at the Crossroads of Metabolism and  Survival: The Role of PGC-1a FoxO is a further target of PKB, belonging to a family of proteins associated with cell cycle regulation by PKB. FoxO also contributes to glucose homeostasis and is an important target of insulin action. FoxO1 promotes hepatic glucose production and regulates the differentiation of cells involved in metabolic ­control (reviewed in [88]). In the liver, FoxOs have been shown to increase gluconeogenesis and triglycerides (TG) metabolism via transcriptional upregulation of phosphoenolpyruvate carboxykinase (PEPCK) and glucose-6-phosphatase (G6Pase), and PPAR-g coactivator 1a (PGC-1a). FoxOs decrease glycolysis and fatty acid/TG synthesis [89]. The results of liver­specific KO of FoxO1 illustrate the pivotal role of this transcription factor in promoting hepatic glucose production and provide details of the interconnection of the cAMP and insulin pathways in the regulation of glucose production [90]. Coactivator PGC-1a is a well-recognized global regulator of liver metabolism in the fasting state that interacts with and recruits transcription factors to gene promoter regions [91]. It is involved in the control of gluconeogenic gene expression by coactivating FoxO1 and HNF-4 and was also shown to be associated with the promoter of the glucose G6Pase gene. It was found recently that PKBb directly phosphorylates and inhibits PGC-1a activity, which suggests a further role for insulin and PKB in controlling lipid catabolism in the liver [92]. FoxOs also protect cells from oxidative stress via upregulation of MnSOD2, catalase, and other enzymes involved in detoxifying reactive oxygen species (ROS). This activity is highly conserved among mammals, in worms, and flies. Overexpression of constitutively active PKB results in increased susceptibility to oxidative stress [93]. In addition, it may also regulate amino acid uptake through the activity of the TSC2/mTORC2 pathway, but the complex network of the PKB/mTOR pathway is still not sufficiently defined.

Role of PKB in Liver Regeneration The liver has an enormous capacity to regenerate after traumatic tissue loss or exposure to hepatotoxic

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substances. It compensates efficiently for large tissue loss as experienced after surgical removal of intrahepatic tumors [94]. The mechanisms of this regeneration are still the subject of intense research and new players like serotonin have been discovered recently [95]. Liver regeneration in response to tissue damage is a complex and well-orchestrated process and many excellent studies cover this topic [96, 97]. After major tissue loss many cytokines and GF (e.g., HGF, EGF, VEGF, or PDGF) are released leading to proliferation of hepatocytes. Within 48 h, the majority of hepatocytes re-enter the cell cycle moving from G0 to S phase. After cessation of proliferation, hepatocytes regain cellular volume and the original liver volume is almost completely restored within 1 week in rodents. PKB is strongly activated by many GF shown to be crucial in hepatic regeneration, e.g., HGF, EGF, or PDGF [96]. During liver regeneration in a partial ­hepatectomy model, a robust and sustained activation of total PKB was observed [98]. Hepatic expression of constitutively active PKBa was associated with a threefold increase in liver mass due to the increase in hepatocyte size without change in cell number [99]. Additionally, PKB-dependent hypertrophy of hepatocytes was sufficient for compensatory recovery of liver mass in liver-specific STAT3 mutant mice and in a thrombocytotic model of liver regeneration [100, 101]. In contrast, total PI3K inhibition with wortmannin, or selective inhibition with siRNA, resulted in a significant decrease in hepatocyte proliferation, especially at the earliest time points [102]. Liver regeneration appears to be dependent on the presence of PKB. However, the expected reduction in regenerative capacity in PKBa or PKBb mutant mice has not yet been demonstrated.

Involvement of the PI3K/PTEN/PKB Pathway in Liver Diseases Liver diseases contribute significantly to human mortality and rising health-care expenditure. Although the liver can compensate for significant injury, there may be a significant decrease in function leading to organ failure. Proper functioning of the PKB network may be needed to maintain organ function and its deregulation is a significant factor in the development of certain hepatic diseases and liver-associated syndromes.

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Insulin Resistance, Nonalcoholic Fatty Liver Disease, and Hepatosteatosis Among liver diseases, nonalcoholic fatty liver disease (NAFLD) is becoming increasingly common; every third adult and tenth child/adolescent in the USA is affected by this condition [103]. NAFLD is considered to be a hepatic manifestation of a metabolic syndrome. The prerequisite feature of NAFLD is increased accumulation of lipids in hepatocytes, which can originate either from increased levels of nonesterified fatty acids (NEFA) circulating in the blood or from enhanced de novo synthesis of lipids in the cells. Whereas, simple steatosis seems to have a relatively benign clinical course, a subgroup of patients develops inflammatory changes, known as nonalcoholic steatohepatitis (NASH) and potentially liver cirrhosis with increased risk for HCC [104]. Increase in intracellular lipid metabolites may lead to activation of the PI3K/PKB pathway. Initial overactivation of the signaling cascade then results in inhibition of IRS-1, which mediates a negative feedback effect on PI3K activation, leading finally to the inhibition of insulin signaling. Patients with NAFLD display lower levels of phosphorylated PKB and an increase in Bax/Bcl-2 ratio [105]. A similar situation has been described for activated PKCe, which led to the decrease in insulin-stimulated IRS-2 phosphorylation and hepatic insulin resistance [106]. Later, insulin resistance involves other peripheral organs, such as skeletal muscle and adipose tissue, eventually becoming systemic. Under physiological conditions, insulin released from the pancreas decreases glucose output from the liver but stimulates glucose uptake by muscle and adipose tissue. In patients with insulin resistance, a syndrome associated with impaired metabolic clearance of glucose, the concentration of NEFA in the bloodstream increases due to the high lipogenic effect of insulin. NEFA act by reducing adipocyte and muscle glucose uptake and promoting hepatic glucose output, which leads to increased blood glucose concentration. Obesity worsens the situation and leads to increased levels of NEFA released directly into the portal vein from visceral adipose. This adds to the vicious cycle leading to even higher insulin resistance. Hepatocellular lipid accumulation concerns about 40% of the population and renders the liver partic­ ularly susceptible to inflammatory cytokines, endo­ toxins, iron accumulation, and oxidative stress [107].

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The latter is caused to a large extent by increased intracellular lipids. Most studied are the effects of lipotoxicity on beta cells [108, 109] and cardiomyocytes [110], but hepatocytes are also affected significantly, via a direct or an indirect mechanism, often displaying mitochondrial dysfunction [111]. Key characteristics of mitochondrial dysfunction are respiratory chain defects that lead to an increase in ROS production. In addition to damaging mtDNA and respiratory chain enzymes, increase in ROS leads to the oxidation of many cytoplasmic enzymes, including PTEN. Thus, it is now clear that ROS are not only deleterious for the cell but are very important with respect to signaling molecules [112]. Their presence at low concentrations helps maintain certain basal levels of kinase activity, namely PI3K/PKB in cells not supplemented with GF. However, when cells are overloaded with ROS, this may lead to hyperactivation of the PI3K/PTEN/PKB pathway, to inhibition of FoxO1 and PGC-1a and the transcription of adipogenic- and lipogenic- as well as beta-oxidation-related genes. This leads to liver steatosis and potentially to hepatic tumorigenesis. The role of cell types other than hepatocytes in insulin resistance should not be neglected. Sinusoidal liver cells are often ignored but they are an important factor in the maintenance of the pathological situation of insulin resistance, exacerbating the oxidative damage of hepatocytes, as well as secreting the proinflammatory cytokines, TNF-a and IL6 [113]. It has been suggested that the degree of insulin sensitivity depends on the state of activation of stellate cells, which have phosphorylated IR and IRS1 when activated and do not respond further to insulin, thus failing in glucose uptake. Their activation brings hepatic tissue one step closer to the development of fibrosis and, ultimately, cirrhosis [113, 114].

Involvement of Phosphatases in Insulin Sensitivity Beneficial for the restoration of insulin sensitivity may be the target of the PI3K/PKB pathway via a decrease in phosphatase activity and the negative regulation of PKB activity. This is indeed the case is shown by the protein tyrosine phosphatase 1 B (PTP1B), which negatively regulates insulin receptor and IRS-1 signaling knock-out mice. Disruption of the PTP1B gene in mice

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leads to enhanced insulin sensitivity and decrease in adipose mass. In mice with polygenic insulin resistance, deficiency in PTP1B results in improved glucose tolerance and a decrease in the occurrence of diabetes [115]. Similarly, muscle-specific KO of PTP1B also has a beneficial effect on insulin sensitivity [114]. Another example of increased insulin sensitivity due to loss of lipid phosphatase activity are PTEN knockout mice. Quite unexpectedly, in addition to the insulin hypersensitive phenotype, liver-specific PTEN knockouts suffer from hepatomegaly and fatty liver, being at the same time overall leaner than their wild type littermates [116]. These mice have low levels of NEFA in the plasma and relative hypoinsulinemia concomitant with the higher PKB activity. They display an increased liver glycogen content and enhanced FA synthesis and secretion. PTEN null mice are in principle a further model of NAFLD that does not involve overnutrition and is not polygenic in nature. These mice are valuable for the absence of obesity/T2D, which is speculated not to be mandatory for insulin resistance but be only an associated complication. PTEN null mice provide a new insight into the in vivo role of PTEN, underlying  the importance of negative regulators of PKB in ­insulin-signaling pathway in liver. HCC, recognized as a complication of NAFLD, is often accompanied by downregulation of PTEN, leading to the poor prognosis in HCC patients [29]. Deregulation of PTEN activity may be due to hypermethylation of its promoter [14], microRNA-21 induced degradation of PTEN mRNA [117], or loss-of-function mutations [118]. Alternatively, PTEN may be the target of increased ROS in cells, leading to its oxidation and blockage of the catalytic cystein residues, important for proper function [119]. In this context, PTEN heterozygous and liver-specific KOs are invaluable models for studying the pathogenesis of NAFLD, that later develops into steatohepatitis and progresses to liver carcinoma without any additional exogenous treatment.

The PI3K/PTEN/PKB Pathway in the Development of HCC Alterations in PI3K/PKB signaling components are generally frequent in many tumors and HCC is not an exception. Various pathway components that provide a balance between cell survival and apoptosis are often

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deregulated. A mutation in the PIK3CA gene, which encodes a p110a subunit of PI3K class IA, is found in 35% of HCC cases, although it may depend on the population [120–122]. This is a gain-of-function mutation that results in enhanced oncogenic activity of PI3K. Other recent data have shown that the phosphorylation status of PKB may serve as a prognostic factor for early disease recurrence and poor prognosis [123]. In addition to altered metabolic regulation upon hyperactivation of PKB, many prosurvival and antiapoptotic activities start to prevail over cell death/apoptosis processes. Among the proapoptotic molecules downregulated in HCC are many PKB targets, including Bid, Bax, and p53. Proapop­ totic molecule Bad was recently shown to be activated upon knock-down of hepatoma-derived growth factor (HDGF) in human HCC cell line HepG2. Tsang et al. reported that downregulation of HDGF leads to inactivation of PKB and ERK and induces Bad expression, leading to activation of the apoptotic pathway followed by the release of cytochrome c and the caspase 3 and 9 cleavage [124]. mTORC1, a further downstream target of PKB that is extremely important in HCC for growth and proliferation, angiogenesis, and resistance to apoptosis, is also affected as a result of PI3K/PKB hyperactivation [125]. Phosphorylation of the mTOR target p70S6 is associated with elevated cyclinD1 levels and decreased overall survival of patients with HCC, indicating the aggressive nature of HCC (p53, apoptosis, and mTOR are described more extensively in corresponding chapters of this book) [125, 126]. Several studies have shown that downregulation of PKB signaling promotes apoptosis and enhances susceptibility of HCC patients to anticancer drugs [127]. Another aspect of PKB involvement in liver carcinoma development and progression is an impact on tumor metabolism. It is known that PKB stimulates the biosynthesis of fatty acids via activation of SREBP, which leads to an increase in the concentrations of cellular fatty acids and PM components [128]. In addition to SREBP activation, PKB also has an impact on FAS expression. FAS has been shown to be significantly upregulated in various types of cancers whereas, except for liver and adipose tissues, FAS levels are usually quite low in normal tissues [129–131]. Many tumors switch their metabolism and start de novo synthesis of fatty acids as an energy component; FAS as one the major enzymes of lipogenesis is consequently upregulated. FAS overexpression is often accompanied by the

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induction of antiapoptotic mechanisms, leading to a selective advantage for tumor cells in survival and cell cycle progression. Indeed, this is the case in HCC, where key lipogenic enzymes such as FAS and ACL are markedly induced [69]. Until recently, the mechanism responsible for this upregulation was unclear. In 2008, the group of Watabe reported that upregulation of the FAS gene occurs under hypoxia conditions via PKB and SREBP-1 activation, and that inhibition of FAS overcomes hypoxia-induced chemoresistance, which is a major clinical problem in many cancer patients [132]. It is worth noting that changes in the metabolism of cancer cells, known as the Warburg effect, or glycolysis under normoxic conditions, lead to changes in redox balance. This, in turn, causes PTEN inactivation and subsequently PKB hyperactivation, all of which later contribute to the induction of FAS and enhancement of lipogenesis.

Viruses, PKB, and Liver Diseases What makes patients with chronic liver diseases more prone to develop HCC? One important feature defining the outcome is PTEN status, which may serve as an independent prognostic marker of HCC associated with HCV patient survival and the presence of viral infections, such as combined hepatitis B and C (HBV/HCV) (for further details, see the corresponding chapters about HBV/HCV in this volume) [118, 133]. Hepatitis viruses are known to cause PKB activation. Thus, HCV protein NS5A may either activate PI3K via binding to its regulatory subunit or inhibit apoptosis by acting on the Bax protein [134, 135]. Furthermore, approximately 50% of patients with a chronic HCV infection develop liver ­steatosis and insulin resistance, mediated by increased ­oxidative stress, the activation of PI3K/PKB and the transactivation of PPARg and SREBP-1/2 [136–138]. On the other hand, insulin resistance may be linked to inhibition of PKB signaling, either through impairment of upstream IRS-1 and PI3K [139], or, as shown recently, through overexpression of protein phosphatase PP2A [140]. Recent analysis of HCC samples revealed that activation and overexpression of PKB and phosphorylation of its target GSK3b are among the most consistent features in HBV-associated HCC [141]. Activation of the PKB pathway as a consequence of viral infection leads to antiapoptotic effects on infected cells. This is

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the case with HBV protein HBx, which activates the PI3K/PKB/Bad prosurvival pathway. HBx was shown to affect PTEN expression by inhibiting the function of p53, an established transcriptional regulator of PTEN [142], and overexpression of PTEN in these cells may reverse signaling modulated by HBx and have a positive effect on apoptosis [143].

Conclusions The serine/threonine protein kinase PKB is major downstream mediator of PI3K. PKB coordinates a constellation of intracellular signals and controls cellular responses to a variety of extrinsic stimuli. PKB regulates cell proliferation, survival, growth, glucose and lipid metabolism, and malignant transfor­ mation. Insulin and other GF are potent PKB activators. In many pathological conditions such as NAFLD or HCC the PI3K/PKB pathway is often overactivated due to aberration in the upstream regulation of the kinase. PKB isoforms were shown to be amplified, overexpressed, or mutated in various types of cancer. The importance of fine tuning of PKB activity can be displayed in one frame that incorporates PI3K/PKB signaling to pathophysiology of insulin resistance associated with other factors like viral infections and subsequent transition to NAFLD/NASH and later to the development and progression of fibrosis, cirrhosis, and finally HCC. Targeting the PI3K/PTEN/PKB/mTOR pathway in order to alleviate burden of associated diseases seems to be an attractive, and, in some cases, successful approach, that could be exploited in targeted therapy.

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2. Protein kinase B/Akt has three isoforms, PKBa, b, g, which (a) Are found in all species, from worms to humans (b) Have low conservation of amino acid sequence, but similar domain organization (c) Have distinct tissue distribution patterns 3. Full activation of PKB requires (a) Phosphorylation on Ser473 within the hydrophobic motif (b) Phosphorylation on Thr308 in the kinase domain, which changes the conformation of kinase, and is followed by Ser473 phosphorylation (c) Dual phosphorylation on Thr308 and Ser473; it is still arguable whether it happens in parallel or consequently 4. Which of the following statements is wrong? (a) PTEN is a lipid phosphatase that regulates the PI3K/PKB pathway. It is overexpressed in many cancer cell lines and its gain-of-function mutations are the hallmark of different cancers (b) PTP1B and PTEN are negative regulators of the PI3K/PKB pathway; mouse models show that decrease in the levels of PTP1B and PTEN may restore insulin sensitivity (c) PKB activity might be regulated via direct dephosphorylation, via interaction with protein other than phosphatases, via upstream modulation of PI3K activity

Multiple Choice Questions

5. The PKBb isoform is (a) The major isoform that regulates cell growth and proliferation; PKBb KO mice are smaller and have increased neonatal lethality (b) Expressed ubiquitously at similar levels in all tissues (c) Expressed at the highest levels in insulin-­ responsive tissues, and its ablation leads to development of insulin resistance and T2D in mice

1. Protein kinase B/Akt phosphorylates its substrates, which leads to (a) Activation of their function only (b) Inhibition of their function only (c) Activation/inhibition of their function and translocation to a different cellular compartment

Acknowledgements  We thank Arnaud Parcellier, Lana Bozulic, Alexander Hergovich, and Patrick King for their ­critical reading of this manuscript. EZ is the recipient of a Swiss  Bridge fellowship. OT is supported by the Gebert Rüf Foundation (GRS 027/06) and Amélie Waring Foundation. The Friedrich Miescher Institute is part of the Novartis Research Foundation.

16  Role of PKB/Akt in Liver Diseases

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Targeting mTOR Signaling Pathways in Liver Disease

17

Hala E. Thomas and Sara C. Kozma

Introduction

mTOR Signaling Pathways

Over the last 15 years, the Ser/Thr protein kinase mammalian target of rapamycin (mTOR) has emerged as a critical regulator of cell growth, proliferation, apoptosis, and metabolism [1]. The diversity of intracellular responses, in which mTOR is implicated, stems from the fact that it integrates input from distinct signaling effectors (growth factors and nutrients) and acts on specific substrates depending on its ­interaction in multienzyme protein complexes termed mTOR complexes [1]. In addition, mTOR signaling is negatively regulated by several tumor suppressors. In a largely selective manner, the macrolide antibiotic rapamycin inhibits the activity of one of the mTOR complexes, mTOR Complex1 (mTORC1, see below). Rapamycin (sirolimus) and its derivatives (AP23573, CCI-779, and RAD001) are used in the clinic as immunosuppressive agents in organ transplantation and as antiproliferative and antiangiogenic agents to prevent coronary restenosis and treat cancer. Here, we will review the molecular mechanisms known to regulate mTOR signaling pathways, illustrate the role of mTOR signaling in liver disease, and report on the preclinical and clinical studies targeting mTOR to treat liver cancer.

mTOR Complexes

S. C. Kozma () Genome Research Institute, University of Cincinnati, 2180 E. Galbraith Rd, Cincinnati, OH 45237, USA e-mail: [email protected]

At present, mTOR is known to be a component of at least two distinct multiprotein complexes: mTORC1 and mTORC2 (Fig. 17.1). In mTORC1 [2, 3], mTOR is associated with the rapamycin-sensitive adaptor protein of mTOR (raptor, [2, 4, 5]), mammalian lethal with SEC13 protein 8 (mLST8, also known as GbL, [3]), and the proline-rich PKB/Akt substrate 40  kDa (PRAS40, [6]). The role of raptor in mTORC1 is to mediate substrate binding of specific targets through their TOR signaling (TOS) motifs [7]. Little is known about the functional role of mLST8, except that it is required for mTORC1 to respond to nutrient and energy inputs [3]. Unlike raptor and mLST8, PRAS40 acts as a negative effector of mTORC1, whose inhibitory effect is released by PKB/Akt phosphorylation, facilitating mTORC1 activation [6]. mTORC1 mediates the phosphorylation and activation of the 40S ribosomal protein S6 kinases (S6K1 and S6K2) at several key residues, including S6K1 Thr389 and S6K2 Thr388 in the highly conserved hydrophobic motif present in the members of the AGC family of protein kinases [8]. In parallel, mTORC1 also mediates the phosphorylation of the 4E binding proteins (4E-BP1-3) at multiple sites, which acts to release the 4E-BPs from the translation initiation factor 4E, allowing 4E to form a productive initiation complex [9]. The second mTOR complex identified, mTORC2, also contains mTOR and mLST8, but instead of raptor and PRAS40, it ­contains three other proteins: rapamycin independent

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_17, © Springer-Verlag Berlin Heidelberg 2010

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companion of mTOR (rictor [10], also known as mAVO3 [5]); mSin1 [11-14]; and protein observed with rictor (protor, [15-17]). This second mTOR complex is largely resistant to rapamycin, controls actin cytoskeleton dynamics [10] [11] and regulates the phosphorylation of the highly conserved hydrophobic motif PKB/Akt Ser473, which is equivalent to S6K1 Thr389 [14] (Fig. 17.1).

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Activation of mTOR Complexes Growth factors and hormones induce mTORC1 and mTORC2 activation via the canonical class 1 phosphatidylinositide-3OH kinase (PI3K) pathway (Fig. 17.1). The model of insulin-like growth factor (IGF) or insulin-induced mTORC1 and mTORC2 activation involves binding of the ligand to the a subunits of the Insulin / IGF1

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Fig. 17.1  Model depicting mTOR signaling pathway in response to nutrients, hormones, and growth factors. Nutrients such as amino acids and glucose induce mTORC1 signaling through class 3 PI3K (hVps34) activity (see also text). The mTORC2/ mTORC1 signaling pathway is also activated by IGF or insulin through IR activation. Ligand binds the insulin receptor and leads to an increase in tyrosine phosphorylation of IRS1. IRS1associated class 1 PI3K increases the production of PIP3, recruiting PDK1 and PKB/Akt to the plasma membrane. PKB/Akt is activated by the concerted action of the rapamycin-insensitive mTORC2 and PDK1. Activated PKB/Akt phosphorylates and subsequently inactivates TSC2, a GTPase-activating protein, leading to an increase in GTP bound Rheb. Rheb-GTP increases mTORC1 activity and further facilitates the phosphorylation of S6K1 and 4E-BP1. The mTOR signaling pathway can sense

energy levels either directly, through mTORC1, or indirectly, through AMPK. Energy stress, such as fasting or lowering cellular ATP concentration, induces AMPK activation, leading to a PKB/Akt-independent TSC2 phosphorylation, which is thought to activate TSC2. This activation of TSC2 results in an increase in GDP bound Rheb and subsequent decrease in mTOR activity. Growth factors such as EGF or PDGF induce mTOR signaling through ligand binding to its respective receptor activating class 1 PI3K either directly or through the ras small GTPase. Ras also leads to mTORC1 signaling through induction of the RAF, MEK, MAPK, and RSK signaling cascade. MAPK and RSK activation results in TSC2 phosphorylation and inactivation, followed by Rheb GTP activation of mTORC1 signaling. Dashed lines indicate an interaction with an unknown mechanism. Red lines indicate inhibitory signal to the mTOR/S6K1 pathway

17  Targeting mTOR Signaling Pathways in Liver Disease

insulin receptor (IR), a tetramer composed of two a and two b subunits. Insulin binding induces intermolecular autophosphorylation of the two b subunits at specific tyrosine residues. These residues then act as docking sites for proteins containing phosphotyrosinebinding (PTB) domains, including the IR substrates, IRS1 and IRS2 [18]. Studies in mice suggest that IRS1 and IRS2 are largely responsible for controlling insulin signaling events implicated in cellular glucose uptake and metabolism [18]. In the liver, IRS1 appears to be more closely linked to glucose homeostasis whereas IRS2 seems to be more involved in lipid metabolism [19]. In turn, binding of IRS proteins to the phosphorylated IR leads to their phosphorylation at specific tyrosine residues by the activated receptor [20]. These phosphorylated residues serve as docking sites to recruit signaling molecules containing phosphotyrosine-binding Src homology 2 (SH2) domains [21]. One of these IRS tyrosine phosphorylation-docking sites recruits the p85 adapter of the p110 catalytic subunit of the class 1 PI3K [22]. The recruitment of PI3K to the IRS proteins stimulates the production of the lipid second messengers PIP3 (PtdIns (3,4,5) P3) and PIP2 (PtdIns (3,4) P2). Increased production of PIP3 leads to the recruitment of the Ser kinase PKB/ Akt to the membrane through its amino terminal PH (pleckstrin homology) domain. PDK1 (phosphoinositide-dependent protein kinase 1) can then phosphorylate PKB/Akt at the T-loop residue Thr308. In parallel, PI3K activates mTORC2 through an unknown mechanism. Activated mTORC2 phosphorylates PKB/Akt at the hydrophobic motif residue Ser473 [14]. PIP3 and PIP2 production is counteracted by the lipid phosphatase PTEN (phosphatase and tensin homologue deleted from chromosome 10), a tumor suppressor gene which is either absent or mutated in a large number of cancers [23]. Activated PKB/Akt phosphorylates tuberous sclerosis complex protein 2 (TSC2), thus inducing degradation of the tumor-suppressor complex made up of TSC1 and TSC2 (Fig. 17.1). In parallel, PKB/Akt phosphorylates PRAS40, releasing mTORC1 from PRAS40 and, thus, facilitating mTORC1 activation [6]. Also, in tumorigenic settings, the small GTPase Ras in the active GTP-bound state may lead to constitutive signaling through distinct proto-oncogenic pathways (Fig.  17.1). Indeed, the binding of growth factors, such as EGF or PDGF, to their respective receptors leads to the activation of Ras that triggers both the PI3K and RAF kinase pathways.

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Ras in the GTP-bound state binds to and activates RAF, leading to the sequential activation of MEK, MAPK, and RSK (Fig. 17.1). Both MAPK and RSK have distinct substrates; however, both have recently been implicated in the phosphorylation of TSC2 and the activation of mTORC1 signaling [24, 25]. It has also been shown that Ras can directly interact with and activate PI3K [26] leading to the stimulation of PKB/ Akt, which phosphorylates TSC2 at distinct sites and inactivates the TSC tumor complex, as described above. In the active state, TSC1/2, through the GTPaseactivating domain of TSC2, drives the small GTPase Rheb (Ras homolog enriched in brain) into the inactive GDP-bound state [27]. The GTP-bound Rheb acts directly on mTORC1 leading to its activation [28] (Fig. 17.1). Activation of mTORC1 (but not mTORC2) is also mediated by nutrients such as glucose and amino acids (AAs), specifically the branched-chain amino acids (BCAAs) exemplified by l-Leu [29-31]. However, contrary to the prevailing view that AAs induce increased mTOR Complex1 signaling through the canonical class 1 PI3K pathway, it was demonstrated that the effects of AAs are mediated through a novel class 3 PI3K, or human vacuolar protein sorting 34 (hVps34), signaling pathway [29, 30]. Activation of hVps34 by AAs leads to an increase in the production of PI3P, which is known to recruit proteins containing FYVE or PX domains to endosomes to create signaling platforms [32] (Fig.  17.1). AAs, or l-Leu alone, induce a rise in intracellular Ca2+ ([Ca2+]i), which triggers mTORC1 and hVps34 activation. Conversely, blocking the rise in [Ca2+]i ablates both responses [33]. These studies also demonstrate that the rise in [Ca2+]i increases the direct binding of Ca2+/calmodulin to an evolutionarily conserved motif in hVps34, which is required for lipid kinase activity, and increased mTOR Complex1 signaling [33]. In parallel, it was recently reported that mTORC1 activation by AAs involves its recruitment to Rheb/Rab7 vesicles by the GTPase proteins RagA, RagB, RagC, and RagD [34, 35]. Rag GTPases regulate mTORC1 localization as heterodimers of Rag A or B in complex with Rag C or D, with GTP loading of RagA/B favoring mTORC1 activation and GTP-loaded RagC/D favoring mTORC1 inactivation [34, 35]. Rab7 has previously been shown to interact with hVps15, a Ser/Thr kinase that forms a complex with hVps34, and whose protein kinase activity is required for hVps34 activity [32]. Whether Rags

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interact with hVps34 through Rab 7 or not has not been resolved as yet. Independent of the results obtained on the AA-induced pathway, it was also shown that intracellular ATP levels [36] and AMPdependent protein kinase (AMPK) [37, 38], as well as phosphatidic acid [39], regulate mTORC1 activity in a manner similar to that of AAs.

Physiological Effects of mTORC1 Activation Positive Anabolic Effects of mTORC1 Signaling The insulin- and nutrient-induced positive anabolic responses associated with cell growth are mediated by the mTORC1 signaling pathway in large part through the downstream effectors S6K1 and 4E-BP1. This is consistent with the role of the first known substrate of S6K1, ribosomal protein S6, an essential ribosomal protein [40] whose phosphorylation had been implicated in increased rates of protein synthesis [41]. Indeed, recent studies employing mutations of S6 Ser235 and Ser236 to either alanines or aspartic acids showed that phosphorylation of S6 at these sites facilitates its recruitment to the mRNA cap-binding complex [42]. This indicates that phosphorylation of S6 may stimulate assembly of the eIF4F translation–initiation complex [43]. A number of S6K1 substrates have been implicated in the cell-growth response, including translation initiation factor 4B, a 5’ UTR-mRNA helicase [44]; eEF2 kinase, a mediator of the phosphorylation of translation elongation factor 2, involved in controlling protein synthesis transit rates [45]; BAD1, a proapoptotic protein [46]; SKAR, a nuclear protein proposed to couple transcription with pre-mRNA splicing [47]; and, most recently, PDCD4 (programmed cell death protein 4), an inhibitor of translation initiation factor eIF4A [48]. As mentioned above, activation of mTOR signaling also increases mRNA translation via inhibition of the 4E-BPs. Multisite phosphorylation of the translational repressor 4E-BP1 results in its dissociation from eIF4E, thereby allowing eIF4E to assemble with eIF4G, facilitating the recruitment of other translation initiation factors to form the eIF4F complex and initiate cap-dependent translation [49]. It has been

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shown in vivo that BCAA supplementation, or l-Leu alone, increases the mass of insulin-responsive tissues including adipose tissue, liver, and skeletal muscle [50]. This comes to play in patients with chronic liver disease who can develop deleterious energy and protein catabolism and present symptoms of protein-caloric malnutrition [51, 52]. Indeed protein-caloric malnutrition leads to a higher rate of complications in chronic liver disease and, overall, an increased mortality rate such that malnutrition is being considered an independent predictor of survival in cirrhotic patients [53]. The use of BCAA supplementation has been recommended and used with evident success to remedy protein-calorie malnutrition in patients with chronic liver disease [54, 55]. Although a causal relationship has not yet been evidenced, the remediation of protein-calorie malnutrition by BCAAs correlates with activation of mTORC1 signaling. Conversely, the anabolic functions of mTORC1 are also subject to exacerbation, as in the case of nutrient overstimulation (see Sect. 17.4.2) and tumor development following either oncogenic activation or loss of tumor suppression (see Sect.  17.5.1). Thus, the mTORC1 inhibitor agent rapamycin and its derivatives are being tested and, in some cases are already in use, in the treatment of pathologies derived from the deregulation of anabolic functions in several tissues including the liver.

Negative Feedback Effect of mTORC1 Signaling Activation of the mTORC1 pathway has also been implicated in a negative anabolic response involving the suppression of insulin signaling. That is, recent studies have shown that, in the case of insulin signaling, activation of S6K1 affects PKB/Akt by a negative feedback loop from S6K1 to IRS1, which suppresses PI3K activation and, consequently, PKB/Akt activation [56-58]. Similarly, in mouse embryo fibroblasts (MEFs) lacking TSC2, or in mammalian cells overexpressing Rheb, mTORC1 is constitutively activated and PKB/Akt activity is suppressed [27, 59]. As with insulin signaling, AA-induced activation of mTORC1 leads to suppression of PKB/Akt activation through IRS1 phosphorylation [56] independent of mitogens [2, 36, 60, 61]. This effect is reversed by the inhibition

17  Targeting mTOR Signaling Pathways in Liver Disease

of mTORC1 by rapamycin, suggesting that the AA inhibitory response is mediated by mTORC1 signaling and S6K1 activation [62]. Notably, in insulin-resistant states of obesity, circulating concentrations of AAs are elevated, particularly BCAAs [63-65] and infusion of AAs in humans, similar to what is observed for lipids, induces insulin resistance in experimental settings [66,  67]. The understanding that mTORC1 activation is implicated in this insulin resistance response comes from the following observations: First, overexpression of a kinase-dead S6K1, but not wild-type S6K1, blocks Rheb-induced PKB/Akt activation, showing that these effects are mediated by S6K1 [58]. Second, insulin resistance, induced in rats by chronic insulin treatment (hyperinsulinemic-euglycemic clamp), is reversed by the administration of rapamycin, an effect that is paralleled by a loss of IRS1/IRS2 phosphorylation and S6K1 inactivation [68]. And third, phosphorylation of a recently identified novel S6K1 phosphorylation site in IRS1 is blocked by siRNAs directed against S6K1, and mutation of the site to alanine potentiates IRS1 tyrosine phosphorylation and PKB/Akt activation [69]. Thus, the mTOR signaling pathway is intimately involved in controlling protein translation and in inducing a negative anabolic effect by downregulation of insulin signaling in response to elevated nutrient concentrations [70].

mTORC1 Regulation in Liver Disease Liver Regeneration In response to acute liver injury, hepatic cells compensate parenchymal liver loss by cell growth and proliferation. A highly exploited model to study liver regeneration is partial hepatectomy in mice or rats, where two thirds of the liver is removed and the remaining liver cells re-enter the cell cycle to replace the lost liver mass. Following partial hepatectomy in the rat, liver cells upregulate ribosome biogenesis by increasing the translation and transcription of ribosomal protein genes as well as the transcription of rRNA genes [71, 72]. It has been shown that the translation of mRNAs encoding ribosomal proteins, all of which contain a 5′-TOP (5′-terminal oligopyrimidine tract), is selectively upregulated during liver regeneration

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[71]. The 5′-TOP mRNAs are characterized by a stretch of pyrimidines at their 5′ end of an average length of 10 nucleotides, and invariably starting with a C [73]. The 5′-TOP sequence is a negative regulator of ribosomal protein mRNA translation, as replacement of pyrimidines with purines results in the constitutive translation of transcripts [74, 75]. Like liver cells following partial hepatectomy, cells in culture also reveal the selective upregulation of 5′-TOP mRNAs translation when stimulated to proliferate. It was shown that rapamycin treatment prevents translational upregulation of 5′-TOP mRNAs in resting cells stimulated with serum [76, 77]. The inhibitory effects of rapamycin were determined to be mediated through the 5′-TOP because the translation of transcripts bearing a mutated 5′-TOP was insensitive to rapamycin treatment [75]. These data are consistent with the finding that, in the livers of l-Leu–fed rats, the mTOR pathway and translation of 5′-TOP mRNAs are activated and rapamycin treatment prevents both responses [78]. Like the translation of ribosomal proteins, transcription of rRNA genes has also been shown to respond to both partial hepatectomy and nutrient availability [79]. The link between transcription of rRNA and nutrients suggests that mTOR, in addition to regulating translation of ribosomal proteins, may also regulate the activity of RNA polymerase I (Pol I). Indeed, mTORC1 most likely regulates Pol I transcription by modulating the activity of its essential cofactor TIF-IA [79]. Rapamycin-induced inhibition of mTOR signaling inactivates TIF-IA by regulating its activity, cellular localization, and phosphorylation state [79]. It remains to be determined as to what extent TIF-IA regulation is influenced by growth factors or nutrients, as well as by liver regeneration following insult or injury. Although the mTOR pathway is activated during liver regeneration, inhibition of mTOR signaling by rapamycin only slows down the regeneration process [80]. Globally, the effect of rapamycin treatment on liver regeneration can be divided in two steps. In the early phase after partial hepatectomy (1–4 days), rapamycin slows down liver regeneration by decreasing the proliferation rate of both hepatocytes and nonparenchymal cells [80, 81]. However, starting from day 7, there is no significant difference in the proliferation of these cells relative to control cells. Liver regeneration is completed by day 14 in rapamycintreated rats, which exhibit no noticeable differences relative to the controls [81]. The transient nature of

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this effect might justify further exploration of the use of mTOR inhibitors in clinical situations that involve liver regeneration.

Metabolic Disorders of the Liver Liver disease of metabolic origin associated with obesity and type 2 diabetes is now recognized as the most prevalent liver disease in Western societies [82, 83]. Because mTORC1 integrates input from nutrients and hormones, its activity is affected by the state of nutrient overload linked to metabolic disorder [70]. Indeed, elevated nutrient concentrations activate mTORC1 and induce the feedback loop, thus decreasing insulin sensitivity in the peripheral tissues and affecting liver physiology [56, 70]. Insulin resistance is commonly observed with obesity. Obese rats on a high-fat diet (HFD) have an increase in mTOR and S6K1 activity in the liver, which potentially mediates insulin resistance in this organ [84]. Double 4E-BP1 and 4E-BP2 knockout mice have higher S6K1 activity in the liver, muscle, and adipose tissue relative to control mice [85]. This results in insulin resistance due to the increased negative feedback of S6K1 on IRS-1. When subjected to HFD, these mice show an increase in liver weight and hepatic triglyceride accumulation, which leads to steatosis [85]. In contrast, S6K1−/− mice have enhanced insulin sensitivity and oxidative phosphorylation, which protects them against diet-induced obesity [56]. Insulin-induced PKB/Akt phosphorylation is suppressed in fat, liver, and muscle of wild-type mice on a HFD, whereas no significant difference is detected in these tissues of S6K1−/− mice [56]. Likewise, liver-specific overexpression of a dominant-negative form of raptor that harbors a C-terminus deletion of the protein rescues insulin resistance in a K/KAy genetic mouse model of obesity and increases both basal and insulininduced phosphorylation of PKB/Akt in the liver [86]. Moreover, rats fed a HFD supplemented with glutamine develop insulin resistance in adipose tissue [87]. This leads to a decrease in the fat mass, thereby causing an increase in insulin sensitivity in the liver and muscles, which had been rendered insulin resistant by the obese status of the animals. In humans, nonalcoholic fatty liver diseases (NAFLD) ranging from benign steatosis to nonalcoholic steatohepatitis (NASH) are strongly linked with

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an underlying insulin resistant state that leads to fat deposition within the hepatic parenchyma. White adipose tissue (WAT) is potentially an important player in the development of this disease [88]. Preliminary results from phosphoproteomic profiles of WAT biopsies from patients with simple steatosis show an increase in the phosphorylation levels of proteins in the insulin pathway, relative to patients with NASH [88]. In addition, it was recently shown that hepatic steatosis can be mediated by alterations of PTEN expression in hepatocytes exposed to high levels of unsaturated fatty acids. Furthermore, the data revealed an interaction between mTOR and NF-kb, suggesting cross-talk between these two pathways [89]. The role of the mTOR signaling pathway in liver pathologies is not limited to metabolic disorders, as mTOR plays a role in liver cancer as well. Indeed, metabolic disorders, over time, increase liver injury and can lead to cancer development. With the increase in nutrient availability and more sedentary lifestyles, the number of obese and diabetic patients is continuously on the rise and has been linked to a 30% increase in the risk of hepatocarcinoma (HCC) development with poor prognosis [83, 90, 91]. Treatment with insulin sensitizers such as metformin, as opposed to treatment with insulin or sulfonylurea, is proposed to improve the metabolic status of these patients while protecting against HCC development [92].

Upregulation of mTOR Signaling in Human HCC Immunohistochemical analyses of HCC samples obtained either from primary liver tumor tissue [9396], xenografts of tumor tissue in SCID mice [97], or human HCC cell lines [98] revealed an increase in the phosphorylation level of mTOR pathway components such as S6K1, 4E-BP1, PKB/Akt, and S6 relative to normal liver tissue. The reports are conflicting as to the clinical relevance of S6K1 phosphorylation in predicting prognosis, survival, and tumor grade [93-95]. However, the discrepancies among these results could be explained by the fact that they were obtained with different antibodies and in most case without ascertaining whether the antibodies specifically recognized activated S6K1. Notably, a recent study reported that an increase in S6 phosphorylation in human HCC tumors

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relative to the surrounding normal or cirrhotic liver tissue is a predictor of poor prognosis [96]. In addition, a correlation was found between disease severity and amplification of rictor DNA, paralleled by an increase in rictor mRNA transcript levels. The data also revealed that risk of tumor recurrence increases in patients who have a concomitant gain in rictor DNA and increased S6 phosphorylation [96]. The negative feedback loop through S6K1 is a potential explanation for the low level of PKB/Akt Ser473 in samples with high S6 phosphorylation [96]. Elevated S6 activity in tumors correlates with an increase in growth factor signaling through EFG and IGF-I in two-thirds of the cases studied [96]. By using microarrays to identify signaling pathways that are altered in human HCC samples with high S6 activity, a number of genes involved in inflammation (NFkb), angiogenesis (VEGFB, VEGFC), transcription, calcium signaling (PLA2G4A), and proliferation were found to be upregulated [96]. Taken together, the data implicate mTOR signaling in HCC progression, but the contribution of mTORC1 activation versus mTORC2 activation is still unclear. While awaiting a better mechanistic understanding, mTOR inhibitors continue to be tested for HCC in both preclinical and clinical settings.

Pharmacological Inhibition of mTOR in HCC Preclinical Studies in Animal Models of HCC as Tools to Decipher Molecular Mechanisms and Clinical Relevance It was reported that oral administration of mTOR inhibitors, such as RAD001 or rapamycin, to SCID mice bearing 0.1  cm3 subcutaneous implants of patient-derived HCC cell lines results in a delay in tumor growth rate and tumor volume for the duration of the treatment [97, 99]. Microvessel density and VEGF expression in the RAD001-treated tumors also decreased relative to control tumors, while no effect was noted on apoptosis, as measured by the amount of cleaved caspase-3 [97, 99]. Combination therapy with a monoclonal antibody against VEGF (bevacizumab), in the same mouse model, resulted in further inhibition of angiogenesis and tumor growth suppression

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relative to treatment with each drug alone [97]. Moreover, a rat orthotopic HCC model involving p53deficient tumor cells revealed a selective effect of combined mTOR inhibition and the anthracyclin doxorubicin [100]. In nude mice with orthotopic implants of a patient-derived metastatic HCC line, treatment with rapamycin and a Raf-specific inhibitor (sorafenib also known as nexavar) resulted in a significant decrease in tumor size, proliferation, and CD31 blood vessel staining compared to single-drug treatment [101]. However, the apoptotic rate in the rapamycintreated mice did not increase further when sorafenib was used in combination [101]. A caveat of these xenograft and orthotopic implant models [96, 97, 99, 101, 102] is that, while they allow for comparisons of chemotherapeutic potency, they do not account for the contribution of important elements in HCC development and progression such as inflammation and cirrhosis. A number of mouse models of HCC have been obtained by chemical carcinogen induction or by genetic engineering and offer the advantage of reproducible hepatocarcinogenic patterns. The question that arises is the extent to which they recapitulate the development of HCC in humans. To assess this point, comparative functional studies were performed to identify “best-fit” mouse models of human HCC obtained from five transgenic or knockout mouse lines and two models involving induction by the chemical carcinogens diethylnitrosamine or ciprofibrate [103]. Gene expression profiles of 68 HCCs from these mouse models were compared to those of 91 human HCCs, which had been divided into two groups depending on the prognosis of the disease [103]. This approach led to a number of key observations. The gene-expression profiles derived from mice treated with ciprofibrate or in which the acox-1 gene had been deleted did not correspond to gene expression profiles of either of the two classes of human HCC suggesting that hepatocarcinogenesis in humans is not likely to be driven by peroxisome proliferators [103]. Secondly, the gene expression profiles corresponding to human HCC with poor prognosis (<3 years survival) were best represented by transgenic mice overexpressing c-Myc in combination with TGFa, or by mice in which tumors were induced by DEN, whose profiles were characterized by the upregulation of genes involved in cell growth and proliferation. Finally, the gene-expression profiles derived from transgenic

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mouse models in which either c-Myc and/or E2F1 was overexpressed corresponded more closely to the gene expression profiles of human HCCs with better prognoses [103]. Mice overexpressing both E2F1 and c-Myc developed liver tumors in which the c-Myb/ COX-2 and PI3K/PKB/Akt/mTOR pathways were upregulated, which is similar to human HCC tumors [98]. The molecular mechanism underlying HCC development in this model is due to E2F1 acting as a suppressor of c-Myc–driven apoptosis [98].

mTOR Inhibitors in Clinical Trials for HCC Because of the promising effects of RAD001 or sirolimus in prolonging disease-free survival of patients with solid tumors, such as metastatic renal small cell carcinomas [104], breast cancers [105], and pancreatic neuroendocrine tumors (PET) [106], mTOR inhibitors, alone or in combination with other therapies, are being tested in various phases in clinical trials [107, 108]. Some of the main adverse effects of the treatment are stomatitis/mouth ulcers, fatigue, rash, thrombocytopenia, hyperglycemia, and hyperlipidemia [104-107, 109]. In some cases, there is also an increase in noninfectious grade 3 pneumonitis [104, 106]. Pharma­codynamic and pharmacokinetic studies conducted in patients to determine the optimal welltolerated dosage regimen of RAD001 showed that daily dosing was a more efficacious antitumor regimen than a higher once-weekly dose [108, 110]. With an average half-life of around 30 h in patients, steadystate serum levels were reached within a week of daily dosing at 5–10  mg, and by the second week in a weekly regimen of 20  mg [107]. S6K activity in peripheral blood mononuclear cells was inhibited for an entire week after administration of a weekly 30 mg dose of RAD001, but for only 48  h after a lower weekly dose of 5 mg [107]. Phase I and II trials of mTOR inhibitors alone or in combination with multikinase inhibitors, such as sorafenib, or with bevacizumab, are ongoing for patients with advanced HCC. A once-daily dose of rapamycin given to 21 patients with advanced HCC resulted in stable disease for at least 5 months in five patients and partial remission in one case as evaluated, primarily, by CT or MRI scans and serum AFP levels

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[111]. No overall serious complications were reported from the therapy except for the development of aphthous ulcers [111]. In a tumor setting, treatment with rapamycin is predicted to lead to the upregulation of PKB/Akt, potentially enhancing tumor survival. Activation of PKB/Akt is known to drive cell growth through increased transcription and translation and to prevent apoptosis by the phosphorylation of a number of key proteins, including BAD and the forkhead and NF-kB transcription factors. In the case of BAD and the forkhead transcription factors, phosphorylation by PKB/Akt blocks their proapoptotic responses. In contrast, PKB/Akt phosphorylation of NF-kB enhances NF-kB-mediated transcription of many antiapoptotic genes [112]. Moreover, activation of PKB/Akt is known to drive glucose uptake and glycolysis, which can protect against apoptosis by maintaining mitochondrial integrity. However, there is, so far, no clear clinical evidence of more aggressive tumor growth due to treatment with mTOR inhibitors, which suggests that other mechanisms exist to counteract the negativefeedback loop that activates PKB/Akt, or that mTOR pathway activation is necessary to drive PKB/Aktdriven tumorigenesis mediated by activation of downstream PKB/Akt substrates.

Conclusion The mTORC1 signaling pathway can play a positive anabolic role in cell growth and proliferation through the upregulation protein synthesis and ribosome biogenesis, but it can also play a negative role through suppression of insulin and growth factor signaling. The importance of both of these roles is underscored in liver pathologies. Solely targeting the mTOR pathway by using inhibitors has resulted in mainly cytostatic effects [1] and is insufficient for treatment of advanced HCC. Nonetheless, the relative tolerability and immunosuppressive properties of mTOR inhibitors coupled with their antiangiogenic and antiproliferative roles puts them at the forefront in the current line of treatments. With this perspective, mouse models are currently being tested for their value in predicting clinical outcome of these therapies. Also, trials combining other therapies with mTOR inhibitors are being tested for their ability to prolong tumor-free survival in patients with HCC.

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Summary

›› The Ser/Thr protein kinase mTOR is a critical ››

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regulator of cell growth, proliferation, apoptosis, and metabolism. The diversity of intracellular responses in which mTOR is implicated stems from the fact that it integrates input from distinct signaling effectors (growth factors and nutrients) and acts on specific substrates depending on its interaction in multienzyme protein complexes termed mTOR complexes. In mTORC1, mTOR is associated with raptor, mLST8 (also known as GbL), and PRAS40; mTORC1 mediates the phosphorylation and activation of the 40S ribosomal protein S6 kinases. The macrolide antibiotic rapamycin inhibits the activity of mTORC1. In mTORC2, mTOR is associated with mLST8, rictor, mSin1, and protor. mTORC2 is largely resistant to rapamycin. mTORC2 controls actin cytoskeleton dynamics and regulates the phosphorylation of the highly conserved hydrophobic motif PKB/Akt Ser473. Growth factors and hormones induce mTORC1 and mTORC2 activation via the canonical class 1 phosphatidylinositide-3OH kinase (PI3K) pathway. Activation of mTORC1 (but not mTORC2) is also mediated by nutrients such as glucose and AAs, specifically the BCAAs. mTOR signaling is negatively regulated by several tumor suppressors. The insulin- and nutrient-induced positive anabolic responses associated with cell growth are mediated by the mTORC1 signaling pathway in large part through the downstream effectors S6K1 and 4E-BP1. In the case of insulin signaling, activation of S6K1 affects PKB/Akt by a negative feedback loop from S6K1 to IRS1, which suppresses PI3K activation and, consequently, PKB/Akt activation. Rapamycin (sirolimus) and its derivatives (AP23573, CCI-779, and RAD001) are used in the clinic as immunosuppressive agents in organ transplantation, as antiproliferative and antiangiogenic agents, to prevent coronary restenosis, and to treat cancer.

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Multiple Choice Questions 1. Inhibition of mTOR complex 1 can lead to PKB/ Akt activation: (a) As a result of the inhibition of the negative feedback loop through S6K1 (b) Due to transcriptional downregulation of raptor (c) Due to an increase in 4EB-P1 phosphorylation which leads to an increase in translation (d) Due to down-regulation of PTEN (e) All of the above statements are true 2. The negative anabolic effects of mTOR in the liver: (a) Involve the suppression of insulin signaling in response to elevated nutrient concentrations (b) Are irreversible by rapamycin treatment (c) Require a decrease in S6K1 activity (d) Are mediated by mTORC2 (e) All of the above statements are true 3. mTORC2 consists of the following proteins in addition to mTOR: (a) Rictor, mSin1, mLST8, and protor-1 (b) Rictor, PRAS40, mLST8, and protor-1 (c) Rictor, raptor, mLST8, and protor-1 (d) Rictor, mSin1, mLST8, and PRAS40 (e) PRAS40, mLST8, and raptor 4. All the following statements are true except (a) The mTOR pathway and translation of 5’-TOP mRNAs are activated in the liver of L-Leu–fed rats (b) mTOR activity in the liver is regulated only by BCAAs and not by insulin (c) Human HCC samples have an increase in the phosphorylation level of mTOR pathway proteins such as S6K1, 4E-BP1, PKB/AKT, and S6 relative to normal liver tissue (d) Rapamycin treatment results in both antiangiogenic and antiproliferative effects (e) Inhibition of mTOR signaling by rapamycin slows down the regeneration process 5. The activity of mTORC1 is affected by: (a) Rapamycin and its derivatives (b) Nutrients such as glucose and BCAAs (c) ATP (d) Phosphatidic acid (e) All of the above

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AMP-Activated Protein Kinase in Liver

18

Louis Hue, Laurent Bultot, and Mark H. Rider

The AMP-activated protein kinase (AMPK) has become an inescapable topic in metabolic disorders. AMPK is a highly conserved eukaryotic protein serine/threonine kinase, which is involved in energy homeostasis both at the cellular and whole-body levels [1–3]. It mediates a nutrient signaling pathway that senses the energy status of the cell and is activated by energy imbalance as a result of decreased ATP production (hypoxia) or increased ATP consumption (muscle contraction). Liver AMPK activity is also modulated by cytokines that regulates whole-body energy homeostasis. At the cellular level, AMPK restores energy charge by switching off anabolic ATP-consuming pathways, while switching on catabolic ATP-generating pathways. It also plays a critical role in systemic energy homeostasis by controlling food intake in the hypothalamus [1–3]. Therefore, AMPK orchestrates a global metabolic response to energy deprivation and can be regarded as a starvation signal. This short article reviews the conditions and consequences of AMPK activation in liver and the potential implication of AMPK in liver diseases.

The AMP-Activated Protein Kinase (AMPK) AMPK is a heterotrimeric complex comprising a catalytic (a) and two regulatory (b and g) subunits. Each subunit has multiple isoforms (a1, a2, b1, b2, g1, g2, g3)

L. Hue () Hormone and Metabolic Research Unit, Université catholique de Louvain, and de Duve Institute Avenue, Hippocrate 75 UCL 7529, B-1200 Brussels, Belgium e-mail: [email protected]

giving twelve possible combinations of holoenzyme with different tissue distribution and subcellular localization [2]. Immunological analysis of liver AMPK complexes indicated that a1 and a2 are equally abundant in rat liver but that a1 is the predominant isoform in human liver; b1 and g1 are the most abundant regulatory subunits, while the other regulatory subunits are present in less than 10% of the complexes ([4]; Guigas and Hue, unpublished results). The N-terminus of the a subunit contains a protein kinase domain. The b subunit acts as a scaffold for the other two subunits. It also contains a glycogen binding domain, whose physiological role remains to be clarified. The g-subunit contains four AMP-binding domains. Crystal structure revealed that two sites contain AMP or ATP, whereas a third site binds a non-exchangeable nucleotide [5]. AMPK can be activated by changes in the intracellular AMP:ATP ratio, as occurs under metabolic stresses leading to ATP depletion [2] or via an increase in intracellular Ca2+ concentration [3, 6]. The upstream kinases, LKB1 (Peutz-Jeghers protein) and calcium/calmodulindependent protein kinase kinase–b (CaMKKb) activate AMPK by phosphorylating Thr172 in the activation loop of the catalytic a-subunit [3,6]. AMP not only allosterically stimulates AMPK activity by binding to the g-subunit, but also prevents Thr172 dephosphorylation by protein phosphatases [7]. Since LKB1 is constitutively active, a rise in AMP leads to AMPK activation by increasing Thr172 phosphorylation. The discovery that CaMKKb could also activate AMPK, independently of AMP, has broadened the potential roles of AMPK in the control of processes that do not necessarily involve a change in cellular energy status [8]. However, the importance of CaMKKb in the control of liver AMPK activity is probably limited, because (1) LKB1 rather than CaMKKb is expressed in liver, and (2) agents such as phenylephrine and vasopressin,

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_18, © Springer-Verlag Berlin Heidelberg 2010

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known to increase cytosolic Ca2+ in hepatocytes, do not affect AMPK activity (our unpublished results). More recently a third AMPK-kinase, the transforming growth factor-b-activated kinase (TAK1), has been found to phosphorylate in vitro AMPK Thr172 [9]. It is not known whether TAK1 has any role to play in AMPK activation in liver or in any other system. It could participate in hepatoblast differentiation, which is in part mediated by transforming growth factor-b [10].

AMPK Activation in Liver Any imbalance between ATP production and consumption activates AMPK via the LKB1 pathway by an increase in the AMP/ATP ratio. Experimentally, such conditions are readily obtained in hepatocytes with metabolic poisons, mitochondrial inhibitors, oxidative or osmotic stress, oxygen or nutrient deprivation, or substrates that decrease ATP by accumulation of phosphate ester and depletion of Pi (e.g., high fructose concentration) [reviewed in [8, 11]]. These can be regarded as abnormal or pathological stresses, because under normal conditions the liver maintains its ATP by adapting ATP supply to metabolic demand. However, liver AMPK can be activated under more physiological conditions, for example, during exercise [12], starvation [13], or in response to cytokines such as adiponectin [14] and IL-6 [15]. The mechanism of AMPK activation by these conditions has not been entirely elucidated. Several substances and drugs activate AMPK. 5-Amino-4-imidazole-carboxamide (AICA) riboside is an analog of adenosine that is phosphorylated in certain cells to AICA ribotide or ZMP, which mimics AMP. AICA riboside has been extensively used in cells or even in vivo to activate AMPK [16–18]. However, several effects of AICA riboside, including the inhibition of liver glucose uptake and of mitochondrial respiration, are independent of AMPK [19, 20]. In addition, AICA riboside decreases intracellular ATP [16, 21, 22], and intracellular ZMP accumulation may directly affect the activity of enzymes that bind AMP, such as glycogen phosphorylase and fructose-1,6-bisphosphatase. Therefore, because of its AMPK-independent effects, AICA riboside is not well suited to investigate the consequences of AMPK activation on cell function. Other more specific tools and/or new pharmacological compounds with AMPK selectivity are required. The development of a new AMPK activator (the Abbott compound, A-769662) is of obvious interest [21, 23, 24].

L. Hue et al.

Metformin, the most prescribed drug for type 2 diabetes, restores glucose homeostasis by increasing glucose disposal and decreasing hepatic glucose production. Although the molecular mechanism of metformin action has long remained elusive [11], experimental evidence now places metformin at the controls of energy homeostasis [103]. Metformin, indeed, induces a mild and specific inhibition of the mitochondrial respiratory chain complex 1 [25, 26] that results in slight but significant decrease in ATP so leading to AMPK activation [27, 103]. Metformin enters the cells via the organic cation transporter (Oct1) [28] and metformin pharmacokinetics is influenced by genetic variations in the expression of this transporter [29]. Interestingly, Oct1 expression is elevated in liver and kidney [30] and its deletion decreased the metformin-induced inhibition of gluconeogenesis [31]. These observations may explain why metformin accumulates and acts preferentially in liver. Thiazolidinediones (TZD), another class of antidiabetic drugs, bind PPAR-g (peroxisome-proliferatoractivated receptor g) and increase insulin sensitivity. Beside their long-term action on lipid metabolism mediated by PPAR-g, TZD also activate AMPK through an increase in AMP/ATP ratio, which could result from an inhibition of mitochondrial respiration [32–35] or from increased adiponectin levels [36]. SIRT1 is a member of the sirtuin family of NADdependent histone/protein deacetylases, which regulate acetylation of lysine residues on histones and transcription factors. SIRT1 is induced by nutrient deprivation and seems to be responsible for longevity by caloric restriction [37]. A mechanism connecting SIRT1 with AMPK has recently been described. LKB1 deacetylation by SIRT1 influences LKB1 intracellular localization, kinase activity and ability to activate AMPK [38]. This indirect mechanism has been suggested for AMPK activation induced by glucose deprivation and resveratrol, a polyphenol constituent of red wine that improves survival in mice kept on a high-caloric diet [38–40]. SIRT1 may thus function as an upstream regulator of the LKB1/AMPK axis and control hepatocyte lipid metabolism [39].

Metabolic Effects of AMPK in Liver Once activated, AMPK decreases ATP-consuming pathways and stimulates ATP-generating processes, in line with the concept that AMPK acts as a metabolic master

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switch that conserves ATP [41]. The regulation involves phosphorylation by AMPK of key metabolic enzymes and transcription factors. In general, lipid, glucose, and protein synthesis, as well as cell growth, are inhibited, whereas fatty acid oxidation and glucose utilization are stimulated [8, 41]. Use of AMPK activating agents and results from animal models have allowed clarification of liver specific effects, which are summarized below (reviewed in [42, 43]).

Glucose Metabolism AMPK activation inhibits hepatic glucose production, explaining the hypoglycaemic effects of metformin, AICA riboside and adiponectin [27, 44–46]. The involvement of liver AMPK in the anti-diabetic effect of metformin is supported by the fact that the blood glucose lowering effect of the compound was abolished in liver specific LKB1-deficient mice [47]. However, the evidence is not entirely convincing, because AMPK is not the sole substrate of LKB1, which also phosphorylates at least twelve other AMPK-related kinases [48], one of which, SIK (salt-activated kinase), is known to inhibit gluconeogenesis [49]. On the other hand, the hypoglycemic effect of AICA riboside is decreased in mice whose livers are deficient in both catalytic subunits of AMPK [42]. Similarly, the hypoglycemic effect of adiponectin seems to require liver AMPK [14,  44]. Taken together, these results confirm the role of liver AMPK in the control of glucose production. There is no simple explanation for the short-term inhibition of gluconeogenesis by AMPK, which is paradoxical, because the other concomitant effect of AMPK to enhance fatty acid oxidation is known to stimulate gluconeogenesis via acetyl-CoA, a known stimulator of the gluconeogenic enzyme pyruvate carboxylase. In the long-term, however, decreased expression of two gluconeogenic enzymes, phosphoenolpyruvate carboxykinase and glucose-6-phosphatase, is involved. This effect is probably mediated by phosphorylation of TORC-2 (the transducer of regulated CREB-binding protein 2), a transcriptional co-activator of CREB (cAMP-response element-binding protein), which controls the expression of the gluconeogenic enzymes. Indeed, TORC-2 phosphorylation by AMPK, or by SIK, sequesters TORC2 in the cytoplasm and prevents it from co-activating CREB [49]. In addition, AMPK directly phosphorylates another gluconeogenic transcription factor, HNF4a (hepatocyte nuclear factor 4a, thereby decreasing its activity and

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promoting its degradation [50, 51]. Lastly, AMPK also inhibits the glycolytic and lipogenic gene induction program induced by glucose and insulin [52, 53]. It decreases the expression of SREBP1c (sterol regulatory elementbinding protein 1c) and ChREBP (carbohydrate response element-binding protein), which mediate the transcriptional regulation of glycolytic and lipogenic genes by glucose and insulin [54–57]. In addition, AMPK directly phosphorylates and inactivates ChREBP [58], and overexpression of a constitutively active AMPK in liver of obese ob/ob mice, restores a normal expression of these genes [42, 46, 55]. AMPK activation in liver does not stimulate glucose uptake, as it does in muscle [59, 60]. The unexpected inhibition of liver glucose uptake by AICA riboside is independent of AMPK and results from ATP depletion and glucokinase sequestration in the nucleus [19]. AMPK activation also inhibits glycogen synthesis by phosphorylating and inactivating liver glycogen synthase (Bultot, unpublished results), as is the case for muscle glycogen synthase [61]. The physiological significance of these changes, which may be related to exercise, warrants further investigation.

Lipid Metabolism AMPK inhibits fatty acid synthesis in liver through phosphorylation and inactivation of acetyl-CoA carbox­ ylase (ACC1). As a consequence, malonyl-CoA concentration falls, thereby relieving inhibition of the carnitine-mediated import of long chain acyl-CoAs into mitochondria and eventually stimulating ATP production through their oxidation. By contrast, inactive AMPK and active ACC1 favor lipogenesis. Therefore, AMPK controls liver fatty acid partitioning between oxidative and lipogenic pathways. The fasted to fed transition reflects these activity changes and corresponds to a progressive inhibition of fatty acid oxidation and ketogenesis, together with fatty acid and triglyceride synthesis [13]. In line with this concept, AMPK deficient mice are hypertriglyceridemic, whereas, AICA riboside administration to obese rats decreased their hypertriglyceridemia [44, 45, 62]. AMPK activation also inhibits cholesterol synthesis by phosphorylating and inactivating hydroxymethyl­ glutaryl-CoA reductase [17]. In AMPKa2 deficient mice, blood levels of cholesterol are normal [42], suggesting that the remaining AMPKa1may compensate. Alternatively, control of liver cholesterol synthesis

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depends more on HMGCoA reductase content than on its phosphorylation state. Indeed, the nutritional and hormonal control of lipid metabolism mediated by SREBP1c, ChREBP, and PPARs is probably crucial in the long-term [54, 56, 57, 63].

Mitochondrial Effects of AMPK Since AMPK is involved in cellular energy homeostasis, one may wonder whether it regulates mitochondrial respiratory chain and ATP production. This does not seem to be the case in the short-term. It does ­however stimulate mitochondrial biogenesis through up-regulation of PGC1a (peroxisome-proliferatoractivated receptor-(co-activator 1a)) [64–67]. Livers of mice deficient in both the catalytic subunits of AMPK have reduced mitochondrial biogenesis, as suggested by decreased expression of PGC1a, cytochrome c oxidase I and IV, and cytochrome c genes [20]. Hepatocytes from these mice do indeed display reduced mitochondrial respiration and contain less ATP than controls [29]. In addition, resveratrol, which is known to activate AMPK, increases mitochondrial number in liver [68]. Clearly, this long-term effect on mitochondrial biogenesis and function contributes to cellular energy homeostasis by AMPK.

Protein Synthesis AMPK activation inhibits protein synthesis and cell growth, both of which are energy-requiring processes. Control of protein synthesis is complex and exerted at initiation and elongation via the (de)phosphorylation of various translation factors and ribosomal proteins, including the mammalian target of rapamycin complex 1 (mTORC1) [69]. mTORC1 activation leads to phosphorylation of the ribosomal protein S6 (rpS6), via phosphorylation and activation of p70 ribosomal protein S6 kinase (p70S6K), and this seems to be required for cell growth. mTORC1 activation also enhances translation initiation of capped mRNAs by phosphorylation of eukaryotic initiation factor-4E-binding protein 1 (4E-BP1). AMPK activation inhibits mTORC1 signaling, possibly via phosphorylation of the tuberous sclerosis

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complex (TSC1-TSC2) [70], which potentiates its ability to inhibit mTORC1. TSC2 acts as a GAP (GTPase activating protein) for the G protein Rheb (Ras homologue enriched in brain), which in its GTP-bound state activates the mTOR kinase. AMPK has also been reported to directly phosphorylate mTOR [71] as well as the mTOR binding partner raptor, as recently reported [72]. Translation elongation is inhibited via the phosphorylation of eukaryotic elongation factor-2 (eEF2) by a dedicated calcium/calmodulin-dependent kinase called eEF2 kinase (eEF2K) [69]. AMPK activation in hepatocytes inhibits protein synthesis as a result of eEF2 phosphorylation via the phosphorylation-induced activation of eEF2K [73]. In these cells, AMPK activation does not decrease mTORC1 signaling, because mTORC1 is already inactive in the absence of insulin or amino acids [73, 74]. Taken together, these observations support the concept that AMPK activation inhibits anabolic ATPconsuming processes including protein synthesis and cell growth.

Non-Metabolic Effects of AMPK Although the primary targets of AMPK are mainly involved in energy homeostasis, converging new evidence indicates that AMPK also controls non-­metabolic processes. AMPK activation controls cytoskeleton organization and cell movement, it suppresses cell ­proliferation and growth and can even trigger apoptosis [8]. This anti-proliferative action of AMPK is mediated by a number of tumor suppressors that are part of  the AMPK signaling pathway, such as LKB1 and TSC1-TSC2 [75, 76]. Indeed, AMPK activation causes cell arrest at G1/S transition by phopshorylation of p53 and p27, independent of mTORC1. This leads to an induction of a p53-dependent metabolic checkpoint to stop cell-cycle progression during starvation [77, 78]. Moreover, apoptosis is induced in hepatocytes submitted to sustained and persistent AMPK activation [79]. In addition, AMPK activation in epithelial cells has also been shown to control tight junction formation and cell migration through reorganization of the actin cytoskeleton ([80, 81]; Horman, Hue and Rider 2008, unpublished results). These observations suggest that AMPK integrates multiple environmental signals to check the availability

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of resources before launching energy-consuming processes and that dysfunction of the LKB1/AMPK axis could lead to metabolic disorders or uncontrolled cell proliferation.

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fail to respond to adiponectin by an increased AMPK phosphorylation [87], suggesting that AdipoR1 mediates adiponectin-induced hypoglycemia. Taken together, these results demonstrate that activation of liver AMPK, either by adiponectin or therapeutic agents, induces a beneficial hypoglycaemic response in these metabolic disorders.

AMPK in Liver Diseases Type 2 Diabetes

Liver Steatosis

As AMPK plays a crucial role in energy homeostasis, its deficiency is expected to cause energy imbalance with excessive storage of nutrient and/or failure to mobilize these stores. This rationale explains the current interest for AMPK in the origin and treatment of Type 2 diabetes, obesity, and the metabolic syndrome. In early type 2 diabetes, insulin resistance leads to hyperglycemia as a result of increased hepatic glucose production and decreased glucose uptake in peripheral tissues. These metabolic disorders are mimicked in mice lacking liver a2 AMPK subunit, which are indeed hyperglycemic and glucose intolerant in the fasted state with an increased hepatic glucose production that is not corrected by adiponectin infusion [42, 44, 62]. Although impressive, these results do not demonstrate that type 2 diabetes results from a deficiency or impairment of AMPK activation. A number of reports support the implication of AMPK in the mechanism of action of anti-diabetic drugs and suggest that AMPK activation could correct  hyperglycemia in animal models. Firstly, AMPK ­activation mediates, independently of insulin, the inhibition of hepatic glucose output exerted by the antidiabetic biguanides [27], and the new Abbott compound A-769662 [23]. Second, expression of constitutively active AMPK in liver suffices to reduce blood glucose levels both in the streptozotocin-induced diabetic, and in the obese ob/ob mice [46]. Third, suppression of AMPK or LKB1 decreases the hypoglycemic effects of metformin and AICA riboside [42, 47]. Lastly, inhibition of hepatic glucose production by adiponectin, whose plasma concentration is decreased in obesity and dia­ betes, is mediated by AMPK activation [14, 44, 82, 83]. Suppression of adiponectin leads to insulin resistance in mice fed with a high lipid and carbohydrate diet, but not in control mice [84–86]. In addition, mice lacking liver adiponectin receptor 1 (adipoR1), but not adipoR2,

As stated above, AMPK controls liver fatty acid partitioning between oxidative and lipogenic pathways. Mice with whole-body or liver-specific deletion of the a2 AMPK subunit exhibit hyperglycemia as well as hypertriglyceridemia [44, 62], suggesting that AMPK deficiency could induce liver steatosis. Since hypertriglyceridemia is linked to insulin resistance [88], one may also wonder whether the lipogenic effects of AMPK deficiency could not contribute to glucose intolerance. Non-alcoholic fatty liver disease (NAFLD) is associated with insulin resistance and type 2 diabetes. It includes discrete and benign triglyceride accumulation but can evolve to non-alcoholic steatohepatitis (NASH), i.e., steatosis with inflammation, which may eventually lead to fibrosis and cirrhosis [89,  90]. Metformin and mainly TZD exert beneficial effects on the evolution of liver steatosis both in humans and animal models [91–93]. In animal models, inhibition by AMPK of the glycolytic and lipogenic gene induction program mediated by SREBP1c and mainly ChREBP, contributes to the beneficial effect of TZD and metformin [27, 46, 57, 63]. ChREBP is indeed an ideal target for treatment because it is a direct substrate of AMPK, which inhibits its transcriptional activity [58]. Although the mechanism of action of TZD may imply AMPK activation in the short-term, it also involves up-regulation of PPARg. All these observations indicate that ChREBP, SREBP1c, and liver AMPK activation are involved in the anti-steatosis action of metformin and TZD in animal models. Whether this conclusion also applies to humans remains to be demonstrated. Liver steatosis may also result from chronic ethanol consumption. The lipogenic effect of ethanol can be attributed to SREBP1c activation as a result of AMPK inactivation. Indeed, chronic administration of

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ethanol to mice decreased their liver AMPK activity and increased their malonyl-CoA content. This effect may be related to the fact that chronic administration of ethanol decrease plasma adiponectin in mice [94,95]. In addition, in hepatoma cell lines, SREBP1 activation by ethanol is exacerbated by dominant-­ negative AMPK and blocked by constitutively active AMPK, metformin or AICA riboside [96]. Inter­ estingly, resveratrol treatment of ethanol-fed mice led to reduced lipid synthesis, increased rates of fatty acid oxidation and prevented alcoholic liver steatosis. This protective action of resveratrol is in whole, or in part, mediated through up-regulation of the SIRT1-AMPK signaling system [97]. Therefore, low AMPK activity leading to SREBP1c activation is probably a determinant factor for the development of alcoholic liver steatosis.

Other Implications: Hepatocellular Carcinoma and Inflammation AMPK inhibits cell growth and proliferation. AICA riboside has been reported to inhibit cell-cycle progression in a human hepatocellular carcinoma cell line [98] and cancer cell proliferation in vivo [76]. Moreover, an epidemiological study has revealed that the incidence of cancer is lower in diabetic patients treated with metformin [99]. Inhibition of cell growth by AMPK is in part at least mediated by inhibition of the mTOR pathway (see above) and inhibitors of this pathway have been tried to inhibit hepatocellular carcinoma growth in vitro and in vivo [100,101]. Skeletal muscle produces and releases cytokines, called “myokines,” in response to intense and sustained physical activity. One of the myokines released is IL-6, whose plasma concentration may increase up to 100-fold after exercise. Sepsis is another condition leading to large increase in IL-6. However, during sepsis a rapid increase in tumor necrosis factor a (TNFa) precedes IL-6 production, whereas during exercise, plasma TNFa levels are less increased and follow the large increase in IL-6 [102]. IL-6 acts in an auto- or paracrine manner in muscle where it activates AMPK. It also activates AMPK in liver by a still unknown mechanism [15]. Whether AMPK activation  is involved in the liver acute phase response to

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inflammation and sepsis, is not known and certainly worth considering.

Conclusions Early studies of the AMPK system established its role in energy sensing and homeostasis both at the cellular and whole-body level. However, in normal liver, a decrease in energy charge is a rare, if not abnormal, phenomenon. Paradoxically, the energy sensitive AMP/ LKB1 axis is the main AMPK activating mechanism in liver, with CaMKKb playing little, if any, role in this organ. Liver AMPK activation mainly depends on extrahepatic changes, such as endurance exercise, variations of plasma cytokines or after administration of certain anti-diabetic drugs. In liver, AMPK activation essentially affects glucose and lipid metabolism, and AMPK activation by metformin counteracts hyperglycemia and hypertriglyceridemia and so contributes to the antidiabetic action of this drug. Conversely, systemic dysfunction of the LKB1/AMPK axis system in animal models exacerbates glucose intolerance, hepatic steatosis and pre-disposition to cancer via specific changes in the liver. However, the changes induced by genetic suppression of the catalytic a AMPK subunits are difficult to interpret, and could result from specific effects of the remaining regulatory subunits, which might conceivably interact with AMPK-related kinases, the other substrates of LKB1. To evaluate the specific effects of a AMPK subunit function, it would be preferable to make kinase-dead a AMPK subunit knock-ins that would keep the normal heterotrimeric structure of AMPK. Considerable progress has been achieved in the elucidation of the 3D structure of AMPK, especially of the b and g subunits. One may expect that the role of the glycogen binding domain in the b subunit will be defined. Similarly, the allosteric stimulation of the a subunit by AMP binding to the g subunit may receive a structural explanation. Thus unraveling AMPK structure and function has been, and still is, a successful story. More non-metabolic targets of AMPK are likely to be identified. AMPK is indeed an ancient eukaryotic kinase encoded by house-keeping genes and its role extends beyond energy homeostasis. Obviously the current continuous flow of papers on AMPK is far from drying up.

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Summary

›› The AMPK is involved in energy homeostasis

›› ›› ›› ›› ›› ›› ››

both at the cellular and whole-body level. When cellular energy is out of balance, AMPK is activated and restores energy charge by promoting ATP production and inhibiting ATP consumption. AMPK is a heterotrimeric complex comprising a catalytic (a) and two regulatory (b and g) subunits. Drugs such as metformin, thiazolidinediones, resveratrol activate AMPK. AMPK activation inhibits hepatic glucose production, glycogen synthesis and lowers gly­ cemia. AMPK inhibits fatty acid synthesis in liver through phosphorylation and inactivation of ACC1. AMPK decreases protein synthesis in part via inhibition of mTORC1 signaling. AMPK has anti-proliferative effects Low AMPK activity leading to SREBP1c activation is a determinant factor for the development of alcoholic liver steatosis

Multiple Choice Questions 1. Which is not a way to activate AMPK (a) LKB1 deacetylation by SIRT1 (b) Phosphorylation of Thr172 in the activation loop of the catalytic a-subunit by CaMKKb (c) AMP prevents Thr172 dephosphorylation by phosphatases (d) AMP allosterically stimulates AMPK activity by binding to the g-subunit (e) Calcium signaling in response to phenylephrine 2. Liver AMPK is activated in response to: (a) Mitochondrial inhibitors (b) Exercise (c) Starvation (d) Adiponectin (e) All the above

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3. Which of the following is not a phosphorylated by AMPK: (a) TORC-2 (b) HNF4a (c) ChREBP (d) SREBP1c (e) mTOR 4. In mice whose livers are deficient in both catalytic subunits of AMPK, what is wrong: (a) The hypoglycaemic effect of AICA riboside is decreased (b) There are hypotriglyceridemic (c) They have decreased expression of PGC1a (d) Their hepatocytes display reduced mitochondrial respiration (e) Specific effects of the remaining regulatory subunits have to be considered 5. Which statement is wrong (a) Chronic administration of ethanol to mice increases liver AMPK activity (b) AMPK controls liver fatty acid partitioning between oxidative and lipogenic pathways (c) Low AMPK activity leading to SREBP1c activation is a factor for the development of alcoholic liver steatosis (d) IL-6 activates AMPK (e) Activation of liver AMPK induces a hypoglycaemic response Acknowledgments  The work carried out in the authors’ laboratory was supported by the Belgian Fund for Medical Scien­tific Research, the Interuniversity Poles of Attraction Belgian Science Policy , the French Community of Belgium (Actions de Recherche Concertées) and the EXGENESIS Integrated Project (LSHM-CT-2004-005272) from the European Commission.

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18  AMP-Activated Protein Kinase in Liver Takahashi  M, Arita Y, Komuro R, Ouchi N, Kihara S, Tochino  Y, Okutomi K, Horie  M, Takeda S, Aoyama T, Funahashi T, Matsuzawa Y (2002) Diet-induced insulin resistance in mice lacking adiponectin/ACRP30. Nat Med 8:731–737 87. Yamauchi T, Nio Y, Maki T, Kobayashi M, Takazawa T, Iwabu M, Okada-Iwabu M, Kawamoto S, Kubota N, Kubota  T, Ito Y, Kamon J, Tsuchida A, Kumagai K, Kozono  H, Hada Y, Ogata H, Tokuyama K, Tsunoda M, Ide  T, Murakami  K, Awazawa M, Takamoto I, Froguel P, Hara  K, Tobe K, Nagai R, Ueki K, Kadowaki T (2007) Targeted disruption of AdipoR1 and AdipoR2 causes abrogation of adiponectin binding and metabolic actions. Nat Med 13:332–339 88. Aguilera CM, Gil-Campos M, Canete R, Gil A (2008) Alterations in plasma and tissue lipids associated with ­obesity and metabolic syndrome. Clin Sci (Lond) 114:183–193 89. Dixon JB, Bhathal PS, O’Brien PE (2001) Nonalcoholic fatty liver disease: predictors of nonalcoholic steatohepatitis and liver fibrosis in the severely obese. Gastroenterology 121:91–100 90. Marchesini G, Brizi M, Bianchi G, Tomassetti S, Bugianesi E, Lenzi M, McCullough AJ, Natale S, Forlani G, Melchionda N (2001) Nonalcoholic fatty liver disease: a  feature of the metabolic syndrome. Diabetes 50:1844–1850 91. Bajaj M, Suraamornkul S, Pratipanawatr T, Hardies LJ, Pratipanawatr W, Glass L, Cersosimo E, Miyazaki Y, DeFronzo RA (2003) Pioglitazone reduces hepatic fat content and augments splanchnic glucose uptake in patients with type 2 diabetes. Diabetes 52:1364–1370 92. Lin HZ, Yang SQ, Chuckaree C, Kuhajda F, Ronnet G, Diehl AM (2000) Metformin reverses fatty liver disease in obese, leptin-deficient mice. Nat Med 6:998–1003 93. Ratziu V, Giral P, Jacqueminet S, Charlotte F, HartemannHeurtier A, Serfaty L, Podevin P, Lacorte JM, Bernhardt C, Bruckert E, Grimaldi A, Poynard T (2008) Rosiglitazone for nonalcoholic steatohepatitis: one-year results of the randomized

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ER Stress Signaling in Hepatic Injury

19

Cheng Ji and Neil Kaplowitz

Abbreviations ALT alanine aminotransferase AMP adenosine monophosphate ARE antioxidant response element ASK1 apoptosis signal regulated kinase 1 ATF activating transcription factor Atg autophagy BHMT betaine homocysteine methyltransferase BI-1 Bax inhibitor-1 Bim a proapoptotic BH3-only member of the Bcl-2 family CBS cystathionine b-synthase CHOP C/EBP-homologous protein CREBH cyclic-AMP responsive element binding protein H eIF2a eukaryotic translation initiation factor 2 alpha subunit EOR ER overload response ERO1 ER oxidase 1 ER endoplasmic reticulum ERAD ER associated degradation ERSE endoplasmic reticulum stress response element Foxa forkhead box protein GCN2 general control of nitrogen protein kinase GRP78 glucose-regulated protein 78 GSH glutathione GSK glycogen synthase kinase

C. Ji () Keck School of Medicine, University of Southern California, 2011 Zonal Avenue, HMR 101, Los Angeles, CA 90033, USA e-mail: [email protected]

HBV hepatitis B virus HCV hepatitis C virus HERP homocysteine-induced ER protein Hcy homocysteine HHcy hyperhomocysteinemia IKK inhibitor of kB kinase IRE inositol requiring enzyme IRS-1 insulin receptor substrate-1 JNK c-jun-N-terminal kinase MHC major histocompatability complex MTHFR 5,10-methylenetetrahydrofolate reductase MTP microsomal triglyceride transfer protein NAFLD nonalcoholic fatty liver disease NASH nonalcoholic steatohepatitis NF-kB nuclear factor kB Nrf-2 NF-E2-related factor-2 NTBC 2-(2-nitro-4-trifluoromethylbenzyol)1,3-cyclohexanedione OASIS old astrocyte specifically induced substance ORP150 oxygen-regulated protein 150 PDI protein disulphide isomerase PEMT phosphatidyl ethanolamine methyl transferase PERK protein kinase ds RNA-dependent-like ER kinase PKB protein kinase B PKR protein kinase dsRNA-dependent PPARa peroxisome proliferator-activated receptor-alpha RT-PCR reverse transcriptase–polymerase chain reaction ROS reactive oxygen species SAH S-adenosylhomocysteine SAM S-adenosylmethionine

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_19, © Springer-Verlag Berlin Heidelberg 2010

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SREBP sterol regulatory element binding protein sXBP1 spliced XBP1 TRAF2 tumor necrosis factor receptor-associated factor-2 TRB3 tribbles 3 TNF tumor necrosis factor TNFR1 TNF receptor 1 TOR target of rapamycin UPR unfolded protein response XBP1 X box binding protein 1

Introduction The endoplasmic reticulum (ER) is an essential membrane-bound organelle for protein synthesis, oxidative protein folding, and posttranslational modifications, most notably the addition of oligosaccharides and the formation of disulfide bonds [1–6]. The ER is also a site for biosynthesis of lipids and sterols and for storing and releasing Ca2+ which is involved in numerous cellular signal transduction pathways. Molecular chaperones in the ER ensure proper folding and targeting of nascent proteins. Unfolded or malfolded proteins (as high as 30% of nascent proteins) are retained in the ER and targeted for retrotranslocation to the cytoplasm by the machinery of ER associated degradation (ERAD), and rapidly degraded through the ubiquitin-­proteosomal pathways [7, 8]. Physiological or pathological conditions such as increased translation of secretory proteins, reduced capacity of folding and proteasomal degradation, alterations of redox state and Ca2+ levels, ATP depletion, and improper posttranslational modifications perturb the homeostasis of ER and cause accumulation of unfolded proteins which stresses the ER leading to an adaptive response (referred to as the unfolding protein response, UPR) to dampen the stress. Prolonged or severe UPR can lead to an attempt to delete the cell which is termed ER stress response [1– 6]. Both responses are critical for the survival of the organism and an intricate relationship exists due to overlap and interplay between the two responses. In this chapter, we highlight the general signaling pathways of UPR and ER stress response, summarize the role of ER stress in a number of experimental or naturally occurring models of liver disease, and discuss our recent advances in alcohol or homocysteine-induced ER stress response and hepatic injury.

C. Ji and N. Kaplowitz

General Signaling Pathways Associated with ER Stress Unfolded Protein Response Several ER-resident transmembrane proteins maintain homeostasis of the ER and monitor the protein folding status and membrane integrity with their ER luminal domains. These proteins are the type I transmembrane protein kinase PERK, the type I transmembrane protein kinase endoribonucleases IRE1a and IRE1b, and the bZIP transcription factors synthesized as type II transmembrane precursors. The bZIP transcription factors include activating transcription factor (ATF6a and ATF6b) as well as OASIS, CREB3/Luman, CREB-H, CREB4, and BBF2H7 that were recently identified in the ER membrane [1–4, 9]. IRE1a, ATF6, and PERK are three major well characterized sensors for the unfolded protein response (Fig. 19.1). They are normally held in an inactive and inhibited state by the binding of intraluminal ER chaperones, especially glucose-­ regulated protein 78 (GRP78 or BiP) [10]. Upon UPR, GRP78 is displaced to deal with the exposed hydrophobic regions of the unfolded proteins, which facilitates folding through conformational change evoked by the hydrolysis of ATP by the ATPase domain. The displacement of GRP78 frees IRE1a, PERK, and ATF6. IRE1a and PERK are self activated by dimerization and/or ­autophosphorylation. ATF6 transfers to the Golgi and is ­activated there by a process called regulated intramembrane proteolysis that requires site 1 and site 2 proteases (S1P and S2P) [1–4]. IRE1a-XBP1 Enhances Folding Capacity and Reduces Protein Load The activated IRE1a functions as a nuclease that splices X box-binding protein 1 (XBP-1) mRNA; the resultant sXBP-l activates transcription through UPRE (UPR element) in promoters of genes which control ERAD. ERAD is a crucial function required to preserve the functionality of the ER under pressure of malfolded proteins. Another function of the nuclease activity of IRE1a is to degrade mRNA of a variety of genes which encode for secretory (e.g., insulin mRNA)

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Fig. 19.1  The unfolded protein response (UPR) is regulated by GRP78, which is displaced from three sensors to allow the activation and transduction of signals that unload the ER, inhibit further loading, and increase folding and antioxidant enzymes

and membrane proteins to reduce the load of newly synthesized proteins that need to be folded [4, 5, 11].

whereas a deficiency in both genes caused death in early murine embryonic development [13].

ATF6 Modulates UPR Transcription

PERK-eIF2a Attenuates mRNA Translation

After regulated intramembrane proteolysis, ATF6 translocates to the nucleus, interacts with ERSE (ER stress response element) which up-regulates chaperones/foldases such as GRP78, GRP94, IRE1a, protein disulfide isomerase (PDI), ERp72, GRP58⁄ERp57, HERP (homocysteine-induced ER protein), calnexin, and calreticulin, all of which increase the capacity of folding in the ER [1–3]. ATF6a is found to be a co-activator of the UPR that interacts with NF-Y and XBP1 and can bind to almost all elements in the promoters of UPRresponsive genes. Recent estimates indicated that ATF6a influences 10–20% of UPR-regulated genes, suggesting a unique role in support of protein folding, ERAD, and general ER function ­[12–14]. In addition, functions of ATF6a and ATF6b may be complementary since neither ATF6a-null nor ATF6bnull mice had a significant pathological phenotype

The activated PERK phosphorylates eukaryotic translation initiation factor 2, alpha subunit (eIF2a) to globally shut down protein translation, halting protein loading while selectively increasing translation of certain mRNAs such as ATF4 which up-regulates chaperones. Activated PERK also phosphorylates NF-E2-related factor-2 (Nrf-2) which promotes antioxidant response and maintains redox homeostasis [15]. In addition, recent studies have shown that the UPR induces autophagy, possibly through an early c-jun N-terminal kinase (JNK) activation or the TOR (target of rapamycin)-Atg (autophagyrelated) signaling pathway which protects the stressed cells by quickly removing superfluous or damaged organelles and portions of cytosol [16]. The protective role of early autophagic response may depend on ER stress response as autophagic response promotes necrosis in apoptosis-deficient cells in response to ER stress [17].

290

Thus, by decreasing the load of nascent proteins, rapid unloading of unfolded proteins, increasing capacity of folding, or clearing out the damaged ER components, the adaptive UPR reverses ER stress when perturbations occur around the ER environment and secretory or membrane proteins accumulate inside the ER (Fig. 19.1).

ER Stress Response The early or low-level chronic activation of the UPR is an adaptive function which dampens ER stress and ensures cell survival. However, prolonged or severe UPR (alternatively called ER stress response) provokes a complex network of interacting and parallel responses leading to pathological consequences including apoptotic cell death, inflammation, and fat accumulation (Fig. 19.2) [1–4].

C. Ji and N. Kaplowitz

ER Stress Response Triggers Cell Death Pathways and Inflammation Sustained activation of IRE1a recruits TNF receptorassociated factor-2 (TRAF2). IRE1a-TRAF2 signaling also activates apoptosis signal regulated kinase 1 (ASK1) and/or JNK which promotes apoptosis ­[18–20]. Under prolonged ER stress, Bax/Bak, two proapoptotic members of the Bcl-2 protein family translocate to the ER membrane causing Ca2+ release leading to calpain activation or causing mitochondrial depolarization resulting in oxidative stress [21–24]. In turn, the oxidative stress enhances ASK1-dependent activation of JNK and p38 MAP kinase and also promotes cell death through oxidizing thioredoxin, a redox-sensitive inhibitor of ASK1 [21]. In addition, Ca2+ activated calpain may activate ER resident caspase 12 in rodents (caspase 4 in humans) causing a cascade of activation of executioner caspases downstream of mitochondria which, along with the activated JNK, modulate apoptosis [3, 25, 26]. Hence, both mitochondrial dependent

Fig. 19.2  Detrimental effects of ER stress response (prolonged UPR) include proapoptotic and proinflammatory responses, oxidative stress, insulin resistance, and steatosis

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and independent cell death pathways can be triggered by the IRE1a-TRAF2 signaling in response to the ER stress. Prolonged activation of PERK and subsequent eIF2a phosphorylation selectively up-regulates ATF4 translation which increases expression of C/EBPhomologous protein (CHOP or GADD153) [1–3]. CHOP is a transcription factor and is of particular interest as its increased expression, along with GRP78, is a hallmark of the UPR and ER stress response. CHOP null cells are resistant to various stimuli of ER stress-induced apoptosis [27, 28]. Increased expression of CHOP is associated with downregulation of antiapoptotic Bcl-2, upregulation of ER oxidase 1 (ERO1) which promotes oxidative stress causing inflammatory response, and upregulation of GADD34 which dephosphorylates eIF2a causing a premature reversal of the translational attenuation by the early UPR, thus intensifying ER stress. CHOP was shown to mediate transcription of Bim, a proapoptotic BH3-only member of the Bcl-2 family which is essential for ER stress induced-apoptosis in a diverse range of cell types both in culture and within the whole animal [29, 30]. In cooperation with ATF4, CHOP increases expression of TRB3 which inhibits Akt and blocks its various protective actions [31]. For instance, Akt phosphorylates GSK-3b and keeps it inactive. TRB3 inhibits Akt which leads to increased activated GSK-3b which promotes ER stress-induced apoptosis [32, 33]. In addition, distinct from ER stress, TNF and dsRNA (viruses) activate PKR and amino acid deprivation activates general control of nitrogen protein kinase (GCN2) [34, 35]. Both PKR and GCN2 phosphorylate elF-2a kinase which blocks translation, promoting apoptosis via the ATF4-CHOP pathway and sensitizing to TNF killing by inhibiting synthesis of NF-kB dependent protective proteins [2, 21]. This is referred to as the integrated stress response [1–3].

ER Stress Response Leads to Lipid Accumulation The prolonged UPR can result in increased production of triglyceride and cholesterol which can be explained by a few possible mechanisms. First, sterol regulatory element binding proteins (SREBP-1c), a transcription factor in control of fatty acids and triglyceride synthetic genes, and SREBP-2, a transcription factor in control of cholesterol synthetic genes, both reside in ER and were found to be activated via regulated intramembrane

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proteolysis in response to ER stress [36]. How ER stress promotes the activation of SREBP is not completely understood. Since SREBP forms a complex with SREBP cleavage activating protein (SCAP), which is retained in the ER by another protein, Insig, and Insig turns over rapidly [37–39], the ER stresscaused slowdown of mRNA translation may lead to a rapid decline in the Insig protein, which would allow SREBP to migrate to the Golgi for activation. Further, ER stress-activated JNK which phosphorylates insulin receptor substrate (IRS-1) and ER stress-induced TRB3 which inhibits Akt signaling lead to insulin resistance. Insulin has been shown to activate SREBP. The effects of insulin on SREBP-1c transcription are opposed in the liver by glucagon [40]. In the insulin resistant state, the high insulin levels drive increased SREBP expression possibly via LXR and PKC/PTEN. Inhibition of PI3K/Akt pathway leads to high levels of glucose which may indirectly enhance the insulin stimulated expression of SREBP and/or ChREBP (carbohydrate response element-binding protein) [41–44]. Second, phosphorylation of eIF2a by the ER stress-activated PERK was found to induce translation of C/EBPa and b which increase expression of PPARg which regulates lipid accumulation, adding another mechanism for ER  stress-induced steatosis [45]. Third, XBP-1 was recently found to directly regulate expression of a subset of lipogenic genes independent of SREBP [46], adding a novel mechanism to hepatic steatosis which might be affected by ER stress response. It appears that multiple signaling pathways are involved in ER stressregulated fat accumulation. Therapeutically, it may be beneficial to intervene to reduce ER stress in order to reduce steatosis.

ER Stress in the Liver Hepatic cells are rich in ER and assume synthesis of a large amount of secretory and membrane proteins. UPR must play a pivotal role in the liver in maintaining ER homeostasis under basal conditions and in adapting to fluctuations in nutrient availability. In early 1980s, stress (temperature, restraint, CCl4, chlorpromazine, DDT, etc.) induced ER damage in the liver began to be recognized in ultrastructural, morphologic, and histological studies [47–49]. However, until recently, very little was known about molecular mechanisms of ER stress signaling in liver disease. In the past several

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Fig. 19.3  ER stress response occurs in a variety of natural and experimental disease models including pathogen infections, genetic defects in secretory proteins, scurvy, drug toxicity, injury of ischemia and reperfusion, and injury by heavy metal, alcohol, excess homocysteine, fatty acid and cholesterol

years, the research field of ER stress and liver disease has moved forward quite rapidly. ER stress response exists in a wide spectrum of liver disorders such as hyperhomocysteinemia of CBS ± mice fed high methionine diet, alcoholic liver injury, insulin resistance/ NAFLD, chronic viral hepatitis, and cholestatic liver injury [3, 4, 50] (Fig. 19.3).

Alcohol-Induced ER and/or Mitochondrial Stress and Liver Injury The pathogenesis of alcoholic liver injury is very complex and involves multiple mechanisms and pathways. Liver steatosis, steatohepatitis, fibrosis, and cirrhosis are characteristic of alcoholic liver disease (ALD). Factors contributing to ALD include, but are not limited to, alteration of redox state, oxidative stress, endotoxin, changes of cytokine milieu and signaling, impaired immune response, and polymorphisms in the genes encoding SOD2, CD14 endotoxin receptor, TNFa, and TGFb, and angiotensinogen inhibition [51– 54]. Recently, alcohol-induced ER stress response has emerged as a novel mechanism in ALD [3, 4, 50, 55].

Evidence of Alcohol-Induced ER Stress Response First, severe steatosis, scattered apoptosis and necroinflammatory foci were observed in intragastric alcohol fed mice [39, 55–57]. Microarray gene expression

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profiling of liver samples from the intragastric alcohol fed versus pair fed mice initially revealed altered expression of a set of genes relating to the UPR or ER stress response. In addition, moderate upregulation of expression of SREBP-1c and SREBP-2 and their responsive genes was detected with RT-PCR and immunoblotting [39, 55]. SREBP-lc knockout mice are protected against triglyceride accumulation but are not protected against increased cholesterol mediated by ethanol induction of SREBP-2. The rapid turnover of Insig during the ER stress-caused attenuation of mRNA translation may contribute to the release and activation of SREBPs in this animal model. CHOP knockout mice fed ethanol exhibited no change in other markers of ER stress, or fatty liver. However, CHOP knockout exhibited minimal alcohol-induced apoptosis [58]. This indicates that CHOP expression in response to ER stress appears to be responsible for apoptosis in this model. Second, in another model of micropigs fed alcohol, liver steatosis and apoptosis were shown to be accompanied by increased mRNA levels of CYP2E1, GRP78 and SREBP-1c, and protein levels of CYP2E1, GRP78, nSREBP, and activated caspase 12 [59]. In addition, the transcripts of lipogenic enzymes fatty acid synthase (FAS), acetyl-CoA carboxylase (ACC) and stearoyl-CoA desaturase (SCD) were elevated in the ethanol-fed micropigs, supporting a correlation of ER stress response with the pathogenesis of ALD. Third, in rat intragastric alcohol infusion model, high dose ethanol consumption was found to cause ER stress as indicated by the induction of P-PERK, CHOP, and TRB3 [60]. Further, recent evidence shows that the UPR is involved in hepatic injury by lipopolysaccharide (LPS) which is interesting as increased LPS has been associated with ALD [61–63]. Rat cirrhotic livers exhibited partial UPR activation as indicated by eIF2a activation in the basal state and full UPR as indicated by activation of IRE1a, ATF-6, and eIF2a after LPS challenge [61]. However, LPS-induced accumulation of NF-kB-dependent antiapoptotic proteins was not seen, suggesting that the UPR sensitized cirrhotic livers to LPS/TNFa-mediated apoptosis. It is conceivable that the sensitization also occurs in alcohol-induced cirrhosis. Fourth, in a Lieber-DeCarli baboon model of ALD, a number of molecular pathways were identified to be altered using cDNA array analysis [64]. Although ER stress mechanism was not recognized in this study, upregulation of calpain 2, calpain p94, and ERD21 and downregulation of eIF2a were among the genes listed. In addition,

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the studies were extended to human ALD by comparison of gene expression profiles in non diseased and cirrhotic liver tissue and upregulation of calpain and downregulation of calreticulin were also among genes revealed by the arrays. Therefore, the above several lines of evidence support that ER stress response contributes to ALD.

Contribution of Alcohol-Induced Hyperhomocysteinemia to ER Stress Homocysteine (Hcy) is an intermediate amino acid involved in methionine metabolism. Nutritional deficiency and mutations in some of the enzymes which remove Hcy lead to hyperhomocysteinemia (HHcy) which is implicated in a variety of diseases including vascular, central nervous system (CNS), and liver disease [3, 50, 65–67]. Recent evidence has linked HHcy to ER stress in diet models and in both knockout mice and humans with altered Hcy metabolism [55, 56, 68–71]. In all models of non-alcoholic HHcy (MTHFR −/−, CBS +/−, high methionine/ low folate diet), hepatic steatosis and ER stress with variable necroinflammation and

Fig. 19.4  Role of homocysteine and ER stress in alcohol-induced liver injury. Alcohol metabolite-acetaldehyde, alcohol-induced reactive oxygen species, and alcohol-induced impairment of homocysteine metabolism contribute to ER stressrelated hepatic cell death and fatty liver

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apoptosis are observed [55, 65–68]. A few potential molecular mechanisms can explain Hcy-induced ER stress. Nitrosylated Hcy interferes with the proofing and editing function of the methionine tRNA synthetase and can be wrongly incorporated into nascent proteins causing malfolding or unfolding. Also the methionine tRNA synthetase converts Hcy to proactive thiolactone which either modifies lysine residues and other free amine groups on proteins (homocysteinylation) or disrupts disulfide bond formation leading to malfolding in the ER. Increased Hcy thiolactone levels have been detected in humans with genetic disorders in Hcy metabolism and in mice fed a high-methionine diet [70–73]. Our work has linked HHcy to ER stress and alcoholic liver injury (Fig. 19.4). The intragastric alcohol feeding exhibited a striking 5–10 fold increase in mouse plasma Hcy [34, 55, 74, 75]. The pathways for Hcy removal are: (1) conversion of Hcy to SAH by SAH hydrolase (which has bidirectional activity); (2) conversion of Hcy to cystathionine by CBS, ultimately leading to cysteine formation (transsulfuration); (3) the remethylation of Hcy to methionine, carried out by methionine synthase (MS) (methyl THF is the methyl donor substrate) and BHMT (betaine is the methyl

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donor substrate) [76–79]. MS is ubiquitous and BHMT is expressed exclusively in hepatocytes and renal cells. Under normal conditions about half of the Hcy produced in hepatocytes is remethylated with MS and BHMT contributing approximately equally [75, 78, 80]. Chronic alcohol-induced disturbance of methionine metabolism appears to contribute to HHcy. In alcohol fed mice and rats decreased MS activity has been observed [80]. This may be due to the effects of acetaldehyde or NO on MS protein [81]. However, MS mRNA is significantly decreased in the intragastric ethanol-fed mouse model [57]. Thus the major factor in causing HHcy in response to alcohol may be decreased MS. In addition, a second factor may also be critical, namely the response of BHMT, which may influence the severity of the Hcy response. The abrogation of ER stress in parallel with decreased ALT and amelioration of liver steatosis and apoptosis were observed when the intragastric alcohol fed mice were fed betaine that provides a methyl-donor to push the conversion of Hcy to methionine [55]. BHMT overexpression in HepG2 cells inhibited Hcy-induced ER stress response, lipid accumulation, and cell death [82]. Suppression of BHMT in primary mouse hepatocytes potentiated Hcy-induced but not tunicamycin-induced ER stress response and cell injury. Transgenic mice expressing human BHMT in organs peripheral to the liver are resistant to HHcy and fatty liver induced by chronic alcohol infusion or a high methionine and low folate diet [74]. In the rat BHMT enzyme is up-regulated by ethanol or high methionine low folate (HMLF) diet thereby minimizing the effect of decreased MS on Hcy levels whereas in the mouse this enzyme is somewhat repressed by these conditions, thereby not suppressing and possibly accentuating the Hcy levels [57, 83, unpublished observations]. These effects on Hcy may determine the extent of ER stress and ALD.

Contribution of Mitochondrial Stress to ALD Mitochondria are the energy storing factories of liver cells. Chronic alcohol consumption-induced impairment in the structure and function of liver mitochondria has often been observed. It is conceivable that alcohol may also induce mitochondrial stress contributing to ALD [50, 51]. Insufficient cellular ATP due to metabolizing alcohol

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and ATP consumption by the UPR in the ER can affect mitochondrial protein folding and degradation which require ATP and molecular chaperones such as heat shock proteins (Hsp60/Hsp10) in mitochondria. In this respect, mitochondrial stress and alterations of mitochondrial chaperones could have pathological consequences. For instance, Hsp60 expression has been linked to cytochrome c release, caspase-3 activation and apoptosis [84–88]. It was also reported that Hsp10 exerted anti-inflammatory activity by inhibiting LPS-mediated Toll-like receptor signaling [88]. Thus, changes in either Hsp60 or Hsp10 will exacerbate ALD as more Bax/Bak are released in response to alcohol-induced ER stress and increased levels of gut-derived LPS are often detected in animals after chronic alcohol feeding [61–63]. In addition, cholesterol accumulation in mitochondria mediated by the ER stress-induced SREBP-2 might have pathological consequences. In HepG2 cells, acetaldehyde induced ER stress and accumulation of cholesterol in mitochondria [89]. The cholesterol increased inner mitochondrial membrane viscosity and impaired mitochondrial GSH transport which led to mitochondrial GSH depletion and increased susceptibility to cell death in response to TNF and FasL signaling pathways [90, 91]. Therefore, the interplay between the ER and mitochondria might contribute to ALD. The question is how the alcohol-induced mitochondrial stress and the interplay between the ER and mitochondria are regulated. A key question is whether mitochondria exhibit a response analogous to ER, i.e., UPR and stress response. This seems to be the case as mitochondrial chaperones and biogenesis are upregulated in response to mitochondrial stress or damage. It is possible that some retrograde signaling pathway(s) from mitochondria to nucleus that involves mitochondria-resident transcriptor(s) and/or co-activator may occur in response to mitochondrial stress which mediates synthesis of mitochondrial chaperones or proliferation of mitochondria. This type of signaling has been found in the yeast which regulates physiological function, nutrient sensing, TOR signaling, and aging [90–92]. Thus, understanding the function and dysfunction of mitochondrial stress response will provide better understanding of how alcohol damages the liver and reveal new preventive and therapeutic strategies.

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ER Stress in Nonalcoholic Fatty Liver Disease and Insulin Resistance The relationship between ER stress, liver steatosis and insulin resistance is complex and a vicious cycle among these disorders exists. Detrimental effects of UPR disrupt lipid homeostasis. In turn, certain components of lipids may disrupt ER membrane integrity and impair UPR exacerbating ER stress response. ER stress-induces JNK signaling leading to insulin resistance, which also contributes to the development of liver steatosis [50].

ER Stress and Liver Steatosis Evidence has emerged showing that fat triggers the ER stress response. In rats, sucrose- and saturated fatenriched diets induced steatosis, characterized by increased liver content of saturated fatty acids, hepatic expression of ER stress marker sXBP-1, GRP78 and CHOP as well as caspase 3 activation. The fat-induced ER stress occurred before obesity and was independent of changes in insulin action [93]. In rat cell culture (McA-RH7777) [94], prolonged exposure of free fatty acid increased cell triglycerides and ER stress which was associated with inhibition of apoB100 secretion. ApoB mediated increase of VLDL secretion could protect the liver from ER stress-induced steatosis. Increased saturated lipid content of ER directly compromised ER morphology and integrity [95]. Interestingly, polyunsaturated fat appears less potent in rats in inducing ER stress. Polyunsaturated fat diets induced hepatic steatosis without increased liver accumulation of saturated fat, did not induce hepatic XBP1 splicing, BiP expression, or liver injury in rats [93]. However, this phenomenon did not occur in mice. Intravenous infusion of oleic acid increased ER stress in a duration dependent manner in mice [94]. Mice deficient in stearoyl-CoA desaturase-1 (Scd1) and being maintained on a very low-fat diet developed ER stress response as indicated by enhanced splicing of XBP1, increased expression of CHOP and ATF3 in parallel with severe loss of body weight, hypoglycemia, and hypercholesterolemia [96]. This suggests that monounsaturated fatty acid synthesis may be important for maintaining metabolic homeostasis in the absence of sufficient dietary unsaturated fat and points

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to a novel cellular nutrient-sensing mechanism linking fatty acid availability and/or composition with the ER stress response. On the other hand, ER stress-induced SREBP disrupts lipid homeostasis enhancing liver steatosis. For instance, ER stress activated SREBP in hepatocytes exposed to homocysteine and thereby induced cholesterol and triglyceride synthesis and steatosis, whereas GRP78 overexpression inhibited SREBP activation and expression of its downstream genes [68]. ATF6 modulates SREBP2 mediated lipogenesis [97]. TNFa activated the UPR in a ROS-dependent manner [21] and also induced steatosis by activating partially the insulin-insig-SREBP signaling pathway in the livers of mice [98]. It has been speculated that disruption of lipid homeostasis by impaired immune response could further enhance steatosis. This is because the liver is enriched with natural killer T (NKT) cells that respond to lipid antigens presented by cluster of differentiation-1 (CD-1) molecules. The properly presented lipid antigens are required for optimal maturation and activation of the NKT cells. ER stress has been shown to decrease CD1 and contribute to NKT cell dysregulation in ob/ob hepatocytes and livers [99]. Thus, a vicious cycle between steatosis and ER stress may facilitate the progression of steatosis to steatohepatitis.

ER Stress and Insulin Resistance ER stress also promotes insulin resistance. ER stressinduces TRB3 which inhibits Akt and induces JNK which inhibits IRS1. Inactivation of either Akt or IRS-1 induces insulin resistance in the liver [100, 101]. TRB3 expression is greatly increased through the cooperative effects of ATF-4 and CHOP in diabetic mice. Obesity (ob/ob or high fat diet) – induced ER stress in the liver is associated with hyperactivation of JNK and inactivation of IRS-1 in cell and mouse models. XBP-1 +/− mice fed high fat diet exhibited insulin resistance and developed type 2 diabetes in the presence of increased ER stress response due to impaired UPR protection [102–104]. Synthetic ER stressors impaired proximal insulin signaling in hepatoma cells by increasing serine and decreasing tyrosine phosphorylation of IRS1 and reducing Akt phosphorylation [102]. The alterations in the insulin signaling cascade were dependent on JNK activation [104–107]. Proteomic analysis revealed differentially upregulated

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proteins of ER stress in fat of obese subjects which were associated with JNK activation and insulin resistance [108]. ER stress and JNK activation were also shown to increase glucose-6-phosphatase activity and glucose output in primary hepatocytes [109], which was mediated by IRE1a probably through TRAF2 recruitment [18, 19]. One of consequences of insulin resistance after ER stress is increased lipid synthesis. This could be mediated by either SREBP, as insulin upregulates it, or the ChREBP. ChREBP gene expression and ChREBP nuclear protein content are significantly increased in the liver of ob/ob mice with insulin resistance [110]. Like SREBP-1c, ChREBP regulates lipogenic gene expression. Liver-specific inhibition of ChREBP markedly improved hepatic steatosis by specifically decreasing lipogenic rates [111].

Proof of Concept from Therapeutic Interventions Experimental tests of therapeutic approaches provide strong support for the interrelations between ER stress, steatosis, and insulin resistance. The therapeutic approaches include increasing expression of chaperones in the ER, utilization of small molecules with chaperone properties, and inhibiting ER stress response and/or insulin resistance. For instance, silencing an ER chaperone, oxygen-regulated protein 150 (ORP150) by antisense ORP150 virus administration in normal mice decreased insulin sensitivity accompanied by loss of IRS-1 tyrosine phosphorylation and increased gluconeogenesis and insulin resistance [112]. Heterozygous deficiency of ORP150 in Akita mice impaired glucose tolerance. However, hepatic overexpression of ORP150 in db/db (leptin receptor deficiency) diabetic obese mice improved insulin sensitivity and glucose tolerance. Hepatic glucose output was suppressed as a result of improved insulin signaling and there was decreased expression of the key gluconeogenic enzymes, phosphoenolpyruvate carboxykinase and glucose-6-phosphatase. Thus, overexpression of specific ER chaperone(s) may be beneficial. Small chemical chaperones and osmolytes that stabilize proteins and improve their folding in and export from the ER have been used to target ER stress related insulin resistance and steatosis [113, 114]. Treatment of ob/ob mice (leptin deficiency) with the chemical chaperones 4-phenyl butyric acid and taurine-conjugated ursodeoxycholic acid restored glucose tolerance,

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insulin sensitivity, and resolved fatty liver in parallel with marked inhibition of PERK, IRE-la, and JNK activation by increased tyrosine phosphorylation of IRS-1 and IRS-2 in liver and fat [115]. Improving insulin resistance by targeting and attenuating ER stress response is promising. For example, chromium attenuates ABCA1, a protein that regulates lipid transport, and plays a key role in carbohydrate metabolism by potentiating the action of insulin [116]. Oral Cr(d-phe)3 treatment reduced hepatic ER stress markers (p-PERK, p-IRE-1, p-eIF2a) that were increased in the obese and insulin-resistant ob/ob mice and subsequently improved glucose intolerance and insulin resistance through reduction of JNK activation. In parallel, chromium treatment was associated with a reduction in liver triglyceride levels and lipid accumulation [116]. Another example of ER stress attenuation is dephosphorylation of eIF2a by its specific phosphatase, GADD34 that enhances glucose tolerance and improves hepatosteatosis in mice [45]. This study not only demonstrated the effectiveness of the therapy that targets the ER stress response but also revealed the involvement of C/EBPa and C/EBPb in ER stressinduced upregulation of lipid synthesis. At present there is strong evidence that ER stress can cause insulin resistance and fatty liver and that fatty liver can cause ER stress. Thus, a vicious cycle of self amplification as well as participation of mitochondria in this cycle is suggested. This is a very fast moving area of research with major health-related relevance. There are striking similarities in the relationship between ER stress and fatty liver in alcohol versus obesity/insulin resistance although the triggers of ER stress in the two conditions may be different.

ER Stress in Viral Infection The UPR in host cells due to viral protein production exerts either a protective role by limiting viral protein translation or a detrimental role by protecting against ER stress-induced cell death allowing continued viral protein accumulation [4, 50, 117]. Evidence from cell culture or transgenic models indicates that some components of hepatitis C virus (HCV) replicon induced expression of UPR genes as well as hepatic injury [117,118]. For instance, HCV replicon induced sXBP-1 but inhibited ERAD in Huh-7 [119, 120].

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HCV gene expression correlates the translocation of ATF6 cytoplasmic domain to the nuclei of cells expressing HCV subgenomic replicons. ATF6 activates the IRE1-XBP1 pathway of the UPR by upregulating the transcription of XBP1. The amount of XBP1 spliced mRNA and sXBP1 protein was elevated in HCV replicon-­expressing cells [119, 120], suggesting that IRE1-catalyzed splicing is enhanced in cells that host HCV gene expression. However, the downstream activity of the XBP1 is somehow repressed in the HCV infected cells which prevents transcriptional induction of the ER degradation-enhancing-mannosidase-like protein (EDEM) and inhibits ERAD [119]. Thus the protective UPR in HCV infected cells may not remove the excessive viral proteins of HCV. Expression of HCV replicon also decreased glycosylation which prevented assembly of MHC-1 and led to continued virus production, avoidance of immune clearance and severe ER stress resulting in apoptosis and liver injury [121–123]. HCV or HBV protein expression in cells induced ER stress response resulting in Ca2+ release from the ER which activated cyclicAMP response element binding protein (CREB). The activated CREB can up-regulate protein phosphatase 2Ac (PP2Ac) which impairs cell-cycle regulation leading to apoptosis and hepatocarcinogenesis [124, 125]. In addition, expression of HCV core in Huh7 or HepG2 and in transgenic mice triggered hyperexpression of GRP78, GRP94, and calreticulin which was followed by Ca2+ depletion [124]. HCV core induced CHOP, BAX translocation to mitochondria, cytochrome c release, caspase 3 and PARP cleavage, all of which promote cell death. Reversal of HCV core induced ER Ca2+ depletion by transfection of sarco/ER calcium ATPase abolished all the effects except CHOP. The uptake of Ca2+ in the mitochondria further induces ROS which may promote activation of multiple signaling pathways, including NF-kB, modulating apoptosis and inflammation, and STAT-3, a transcription factor that controls cell survival, proliferation, and differentiation [117, 118]. In the liver of transgenic mice, conditional expression of HCV structural proteins for about one month increased levels of cleaved caspase-3 and CHOP and hepatic apoptosis. This study suggests an in vivo correlation between HCV structural protein expression, ER stress and hepatic injury [126]. Clearly, viral proteins interfere with various aspects of ER stress response in experimental systems which suggest main effects leading to prevention of virus

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eradication. However, verification of these effects in HCV infected human liver is needed to determine whether the ER stress signaling is inhibited or enhanced by HCV and if these effects promote or inhibit liver injury or viral replication. Recent in vitro evidence has demonstrated that HCV induced UPR response which was followed by an incomplete autophagic response, suggesting a possible utilization of host cellular response by HCV in enhancing its own replication [127]. It is tempting to speculate that the UPR/ER stress response could also lead to evasion of viral eradication and promotion of injury in the infected human liver as a rapidly progressive liver injury accompanying massive hepatic viral loading has been observed in post-OLT or with concomitant HIV infection [3, 4, 50].

ER Stress in Other Liver Diseases Genetic Disorders Genetic disorders that link to UPR/ER stress and liver disease include defects of fibrinogen, fumarylacetoacetate hydrolase, and al-antitrypsin (a1-AT). Patients with variant fibrinogens develop accumulation of misfolded proteins in the ER and liver cirrhosis [128]. Aberrant accumulation of fumarylacetoacetate due to fumarylacetoacetate hydrolase deficiency induced ER stress in cell culture [129]. This deficiency led to hereditary tyrosinemia type 1 disease and both ER stress and apoptosis were observed in mice in the absence of NTBC therapy. In a1-AT deficiency which can result in significant liver disease in 10% of all affected patients, non-polymer forming mutants of a1-AT induced the UPR without detectable hepatic injury whereas the polymer-forming common mutant protein (a1-ATZ) did not induce the UPR but elicited injurious components of the ER stress response such as PERK/CHOP and EOR (NF-kB activation) [130]. Transgenic mice expressing the polymer-forming mutant protein had increased expression of CHOP and were more susceptible to liver fibrosis induced by cholestasis from bile duct ligation [131, 132]. However, in the hepatocytes of individuals carrying the polymer-forming mutant, a1-ATZ localizes both to the ER and to membrane-surrounded inclusion bodies (IBs) that are negative for autophagosomal and

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lysosomal markers, and contained several ER components [133]. This evidence suggests that segregation of a1-AT from the ER to the IBs may initially be a protective cell response to maintain a functional secretory pathway.

Drug-Induced Liver Disease Acetaminophen (APAP) is a widely used drug with analgesic and antipyretic effects. APAP overdose is a major and frequent cause of acute liver failure and the precise mechanism is not fully understood. Perturbation of redox balance of the ER and consequential activation of the ER stress signaling and proapoptotic events may be involved in hepatocellular damage caused by APAP overdose. In fact, APAP overdose resulted in a decrease in microsomal total glutathione (GSH) and in the GSH/GSSG ratio [134]. Redox state of thiols of ERp72 and PDI was shifted towards the oxidized form. ATF6, CHOP, and caspase-12 were also activated followed by an increased apoptosis of hepatocytes, all of which suggest a possible role of ER stress in APAP induced liver injury. Other drugs that induce typical ER stress are methapyrilene and HIV protease inhibitors (PIs). Methapyrilene elicited hepatic damage that increased in severity with the number of doses. High-dose methapyrilene elicited thousands of gene changes including genes associated with ER stress [135, 136]. HIV PIs including amprenavir, atazanavir, and ritonavir are used in treatment of HIV-infected patients. However, atazanavir and ritonavir cause serious lipid disturbances which are associated with activation of the UPR signaling, increased levels of active SREBPs, induction of apoptosis, and formation of foam cell in macrophages [137]. How protease inhibitors cause ER stress is not completely understood. A number of possible cellular stress signals, such as depletion of ER calcium stores, increased cholesterol in ER membranes, deprivation of glucose, or inhibition of proteasome activity are proposed to activate the UPR [137–139]. Interestingly, different HIV PIs exert different effects on ER stress response in liver cells. This may provide useful information to help predict clinical adverse effects in the development of new HIV PIs and also provide a better understanding of the cellular mechanism of HIV PI-induced ER stress and dyslipidemia.

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Ischemia-Reperfusion Injury Hepatic ischemia-reperfusion injury (IRI) is a ­multistep process that involves oxidative stress, ­inflammation, and cell death. Pathophysiological interplay between mitochondria and ER may be essential in IRI. Mitochondrial responses in IRI involve ROS, NF-kB activation, increased activity of caspase 9 and caspase 3. Mitochon­ drial dysfunction also leads to ATP depletion which causes Ca2+ leakage from the ER [140]. ER stress response in IRI is associated with increased GRP78, CHOP, sXBP-1 and PERK. In addition, Bax inhibitor-1 (BI-1) resides in the ER and suppresses ER stress-induced apoptosis [141]. BI-1 knockout mice exhibited enhanced ER stress response and IRI [142, 143]. The small molecule chemical chaperone, sodium 4-phenylbutyrate protected against IR induced ER stress-mediated apoptosis in the liver [144]. Thus, dysfunction in both mitochondria and the ER may play a role in IRI that may affect liver graft viability and functional integrity.

Cholestasis and Bile Acid-Induced ER Stress Bile salts are natural detergents produced in the liver from cholesterol and function to emulsify fats as an aid to digestion. However, high concentrations of certain bile salts, e.g., hydrophobic bile acids, are cytotoxic and induce apoptosis in hepatocytes in chronic cholestatic liver diseases. ER stress-mediated apoptotic pathways appear to contribute to the cytotoxicity by bile acids [145]. Increased intracellular calcium ion, activations of calpain and caspase-12, and increased expression of Bip and Chop mRNA were observed in rat hepatocytes [146] and HepG2 cells [147] after treatment with glycochenodeoxycholic acid (GCDCA). Further, CHOP deficiency attenuates GCDCA or bile duct ligation-induced cholestasis and liver fibrosis by reduction of hepatocytic cell death, confirming that downstream activation of CHOP by ER stress and subsequent cell death by CHOP contributes to bile acidinduced damages [148]. In addition, hepatocyte-specific ablation of Foxa2 which altered bile acid homeostasis upstream resulted in ER stress response and cholestatic liver injury [149]. Thus, strategies to maintain FOXA2 expression in individuals with cholestasis may be a new therapeutic goal.

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Scurvy and Damages by H2O2 and Heavy Metals Nutritional deficiency in ascorbate (vitamin C) causes scurvy in certain species deficient in gulonolactone oxidase that catalyzes the last step of ascorbate synthesis. Chronic ascorbate deficiency leads to ER stress, UPR, and apoptosis in the liver of guinea pigs, implying that impaired protein processing may take part in the pathology of scurvy [150]. How ascorbate deficiency induces ER stress is not clear but malfunction of microsomal ascorbate oxidase due to ascorbate depletion can lead to enhanced oxidation of protein thiol groups which results in malfolding of proteins. In addition, ascorbate can react with oxygen in a oneelectron transfer which influences intracellular generation of ROS and redox balance and thus affects proper protein folding in the ER. Recent evidence shows that generation of H2O2 from the glucose oxidase-treated HepG2 cells induced dysfunction of the ER and resulted in formation of Mallory bodies, supporting a link between ROS and ER stress [151]. As co-factors, proper amount of heavy metals is essential in mediating enzymatic reactions. However, heavy metal accumulation in organs, especially in the liver and kidney, via drinking water, foods, alcohol consumption, and cigarette smoking causes injury. Recently, ER stress was shown to be involved in heavy metal-induced injury [152–154]. Systemic exposure of mice to cadmium, nickel or cobalt caused rapid, transient and reversible induction of ER stress in vivo. Thus, therapeutic agents targeting the ER should be considered in dealing with heavy metalinduced liver injury.

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directed at questions about the physiological significance of the various ER-stress signaling pathways in mediating cell responses and liver injury. Hopefully, these studies should identify novel therapeutic targets that may have impact in the treatment of liver disease.

Summary

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and cytosolic compartment where secretory and membrane proteins are synthesized and folded. The ER is directly continuous with the nucleus, which enables them to share and commute information in a very efficient manner. When the ER undergoes functional problems and unfolded proteins accumulate, the ER quickly sends signals to the nucleus. The nucleus responds by slowing mRNA translation and increasing production of chaperones thereby giving the ER extra time to catch up on its protein folding, thus maintaining cellular health. This process is called UPR. Prolonged UPR is hazardous to the cell. Hepatocytes are rich in ER and are naturally vulnerable to any stress from the ER. A wide spectrum of liver disorders such as hyperhomocysteinemia, alcoholic liver injury, insulin resistance/NAFLD, chronic viral hepatitis, and cholestatic liver injury are associated with ER stress response. Strategies to maintain homeostasis in the ER in individuals with liver disease should be a new therapeutic goal.

Conclusions In the past decade, a growing body of research has revealed that ER stress contributes to liver disease in a variety of natural and experimental models such as hyperhomocysteinemia of CBS +/− mice fed high methionine diet, alcoholic liver injury, insulin resistance/ NAFLD, chronic viral hepatitis, cholestatic liver injury, and hepatic ischemia-reperfusion injury. Despite this, the specific roles of each of the UPR sensors IRE1a, ATF6, and PERK in ER stress-induced liver cell injury is still not fully understood. Future studies will be

Multiple Choice Questions 1. Functions of the endoplasmic reticulum of hepatocytes are: (a) Synthesis and modifications of secretory and membrane proteins (b) Synthesis of triglycerides and cholesterol (c) Storing and releasing calcium (d) Detoxifications of certain drugs (e) All of the above

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2. What are the three major sensors on the ER membrane that mediate the unfolded protein response? (a) ATF6, OASIS, and CREB (b) IRE1, ATF6, and GRP78 (c) IRE1, ATF6, and PERK (d) OASIS, GRP78, ATF6 (e) XBP-1, PERK, IRE1 3. The PERK-eIF2a pathway: (a) Enhances folding capacity (b) Modulates UPR transcription (c) Attenuates mRNA translation (d) Degrades mRNA of genes which encode for secretory and membrane proteins (e) Up-regulates calnexin and calreticulin 4. Liver ER stress response is caused by: (a) Alcohol, homocysteine, or drugs (b) Genetic protein folding disorders, or insulin resistance (c) Pathogen infection, heavy metals, or scurvy (d) Ischemia/reperfusion (e) All of the above 5. Which of the following does not contribute to ER stress-induced apoptosis? (a) Increased CHOP (b) JNK acivation (c) Caspases (d) Activation of SREBP (e) Activation of ASK1 Acknowledgments  This work was supported by NIH grants R01 AA014428, R01 AA018612, P50 AA11999, and P30 DK48522.

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C. Ji and N. Kaplowitz 142. Chae H, Kirn H, Xu C, et al BI-1 regulates an apoptosis pathway linked to endoplasmic reticulum stress. Mol Cell. 2004;15:355–366 143. Reimers K, Choi CY, Bucan V, Vogt PM. The Bax Inhibitor-1 (BI-1) family in apoptosis and tumorigenesis. Curr Mol Med. 2008;8(2):148–156 144. Vilatoba M, Eckstein C, Bilbao G, et al Sodium 4-­phenybutyrate protects against liver ischemia reperfusion injury by inhibition of endoplasmic reticulum-stress mediated apoptosis. Surgery. 2005;138:342–351 145. Bernstein H, Payne CM, Bernstein C, Schneider J, et al Activation of the promoters of genes associated with DNA damage, oxidative stress, ER stress and protein malfolding by the bile salt, deoxycholate. Toxicol Lett. 1999;108(1):37–46 146. Tsuchiya S, Tsuji M, Morio Y, Oguchi K. Involvement of endoplasmic reticulum in glycochenodeoxycholic acidinduced apoptosis in rat hepatocytes. Toxicol Lett. 2006; 166(2):140–149 147. Iizaka T, Tsuji M, Oyamada H, Morio Y, Oguchi K. Interaction between caspase-8 activation and endoplasmic reticulum stress in glycochenodeoxycholic acid-induced apoptotic HepG2 cells. Toxicology. 2007;241(3):146–156 148. Tamaki N, Hatano E, Taura K, Tada M, et al CHOP deficiency attenuates cholestasis-induced liver fibrosis by reduction of hepatocyte injury. Am J Physiol Gastrointest Liver Physiol. 2008;294(2):G498–G505 149. Bochkis IM, Rubins NE, White P, Furth EE, et al Hepatocyte-specific ablation of Foxa2 alters bile acid homeostasis and results in endoplasmic reticulum stress. Nat Med. 2008;14(8):828–836 150. Margittai E, Bánhegyi G, Kiss A, Nagy G, Mandl J, Schaff Z, et al Scurvy leads to endoplasmic reticulum stress and apoptosis in the liver of Guinea pigs. J Nutr. 2005; 135(11): 2530–2534 151. Hanada S, Harada M, Kumemura H, et al Oxidative stress induces the endoplasmic reticulum stress and facilitates inclusion formation in cultured cells. J Hepatol. 2007;47 (1):93–102 152. Hiramatsu N, Kasai A, Du S, et al Rapid, transient induction of ER stress in the liver and kidney after acute exposure to heavy metal: evidence from transgenic sensor mice. FEBS Lett. 2007;581(10):2055–2059

PPARa, A Key Regulator of Hepatic Energy Homeostasis in Health and Disease

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Nicolas Leuenberger and Walter Wahli

Introduction Liver participates in the metabolism of almost all ­nutrients and contributes to maintaining balanced levels of energy-providing molecules in the circulating plasma. This homeostatic regulation plays a central role during the times of food deprivation, for instance, at night for animals feeding during the daytime, and when plenty of food has been ingested. For their contribution to the maintenance of this energy balance, liver cells depend on a variety of transcription factors that control gene expression. Among those, the peroxisome proliferator activated receptors (PPARs) and more particularly the PPARa isotype are the key actors in the regulation of hepatic functions. Below, we review the roles of PPARa in the liver, in both health and disease.

Peroxisome Proliferator Activated Receptors (PPARs) PPARs, as well as the more widely known estrogen, glucocorticoid, or thyroid hormone receptors, belong to the same family of transcription factors, which comprise 49 members(In other chapters on PXR and FXR, this family is put as there are 49 members in mouse and 48 members in human; is there a new addition?). They display the canonical nuclear receptor domain structures, with each of their domains (DNA binding, ligand binding, transactivation) modulating their gene regulatory W. Wahli () Center for Integrative Genomics, National Research Center Frontiers in Genetics, University of Lausanne, Genopode Building, 1015 Lausanne, Switzerland e-mail: [email protected]

potential [1]. Three PPAR isotypes have been identified: PPARa (NR1C1), PPARb/d (NR1C2), and PPARg (NR1C3) [2]. The PPARa was first cloned from rodents as a nuclear receptor activated by agents who cause peroxisome proliferation in their liver [3]. However, such an effect has not been observed in humans [4]. The PPARs are activated by ligands, as the majority of nuclear receptors. The endogenous PPAR ligands are fatty acids and fatty acid derivatives, such as leukotrienes and postaglandins [5–7]. Because they sense these lipid signals, PPARs have been proposed to act as “lipostats.” In addition, several synthetic lipid-lowering agents, including the hypolipidemic drugs fenofibrate (TriCor®), gemfibrozil (Gemcor®, Lopid®), and clofibrate (Atromid-S®), activate PPARa, while the insulin sensitizers rosiglitazone (Avandia®) and pioglitazone (Actos®) activate PPARg [8, 9]. PPARs act on gene regulation in several stages, including activation by a ligand, binding to DNA, co-repressor dismissal and co-activator recruitment, and finally ­transcription regulation. Ligand-independent actions, as well as non-genomic effects, have also been reported [10]. DNA binding requires PPARs to heterodimerize with their designated partner, the retinoid X receptor (RXR; receptor for 9-cis retinoic acid, NR2B) [11–13], and bind to their own specific sequence element in the regulatory regions of target genes, which is called the peroxisome proliferator response element (PPRE) [14–16]. To activate transcription, PPARs need co-factors [17], some of which having chromatin modifying activity, such as the histone acetyl transferase (HAT) p300 [18]. To regulate specific target genes in a given tissue, coexpression of PPARs and a defined set of their co-regulators in the same cell are required. In addition to this classic ligand-dependent control of gene expression, the transcriptional activity of PPARs is fine-tuned by post-translational modifications including

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phosphorylation [19] and sumoylation (SUMO1; a small ubiquitin-like modifier), the latter being involved in the gene repression activity of PPARs [20]. The expression patterns of the three PPAR isotypes are different. PPARa is highly expressed in tissues with high fatty acid catabolism activity. In rodents, PPARa is highly expressed in liver, where its maximal levels are in parenchymal cells, while endothelial, Kupffer, and stellate cells express much less, if any, PPARa messenger RNA (mRNA) [21, 22]. PPARb/d is ubiquitously expressed, but at different levels according to cell types, and plays a role in lipid metabolism, tissue repair, and embryonic development [23]. In the liver, PPARb/d is well expressed in stellate cells, but is poorly expressed in parenchymal cells [24]. PPARg is highly expressed in adipose tissue, where it regulates adipogenesis and adipose tissue integrity [25, 26]. In the liver, PPARg is normally poorly expressed, but its levels increase significantly during lipid accumulation in both parenchymal and stellate cells [27]. Because of their presence and action in organs involved in nutrient processing and energy homeostasis pathways, PPARs have become pharmaceutical targets in the treatment of dyslipidemia, cardiovascular diseases, and type 2 diabetes.

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a­ ctivity and remnant clearance that was observed in both humans and rodents [30–32]. Moreover, expression of apoA-V, a potent activator of lipolysis, is upregulated by PPARa agonists [33].

PPARa and Cellular Lipid Uptake and Transport To benefit from fatty acids as a source of energy, cells have to take them up from the extra-cellular space. The fatty acid transporter 1 (FATP1), which participates in the uptake of long-chain fatty acids and oxidized lowdensity lipoproteins, and the well-known fatty translocase (FAT/CD36) are encoded in genes that are PPARa targets [34, 35]. Once within the cells, fatty acids bind to proteins called fatty acid binding proteins (FABPs) which are involved in shuttling them to the cell nucleus, where they bind to PPARs and enhance target gene expression [36]. Interestingly, PPARa ligands are known to stimulate the expression of FABPs [37]. Thus, PPARa influences nuclear fatty acid availability and, therefore, its own level of activation.

The Role of PPARa in Lipid Metabolism

PPARa and Fatty Acid b-Oxidation

In the liver, PPARa is the most highly expressed isotype, especially in rodents that are often used as experimental models. Therefore, the focus below is on this isotype.

A functional link between fatty acid oxidation and PPARa was first established in 1992, when it was found that the peroxisomal acyl-coenzyme A (CoA) oxidase gene, which is involved in peroxisomal fatty acid b-­ oxidation, is regulated by PPARa [15, 16]. Since then, numerous genes involved in hepatic fatty acid catabolism were shown to be induced by peroxisome proliferators and PPARa [14]. Hepatic fatty acid catabolism involves several distinct pathways. Fatty acids can be provided from the diet or are generated by the lipolysis of TGs in adipose tissues before circulating in the blood plasma and being taken up by the liver. In hepatocytes, fatty acids are activated to fatty acyl-CoAs and are subsequently translocated into peroxisomes or mitochondria for degradation to acetyl-CoA via b-oxidation. These pathways are regulated by PPARa. The fatty acid activation step is controlled by the long-chain fatty acyl-CoA synthetase, whose transcriptional rate

PPARa and Lipoprotein Metabolism In humans, the therapeutic benefits of fibrates, mediated by PPARa, result from their ability to efficiently reduce  VLDL (very low-density lipoprotein) production, enhance the catabolism of triglycerides (TGs), and increase the plasma levels of high-density lipoprotein (HDL) [28]. One of the major effects of PPARa on VLDL clearance is through the regulation of the lipoprotein lipase (LPL) [29]. Another mechanism behind the lipid lowering effect of fibrates is the down-­regulation of apoliprotein ApoC-III, an inhibitor of both LPL

20  PPARa, A Key Regulator of Hepatic Energy Homeostasis in Health and Disease

is regulated by PPARa [9]. In short, PPARa promotes the transport of fatty acids across the cell membrane, their accumulation within the cell, and their activation into a metabolically active form that serves as a substrate for further metabolism [38]. Once activated, fatty acids translocate into mitochondria through transporters called carnitine palmitoyltransferases (CPTs): CPT-I helps to cross the outer membrane, and CPT-II the inner membrane. b-oxidation begins once fatty acids have been imported into the mitochondrial matrix. It is the major source of cellular ATP. PPARa stimulates the genes for the proteins involved in all the above-mentioned steps [39, 40]. Whereas mitochondria oxidize short-, medium- and long-chain fatty acids, peroxisomes oxidize mostly very-long-chain polyunsaturated fatty acids. As in mitochondria, PPARa controls all steps of the cascade of peroxisomal fatty acid b-oxidation [14]. Because of the key implication of PPARa in lipid utilization, the development of novel PPARa activators, specifically targeting this function, is of interest to the pharmaceutical industry.

The Role of PPARa in Glucose Metabolism The role of PPARa in glucose metabolism emerged from several studies showing that PPARa deficient mice suffer from pronounced hypoglycemia [41]. Plasma glucose levels are maintained during fasting by the combination of processes including glycogen breakdown (glycogenolysis), de novo glucose synthesis (gluconeogenesis), and glucose utilization by peripheral organs. PPARa induces the conversion of glycerol into glucose through direct stimulation of the expression of several hepatic enzymes, such as glycerol-3-phosphate dehydrogenase (GPDH) and glycerol kinase (GK), and of the glycerol transporters aquaporins 3 and 9 [42]. GPDH deficiency in mice and humans leads to hypoglycemia, underlining the important role of glycerol as a substrate for glucose homeostasis [38]. Furthermore, PPARa participates in the regulation of gluconeogenesis by increasing the expression of the gene encoding pyruvate dehydrogenase kinase 4 (PDK4) [43]. This enzyme catalyzes the phosphorylation, and thereby, the inactivation of the pyruvate dehydrogenase complex. It results in the utilization

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of pyruvate for glyconeogenesis instead of for fatty acid synthesis. Another pathway that may be influenced by PPARa, with respect to glucose utilization, is the Akt/protein kinase B (Akt/PKB) cascade, which is a positive regulator of the cellular responses to insulin, through the mammalian tribbles homolog ­TRB-3. In the liver, PGC-1 promotes insulin resistance through PPARa-dependent induction of TRB-3, which leads to the inhibition of Akt/PKB [44, 45]. Therefore, upregulation of TRB-3 expression by PPARa may repress hepatic insulin signaling, and interfere with glucose homeostasis. These findings identify PPARa as a key player, not only in liver fatty acid catabolism, but also in hepatic glucose homeostasis.

The Role of PPARa in Amino Acid Metabolism Not only does PPARa govern lipid and glucose metabolism, but it is also associated with the control of hepatic amino acid metabolism [46]. PPARa down-regulates genes involved in major pathways of amino acid metabolism, including transamination, deamination, the urea cycle, oxidation of alpha keto acids, amino acid inter-conversions, and the synthesis of amino acid-derived products. In support of this gene expression control, there are increased plasma urea levels in PPARa deficient mice. Although the molecular mechanism by which PPARa exerts the coordinated down-regulation of all these genes is unknown, the benefit for the organism during fasting is thought to be the use of lipids, rather than amino acids derived from muscle protein degradation, as an initial energy source. This hierarchy in the utilization of energy-rich molecules may prevent rapid weakening of the organism because of a reduction of the muscle mass.

The Role of PPARa During Fasting The overall metabolic response to fasting operates at several levels. One important metabolic feature of starvation is the shift in fuel utilization from carbohydrates and fat in the fed state, to almost exclusively fat after a day of fasting in humans. This adaptation is

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particularly important in the brain, a major glucose consumer in the fed state, which is able to acquire energy predominantly from ketone bodies after prolonged fasting [47]. In connection with its fatty acid sensor function, PPARa is a key factor in the complex network of signaling pathways that operates in the liver during fasting. Most importantly, it stimulates fatty acid oxidation, which produces substrates such as ketone bodies that can be metabolized by periph­ eral organs to acquire the necessary energy [41] (Fig.  20.1). The PPARa-mediated regulation of the mitochondrial HMG-CoA synthase (HMGCS2), a key enzyme in ketogenesis, is essential in this process [48]. Furthermore, the fibroblast growth factor 21

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(FGF21), which is directly stimulated by PPARa in response to fasting, has emerged as a central factor in the biology of fasting [49, 50]. It enhances lipolysis in adipose tissue and ketogenesis in the liver, reduces physical activity, and promotes hypothermia that conserves energy [49, 50]. By stimulating lipolysis, FGF21 also provides fatty acids to the liver, which, in addition to being PPARa ligands, are also used for ketogenesis. Collectively, these findings unveiled major roles for the PPARa and FGF21 duo of proteins in regulating metabolic pathways and behavior, which participate in the adaptive responses to food deprivation.

PPARa and the Hepatic Circadian Clock

Fig. 20.1  PPARa-FGF21 effects during fasting. PPARa regulates the expression of FGF21 that enhances lipolysis in adipose tissue and ketogenesis in liver. In brain, increased FGF21 levels cause a reduction of physical activity and promote hypothermia. By stimulating lipolysis, FGF21 also provides fatty acids to the liver, which in addition to being used for ketogenesis act as PPARa ligands as well

Many physiological processes display day–night rhythms. Feeding behavior, lipid and carbohydrate metabolism, and detoxification are only a few examples of processes subjected to daily variations [51]. Oscillations in the expression of nuclear receptors that serve as sensors of dietary lipids in key metabolic tissues may contribute to a circadian rhythm of nutrient metabolism and energy homeostasis [52]. In fact, PPARa follows a circadian pattern with a pick of expression at the end of the day, when mice become active (rodents are active during the night and rest during the day) [53]. A similar pattern is seen for the plasma levels of glucocorticoids that stimulate the expression of PPARa [54]. Because both a high concentration of plasma glucocorticoids, and an increased level of PPARa expression occur during fasting, one may consider that rodents are in a pseudo-fasted state in the late afternoon. In support of this model, genes encoding enzymes and regulatory proteins involved in food processing and energy metabolism, such as cholesterol metabolism and fatty acid b-oxidation, show increased levels of expression at the end of the day in anticipation of feeding [55]. Not surprisingly, there is growing evidence for a tight link between metabolic diseases and the circadian clock [56, 57]. It is likely that chronobiological and chronopharmacological studies both at the bench and in clinics will contribute to improving or developing new chronotherapies [58, 59]. Temporal drug delivery strategies that take into consideration the time of day at which

20  PPARa, A Key Regulator of Hepatic Energy Homeostasis in Health and Disease

the activity of drugs is optimal will be directly on the basis of the results of such studies. For instance, the effect of hypolipidemic drugs as ligands for PPARs could be maximized by taking the circadian expression pattern of PPARs into account.

The Role of PPARs in Disease Chronic diseases, such as diabetes, obesity, atherosclerosis, and cancer, are responsible for most deaths in developed countries. They have many causes comprising genetic, environmental, and nutritional factors. There is evidence that PPARs are implicated in these diseases. Furthermore, PPAR activity can be modulated by drugs such as fibrates and glitazones, a fact which has stimulated research into the possible roles of PPARs in several diseases (Fig. 20.2).

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PPARs and the Metabolic Syndrome The metabolic syndrome, also called syndrome X, is characterized by an aggregation of disorders that include excess weight and obesity, dyslipidemia, hypertension, and insulin resistance. The occurrence of all these conditions in the same patient is associated with an elevated risk of developing cardio-vascular diseases and their dramatic consequences. More than 60% of the U.S. population is overweight (a body mass index of >25), and 31% meet the criteria for obesity (a body mass index of >30) [60]. The high-circulating levels of fatty acids associated with obesity are crucial in the development of insulin resistance. Because circulating fatty acid levels are important determinants in promoting or exacerbating insulin resistance, an improvement of this condition can be predicted by increased fatty acid oxidation in the liver [61]. Therefore, PPARa could be a target for restoring insulin sensitivity in patients suffering from dyslipidemia, but this has not been rigorously examined so far. On the contrary, the role of glitazones, which are PPARg ligands, in improving insulin resistance in human has been extensively studied [62, 63]. Activation of PPARg enables the retrieval of lipids that have unduly accumulated in the circulation and in non-adipose tissues, by increasing the ability of the adipose tissue, especially subcutaneous white adipose tissue, to store this excess fat [61].

PPARs and Hepatic Inflammation

Fig.  20.2  PPARa is involved in multiple hepatic functions. PPARa promotes nutrient processing, including lipid, glucose, and amino acid metabolism. Furthermore, PPARa represses lipid related hepatic diseases, such as NAFLD and inflammation. In rodents, PPARa promotes peroxisomal proliferation that leads to hepatocarcinogenesis

A critical role of PPARa in inflammation was first reported in a study published in 1999, which provided evidence for an extended duration of inflammatory responses in a PPARa knock-out mouse [64]. The liver is the site of production of acute phase reactants, such as serum amyloid A, fibrinogen, and C-reactive protein (CRP). The acute phase response consists of the secretion of a large number of acute phase proteins by the liver in response to injuries and various stimuli, thereby reducing inflammatory cytokines. To date, there is a large body of literature concerning the role of PPARa in limiting inflammatory responses in the liver [65]. Fibrates decrease the levels of CRP [38], and the expressions of fibrinogen-a/b and serum

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amyloid A are also repressed by PPARa [66]. At the transcriptional level, the reduction effects may be generated by different mechanisms. Activation of PPARa stimulates the expression of the protein inhibitor of kappaB (IkB) a, which sequesters the NF-κB subunits in the cytoplasm and consequently reduces their DNAbinding activity. As a result, the amount of nuclear p50-NFkB ~ C/EBPb complexes, which are crucial for inflammatory gene transcription, is decreased [67]. In parallel, the expression of both these factors is also lowered. The biological and therapeutic activities of PPARa are therefore the result of a combination of both the trans-activating and trans-repressive properties of this receptor. In addition, post-translational modifications, as well as promoter architecture and ligand structure, are important regulatory determinants. This raises the possibility of designing ligands with dissociated transactivating and trans-repressive activities, thus enabling specific targeting of gene subsets.

PPARa and Hepatocarcinogenesis (HCC) PPARa agonists cause liver tumors in rodents via PPARa-dependent mechanisms [68]. In support of this role of PPARa in rodent hepatocarcinoma, no hepatocellular proliferation or liver tumors were observed in PPARa-null mice fed with the potent PPARa agonist Wy-14643 for 11 months, while all wild-type mice developed liver tumors [69]. PPARa agonists were shown to increase oxidative stress through overproduction of reactive oxygen species and increased hepatic cell proliferation, both of which are critical for the development of HCCs [70, 71]. Thus, it is plausible that the increasing concentrations of H2O2 may generate DNA damage and participate in the tumorigenic effect of PPARa activators in mice. Besides the oxidative stress, a new signaling mechanism regulated by PPARa has been proposed, which implicates micro-RNAs (miRNAs) in PPARa inducedhepatocarcinogenesis [72]. MiRNAs represent a large class of non-coding RNAs that are transcribed in the nucleus as single primary transcripts or large polycistronic transcripts encoding several miRNAs. Increasing evidence involves several miRNAs in tumorigenesis [73, 74]. The above-mentioned study demonstrated the

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role of let-7C miRNA in the down-regulation of the expression of c-myc, an oncogene known to be induced after PPARa agonist treatment [72]. Thus, following Wy-14643 treatment, PPARa inhibited let-7C, leading to an increase of c-myc expression. In turn, c-myc induced the expression of miRNA polycistronic clusters, known to promote HCCs. Another role of PPARa in HCC has emerged on the basis of the interference of the hepatitis C virus (HCV) with lipid metabolism [75]. HCV is one of the major causes of chronic hepatitis; patients with persistent HCV infection have a high incidence of HCC [76]. Mice engineered to over-express the HCV core protein exhibited hepatic steatosis [77]. Coincident with the role of PPARa in lipid metabolism, this isotype was shown to be essential for the HCV core protein-induced hepatic steatosis and HCC [75]. In fact, persistent and potent PPARa activation was a prerequisite for the development of the severe steatosis and HCC induced by the HCV core protein. The link between HCV infection and PPARa offers clues in the search for new therapeutic and nutritional management options for chronically HCV-infected patients. PPARs have been implicated in numerous cancer types, where they regulate autonomous processes in tumor cells, such as apoptosis, proliferation, and differentiation, by interacting with major pathways involved in carcinogenesis. These points have been addressed recently, as well as the action of PPARs on the tumor cell environment and their ability to modify angiogenesis, inflammation, and immune cell functions [78, 79].

PPARs and Liver Fibrosis Chronic liver diseases remain an important cause of mortality and morbidity. Recurring or chronic injuries and inflammation trigger tissue remodeling pathways that can lead to severe fibrosis and eventually end-stage cirrhosis [80, 81]. Activation of hepatic stellate cells (HSC) plays an important role in the regulation of liver repair. In the damaged areas, the transition of normally quiescent HSCs to proliferative myofibroblasts, through the action of cytokines and oxidative stress, is accompanied by the production of collagen and extra-cellular matrix components [81]. Activation of PPARa produces antifibrotic effects in liver in different models [82, 83]. Moreover, there is a decreased expression of PPARa

20  PPARa, A Key Regulator of Hepatic Energy Homeostasis in Health and Disease

and liver fatty acid binding protein (L-FABP) after partial hepatectomy of rats and mice [84]. In addition, liver regeneration in PPARa-null mice is transiently impaired and is associated with the altered expression of genes involved in cell cycle control, cytokine signaling, and fat metabolism. Collectively, the results from rodent models support the hypothesis that PPARa is necessary for cell cycle progression, possibly via mechanisms involving the small GTPases Ras and RhoA [84–86]. The PPARg isotype suppresses the growth and fibrotic activity of HSCs by the down-regulation of collagen and components of the extra-cellular matrix. At the transcriptional level, the reversal of the activated HSC to a more quiescent phenotype involves suppression of AP-1 and NF-1 dependent transcription activities [87, 88]. On the contrary, in a model of carbon tetrachloride-induced acute liver damage, PPARb/d activation induced HSC proliferation during early fibrogenesis and enhanced expression of fibrotic markers [24]. Collectively, these observations suggest conflicting effects of both PPARa and PPARg versus PPARb/d. Thus, pharmacological manipulation of the balance of PPAR isotype expression in HSC, if made possible, would be promising for the attenuation of liver fibrosis progression in the damaged liver.

PPARs and Nonalcoholic Fatty Liver Disease Nonalcoholic fatty liver disease (NAFLD), defined by a liver fat content exceeding 5%, is a major cause of liver disease. Its prevalence is high, as an estimated 30% of adults and 10% of children and adolescents in the USA suffer from NAFLD [89]. It can progress to nonalcoholic steatohepatitis (NASH) and eventually to cirrohosis and HCC. NAFLD is associated with type 2 diabetes, hypertriglyceridemia, and obesity, and it represents the hepatic signature of the metabolic syndrome [89]. The pathogenesis of NASH is often conceptualized as a two-step process, consisting of hepatic triglyceride accumulation, followed by the development of oxidative stress and cytokine expression leading to steatohepatitis [90]. As mentioned earlier in this chapter, PPARa and PPARg modulate hepatic TG accumulation, which is the primary cause of NAFLD development. Most likely because of the anti-inflammatory and anti-fibrotic roles of PPARg, its ligands reduce liver damage in patients

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with NASH [91]. Therefore, it is not surprising that a growing number of observations involve PPARa and PPARg in the pathogenesis and treatment of NAFLD. The data that link these two PPARs to NAFLD/NASH have been reviewed recently, as well as the potential use of PPAR ligands for the treatment of NASH [92].

Conclusion The analysis of PPAR functions in health and disease, underscores a critical role of PPARs and their ligands in the fine-tuned adaptive response to fluctuating nutrient supplies. In addition to functions in nutrient ­metabolism, PPARs are also involved in other energy-­demanding vital processes, such as immune response and cell proliferation, which promote inflammation and cancer when ill-controlled [60] (Fig. 20.2). For many years, the use of the PPAR agonists, fibrates, and glitazones has confirmed the wide range of actions of PPARs on lipid metabolism and glucose homeostasis. An optimized clinical application of these drugs would promote the effective management of a wide spectrum of chronic metabolic disorders such as dyslipidemia, type 2 diabetes, and metabolic syndrome. However, the whole potential of the protective effect of PPARs in acute liver injuries and inflammation has not, as yet, been fully explored. Despite the proven benefits of targeting PPARs, safety concerns have recently led to late stage development arrests of new specific PPARg agonists and dual PPARα/γ agonists [93]. These concerns are based on signs of myopathy and rhabdomyolysis, an increase in plasma homocysteine and creatinine, fluid retention, weight gain, peripheral edema, potentially increased risk of cardiac failure and, finally, potential carcinogenicity in rodents [93]. Today the preclinical and clinical adverse events of PPAR agonists that could be of concern are well characterized and should help the development of new PPAR ligands. For instance, the inhibitory function of PPARs that could be exploited for the development of PPAR modulators with repressive properties has not been much explored. The production of new selective PPAR modulators (SPPARMs) with a broad spectrum of activities, ranging from inhibition to stimulation of gene expression, and reducing adverse effects, would represent an important extension of the therapeutic possibilities of PPARs in the future, especially in the context of inflammation and cancer, in addition to metabolism.

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N. Leuenberger and W. Wahli

Summary

›› PPARs are ligand-activated transcription factors ›› ›› ›› ››

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that are members of the nuclear receptor superfamily. Three PPAR isotypes have been identified: PPARa (NR1C1), PPARb/d (NR1C2), and PPARg (NR1C3). In rodents, PPARa is highly expressed in liver, especially in parenchymal cells. Ligands for PPARa comprise lipid-lowering fibrate drugs, long-chain unsaturated fatty acids, and eicosanoids. Because of its crucial role in lipoprotein, cellular fatty acid transport, and fatty acid b-oxidation, PPARa has beneficial effects in lipid-related diseases such as dyslipdemia and NAFLD. In contrast to their beneficial effects in humans, PPARa agonists cause liver tumors in rodents. However, this negative effect has not been observed in humans. Because of its multiple connections with hepatic pathways, the whole potential of PPARa as an important pharmaceutical target in the treatment of hepatic diseases has not, as yet, been exploited.

Multiple Choice Questions 1. Which type of compounds can dock into the PPARa ligand binding domain? (a) Triglycerides (b) Fatty acids (c) Retinoic acid (d) Cholesteryl esters (e) Phospholipids 2. In the liver, PPARb/d mRNA is well expressed in: (a) Stellate cells (b) Kupffer cells (c) Hepatocytes (d) Biliary epithelial cells (e) Endothelial cells

3. During fasting, PPARa-null mice suffer from: (a) Hypoglycemia (b) Hyperketonemia (c) Hypoinsulinemia (d) Hypercholesterolemia (e) Hypothermia 4. Activated PPARa increases the expression of the gene encoding: (a) CRP (b) FGF21 (c) Fibrinogen (d) Serum Amyloid A (e) FGF2 5. Which type of compounds are ligands for PPARg? (a) Fibrates (b) Biguanides (c) Glitazones (d) Corticosteroids (e) Retinoids Acknowledgments  The authors thank Dr. Liliane Michalik for discussions and stimulating comments on the manuscript. They also acknowledge grant support from the Swiss National Science Foundation and the Etat de Vaud.

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21

Bile Acids and Their Receptors Thierry Claudel and Michael Trauner

Introduction In animals, cholesterol is an essential molecule for membrane formation and synthesis of hormones and bile acids (BAs). Excess of cholesterol, either due to food consumption or endogenous synthesis, leads to gallstone formation and atherosclerosis. BAs produced from cholesterol and free cholesterol are secreted into bile and subsequently eliminated via feces, the only route to eliminate excess of cholesterol. As a major secretory pathway, the different steps of bile formation are precisely controlled and coordinated mostly via a complex network of nuclear receptors. Nuclear receptors (NRs) are transcription factors that, upon ligand binding and cofactors recruitment, modulate polymerase II activity and therefore gene expression, after binding to highly specific DNA response elements located in gene promoters. The Farnesoid X Receptor (FXR, NR1H4) [1] is a bile acid activated nuclear receptor [2–4], regulating several key steps of hepatic physiology such as bile formation, phase I/II metabolism, and glucose, lipid, and lipoprotein metabolisms [5]. Moreover, other NRs like Pregnane X Receptor (PXR; NRI2) [6, 7], Vitamin D Receptor (VDR; NR1I1) [8], and Constitutive Androstane Receptor (CAR; NR1I3) [9, 10] were identified as additional bile acid responsive NRs. In addition to NRs, BAs can also activate a membrane receptor for BAs (TGR5/BG37) [11, 12], a step which does not require

M. Trauner () Laboratory of Experimental and Molecular Hepatology, Division of Gastroenterology and Hepatology, Department of Internal Medicine, Medical University, Graz, Stiftingalstrasse 24, 8010 Graz, Austria e-mail: [email protected]

bile acid uptake into target cells. Both FXR and TGR5 may play an important role in the pathogenesis and treatment of a variety of hepatic and extrahepatic metabolic disorders including cholestasis, fatty liver, diabetes, dylipidemia, and atherosclerosis.

Role of Bile Acid Activated Nuclear Receptors for Bile Acid Metabolism, Bile Secretion and Cholestasis Background Hepatocytes secrete bile at the canalicular level, which is further modified by the bile duct epithelium. Hepatocellular secretion of BAs secretion is mediated by ATP-binding cassette (ABC) transporters and is the rate limiting step in bile formation [13]. Water movement via aquaporins follows the canalicular excretion of BAs and constitutes the “bile acid-dependent bile flow.” Moreover, BAs promote canalicular secretion of phospholipids and cholesterol to form mixed micelles with them [14]. Canalicular excretion of reduced glutathione and bicarbonate accounts for the major components of the “bile acid-independent” fraction of bile flow [15]. “Canalicular bile” is further modified by secretory and absorptive processes along the bile ductules and ducts (“ductal bile” secretion) [16–18]. BAs secreted into bile are reabsorbed in the intestine, taken up again by the liver and re-secreted into bile in an “enterohepatic circulation.” Since intestinal BA extraction is efficient, only 0.5 g of BAs are lost through fecal excretion per day and must be replaced by de novo synthesis accounting for 3–5% of the BAs excreted into bile [13, 14, 19]. BAs escaping the ­first-pass clearance by

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_21, © Springer-Verlag Berlin Heidelberg 2010

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the liver are filtered at the glomerulus and excreted into urine, where they are also actively reabsorbed via transporters in the brush border of the proximal ­convoluted tubule [19, 20]. BAs may also undergo “­cholehepatic shunting” from the bile duct lumen, via cholangiocytes and the periductular capillary plexus back to hepatocytes [20]. Although this pathway plays a minor role for conjugated BAs under normal physiologic conditions, it may become an important escape route under cholestatic conditions [20].

Regulation of Bile Acid Synthesis by NRs BAs are detergents for lipid digestion and absorption and have a broad spectrum of signaling properties beyond bile formation including effects on lipid and glucose homeostasis, thermogenesis, liver regeneration, fibrosis, immuno-modulatory effects, and carcinogenesis [5, 21, 22] all requiring a tight control of BA concentration. BAs are synthesized exclusively in the liver from cholesterol, [23] via the rate limiting enzyme cholesterol 7a-hydroxylase (CYP7A1), which is mainly regulated at the transcriptional level in a positive feed-forward fashion by cholesterol and negatively by BAs [24]. In rodents, the NR cholesterol sensor Liver X Receptor alpha (LXRa; NR1H3) directly mediates feed-forward regulation of CYP7A1 via a bile acid response element (BARE I) in the CYP7A1 promoter resulting in increased bile acid synthesis [25, 26] (Fig.  21.1). The human CYP7A1 promoter lacks this LXR/BARE I binding site and regulation is only mediated via negative feedback mechanisms, although one study showed that high cholesterol consumption in human expands the bile pool without increasing serum cholesterol levels [27]. Until recently, BAs were supposed to repress their own synthesis via a negative feedback mechanism by activation of FXR and induction of the Small Heterodimer Partner (SHP; NR0B2), which in turn negatively interacts with Liver Receptor Homolog 1 (LRH-1; NR5A2) and suppresses CYP7A1 gene transcription [28, 29]. Therefore, Hepatocyte Nuclear Factor 4 alpha (HNF4a; NR2A1) and LRH-1 binding to the upstream BARE II of the promoter were thought to be essential for CYP7A1 baseline expression and negative interference with either HNF4a or LRH-1 to be critical for reducing CYP7A1 transcription [24, 30,

T. Claudel and M. Trauner

FXR�? HNF4� LXR� LRH-1

Cholesterol

Liver

Cyp7a1

SHP ? JNK

Cyp8b1

BAs

PPAR� PXR

FGFR4

B k l o t h o

FGF19/15 BAs FGF19/15

FXR�

Intestine

Fig.  21.1  Negative and positive feedback regulation of key enzymes controlling BAs production.FGF15/19, synthesized in intestine upon FXR activation by BAs, activate FGFR4/bKlotho in liver that in turn results in CYP7A1 repression via JNK and possibly SHP. Nuclear receptors HNF4a, LXRa and LRH-1 are transcriptional activators of CYP7A1 and CYP8B1 respectively. PXR and PPARa are repressors of CYP7A1 expression

31]. However, HNF4a-deficient mice have low CYP7A1 expression [30] and LRH-1-deficient mice do not display a dysregulation of CYP7A1, but rather a marked decrease of CYP8B1 expression, despite very low levels of SHP and unchanged HNF4a expressions [32, 33]. Moreover, LXR induction of target genes is unchanged in LRH-1-deficient mice, which is notable since LRH-1 was previously described as a LXR competent transcription factor [32]. LRH-1deficient mice had a similar phenotype as CYP8B1deficient animals [34]. Together with the weak bile phenotype of SHP-deficient mice [35, 36], these novel insights from LRH-1 deficient mice suggest that other hepatic or extrahepatic (e.g., intestinal) factors may be more relevant (Fig. 21.1) [37]. Several FXR and SHP independent mechanisms regulating CYP7A1 transcription have also been identified. BAs can directly decrease HNF4a promoter activity and gene expression [38], but also impair HNF4a-mediated activation of the CYP7A1 promoter by blocking the recruitment of co-activators, such as peroxisome proliferator activated receptor

21  Bile Acids and Their Receptors

gamma coactivator (PGC-1a) and cAMP response element binding protein-binding protein (CBP) to HNF4a [39]. Furthermore, PXR, which is activated by hydrophobic bile acids can also impair HNF4a binding to the CYP7A1 promoter by reducing the interaction of PGC-1a with HNF4a, thus leading to inhibition of human CYP7A1 gene transcription in vitro [40] (Fig. 21.1). Likewise, peroxisome proliferator activated receptor alpha (PPARa; NR1C1) reduces CYP7A1 transcription via reduced HNF4a binding, effects which might contribute to the increased risk of gallstone formation after treatment with fibrates (PPARa activators) [41–43] (Fig. 21.1). In addition, bile acidstimulated cytokine release from macrophages can decrease CYP7A1 transcription via activation of the c-Jun terminal kinase (JNK) pathway and the reduction of HNF4a binding in vitro [44, 45]. In addition to hepatic factors, intestinal (ileal) factors also have a profound impact on CYP7A1 gene [46]. FXR-induced-ileal fibroblast growth factor 15 (FGF15) in mouse [47] (FGF19 in humans [48]) signals via either the portal vein or the lymphatic vessels to the liver, where it binds with FGFR4, a widely distributed receptor with tyrosine kinase activity [49] and represses CYP7A1 through a JNK-dependent pathway [47], (Fig. 21.1). The interactions between FGF19 and FGFR4 are mediated via the membrane bound protein bKlotho [50], which allows a tissue specific fine-tuning of the ligand-receptor interaction [51–53]. Experiments with liver and ileum specific FXRdeficient mice, suggest that the ileal route of CYP7A1 repression via the FGF15 pathway dominates over hepatic negative feedback pathways, indicating that a functional gut-liver signaling is a pre-requisite for bile acid homeostasis [37] (Fig.  21.1). As such, patients with bile acid malabsorption may suffer from excessive fecal bile acid excretion and subsequent bile acid induced diarrhea and steatorrhea as a result of interrupted ileal negative feedback regulation of bile acid synthesis. Moreover, FXR agonists or FGF19 peptide could be used therapeutically to interrupt the cycle of excessive bile acid production in patients with bile acid malabsorption [54]. Another enzyme, CYP8B1, controls the relative hydrophobicity of the bile pool through chenodeoxycholic acid (CDCA) synthesis and CYP8B1-deficient mice lack the negative feedback on CYP7A1, probably as a consequence of cholic acid (CA) elimination, despite preserved hepatic FXR-target gene expressions

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[34]. It is tempting to speculate that in CYP8B1deficient mice, FXR may fail to induce FGF15 as a consequence of the lack of its activator CA in the intestine [34]. It was suggested that HNF4a could be inhibited by SHP by mechanisms similar to LRH-1/CYP7A1 [55]. However, LRH-1-deficient mice have reduced CYP8B1 expression and low CA levels in their bile pool composition, with normal levels of HNF4a and its target genes, as well as very low levels of SHP. Therefore, HNF4a activity may not be dramatically important for CYP8B1 and LRH-1 appears to be the central regulator of CYP8B1 expression [32, 33], thus, indirectly controlling CYP7A1 expression by generating intestinal FXR activators (Fig. 21.1).

Regulation of Phase I and II Bile Acid Metabolism Phase I hydroxylation/phase II conjugation renders BAs more hydrophilic and amenable for urinary excretion, which is an additional elimination route for BAs under cholestatic conditions [56, 57] (Fig. 21.2). Human CYP3A4 and the rodent CYP3A11 are the main cytochrome P450 enzymes for BA metabolism as well as for more than 50% of xenobiotic hydroxylation [58, 59]. Expression of CYP3A4 is positively regulated by several NRs like FXR [60], PXR [6, 7], VDR [61], and CAR [62] (See also chapter on PXR and CAR). Administration of ligands for these receptors (xenobiotics, drugs but also BAs) induce CYP3A4 expression and subsequent phase I detoxification reactions [59]. Thus, BAs – being both activators and substrates of CYP3A4 – can initiate a feed-forward mechanism to minimize hepatocellular damage. BAs conjugations with sulfate or glucuronidate are additional mechanisms of BA detoxification. Dehydroepiandrosterone-sulfotransferase (SULT2A1) catalyzes sulfo-conjugation of a broad range of endogenous compounds including BAs turning their substrates into more water soluble compounds [63, 64]. In human bile acid, sulfation predominantly occurs under cholestatic conditions as reflected by the appearance of sulfated BAs in serum and in urine of patients with cholestatic liver diseases [65–67]. FXR, PXR, VDR, and CAR positively regulate SULT2A1 expression

320 Fig. 21.2  Nuclear receptors control BAs detoxification. Oxidation is the first step of BA detoxification mainly accomplished by CYP3A11/ CYP3A4 and CYP2B10 under the control of CAR, PXR, VDR and FXR. BAs are then glucuronidated via UGT2B4 or sulfated via SULT2A1. FXR up-regulates UGT2B4, while FXR, VDR, CAR and PXR up-regulate SULT2A1. Finally, BAs are excreted via MRPs and BSEP in the blood or in the bile and eliminated in urine or feces respectively

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[68–73] (Fig.  21.2). Moreover, cholestatic patients also display a high proportion of glucuronidated BAs in urine [74, 75]. Glucuronidation of bile acids is catalyzed by the UDP-glucuronosyltransferases UGT2B4 and UGT2B7, which again facilitate their renal elimination [76, 77]. BAs induce human UGT2B4 via activation of FXR [76], but UGT2B4 promoter also contains a PPAR response element and is activated by the PPARa agonist fenofibrate [77]. A cross-talk between FXR and PPARa pathways was provided by identification of PPARa as an FXR target gene [78]. Thus, BAs might induce UGT2B4 expression directly via activation of FXR and indirectly via FXRdependent induction of PPARa. In contrast to the UGT2B4, UGT2B7 seems to be repressed by hydrophobic bile acids through a negative FXR response element in the UGT2B7 promoter in vitro [79].

Regulation of Hepatobiliary Transport by NRs Basolateral Hepatocellular Bile Acid Uptake BAs hepatic uptake is mediated by a high-affinity Na+dependent bile acid transporter NTCP (SLC10A1) and a family of multi-specific organic anion transporters (OATPs; SLC21A) that mediate Na+-independent uptake of amphipathic compounds, including conjugated or free BAs or bilirubin. Na+-independent BA uptake is

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quantitatively less important than Na+-dependent uptake and is mediated by facilitated exchange with intracellular anions [13]. Regulation of NTCP by BAs differs considerably among humans, mice, and rats [80]. So far, the best studied rat NTCP promoter is trans-activated by several positive regulating elements including HNF-1a, Retinoic acid receptor alpha/ Retinoic X receptor alpha (RARa; NR1B1/RXRa; NR2B1) heterodimer or the signal transducer, and trans-activator 5 (Stat 5) [81] (Fig. 21.3). Negative feedback inhibition of NTCP is mediated via FXR-SHP dependent and independent mechanisms, and thus limits hepatocellular bile acid uptake [82, 83]. Induction of SHP by bile acid-activated FXR interferes with RXRa/RARa mediated activation only of the rat NTCP promoter [84]. Moreover, bile acid-induced SHP inhibits the activity of HNF4a [38, 55, 85]. SHP-independent mechanisms could involve activation of the JNK signaling pathway by BAs [36], which leads to RXRa phosphorylation and then reduced binding of RXRa/RARa to the rat NTCP promoter [86]. FXR seems also to play a major role in NTCP regulation at least in mice, since CA-fed and bile duct-ligated FXR knock-out mice fail to down-regulate NTCP [87]. In humans, the NTCP promoter lacks the RXRa/RARa and HNF4a response elements [80], and SHP acts mainly by suppressing the glucocorticoid receptor (GR; NR3C1)-mediated activation of human NTCP [88], which might account for NTCP down-regulation in human cholestatic liver ­diseases [89–91]. Similar to NTCP, repression of the

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Fig.  21.3  Nuclear receptors and transcriptional regulation of hepatocellular BA transport. Uptake: HNF1a positively controlled by HNF4a, regulates basolateral Na+-dependent (NTCP) and independent (OATP1B1) BA uptake. HNF4a may be under negative control of the FXR-SHP or directly inhibited by BAs. NTCP expression is increased by STAT5, GR and RAR. FXR

up-regulates only human OATP1B3. Canalicular: All canalicular transporters involved in bile formation (BSEP, MDR2, MRP2, ABCG5/8) are up-regulated by FXR. RARa/RXRa also positively regulate MRP2 and PPARa regulates MDR2. Alternative Export: CAR positively regulates MRP3/4, while PXR and VDR up-regulated MRP3. FXR only induced OSTa/b

sodium-independent bile acid uptake system in humans, OATP1B1 is also mediated by FXR involving SHP, HNF4a, and HNF1a, the latter being a strong activator of the OATP1B1 promoter [92]. In contrast, the multi-specific uptake system for organic anions, xenobiotics and potentially bile acids OATP1B3 is activated by FXR [93] even under cholestatic conditions when other uptake systems are repressed. This could preserve the metabolic functions of the liver in xenobiotic disposal during cholestasis. Alternatively, OATP1B3 could also function as a basolateral export system, since OATPs act as anion exchangers, and bi-directional transport of bromosulphthalein and taurocholate has been observed [94]. Taken together, BAs regulate their hepatocellular supply on demand, mainly by negative feedback regulation of uptake transporters via FXR.

Canalicular Bile Acid Excretion Canalicular excretion of BAs and non-bile acid organic anions via ABC transporters represents the rate limiting step in bile formation. Monovalent BAs such as ­glycine- or taurine-amidates of CA, CDCA and ursodeoxycholic acid (UDCA) are excreted into the bile canaliculus via the bile salt export pump BSEP (ABCB11) [95, 96] (Fig.  21.3). Divalent BAs with two negative charges such as sulfated tauro- or glycolithocholate are transported by multidrug resistanceassociated protein  MRP2 (ABCC2) [97]. MRP2 also mediates the excretion of a broad range of non-bile acid organic anions, mostly conjugates with glutathione, glucuronidate, and sulfate formed by phase II conjugation in the hepatocyte and of reduced glutathione (GSH) [97–100]. Additional transport systems in

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the canalicular membrane include a multidrug export pump (MDR1) for various drugs, a phospholipid floppase (MDR3/Mdr2 in rodents) for phosphatidylcholine translocation, the cholesterol two half-transporters ABCG5/8 for sitosterol and cholesterol export, a P-type ATPase (Fic1/FIC1; ATP8B1) mutated in hereditary cholestasis, whose substrate/function is still unclear and an Cl−/HCO3− anion exchanger 2 (SLC4A2/ AE2), all of them involved in bile formation (for review see [101–104]). BAs can promote their own biliary elimination by increasing both BSEP and MRP2 in a feed-forward manner [105, 106] (Fig. 21.3). Human, rat, and mouse BSEP promoters are transcriptionally activated by FXR [107–109] and BSEP baseline-expression is reduced in FXR knock-out mice [106, 110, 111]. A role for VDR in BSEP repression via direct VDR-FXR interaction has been postulated in vitro [112]; but given the very low levels of VDR expression in hepatocytes [113], a major contribution to negative feedback regulation of BSEP via this mechanism seems unlikely. In contrast to BSEP, transcriptional activation of MRP2 involves several NRs, reflecting the diverse substrate spectrum of MRP2. FXR binds to response elements in the promoter that are shared with CAR and PXR [114] (Fig. 21.3). Thus, BAs, as well as several CAR and PXR ligands, induce human and rodent MRP2 expression [92]. FXR also has been shown to enhance human MDR3 transcription, while PPARa stimulates the expression of rodent Mdr2 [102, 115, 116]. Taken together, orthograde canalicular bile acid efflux is mainly mediated via feed-forward regulation, involving FXR as a critically transcription factor.

Alternative Basolateral Bile Acid Export While BAs are excreted into canalicular bile under normal conditions, basolateral BA excretion back into portal blood may represent an alternative elimination route for BA during cholestasis, when canalicular excretion is impaired. Alternative basolateral bile acid export is mediated by MRP3 and MRP4, and the organic solute transporters, OSTa and OSTb [92] (Fig. 21.3). These export systems are normally expressed at very low levels at the basolateral membrane under normal conditions but can be significantly up-regulated in cholestasis [20, 92]. Since MRP3, MRP4, and OSTa/OSTb are

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able to transport sulfated as well as glucuronidated bile acids, the induction of these transporters may explain the shift towards renal excretion of these BAs as a major mechanism for bile acid elimination in patients with chronic cholestasis [92]. Induction of both MRP3 and MRP4 is independent of FXR in rodent models of cholestasis [106, 111, 117, 118] (Fig. 21.3), but PXR and VDR are able to induce human and rodent MRP3 expression [119, 120] while CAR ligands can induce both human and rodent MRP3 and MRP4 expression [10, 71, 121–123]. The bile acid transporter OSTa/b is induced by FXR and baseline OSTa/OSTb expression is reduced in FXR knock-out mice [124–126]. Taken together, a complex picture is emerging where multiple NRs (including FXR, PXR, VDR, and CAR) are required for coordination of adaptive basolateral bile acid efflux under bile acid load and cholestatic conditions.

Therapeutic Principles Targeting Nuclear Receptors in Cholestasis Counteracting cholestatic disorders should be a multilevel approach to target impaired physiological ­functions and pathophysiological mechanisms of bile acid/bilirubin transport and metabolism and to create/ support adaptive rescue pathways for potential toxic accumulating biliary compounds. Therapeutic strategies may therefore be aimed at NRs and will also target consequences of cholestasis such as inflammation and fibrosis. So far, the only approved drug for treatment of cholestatic disorders is UDCA [127]. The effects of UDCA are mediated via transcriptional and post-transcriptional mechanism but no unique definite nuclear receptor for UDCA has been found. Several studies have shown that UDCA was not a FXR ligand [2–4], while one study reported UDCA as a gene-selective partial FXR agonist inducing BSEP and down-regulating CYP7A1. Moreover, it was speculated that UDCA might activate PXR [118] and GR [128, 129], the latter in turn was able to increase CAR transcription via a glucocorticoid response element in the CAR promoter [130]. In line with its effects on PXR and CAR, UDCA stimulates bile acid hydroxylase CYP3A4/CYP3A11 in human and rodents respectively [118, 131] and, in

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accordance with its potential role as partial FXR agonist, down-regulates the key enzyme of bile acid synthesis CYP7A1 in vitro [132, 133]. Thus, UDCA appears to act on the metabolic level by induction of detoxification pathways and on transporter level by restoring defective transporters and generating alternative overflow-systems for accumulating biliary compounds [106]. At a post-transcriptional level UDCA may also directly activate canalicular transport function by inducing phosphorylation of ABC transporters, stimulating vesicular exocytosis, and inserting canalicular transport systems into the canalicular membrane [134–136]. In addition, UDCA also has antiapoptotic and antifibrotic properties contributing at least to some of the beneficial effects under UDCA treatment [127]. FXR agonists (e.g., 6 ECDCA) are promising treatment options for cholestasis and phase II studies have already been initiated in PBC. FXR agonists could overcome the reduction of bile flow in cholestasis via stimulation of BSEP (increasing bile acid-dependent bile flow) and MRP2 (increasing bile acid-independent bile flow) [137–139]. However, it may be simple to treat cholestasis by the stimulation of bile flow, particularly in the presence of bile duct injury. FXR agonists may also support some adaptive reactions of the cholestatic hepatocyte which would be predicted to limit the hepatocellular bile acid burden, such as downregulation of bile acid import via NTCP and OATP1A1, upregulation of basolateral overflow systems via OSTa/b, and reducing endogenous bile acid synthesis via down-regulation of CYP7A1 and CYP8B1 [20, 81, 92]. In addition, stimulation of the canalicular phospholipid floppase MDR2/MDR3 is predicted to change the intrabiliary bile composition by rendering bile less aggressive [115, 140]. In addition, the wider therapeutic use of FXR agonists could be jeopardized by interfering with FXR functions in lipid homeostasis such as HDL and triglyceride metabolism [5]. Drugs empirically used to treat pruritis, such as phenobarbital and rifampicin, may act as CAR and PXR agonists [20]. As such, animal experiments showed the induction of phase I, II, and III genes facilitating detoxification and elimination of BAs [121]. One can speculate that specific CAR and PXR modulators – separately or in combination with other drugs – could constitute a future therapeutic option for cholestasis. The future should bring us gene-selective agonists that ­specifically target subsets of genes and separate “desired” from “unwanted” effects.

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Role of Bile Acid Activated Nuclear Receptors for Lipid and Glucose Metabolism HDL Metabolism HDL-cholesterol transports excess cholesterol from the periphery back to the liver where it can be secreted as free cholesterol or as BAs into the bile and subsequently eliminated in the feces in a process called “reverse cholesterol transport.” As such, surgical ileum exclusion was shown to promote excretion of BAs and subsequently to increase HDL-cholesterol [141, 142] and apoA-I levels [143]. Further bile acid manipulations using sequestrants/resins such as cholestyramine, colestipol, or colesevelam, which bind to BAs in the intestine and induce bile acid malabsorption, increases HDL-cholesterol [144–146]. Conversely, dietary bile acid supplementation in human lowered HDL-cholesterol [147, 148] and finally accumulation of intrahepatic BAs in cholestatic patients lowered apoA-I serum levels [149, 150], whereas biliary diversion in PFIC patients increased their HDL-cholesterol levels [151]. Taken together, these results lead to the concept that bile acid-mediated FXR activation lowers HDL-cholesterol, while FXR deactivation increases HDL-cholesterol levels. Indeed, FXR activation by BAs in human apoAI transgenic mice lowers HDL-cholesterol levels by repressing apoA-I gene expression via a negative FXR response element located in the C-site of the promoter [150]. In conclusion, BAs lower HDL-cholesterol by impacting on key structural apolipoprotein (apoA-I) and by modulating enzyme activities involved in the modeling such as hepatic lipase (Fig.  21.4) [5]. Therefore, FXR agonists designed for long term treatment of human disorders like cholestasis must integrate the fact that broad nonselective FXR activation will most likely have severe detrimental cardio-vascular effects, through a decrease in HDL-cholesterol.

Triglyceride and Fat Metabolism Hypertriglyceridemia is a well-known risk factor for cardio-vascular disease [152–155]. Bile acid supplementation in normal, or hypertriglyceridemic patients

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resulted in lowering of triglycerides [156–159]. Bile acid malabsorption, due to either ASBT deficiency in the ileum and its treatment with sequestrants, or ileum resection was shown to reduce triglyceride levels [143, 160–164]. A more direct connection became apparent when hypertriglyceridemia was found in CYP7A1deficient patients [165]. FXR deficient mice were also shown to be hypertriglyceridemic due to the increase in apoCIII and decrease in apoCII [110, 166], while very low density lipoprotein receptor and SREBP1C were identified as FXR-induced and -repressed target genes respectively [5] (Fig. 21.4). In conclusion, BAs are hypotriglyceridemic molecules that impact on both production and clearance at multiple steps (Fig. 21.4). Since, triglycerides and apoCIII are independent risk factors for development of cardiovascular disease, bile acid like compounds modulating FXR and retaining this property could be of major pharmaceutical interest.

Glucose Metabolism Several lines of evidences indicate the role for BAs in regulating glucose metabolism. FXR expression was

shown to be induced by glucose in rodents, probably via the pentose phosphate pathway [167], a result that could explain why glucose enriched diet in normal [168] or in hypercholesterolemic patients lowers bile synthesis [169]. Since, total parenteral nutrition (TPN) patients have low BA synthesis when they are not feed by mouth [170]; it is tempting to speculate that interruption of entero-hepatic circulation and induction of FXR expression by glucose are likely to explain CYP7A1 repression in these patients, perhaps via FXR stimulation of hepatic FGF19 expression [48]. Several groups have examined the effects of FXRdeficiency and/or activation in mouse model of diabetes. As such, FXR deletion resulted in glucose intolerance and insulin resistance, while treatment using FXR synthetic agonist GW4064, or liver adenovirus over-expression of FXR, lowered blood glucose levels by repressing hepatic gluconeogenesis and enhancing glycogen synthesis and storage (Fig. 21.4) [171]. FXR mediated insulin-resistance was also shown to correlate with impaired peripheral disposal due to high levels of free fatty acids together with high glucose production in liver [172]. Taken together, the loss of FXR function results in insulin resistance and glucose intolerance (Fig. 21.4).

HDL metabolism

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Fig. 21.4  FXR and TGR5 control lipid and glucose homeostasis.BAs activation of FXR results in lowering of hepatic lipase and apoA-I and increase of PLTP (phospholipid transfer protein) expressions that altogether decrease HDL-cholesterol levels. Moreover, BAs increase apoCII and lower apoCIII and SREBP1c expression, resulting in lowered triglyceride levels. FXR activation by increasing glycogen synthesis and lowering gluconeogenesis results in insulin resistance. TGR5 activation, by increasing energy expenditure and GLP1 synthesis, increases insulin sensitivity. Different BAs are activating FXR and TGR5 but a cross-talk between the two signaling pathways is unknown

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In diabetic patients, beneficial effects of BAs binding resin may be due to depletion of hepatic cholesterol concentration as a result of increasing CYP7A1 activity on one hand by interfering with intestinal absorption capacity, on the other hand by normalizing dyslipidemia and improving glucose control in these patients. Additionally, since BAs control the release of the potent antidiabetic glucagon like peptide 1 ­(GLP-1) in intestinal enterocrine cells, one can speculate that BAs absorption modifications, could stimulate GLP-1 release and therefore impact directly on glucose metabolism. Future studies will have to clarify these issues.

Non Alcoholic Fatty Liver Disease Non alcoholic fatty liver disease (NAFLD), characterized by fat accumulation in the liver is associated with risk factors like obesity [173], diabetes [174], hypertension [175] – all known to define the metabolic syndrome. NAFLD is generally believed to have a benign course [176], because only 10% of NAFLD patients develop non alcoholic steatohepatitis (NASH), from which up to one third will ultimately progress to advanced liver fibrosis and cirrhosis [177]. However, NAFLD patients have a higher total mortality rate [178] as a result of increased cardiovascular mortality [179, 180]. SHP also plays an important role in NAFLD, since SHP expression was found to be induced in leptin deficient (OB−/−) mice and high-sucrose/high fat diet models of NAFLD, while SHP deficiency in OB−/−/SHP−/− double knock-out mice prevented fatty liver. SHP deletion increased serum TG levels in OB−/−/SHP−/− mice, via higher rates of hepatic VLDL-TG secretion due to increased expression of microsomal transfer protein [181]. In addition, OB−/−/SHP−/− mice also showed a reduced expression of FA uptake and de novo FA synthesis genes, which could contribute to protection against steatosis, whereas no effects on hepatic FA oxidation were observed [181]. The protective effect of SHP deficiency against obesity and NAFLD in mice contrasts the reported association of SHP haploinsufficiency with increased body weight in Japanese [182] and a possible link between decreased SHP activity and body weight in the European population [183, 184], although it does not appear to be a common cause of obesity [185].

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G-Protein-Coupled-Receptor for Bile Acid In 2002 and 2003, two independent groups cloned a G-protein-coupled receptor (GPCR) named TGR5 or BG37, and showed that it was a bile acid responsive membrane receptor [11, 12]. GPCRs belong to a family of 720 members in human genome, that transduce extracellular signals including BAs, fatty acids or phospholipids into an intra-cellular response involving generation of cyclic AMP (cAMP) or phosphorylation via MAP kinase. TGR5 was shown to be maximally activated by bile acids like LCA or DCA with a potency differing notably from FXR [11, 12]. Moreover, TGR5 is expressed in various tissues in which FXR is either absent or low as in leukocytes, spleen, lung, or white and brown adipose tissues [11, 12], with the highest expression found in gallbladder and colon [186, 187]. Mice receiving high fat diet containing BAs show a wasting phenotype characterized by lower body weight despite preserved food intake and increased energy expenditure [188]. The latter is due to bile acid stimulation of TGR5, resulting in the production of cAMP with subsequent induction of deiodinase 2 (D2) expression, converting inactive thyroid hormone T4 into active T3 form. T3, by activating nuclear receptors thyroid hormone receptors (TRa1, NR1A1), subsequently induces uncoupling protein 1 (UCP1) expression and therefore increases energy expenditure in brown adipose tissue (BAT) but not in white adipose tissue (WAT) [188]. Since TGR5/D2/UCP1 axis was found in BAT, the potential relevance in humans may be questioned. However, the TGR5/D2/UCP1 axis was also identified in vitro in a human muscle cell line, possibly permitting the extrapolation to the situation to humans [188]. Although adipose tissue in adult humans is mainly made of WAT and not BAT [189], UCP1 expression data suggest that up to 1/200 adipocytes in WAT may in fact be a brown adipocyte [190]. Moreover, the amount of BAT is induced in outdoor workers in cold countries [191], or during pathological situations like hibernomas [192] and pheochromocytoma [193, 194], but the overall contribution to energy dissipation may be negligible [195]. Proof in humans that TGR5 activation by bile acids, increases energy expenditure is therefore still lacking. Finally, since TGR5 is activated by high concentrations of bile acids, it is of interest to note that such quantities are achieved either transiently during a post-prandial

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state, under theapeutic bile acid feeding/therapy or ­during pathological conditions such as cholestasis and portosystemic shunting (bypassing the liver). TGR5 expression in leukocytes could also explain why cholestasis was shown to favor bacterial infection, since TGR5 stimulation in macrophages has been shown to suppress cytokine production as a consequence of high cAMP levels [196]. However, VDR activation by bile acids may also contribute to this effect [8].

Summary

›› BAs ›› ›› ›› ›› ›› ›› ›› ››

activate nuclear receptor FXR and G-coupled membrane receptor TGR5. BAs repress their own synthesis by a negative feedback on CYP7A1 involving nuclear receptor pathways. Oxidation, conjugation, and excretion of BAs are the three phases involved in BAs detoxification. NTCP and BSEP are the main BAs transporters involved in uptake and excretion in hepatocytes. FXR, PXR, CAR, and VDR are the NRs controlling BAs synthesis, transport, and detoxification. BAs lower apoA-I and HDL cholesterol via nuclear receptor signaling. BAs lower triglyceridemia by increasing clearance and decreasing production. BAs activation of FXR increases insulin resistance. BAs activation of TGR5 results in increase energy expenditure and GLP1 secretion

Multiple Choice Questions 1. How do BAs activate their uptake and excretion? (a) By up-regulating NTCP and BSEP (b) By repressing NTCP and BSEP (c) By up-regulating NTCP and repressing BSEP (d) By repressing NTCP and up-regulating BSEP (e) By up-regulating SHP and repressing CYP7A 2. Which nuclear receptor(s) is/are activated by BAs? (a) FXR, RXR, RAR (b) HNF4, RXR, FXR (c) FXR, PXR, VDR

(d) FXR, SHP, RXR (e) FXR, SHP, LRH-1 3. Which intestinal factor represses hepatic CYP7A1 expression? (a) FGF15/19 (b) beta Klotho (c) SHP/LRH-1 (d) FGFR4 (e) FXR 4. How does FXR activation affect HDL-cholesterol? (a) Lowers HDL via repression of apoAI (b) Increases HDL via induction of apoAI (c) Lowers HDL via repression of PLTP (d) Lowers HDL by inducing apo CIII (e) Does not affect HDL metabolism 5. What are the metabolic consequences of TGR5 activation? (a) Repression of CYP7A1 (b) Increased food intake and body weight (c) Increased energy expenditure and insulin sensitivity via GLP-1 (d) Lowered insulin sensitivity and energy expend­ iture (e) Increased energy expenditure and lowered insulin sensitivity

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21  Bile Acids and Their Receptors 138. Yu J, Lo JL, Huang L et al (2002) Lithocholic acid decreases expression of bile salt export pump through farnesoid X receptor antagonist activity. J Biol Chem 277(35):31441–31447 139. Claudel T, Sturm E, Kuipers F, Staels B (2004) The farnesoid X receptor: a novel drug target? Expert Opin Investig Drugs 13(9):1135–1148 140. Moschetta A, Bookout AL, Mangelsdorf DJ (2004) Prevention of cholesterol gallstone disease by FXR agonists in a mouse model. Nat Med 10(12):1352–1358 141. Grundy SM, Ahrens EH Jr, Salen G (1971) Interruption of the enterohepatic circulation of bile acids in man: comparative effects of cholestyramine and ileal exclusion on cholesterol metabolism. J Lab Clin Med 78(1):94–121 142. Heaton KW (1977) Disturbances of bile acid metabolism in intestinal disease. Clin Gastroenterol 6(1):69–89 143. Buchwald H, Varco RL, Matts JP et al (1990) Effect of partial ileal bypass surgery on mortality and morbidity from coronary heart disease in patients with hypercholesterolemia. Report of the Program on the Surgical Control of the Hyperlipidemias (POSCH). N Engl J Med 323(14): 946–955 144. Shepherd J, Packard CJ, Morgan HG, Third JL, Stewart JM, Lawrie TD (1979) The effects of cholestyramine on high density lipoprotein metabolism. Atherosclerosis 33(4):433–444 145. Brensike JF, Levy RI, Kelsey SF et al (1984) Effects of therapy with cholestyramine on progression of coronary arteriosclerosis: results of the NHLBI Type II Coronary Intervention Study. Circulation 69(2):313–324 146. Levy RI, Brensike JF, Epstein SE et al (1984) The influence of changes in lipid values induced by cholestyramine and diet on progression of coronary artery disease: results of NHLBI Type II Coronary Intervention Study. Circulation 69(2):325–337 147. Kuriyama M, Tokimura Y, Fujiyama J, Utatsu Y, Osame M (1994) Treatment of cerebrotendinous xanthomatosis: effects of chenodeoxycholic acid, pravastatin, and combined use. J Neurol Sci 125(1):22–28 148. Leiss O, von Bergmann K (1982) Different effects of chenodeoxycholic acid and ursodeoxycholic acid on serum lipoprotein concentrations in patients with radiolucent gallstones. Scand J Gastroenterol 17(5):587–592 149. Iglesias A, Arranz M, Alvarez JJ et al (1996) Cholesteryl ester transfer activity in liver disease and cholestasis, and its relation with fatty acid composition of lipoprotein lipids. Clin Chim Acta 248(2):157–174 150. Claudel T, Sturm E, Duez H et al (2002) Bile acid-activated nuclear receptor FXR suppresses apolipoprotein A-I transcription via a negative FXR response element. J Clin Invest 109(7):961–971 151. Melter M, Rodeck B, Kardorff R et al (2000) Progressive familial intrahepatic cholestasis: partial biliary diversion normalizes serum lipids and improves growth in noncirrhotic patients. Am J Gastroenterol 95(12):3522–3528 152. Albrink MJ, Man EB (1959) Serum triglycerides in coronary artery disease. Archives of Internal Medicine. 103: 4–8 153. Goldstein JL, Schrott HG, Hazzard WR, Bierman EL, Motulsky AG (1973) Hyperlipidemia in coronary heart disease II Genetic analysis of lipid levels in 176 families and delineation of a new inherited disorder, combined hyperlipidemia. J Clin Invest 52(7):1544–1568

331 154. Brunzell JD, Schrott HG, Motulsky AG, Bierman EL (1976) Myocardial infarction in the familial forms of hypertriglyceridemia. Metabolism 25(3):313–320 155. Genest JJ Jr, Martin-Munley SS, McNamara JR et al (1992) Familial lipoprotein disorders in patients with premature coronary artery disease. Circulation 85(6):2025–2033 156. Bateson MC, Maclean D, Evans JR, Bouchier IA (1978) Chenodeoxycholic acid therapy for hypertriglyceridaemia in men. Br J Clin Pharmacol 5(3):249–254 157. Begemann F (1978) Influence of chenodeoxycholic acid on the kinetics of endogenous triglyceride transport in man. Eur J Clin Invest 8(5):283–288 158. Camarri E, Fici F, Marcolongo R (1978) Influence of chenodeoxycholic acid on serum triglycerides in patients with primary hypertriglyceridemia. Int J Clin Pharmacol Biopharm 16(11):523–526 159. Camarri E, Marcolongo R, Zaccherotti L, Marini G (1978) The hypotriglyceridemic effect of chenodeoxycholic acid in type IV hyperlipemia. Biomedicine 29(6):193–198 160. Duane WC (1995) Abnormal bile acid absorption in familial hypertriglyceridemia. J Lipid Res 36(1):96–107 161. Duane WC, Hartich LA, Bartman AE, Ho SB (2000) Diminished gene expression of ileal apical sodium bile acid transporter explains impaired absorption of bile acid in patients with hypertriglyceridemia. J Lipid Res 41(9): 1384–1389 162. Angelin B (1995) 1994 Mack-Forster Award Lecture Review Studies on the regulation of hepatic cholesterol metabolism in humans. Eur J Clin Invest 25(4):215–224 163. Angelin B, Einarsson K, Hellstrom K, Leijd B (1978) Effects of cholestyramine and chenodeoxycholic acid on the metabolism of endogenous triglyceride in hyperlipoproteinemia. J Lipid Res 19(8):1017–1024 164. Molgaard J, von Schenck H, Olsson AG (1989) Comparative effects of simvastatin and cholestyramine in treatment of patients with hypercholesterolaemia. Eur J Clin Pharmacol 36(5):455–460 165. Pullinger CR, Eng C, Salen G et al (2002) Human cholesterol 7alpha-hydroxylase (CYP7A1) deficiency has a hyper­ cholesterolemic phenotype. J Clin Invest 110(1): 109–117 166. Claudel T, Inoue Y, Barbier O et al (2003) Farnesoid X receptor agonists suppress hepatic apolipoprotein CIII expression. Gastroenterology 125(2):544–555 167. Duran-Sandoval D, Mautino G, Martin G et al (2004) Glucose regulates the expression of the farnesoid X receptor in liver. Diabetes 53(4):890–898 168. DenBesten L, Reyna RH, Connor WE, Stegink LD (1973) The different effects on the serum lipids and fecal steroids of high carbohydrate diets given orally or intravenously. J Clin Invest 52(6):1384–1393 169. Stacpoole PW, Grundy SM, Swift LL, Greene HL, Sloni  AE, Burr IM (1981) Elevated cholesterol and bile acid synthesis in an adult patient with homozygous familial hypercholesterolemia Reduction by a high glucose diet. J Clin Invest 68(5):1166–1171 170. Dawes LG, Laut HC, Woodruff M (2007) Decreased bile acid synthesis with total parenteral nutrition. Am J Surg 194(5):623–627 171. Ma K, Saha PK, Chan L, Moore DD (2006) Farnesoid X receptor is essential for normal glucose homeostasis. J Clin Invest 116(4):1102–1109

332 172. Zhang Y, Lee FY, Barrera G et al (2006) Activation of the nuclear receptor FXR improves hyperglycemia and hyperlipidemia in diabetic mice. Proc Natl Acad Sci U S A 103(4):1006–1011 173. Ludwig J, Viggiano TR, McGill DB, Oh BJ (1980) Nonalcoholic steatohepatitis: Mayo Clinic experiences with a hitherto unnamed disease. Mayo Clin Proc 55(7): 434–438 174. Shibata M, Kihara Y, Taguchi M, Tashiro M, Otsuki M (2007) Nonalcoholic fatty liver disease is a risk factor for type 2 diabetes in middle-aged Japanese men. Diabetes Care 30(11):2940–2944 175. Bedogni G, Miglioli L, Masutti F, Tiribelli C, Marchesini G, Bellentani S (2005) Prevalence of and risk factors for nonalcoholic fatty liver disease: the Dionysos nutrition and liver study. Hepatology 42(1):44–52 176. Bedogni G, Miglioli L, Masutti F et al (2007) Incidence and natural course of fatty liver in the general population: the Dionysos study. Hepatology 46(5):1387–1391 177. Neuschwander-Tetri BA, Caldwell SH (2003) Nonalcoholic steatohepatitis: summary of an AASLD Single Topic Conference. Hepatology 37(5):1202–1219 178. Adams LA, Lymp JF, St Sauver J et al (2005) The natural history of nonalcoholic fatty liver disease: a populationbased cohort study. Gastroenterology 129(1):113–121 179. Ekstedt M, Franzen LE, Mathiesen UL et al (2006) Longterm follow-up of patients with NAFLD and elevated liver enzymes. Hepatology 44(4):865–873 180. Targher G, Bertolini L, Poli F et al (2005) Nonalcoholic fatty liver disease and risk of future cardiovascular events among type 2 diabetic patients. Diabetes 54(12): 3541–3546 181. Huang J, Iqbal J, Saha PK et al (2007) Molecular characterization of the role of orphan receptor small heterodimer partner in development of fatty liver. Hepatology 46(1): 147–157 182. Nishigori H, Tomura H, Tonooka N et al (2001) Mutations in the small heterodimer partner gene are associated with mild obesity in Japanese subjects. Proc Natl Acad Sci U S A 98(2):575–580 183. Echwald SM, Andersen KL, Sorensen TI et al (2004) Mutation analysis of NR0B2 among 1545 Danish men identifies a novel c.278G>A (p.G93D) variant with reduced functional activity. Hum Mutat 24(5):381–387 184. Hung CC, Farooqi IS, Ong K et al (2003) Contribution of variants in the small heterodimer partner gene to birth-

T. Claudel and M. Trauner weight, adiposity, and insulin levels: mutational analysis and association studies in multiple populations. Diabetes 52(5):1288–1291 185. Mitchell SM, Weedon MN, Owen KR et al (2003) Genetic variation in the small heterodimer partner gene and youngonset type 2 diabetes, obesity, and birth weight in U.K. subjects. Diabetes 52(5):1276–1279 186. Maruyama T, Tanaka K, Suzuki J et al (2006) Targeted disruption of G protein-coupled bile acid receptor 1 (Gpbar1/ M-Bar) in mice. J Endocrinol 191(1):197–205 187. Vassileva G, Golovko A, Markowitz L et al (2006) Targeted deletion of Gpbar1 protects mice from cholesterol gallstone formation. Biochem J 398(3):423–430 188. Watanabe M, Houten SM, Mataki C et al (2006) Bile acids induce energy expenditure by promoting intracellular thyroid hormone activation. Nature 439(7075): 484–489 189. Lean ME, James WP, Jennings G, Trayhurn P (1986) Brown adipose tissue uncoupling protein content in human infants, children and adults. Clin Sci (Lond) 71(3): 291–297 190. Oberkofler H, Dallinger G, Liu YM, Hell E, Krempler F, Patsch W (1997) Uncoupling protein gene: quantification of expression levels in adipose tissues of obese and nonobese humans. J Lipid Res 38(10):2125–2133 191. Huttunen P, Hirvonen J, Kinnula V (1981) The occurrence of brown adipose tissue in outdoor workers. Eur J Appl Physiol Occup Physiol 46(4):339–345 192. Zancanaro C, Pelosi G, Accordini C, Balercia G, Sbabo L, Cinti S (1994) Immunohistochemical identification of the uncou­­ pling protein in human hibernoma. Biol Cell 80(1): 75–78 193. Lean ME, James WP, Jennings G, Trayhurn P (1986) Brown adipose tissue in patients with phaeochromocytoma. Int J Obes 10(3):219–227 194. Ricquier D, Nechad M, Mory G (1982) Ultrastructural and biochemical characterization of human brown adipose tissue in pheochromocytoma. J Clin Endocrinol Metab 54(4): 803–807 195. Cannon B, Nedergaard J (2004) Brown adipose tissue: function and physiological significance. Physiol Rev 84(1): 277–359 196. Yoshimura T, Kurita C, Nagao T et al (1997) Inhibition of tumor necrosis factor-alpha and interleukin-1-beta production by beta-adrenoceptor agonists from lipopolysaccharide-stimulated human peripheral blood mononuclear cells. Pharmacology 54(3):144–152

Signaling Pathways in Liver Diseases: PXR and CAR

22

Catherine A.M. Stedman, Michael Downes, and Christopher Liddle

Nuclear Hormone Receptor Family Nuclear hormone receptors (NHRs), of which there are 48 unique members in humans and 49 members in mouse, function as ligand-activated transcription factors and have critical roles in diverse cellular processes ranging from mammalian development and differentiation to metabolic homeostasis [1]. NHRs bind to the sequence-specific DNA response elements on target gene promoters as homodimers, heterodimers, or monomers. Structural and functional analyses of the NHR family have demonstrated that these receptors are comprised of functional modular domains. The DNA binding domain (DBD) consists of a well-characterized zinc finger motif that recognizes a degenerate six to eight nucleotide sequence on the target DNA. The ligand-binding domain (LBD) resides in the C-terminal portion of the protein and shares a common, predominantly alpha helical fold [1]. As implied, this domain of the receptor is where cognate ligands of the receptors interact and induce conformational changes associated with transcriptional activation. Many of the known ligands for these receptors are essential metabolic products including retinoids, thyroid hormone, vitamin D3, bile acids, oxysterols, and prostenoids that act through their cognate receptors to control metabolic homeostasis [2, 3]. The transcriptional activity of NHRs is regulated by associated factors, specifically, co-activators and co-repressors that serve as scaffolding proteins to recruit chromatin

C. Liddle (*) Department of Clinical Pharmacology, Storr Liver Unit, Westmead Millennium Institute and University of Sydney, Westmead Hospital, Westmead NSW 2145, Australia e-mail: [email protected]

remodeling complexes which repress transcription through limiting access to gene targets, or activate transcription via unwinding chromatin. The association of co-regulators is determined by the liganded state of the receptor, as a ligand-induced conformational change promotes interaction of NHR co-­activators such as the p160 gene family [4] and P300, which recruit enzymes such as histone acetylases. Conversely, in the unliganded state the receptor is bound to scaffold corepressors such as silencing mediator of retinoic acid and thyroid hormone receptor (SMRT) and nuclear receptor co-repressor (NCoR) which recruit histone deacetylases [5]. In addition, NHRs are also instrumental in the ability of the body to respond and adapt to complex environmental cues. In particular, pregnane X receptor (PXR, NR1I2) and constitutive androstane receptor (CAR, NR1I3) function as master regulators of the body’s response to environmental stimuli and a subset endobiotics. PXR was discovered in 1998 [6–8] and it was almost immediately apparent that this receptor was an important regulator of genes intimately involved in drug metabolism, as exemplified by cytochrome P450 (CYP) 3A genes. Human CAR was discovered in 1994 [9], at which time its purpose was unknown, and in many ways it still remains an enigmatic receptor. It was not until 1998 that CAR was recognized as the transcription factor mediating the induction of CYP2B gene expression by phenobarbital [10, 11]. Thus, both PXR and CAR were initially characterized as transcription factors regulating the hepatic genes that encode xenobiotic metabolism [12], and therefore, commonly have been referred to as “xenoreceptors” or “xenosensors.” They are liver and gut-predominant in their expression patterns and exhibit little expression in other tissues (Fig. 22.1).

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_22, © Springer-Verlag Berlin Heidelberg 2010

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Fig. 22.1  Mouse tissue expression patterns of mRNA for the nuclear hormone receptors PXR and CAR demonstrating that these receptors are predominantly expressed in the liver and

PXR Biology As a primary xenobiotic sensor, PXR is the most promiscuous of all the NHR family in its ability to bind

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gut. Figures courtesy of the Nuclear Receptor Signaling Atlas project: (www.nursa.org/10.1621/datasets.02001)

and be activated by a chemically diverse range of molecules due to its large flexible ligand-binding pocket [13, 14]. Indeed, this ability of PXR to bind a wide variety of ligands has sometimes made development of

22  Signaling Pathways in Liver Diseases: PXR and CAR Fig. 22.2  Structures of the prototypical ligands for PXR and CAR that have been used to explore the functions of these receptors. Pregnenolone 16a-carbonitrile is a mouse and rat PXR ligand while Rifampin is a human PXR ligand. TCPOBOP is a mouse CAR ligand while CITCO is a human CAR ligand. Phenobarbital is an indirect CAR activator that causes CAR to undergo nuclear translocation and exert transcriptional activity without interacting with the ligand binding domain of the receptor, as described in the text

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synthetic ligands to other nuclear receptors challenging, as all too often they act as PXR agonists, something that is invariably undesirable due to the propensity of PXR to cause drug–drug interactions (see below). PXR has a major on impact human health through its regulation by many pharmaceutical ligands, including antibiotics such as rifampin, cholesterol lowering drugs of the statin class, the anti-neoplastic drug paclitaxel, and the commonly used herbal remedy – St John’s wort – to name a few [15]. In addition, protection from environmental contaminants is achieved through PXR activation, as exemplified by the endocrine disrupting chemicals such as non-planar polychlorinated biphenyls (PCBs) and organo-chloride pesticides such as trans-nonachlor and chlordane [16]. A role in the whole body homeostasis has also been demonstrated through PXR’s activation by and regulation of circulating levels of endobiotics including bile acids and steroid hormones [6]. Environmental and endobiotic exposure to varied chemical entities is often species specific due to

CI

N CITCO

variations in diet, environment, and physiology. PXR is an evolutionarily adaptive gene and the activation of this receptor protein by different stimuli can be species specific. For example, rifampin readily activates human PXR but fails to bind to mouse or rat PXR, while pregnenolone 16a-carbonitrile activates the latter species (Fig. 22.2). This species’ specificity has drawn intense interest from the pharmaceutical industry as it highlights the limitations of animal models for drug testing [6, 17].

CAR Biology Similar to PXR, CAR target genes include members of the phase I and phase II drug metabolism enzymes and transport pathway proteins that can either overlap with or be distinct from PXR targets. CAR was initially isolated in a yeast-2-hybrid screen as a factor that interacts with RXR and was termed “constitutive androstane receptor” as it was active in non-hepatic cells and could

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be repressed by compounds such as androstanol and andostenol [18]. CAR is unique in the NHR family, as its name implies, in non-hepatic cells the receptor resides in the nucleus bound to co-activators and is able to ­activate target genes in the absence of ligand. However, in hepatic cells, a major site of CAR expression, unliganded CAR is retained in the cytoplasm bound to the chaperones CAR retention protein (CCRP) and heat shock protein 90 (HSP90). Activation in hepatic cells is achieved in either of two mechanisms [19–21]. Direct binding of ligands such as 1,4-Bis[2-(3,5 dichloropyridyloxy)] benzene (TCPOBOP) to the mouse receptor [22] and 6-(4-Chlorophenyl)imidazo[2,1-b]thiazole-5carbaldehydeO-(3,4-dichlorobenzyl)oxime (CITCO) to human CAR [23] induces the translocation of the liganded receptor into the nucleus where it initiates the transcription of target genes, including CYP2B subfamily CYPs [24, 25] (Fig. 22.2). However, most activating compounds, as exemplified by phenobarbital, trigger CAR activity in hepatic cells through changes in the phosphorylation state of the LBD or associated chaperones that results in their dissociation, allowing CAR to translocate to the nucleus where its constitutive activity mediates expression of target genes [26]. No bona fide endogenous CAR ligands have been discovered to date leaving open the question as to the primary function of this receptor. Androstanol and andostenol work as antagonists for CAR by binding the LBD and dissociating the chaperones to facilitate nuclear translocation and subsequent recruitment of co-repressors [18].

Gene Regulation by PXR and CAR Response Elements in Target Genes PXR and CAR have been found to transcriptionally activate target genes by binding to conserved DNA response elements as heterodimers with RXRa. Both PXR and CAR efficiently bind direct or everted repeats of the core hexad AG(G/T)TC(A/C). As the CYP3A4 gene is a major contributor to inductive drug–drug interactions and is highly induced by a range of drugs, an understanding of cis-acting response elements within this gene was a logical starting point to understanding the mechanisms involved in the inductive process. PXR/ RXRa heterodimers were initially found to interact with an everted repeat with a six base spacer (ER-6) in the

C. A. M. Stedman et al.

human CYP3A4 proximal promoter [6, 7, 27]. However, in the context of the native CYP3A4 promoter, the proximal ER-6 element has no inherent ability to promote PXR-mediated transcription in experiments performed in liver-derived cell lines [28] or in transgenic mice [29]. PXR-mediated induction of the CYP3A4 gene is dependent on a distal xenobiotic-responsive enhancer module (XREM) located approximately eight kilobases upstream of the transcription initiation site [28]. The XREM region contains an additional high-affinity PXRRXRa binding site (a DR-3 element) as well as low affinity elements (including an ER-6 element), which work in a coordinate manner with the ER-6 element in the proximal promoter of CYP3A4 (Fig.  22.3). It has also been demonstrated that the proximal ER-6 and distal DR-3 response elements in CYP3A4, efficiently bind CAR/RXRa heterodimers and that CAR can transcriptionally activate CYP3A4 expression [30]. The CYP2B6 gene is also highly inducible by some xenobiotics, particularly phenobarbital. Two adjacent DR-4 elements separated by 16 base pairs are located 1.7 kilobases upstream of the transcription initiation site of CYP2B6, in a region referred to as the phenobarbitalresponsive enhancer module (PBREM) [11] (Fig. 22.3). These elements are capable of interacting with both CAR and PXR [31, 32]. Interestingly, the layout of the response elements in the CYP2B6 favors transactivation by CAR, despite the observation that DR-4 elements also efficiently bind PXR and VDR. In contrast, the widely spaced response elements in CYP3A4 appear to favor PXR-mediated transactivation. The broad specificity cellular efflux ABC transporter ABCB1 is also recognized to be transcriptionally induced by xenobiotics that are ligands for PXR. The layout of response elements in MDR1 resembles that observed in CYP3A4, with a cluster of elements at approximately 7.8 kilobases. Electromobility shift assays and site directed mutagenesis have shown that an overlapping DR-4/ER-6 response element binds PXR/RXRa with high affinity [33] (Fig. 22.3).

Spectrum of PXR and CAR Target Genes While CYP3A and CYP2B genes were the gene targets initially used to explore the function of PXR and CAR respectively, the advent of gene array technology has given an in-depth understanding of the spectrum of

22  Signaling Pathways in Liver Diseases: PXR and CAR

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R X R -7817bp

Fig.  22.3  Organization of DNA response elements capable of binding PXR/RXRa or CAR/RXRa heterodimers in the regulatory 5’-flanking regions of human genes that are transcriptionallyinduced by activation of these receptors. Many of these elements

are capable of binding either PXR or CAR, while some are selective, such as the PXR response element in the ABCB1 gene. PBREM, Phenobarbital-responsive enhancer module; XREM, xenobiotic-responsive enhancer module

genes regulated by PXR and CAR. Using both mouse and human hepatocytes in culture treated with activators of either PXR or CAR, Maglich et al [34] showed there was considerable overlap in the spectrum of genes transcriptionally induced by PXR and CAR, notably CYPs, glutathione S-transferases, uridine glucuronosyltransferases, ATP-binding cassette transporters, aldehyde dehydrogenease 1A1, amino levulinic acid synthase 1 and the aryl hydrocarbon receptor. While many of these genes would be immediately familiar to anyone working in the drug development field as major players in the process of drug elimination, some of these genes are also responsible for the metabolism and elimination of endobiotics. However, significant differences are also apparent in the spectrum of genes regulated by these receptors. For example, sulphotransferases involved in the sulphation of bile acids appear to be selective targets for CAR [35]. Thus, PXR and CAR are capable of upregulating genes in the liver that are able to sequentially

remove both xeno- and endobiotics from the body, from hepatocyte sinusoidal uptake transporters, oxidative and conjugating enzymes, through to apical biliary canalicular transporters that excrete compounds into bile. NHRs are also known to be able to repress transcription. For example, when mice are administered the CAR activating ligand TCPOBOP, there is strong repression of Cyp4a genes involved in the oxidative metabolism of fatty acids [36]. More work is needed to understand the extent and role of PXR and CAR as transcriptional repressors.

Xenobiotic Metabolism As discussed above, PXR and CAR regulate genes involved in all three phases of drug disposition as illustrated by phase I CYP enzymes, phase II transferases,

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and phase III transporters. Therefore, it is not surprising that activation of these receptors has an impact on the clearance of many commonly used drugs. Human PXR ligands including rifampin, phenytoin, carbamazepine [13, 27], and hyperforin (a component of the popular herbal medication, St John’s Wort) [37] are well recognized to cause inductive drug–drug interactions, particularly for co-administered drugs that are substrates of CYP3A and/or CYP2C CYP enzyme subfamilies. The degree of induction of drug metabolizing enzymes after exposure to PXR ligands can occasionally be massive, rendering some co-administered drugs completely ineffective. In contrast, CAR has a more restricted range of xenobiotic ligands, and other than phenobarbital, which is an indirect activator of CAR, there appears to be little contribution of this receptor to  the inductive drug–drug interactions, commonly encountered in clinical medicine. From a practical standpoint, mechanistic knowledge of the roles of PXR and CAR is regulating drug metabolism has allowed the development of systems that can determine if newly developed drugs have the propensity to cause inductive drug–drug interactions in humans, well before clinical studies are undertaken [17].

Endobiotic Homeostasis and Disease

C. A. M. Stedman et al.

regulated, with the induction of the hepatocyte basolateral uptake transporter Oatp2 by both PXR and CAR agonists, and induction of the hepatocellular canalicular efflux pumps Mrp3 and Mrps2–4 by PXR and CAR, respectively, promoting bile acid efflux from the hepatocyte. This nuclear receptor-mediated regulation of bile acid detoxification pathways has a marked impact on the development of hepatic damage in cholestasis, and this is most clearly demonstrated in nuclear receptorknockout mice subjected to various models of cholestasis and/or bile acid overload. Mice with deletion of PXR or CAR have an increase in the areas of hepatic necrosis and bile infarcts after injection of lithocholic acid (LCA) [38, 39], or bile duct ligation [42]. Conversely, PXR activation by pregnenolone 16a-carbonitrile protects wild-type mouse livers against necrosis caused by administration of LCA [38, 39]. Key enzymes involved in bile acid synthesis include Cyp7a1, Cyp7b1, and Cyp8b1. Elevated bile acid concentrations are able to repress these enzymes in a negative feedback mechanism; for example, Cyp7a1 is repressed via induction of SHP by activated FXR [43]. However, PXR and CAR have also been shown to play some role in coordinate repression of these genes, with loss of repression of Cyp7b1 and Cyp8b1 after bile duct ligation in PXR or CAR knockout mice compared to wild-type mice [42].

Bile Acids

Bilirubin

The interaction between bile acids and nuclear receptors has been covered in the chapter on bile acids and their receptors. Although the farnesoid X receptor (FXR) was originally characterized as the “bile acid receptor,” PXR and CAR both play important roles in the regulation of hepatic detoxification of bile acids. This has important clinical implications for the cholestatic liver disorders that are characterized by impaired hepatocellular secretion of bile, resulting in accumulation of bile acids, bilirubin, and cholesterol, subsequently leading to liver injury. PXR is directly activated by some bile acids and bile acid precursors as a low affinity receptor [38–40]. PXR and CAR agonists stimulate the hepatic phase I bile acid-detoxifying enzymes Cyp3a11 and Cyp2b10, and CAR agonists induce phase II conjugation by sulphation (Sult2a1) [41]. Transport (phase III) systems are also

Bilirubin is an end product of hemoglobin breakdown that is excreted into bile, and like bile acids it also accumulates in cholestatic liver disease. CAR has been implicated as a regulator of bilirubin clearance. Activation of CAR has been shown to increase hepatic expression of components of the bilirubin clearance pathway, including Oatp-c (Slc21A6), GSTA1, UGT1A1, and MRP2 [44]. In wild-type, but not CAR knockout mice, activation of CAR results in increased clearance of an acute dose of bilirubin, and activation of PXR increases bilirubin clearance from hepatocytes [45]. In mouse models of cholestasis, CAR agonists (and to a lesser extent PXR agonists) induce Ugt1a1 (selectively conjugating bilirubin) and the bilirubin conjugate transport systems Mrp2 and Mrp3, accompanied by a reduction in serum bilirubin levels in both cholestatic and healthy mice [41]. In PXR and CAR

22  Signaling Pathways in Liver Diseases: PXR and CAR

knockout mice with cholestasis, bile acid-induced repression of Oatp-c (mediating bilirubin influx from serum into the hepatocyte) was dependent on both PXR and CAR [42]. Therefore both CAR, and to a lesser extent, PXR play an important role in regulating bilirubin clearance pathways in vivo in both normal physiology and the pathological state of cholestasis.

Steroids and Thyroid Hormone Glucocorticoids consistently show stimulatory effects on CAR activity; however, this effect is mutual as CAR potentiates glucocorticoid receptor (GR) signaling. Thus, GR and CAR can synergize to induce target genes. For particular CYPs, such as CYP3A subfamily enzymes, PXR can also contribute to GR-dependent regulation of these genes, but the precise physiological relationship between these three receptors is yet to be fully defined [19]. Sex hormones may also regulate CAR activity, and estrogens have been found to activate CAR to modest levels, while progesterone and androgens repress CAR activity [46]. However, there remains a relative paucity of in vivo data in this area, so the relevance of these observations to endocrine homeostasis is yet to be determined. Drugs such as phenytoin and phenobarbital have been recognized to influence thyroid hormone levels [47], and CAR has been shown to influence thyroid hormone activity in vivo by directly regulating the ­thyroid hormone-activating Dio 1 gene in partially-­ hepatectomized mice [48]. Drug activation of CAR has also been shown to decrease serum level of total T4, but not T3 in Car+/+ mice. However, at present it is not clear whether CAR plays any role in the regulation of thyroid hormone activity in normal adult mice or humans [49].

Hepatic Energy Homeostasis Lipid Metabolism Hepatic lipid homeostasis is tightly maintained by a balance between lipid formation (lipogenesis), catabolism (oxidation), lipid uptake, and secretion. Many drugs that are CAR and /or PXR activators affect lipid

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metabolism and induce hepatic lipid accumulation giving the histological picture of steatosis. For example, rifampin (a PXR activator) can induce hepatic steatosis in tuberculosis patients, and phenobarbital (a CAR activator) has shown significant changes in patient’s hepatic and plasma lipid profiles [50]. Several mechanisms have been implicated in these effects. CAR and PXR activation provokes a decrease in b-oxidation-related gene expression, via interference with Fox (Fork head box) A2 and HNF-4a, positive regulators of the carnitine palmitoyltransferase 1 (CPT1) gene, resulting in repression of this gene [49–51]. PXR activates the CD-6 free fatty acid transporter gene, associated with marked hepatic accumulation of triglycerides in “humanized” PXR transgenic mice [52]. PXR also increases hepatic expression of transcription factors and enzymes involved in lipogenesis, including peroxisome proliferator-activated receptor g (PPAR g), stearylCoA desaturase (SCD1), and fatty acid elongase [51]. These findings may have significant implications for the management of lipid disorders, obesity, and fatty liver disease; however, there is an important species-specificity to many of these findings, and so the relevance for human disease has yet to be determined.

Glucose Metabolism Hepatic gluconeogenesis is tightly controlled by insulin and glucagon and has an important role in the survival during fasting. Drugs that activate PXR and CAR are known to repress gluconeogenic enzymes and genes. For example, phenobarbital, an indirect CAR activator, decreases plasma glucose and improves insulin sensitivity in diabetic patients. The Pepck1 and G6Pase genes are down-regulated in transgenic mice expressing constitutively activated PXR [52]. It seems likely that CAR and PXR actively repress the glucogenic pathway by interfering with transcription factors or cofactors involved in the transcriptional regulation of gluconeogenic enzymes [51]. Both CAR and PXR bind directly to FoxO1, preventing FoxO1 binding to its response element, the insulin response sequence (IRS), and this interaction appears to be the underlying mechanism repressing the G6Pase and PEPCK1 genes in response to xenobiotics [49]. Similar to the cross-talk with FoxO1, PXR also interacts with the cAMP-response element binding protein (CREB) to repress G6Pase,

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antagonizing glucagon activation. The PPAR g co-activator (PGC1)a is also a glucagon-activated gene and binds to and co-activates HNF-4a-mediated transcription. Drug-activated PXR and/or CAR have been shown to dissociate PGC1a from the HNF-4a complex, thus repressing transcription of PEPCK and G6Pase. Therefore, CAR and PXR repress glucose production by directly binding to and interfering with the action of several transcription factors that activate gluconeogenesis [49]. Aspects of these pathways may represent potential novel targets for treatment of diseases such as diabetes.

C. A. M. Stedman et al.

Summary

›› PXR and CAR are closely related nuclear hor›› ››

›› Prospects for Use of PXR and CAR as Therapeutic Targets

›› As both PXR and CAR are potent inducers of adaptive transcriptional programs that favor the detoxification and elimination of both xeno- and endobiotics, they appear, at least on the surface, to be attractive therapeutic targets for liver diseases where detoxification and excretion of hydrophobic molecules is impaired, as exemplified by retention of bile acids in cholestasis. However, to date it is mainly data derived from animal models that support the use of PXR and CAR activators in this context. While there are many existing drugs that are ligands for PXR, their propensity to cause drug interactions places limitations on their usefulness, though previous experience with rifampin in the treatment of severe cholestasis has shown long term benefits in the control of troubling symptoms such as pruritus [53]. There has been recent interest in the role of PXR in the pathogenesis of inflammatory bowel disease (IBD) with both low expression of PXR in colonic mucosa of ulcerative colitis patients [54] and polymorphisms of the PXR gene being linked to susceptibility to IBD [55], however, the clinical utility of these observations is presently unknown. The use of CAR as a therapeutic target has to be approached with caution. Based upon effects of CAR on lipid metabolism and regulation of detoxification pathways, a recent study examined the impact of the CAR ligand TCPOBOP on a mouse model of nonalcoholic steatohepatitis and found that the liver disease was exacerbated [56]. This effect may be linked in part to suppression of CYP-mediated fatty acid oxidation by CAR [36].

››

››

››

››

››

mone receptors that heterodimerize with RXRa. They are receptors for xenobiotics, though their function extends well beyond regulation of xenobiotic elimination. Both PXR and CAR are predominantly expressed in the liver and intestinal mucosa, their anatomical distribution suggesting, in part, a protective role from ingested chemicals. They trigger adaptive programs that favor the metabolism and elimination of a wide range of xeno- and endobiotics, such as therapeutic drugs and bile acids. Target genes include phase I CYP enzymes, phase II transferases, and phase III exceretory transporters. Transcriptional induction of target gene expression is accomplished through binding as heterodimers with RXRa to tandem repeats of a core hexad DNA sequence in gene promoters/enhancers and recruitment of co-activator proteins. PXR has many known ligands while CAR is most often indirectly activated, without ligand binding, through processes that allow it to translocate to the cell nucleus and exert constitutive transcriptional activity. While PXR and CAR were originally identified as regulators of xenobiotic elimination, it is now apparent that they have roles in the homeostasis of bile acids, bilirubin, lipids, glucose, and energy utilization. The identification of lithocholic acid as a ligand for PXR demonstrated a direct role for this receptor in endobiotic homeostasis and CAR has been shown to be a regulator of bilirubin elimination. Both PXR and CAR may have potential as therapeutic drug targets, though the activation of these receptors usually has unwanted consequences, not least of which is increased clearance and hence reduced effect of many co-administered therapeutic drugs.

22  Signaling Pathways in Liver Diseases: PXR and CAR

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Multiple Choice Questions

References

1. The pregnane X receptor (PXR) and constitutive androstane receptor (CAR) belong to which gene family? (a) Leucine zipper (b) Nuclear hormone receptor (c) Cytokine receptor (d) Toll-like receptor (e) Cytochrome P450

  1. Mangelsdorf DJ, Thummel C, Beato M et al (1995) The nuclear receptor superfamily: the second decade. Cell 83: 835–839   2. Gudas LJ (1994) Retinoids and vertebrate development. J Biol Chem 269:15399–15402   3. Barish GD, Downes M, Alaynick WA et al (2005) A nuclear receptor atlas: macrophage activation. Mol Endocrinol 19: 2466–2477   4. Lonard DM, O’Malley BW (2007) Nuclear receptor coregulators: judges, juries, and executioners of cellular regulation. Mol Cell 27:691–700   5. Ordentlich P, Downes M, Evans RM (2001) Corepressors and nuclear hormone receptor function. Curr Top Microbiol Immunol 254:101–116   6. Blumberg B, Sabbagh W, Juguilon H et al (1998) SXR, a novel steroid and xenobiotic-sensing nuclear receptor. Genes Dev 12:3195–3205   7. Bertilsson G, Heidrich J, Svensson K et al (1998) Identification of a human nuclear receptor defines a new signaling pathway for CYP3A induction. Proc Natl Acad Sci U S A 95:12208–12213   8. Kliewer SA, Moore JT, Wade L et al (1998) An orphan nuclear receptor activated by pregnanes defines a novel steroid signaling pathway. Cell 92:73–82   9. Baes M, Gulick T, Choi HS et al (1994) A new orphan member of the nuclear hormone receptor superfamily that interacts with a subset of retinoic acid response elements. Mol Cell Biol 14:1544–5152 10. Honkakoski P, Zelko I, Sueyoshi T, Negishi M (1998) The nuclear orphan receptor CAR-retinoid X receptor heterodimer activates the phenobarbital-responsive enhancer module of the CYP2B gene. Mol Cell Biol 18:5652–5658 11. Sueyoshi T, Kawamoto T, Zelko I, Honkakoski P, Negishi M (1999) The repressed nuclear receptor CAR responds to phenobarbital in activating the human CYP2B6 gene. J Biol Chem 274:6043–6046 12. Sonoda J, Rosenfeld JM, Xu L, Evans RM, Xie W (2003) A nuclear receptor-mediated xenobiotic response and its implication in drug metabolism and host protection. Curr Drug Metab 4:59–72 13. Moore LB, Parks DJ, Jones SA et al (2000) Orphan nuclear receptors constitutive androstane receptor and pregnane X receptor share xenobiotic and steroid ligands. J Biol Chem 275:15122–15127 14. Watkins RE, Wisely GB, Moore LB et al (2001) The human nuclear xenobiotic receptor PXR: structural determinants of directed promiscuity. Science 292:2329–2333 15. Goodwin B, Redinbo MR, Kliewer SA (2002) Regulation of cyp3a gene transcription by the pregnane x receptor. Annu Rev Pharmacol Toxicol 42:1–23 16. Schuetz EG, Brimer C, Schuetz JD (1998) Environmental xenobiotics and the antihormones cyproterone acetate and spironolactone use the nuclear hormone pregnenolone X receptor to activate the CYP3A23 hormone response element. Mol Pharmacol 54:1113–1117

2. Which of the following bile acids is a ligand for the pregnane X receptor (PXR)? (a) Hyocholic acid (b) Chenodeoxycholic acid (c) Ursodeoxycholic acid (d) Cholic acid (e) Lithocholic acid 3. Interspecies differences in drugs capable of inducing cytochrome P450-mediated drug metabolism are the result of? (a) Differences in drug absorption (b) Differences in drug transport across cell membranes (c) The ligand specificity of xenobiotic-sensing nuclear hormone receptors (d) Differences in P450 substrate specificity (e) The rate of metabolism of the inducing drug 4. Activation of the pregnane X receptor (PXR) or the constitutive androstane receptor (CAR) by either xenobiotics or endobiotics increases the expression of genes involved in? (a) Phase I drug metabolism (b) Phase II drug metabolism (c) Drug transport (d) All of the above (e) None of the above 5. Constitutive androstane receptor (CAR) knockout mice exhibit impaired elimination of? (a) Sodium (b) Uric acid (c) Bilirubin (d) Potassium (e) Cholesterol

342 17. Liddle C, Robertson GR (2003) Predicting inductive drugdrug interactions. Pharmacogenomics 4:141–152 18. Forman BM, Tzameli I, Choi HS et al (1998) Androstane metabolites bind to and deactivate the nuclear receptor CARbeta. Nature 395:612–615 19. Timsit YE, Negishi M (2007) CAR and PXR: the xenobiotic-sensing receptors. Steroids 72:231–246 20. Kobayashi K, Sueyoshi T, Inoue K, Moore R, Negishi M (2003) Cytoplasmic accumulation of the nuclear receptor CAR by a tetratricopeptide repeat protein in HepG2 cells. Mol Pharmacol 64:1069–1075 21. Kawamoto T, Sueyoshi T, Zelko I et al (1999) Phenobarbitalresponsive nuclear translocation of the receptor CAR in induction of the CYP2B gene. Mol Cell Biol 19:6318–6322 22. Tzameli I, Pissios P, Schuetz EG, Moore DD (2000) The xenobiotic compound 1, 4-bis[2-(3, 5-dichloropyridyloxy)] benzene is an agonist ligand for the nuclear receptor CAR. Mol Cell Biol 20:2951–2958 23. Maglich JM, Parks DJ, Moore LB et al (2003) Identification of a novel human constitutive androstane receptor (CAR) agonist and its use in the identification of CAR target genes. J Biol Chem 278:17277–17283 24. Xu RX, Lambert MH, Wisely BB et al (2004) A structural basis for constitutive activity in the human CAR/RXRalpha heterodimer. Mol Cell 16:919–928 25. Suino K, Peng L, Reynolds R et al (2004) The nuclear xenobiotic receptor CAR: structural determinants of constitutive activation and heterodimerization. Mol Cell 16:893–905 26. Hosseinpour F, Moore R, Negishi M, Sueyoshi T (2006) Serine 202 regulates the nuclear translocation of constitutive active/androstane receptor. Mol Pharmacol 69:1095–1102 27. Lehmann JM, McKee DD, Watson MA et al (1998) The human orphan nuclear receptor PXR is activated by compounds that regulate CYP3A4 gene expression and cause drug interactions. J Clin Invest 102:1016–1023 28. Goodwin B, Hodgson E, Liddle C (1999) The orphan human pregnane X receptor mediates the transcriptional activation of CYP3A4 by rifampicin through a distal enhancer module. Mol Pharmacol 56:1329–1339 29. Robertson GR, Field J, Goodwin B et al (2003) Transgenic mouse models of human CYP3A4 gene regulation. Mol Pharmacol 64:42–50 30. Goodwin B, Hodgson E, D’Costa DJ, Robertson GR, Liddle C (2002) Transcriptional regulation of the human CYP3A4 gene by the constitutive androstane receptor. Mol Pharmacol 62:359–365 31. Goodwin B, Moore LB, Stoltz CM, McKee DD, Kliewer SA (2001) Regulation of the human CYP2B6 gene by the nuclear pregnane X receptor. Mol Pharmacol 60:427–431 32. Makinen J, Frank C, Jyrkkarinne J et al (2002) Modulation of mouse and human phenobarbital-responsive enhancer module by nuclear receptors. Mol Pharmacol 62:366–378 33. Geick A, Eichelbaum M, Burk O (2001) Nuclear receptor response elements mediate induction of intestinal MDR1 by rifampin. J Biol Chem 276:14581–14587 34. Maglich JM, Stoltz CM, Goodwin B et al (2002) Nuclear pregnane X receptor and constitutive androstane receptor regulate overlapping but distinct sets of genes involved in xenobiotic detoxification. Mol Pharmacol 62:638–646 35. Saini SP, Sonoda J, Xu L, Toma D et al (2004) A novel constitutive androstane receptor-mediated and CYP3A-independent

C. A. M. Stedman et al. pathway of bile acid detoxification. Mol Pharmacol 65(2): 292–300 36. Maglich JM, Lobe DC, Moore JT (2009) The nuclear receptor CAR (NR1I3) regulates serum triglyceride levels under conditions of metabolic stress. J Lipid Res 50: 439–445 37. Moore LB, Goodwin B, Jones SA et al (2000) St. John’s wort induces hepatic drug metabolism through activation of the pregnane X receptor. Proc Natl Acad Sci U S A 97:7500–7502 38. Staudinger JL, Goodwin B, Jones SA et al (2001) The nuclear receptor PXR is a lithocholic acid sensor that protects against liver toxicity. Proc Natl Acad Sci U S A 98: 3369–3374 39. Xie W, Radominska-Pandya A, Shi Y et al (2001) An essential role for nuclear receptors SXR/PXR in detoxification of cholestatic bile acids. Proc Natl Acad Sci U S A 98:3375–3380 40. Goodwin B, Gauthier KC, Umetani M et al (2003) Identification of bile acid precursors as endogenous ligands for the nuclear xenobiotic pregnane X receptor. Proc Natl Acad Sci U S A 100:223–228 41. Wagner M, Halilbasic E, Marschall HU et al (2005) CAR and PXR agonists stimulate hepatic bile acid and bilirubin detoxification and elimination pathways in mice. Hepatology 42:420–430 42. Stedman CA, Liddle C, Coulter SA et al (2005) Nuclear receptors constitutive androstane receptor and pregnane X receptor ameliorate cholestatic liver injury. Proc Natl Acad Sci U S A 102(6):2063–2068 43. Chiang JY (2004) Regulation of bile acid synthesis: pathways, nuclear receptors, and mechanisms. J Hepatol 40:539–551 44. Huang W, Zhang J, Chua SS et al (2003) Induction of bilirubin clearance by the constitutive androstane receptor (CAR). Proc Natl Acad Sci U S A 100:4156–4161 45. Xie W, Yeuh MF, Radominska-Pandya A et al (2003) Control of steroid, heme, and carcinogen metabolism by nuclear pregnane X receptor and constitutive androstane receptor. Proc Natl Acad Sci U S A 100:4150–4155 46. Kawamoto T, Kakizaki S, Yoshinari K et al (2000) Estrogen activation of the nuclear orphan receptor CAR (constitutive active receptor) in induction of the mouse Cyp2b10 gene. Mol Endocrinol 14:1897–1905 47. Curran PG, DeGroot LJ (1991) The effect of hepatic enzymeinducing drugs on thyroid hormones and the thyroid gland. Endocr Rev 12:135–150 48. Tien ES, Matsui K, Moore R et al (2007) The nuclear receptor constitutively active/androstane receptor regulates type 1 deiodinase and thyroid hormone activity in the regenerating mouse liver. J Pharmacol Exp Ther 320:307–313 49. Konno Y, Negishi M, Kodama S (2008) The roles of nuclear receptors CAR and PXR in hepatic energy metabolism. Drug Metab Pharmacokinet 23:8–13 50. Calandre EP, Rodriquez-Lopez C, Blazquez A et al (1991) Serum lipids, lipoproteins and apolipoproteins A and B in epileptic patients treated with valproic acid, carbamazepine or phenobarbital. Acta Neurol Scand 83:250–253 51. Moreau A, Vilarem MJ, Maurel P et al (2008) Xenoreceptors CAR and PXR activation and consequences on lipid metabolism, glucose homeostasis, and inflammatory response. Mol Pharm 5:35–41 52. Zhou J, Zhai Y, Mu Y et al (2006) A novel pregnane X receptor-mediated and sterol regulatory element-binding

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23

p53 Wen-Wei Tsai and Michelle Craig Barton

Introduction The p53 tumor suppressor is well known as the major target of mutation in human cancers and plays a primary role in protecting cells in the face of genotoxic stresses and challenges to genomic stability. The principal responsibilities of p53 include regulation of genes that promote either arrest of cell cycle or apoptosis, both of which inhibit cellular propagation of DNA damage and tumor development [1–3]. The gene encoding human p53 (TP53) is mutated in more than 50% of all types of human cancers; however, studies of tumor progression in the liver show that mutation of TP53, in the absence of environmental influences discussed below, is a relatively late event in development of hepatocellular carcinoma (HCC) and other cancers of this tissue [4]. In this chapter, we will discuss multiple ways in which dysfunction in p53-signaling occurs, even when TP53 itself is not mutated, in relationship with the biology of p53, its protein domains and specific functions, the influ­ences of p53-family members, and cross-talk with other signaling pathways. Tumor suppressor p53 is primarily known for its role in maintaining genomic stability and guarding against tumor development. More recently, functions of p53 in normal cells, including regulation of specific, developmentally regulated genes [5, 6], cellular progression to senescence, and aging [7], have been shown. The apparent viability of the p53-null mouse suggests that p53 has no functions in development; however, its roles in tumor suppression are clear as p53-null and -heterozygous mice M. C. Barton () Department of Biochemistry and Molecular Biology, University of Texas M.D. Anderson Cancer Center, 1515 Holcombe Blvd., Houston, TX 77030, USA e-mail: [email protected]

develop tumors at an early age, 4–6 months and 6–8 months, respectively [8–10]. Confusion regarding p53 functions during development or in normal cells may be due to compensatory roles played by members of the p53-family. p53 is the founding member of a family of proteins encompassing p53-, p63-, and p73-­isoforms, defined by their conserved domains and sequence homology. Multiple differences among the family members exist due to alternative splicing and/or divergent promoter usage at TP53, TP63, and TP73 genes to yield several protein isoforms [11–13]. Unlike p53, mice genetically engineered for loss of p63 or p73 have profound developmental phenotypes, leading to early death [11, 14–16]. Expression of transactivating (TA)-isoforms of p63/p73 activates transcription of some, but not all, p53-regulated genes with functions in cell cycle arrest and apoptosis, as well as genes not regulated by p53 [17, 18]. These activities likely underpin the described roles of p63 and p73 in tumor suppression [19, 20]. Although this chapter focuses on p53, the reader should bear in mind that intra-family influences on p53-regulation and activities may occur and add complexity in a variable, cell-type specific manner [6].

The Transcription Factor p53 First and foremost, p53 is a transcription factor with the expected activities of binding to a sequence-specific response element (p53RE) within the regulatory regions of target genes and effecting either activation or repression of target gene expression. Transcription response of either activation or repression is dictated by a number of interrelated variables, including protein–protein interactions between p53 and specific co-repressors or co-activators, the chromatin structure of target genes, the

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specific upstream signals received by p53, and the modifying influences of intersecting signaling pathways. The vast majority of studies of p53 as a regulator of transcription focus on activation of transcription. This is readily understandable as p53-mediated response to stress activates expression of p21 (CDKN1A), which leads to cell cycle arrest, or PUMA, BAX, and others, which induce apoptosis, and plays a clear role in tumor suppression [3]. Functions of p53 in repression of transcription are less understood and include target genes encoding tumor marker alpha-fetoprotein (AFP) [21], mitotic progression kinase CDC25c [22], promoter of proliferation myc [23], and antiapoptotic survivin [24]. Repression of transcription may have a greater effect on genes involved in apoptosis, rather than cell cycle arrest, although this is not fully delineated [25, 26]. The most striking feature of p53-signaling, in response to inductive signals, is the highly regulated process of posttranslational modification of p53, which in turn dictates its interactions with proteins that control (1) the levels of p53, (2) the ability of p53 to bind DNA, (3) the subcellular localization of p53, (4) interactions with regulatory proteins, and (5) the activities of p53 in ­transcription-dependent and transcription-independent responses [27, 28]. In order to understand how upstream signaling to p53 is controlled and how p53 in turn regulates downstream gene targets, whether in activation or repression of transcription, some discussion of p53 protein structure is needed. This knowledge will underscore the impact of mutations that arise in TP53 during tumor progression.

W.-W. Tsai and M. C. Barton

Transcription factors are highly modular, and p53 is no exception to this rule. Its protein structure is divided into three major domains responsible for transcription

activation (TA, approximately amino acids 1–100), DNA binding (DBD, amino acids 101–300), and a tetramerization/regulatory domain (TD, amino acids 301–393); Fig. 23.1 [29–31]. Post-translational, enzymatic addition of moieties, e.g., phosphorylation, acetylation, methylation, ubiquitylation, and others, is targeted to specific amino acids by numerous upstream signaling pathways [28]. Critical amino acids of p53 may be mutated to disrupt modification by upstream regulators of p53 and, in turn, downstream signaling by p53. However, a large number of amino acid targets of post-translational modification are not found because of mutations in human tumors. This complexity may be due to response specificity, in stress or cell type, or redundancies that suggest function dictated by structural integrity rather than by specific amino acid sequence. Additionally, most analyses of amino acid residues of p53 and their modifications are experimentally determined in vitro, using cultured cells where p53-­functions  are likely compromised. Therefore, some caution should  be exercised in strict interpretation of how post-­translational modification of p53 dictates its regulatory activities [32]. The TA domain: The N-terminal domain of p53 encom­ passes two functionally separable regions, AD1 (amino acids 1–42) and AD2 (amino acids 43–92), Fig.  23.1. Deletion of amino acid residues 20–42 or double mutation of amino acids L22 and W23 [33, 34], but no single point mutation, disrupts transcription activation by p53 [29]. Genomic targeting by homologous recombination of the p53-encoding gene (Trp53) in mice (knock-in mice) was used to create the p53QS (murine L25Q/W26S, homologous to human L22Q/W23S) mouse, which dies during embryonic development [35]. As p53-null mice generally survive embryonic development [8], these results suggest that mutant forms of p53 exhibit gain-offunction, deleterious to development. Further analysis showed that p53QS is stable, like many mutant p53 proteins, due to loss of interaction with Mdm2; however,

Fig. 23.1  Structural domains of p53. The protein structure of p53 is divided into three major domains: transcription activation (TA, approximately amino acids 1–100), the DNA binding domain (DBD, amino acids 101–300), and a tetramerization/regulatory domain (TD, amino acids 301–393). The TA domain is further divided by function and sequence conservation into the AD1, the

AD2, and the proline rich domain (PXXP). The DBD is the primary site of tumor-derived mutations in the p53-encoding gene. This “hotspot” of mutation frequencies is represented by red bars in this figure. At the C-terminus of p53, the TD/Reg domain also features a NLS and a NES, which were characterized by conserved sequence motifs and functionally

Protein Structure of p53

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p53QS lacks the ability to activate transcription in response to DNA damage [36]. Overall, the AD1 region appears to act predominantly in p53-mediated cell cycle arrest, while the AD2 domain has a greater role in p53-regulated apoptosis. Deletion of AD2 (residues 43–62 or 62–91), or mutation of hydrophobic residues W53Q/F54S, abolishes p53-mediated apoptosis but only partially affects p53mediated cell cycle arrest [37, 38]. The TA domain of p53 contains several serine and threonine residues and is a major site for p53 phosphorylation (S6, 9, 15, 20, 33, 37, 46, and T18), often targeted at the same amino acid by multiple enzymes, e.g., casein kinase, PI3Kkinases (ATM, ATR, and DNA-PK), Chk2 kinase, and the MAP kinase family (p38, ERK 1/ 2, and JNK) [27]. Within the AD2 region, S46 is modified by p38 kinase, T55 by ERK2 kinase, and T81 by JNK kinase. Phosphorylation of S46 is implicated in p53-mediated apoptosis, while phosphorylation of T81 is induced in both p53-mediated responses of cell cycle arrest and apoptosis [27]. A proline-rich region, consisting of PXXP motifs (amino acids 64–92), lies within the AD2 domain and is implicated in growth suppression and apopotic activities of p53 [39, 40]. The PXXP region is not tightly conserved in sequence throughout evolution or required in vivo for trans-activation of p53-regulated genes in the mouse [32]. However, several mutations within the proline-rich region are found as spontaneous mutations (P85S and P89S) and as mutations in Li-Fraumeni syndrome patients (P82L) [39, 41]. Trans-activation of specific, p53-target genes is characterized by interactions between AD1/AD2 regions and RNA polymerase II, p300/CBP, and other co-activators. However, a co-repressor complex of mSin3a/histone deacetylase (HDAC) interacts with residues 61–75 within the AD2 region of p53 in response to hypoxic stress and induction of apoptosis [25, 42]. Specific interactions with co-repressors or co-activators of transcription, many of which are modifiers of chromatin structure, dictate whether p53 acts as a repressor or an activator of target genes in downstream signaling [26, 43]. Identi­ fying the determinants of these interactions with co-regulators of p53-function is an area of active research. The central DBD domain: The vast majority of p53regulated functions requires interaction between p53 and chromatin, where p53 binds to its specific regulatory element, the p53RE. The canonical p53RE consists of two direct repeats of a ten base pair (bp) half-site “PuPuPuC(A/T)(A/T)GPyPyPy” (Pu: purine,

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Py: pyrimidine), either directly juxtaposed or separated by a nucleotide spacer of 1–13 bps [44]. The frequency of point mutations within TP53, revealed by sequencing of DNA from human tumors and tumorderived cells in culture, underscores the importance of DNA binding by p53 to effect tumor suppression. The DBD is a “hot spot” target (Fig. 23.1), where 80% of tumor-derived mutations in TP53 arise to disrupt the ability of p53 to bind DNA [45, 46]. The most frequently mutated residues of the DBD domain are R248 and R273, which are critical in p53-DNA interaction, and R175, G245, R249, and R282, which are required to maintain the DBD domain structure. Site-specific mutations in the “hot spot” domain of p53 are known to occur in response to high levels of exposure to aflatoxin, a fungal toxin that is a causative agent in HCC [47, 48]. Analysis of patients afflicted with HCC in areas with high levels of exposure to aflatoxin reveals that a specific G-to-T transversion at codon R249 of p53 occurs [49, 50]. However, correlation between development of HCC and mutation of specific codons of TP53 is limited in the absence of aflatoxin exposure. Loss of interactions between the DBD of p53 and DNA also occurs without mutation of TP53. The large T antigen of SV40 virus binds the p53 DBD to inactivate p53 and promote cellular transformation [51, 52]. Phosphorylation of S215 by Aurora kinase A inhibits p53 transcriptional activity, and is one of the few posttranslational modifications that occur in the DBD domain to modulate p53 functions [27]. Other proteins, such as 53BP1 [53], ASPP1/ASPP2 [54], HIF-1alpha, Bcl-XL, and Rad51 [55, 56], interact with the DBD of p53 to oppose or promote p53-response, many in ways that remain poorly understood. The C-terminal TD/Reg domain: The consequences of mutations within the DBD extend beyond the encoded mutant p53, due to the presence of the TD or tetramerization domain. As its name implies, this domain is a platform for p53–p53 interactions and formation of a functional tetramer [57–59]. A single monomer of mutant p53, when unable to bind to DNA, compromises the ability of the tetrameric complex to bind DNA and regulate transcription, thereby having a “dominant negative” effect on transcription [60, 61]. Nonproductive p53-tetramers may maintain interactions with co-­regulatory proteins, and mutant p53 that is stable and expressed at high levels causes further regulatory disruption by effectively sequestering coregulatory proteins required for multiple pathways.

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Several important regulatory motifs and amino acids, important in control of p53 levels and activity, lie within the TD domain (Fig. 23.1). These include the nuclear localization signal (NLS, approximately amino acids 305–322), nuclear export signal (NES, amino acids 340–351), and negative regulatory region (REG, amino acids 364–393). The NLS and NES motifs function in subcellular localization of p53, as part of a highly regulated nuclear-cytoplasmic switch [62], discussed in more detail below. The REG region is an unstructured domain, at the most C-terminal end of p53, and is rich in serine and lysine residues. These residues are targets of modifying enzymes and major sites of post-translational modifications, e.g., phosphorylation, acetylation, methylation, and others. In general these post-translational modifications are induced by stress stimuli, and affect p53 activities by multiple mechanisms, e.g., induc­ing p53 stability, exposure of the DBD, promoting interactions with coactivators, or repressing interactions with co-repressors [27, 63–65]. Deletion of the most C-terminal, 30 amino acids of p53 generates a protein that is constitutively active [66]. The importance of the C-terminus of p53, as a platform for regulatory modifications and/or protein interactions, is supported in vivo by genetically engineered mice that express C-terminally truncated p53 and exhibit premature aging [67]. In contrast to this induction of aging, full-length p53 ectopically expressed from multiple copies of a p53-transgene and regulated normally, does not cause premature aging but rather confers protection from tumor development in transgenic mice [68].

W.-W. Tsai and M. C. Barton

TP53 in several human tumors was reported [74]. The role of p53 as a tumor suppressor, rather than a tumor promoter, was further supported by several studies showing that wild-type p53 inhibited tumor formation [75] and, in response to DNA damage, is induced by multiple signal transduction pathways [76]. One reason behind the confusion in identifying p53 as an oncoprotein, rather than a tumor suppressor, became clear when investigators found that p53 is normally held at very low basal levels, but mutant p53 often exhibits increased protein stability and is readily isolated and cloned from tumor-derived cells [1, 77]. The levels of p53 protein in any given cell are tightly controlled by an intricate regulatory network of proteins, which elevates or reduces p53 levels [78]. Disruption within any of the arms of this network may alter or disable the surveillance powers of p53, even in the absence of mutation in TP53, Fig. 23.2. Chief among the negative regulatory proteins that control p53 levels is Mdm2. Mdm2 controls p53 in

Regulation of p53 Protein Levels Interestingly, p53 was first identified in 1979 as an oncoprotein rather than a tumor suppressor protein, as it was detected in several Simian virus 40 (SV40)transformed cell lines and in sarcomas, chemically induced in mice, but not in primary cells generated from adult mouse tissues [69–72]. The mouse gene encoding p53 was cloned soon after protein identification and its name was abbreviated as Trp53 for ­transformation-related protein p53 [73]. For years p53 was considered a biomarker of tumor cells, until 1989 when the detection of multiple mutations or deletion of

Fig. 23.2  Levels of p53 are tightly controlled by multiple regulatory proteins. Negative regulators of p53 levels (dark blue) act as E3-ubiquitin ligases to ubiquitylate p53 and promote its degradation by the proteasome. Mdm4 primarily modulates Mdm2 activity but does not have this E3-capability. In response to stress, negative regulation of p53 is disrupted and p53 levels increase. Autoregulation is established by p53-mediated activation of genes encoding these negative regulators, which restores normal, low levels of p53 at the termination of the stress-induced response

23  p53

multiple ways: by interaction with the N-terminal TA domain of p53 to block functions of p53 in transcription and by targeting p53 for ubiquitylation, as an E3-ubiquitin ligase, and protein degradation [79, 80]. Expression of Mdm2 (or HDM2 in humans) is regulated by p53 in an auto-regulatory loop. In this way activated p53 induces expression of its target genes, including effectors of cell cycle arrest or apoptosis and those, such as Mdm2, that terminate the p53response and restore basal, low levels of p53 [81]. The importance of Mdm2-mediated regulation of p53 is illustrated by deletion of Mdm2 in mice, which is early embryonic lethal at the peri-implantation stage [80, 82]. Levels of p53 protein increase unchecked in cells of the mdm2-/- embryo and cause unregulated apoptosis, a phenotype that is rescued by deletion of Trp53 [82]. Other negative regulators of p53 protein stability, e.g., PirH2, ARF-BP1, and Cop1, have been ­identified, and they also function as p53-regulated, E3-ubiquitin ligases. Specific proteins that regulate p53 by control of protein stability may function in specific cell types or under particular conditions, but these determinations await in vivo analysis. Over expression of negative regulators of p53 is found in a number of cancers and tumor-derived cells, and is a major mechanism whereby p53-signaling is disrupted without mutation of TP53. Mdm2 interacts with the AD1 region (TA-domain) at the N-terminus of p53, in the absence of stress or to terminate p53-response, and promotes degradation of p53 through the 26S proteasome [83, 84]. The RING domain of Mdm2 is critical for Mdm2 functions as an E3-ubiquitin ligase, which targets p53 at an FWL motif (residues 19–26, FSDLWKLL) for subsequent ubiquitylation of lysine residues within the C-terminus of p53 [27]. Interaction between p53 and Mdm2 not only mediates p53 degradation but also prevents p53 binding to several protein complexes, e.g., members of RNA polymerase II transcription complexes: TBP [85, 86], TAFII40 (TAF11), and TAFII60 (TAF6) [87]. Additionally, Mdm2 blocks histone acetyltransferases p300/CBP, important co-regulators of p53 [88], in their binding to an LXXLL motif (residues 22–26) in the AD1 domain of p53 [89]. However, E3-ubiquitin ligase activity is probably the primary mechanism by which Mdm2 controls p53-response. A mutated form of Mdm2 that disrupts E3-ligase function but allows protein–protein interactions between p53 and Mdm2,

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Fig. 23.3  Threats to genomic stability induce p53-activated cell cycle arrest. This simplified version of a subset of pathways that activate p53 and downstream responses to this activation shows p53-mediated arrest at G1/S and G2/M checkpoints. Primarily, p53 works through induction of p21 expression, a cyclin-dependent kinase inhibitor that can function in both G1/S and G2/M. Additional inhibitors acting in arrest include Gadd45 and 14-3-3s. Activation of these genes by p53 is further augmented by p53-mediated, direct repression of Cdc25c transcription

engineered in MEFs, no longer controls levels of p53 or regulates its activities in transcription [90]. Disruption of protein–protein interactions between p53 and its negative regulators allows p53 protein stabilization and is well documented in response to stress, such as DNA damage (Fig.  23.3). Rapid response to such stimuli is marked by p53 phosphorylation and degradation of Mdm2, which itself is a target of ubiquitylation [79]. Post-translational modifications within the TA domain may further promote interactions with protein partners of p53, which act in cell cycle arrest or apoptosis, but more likely serve mainly to disrupt Mdm2–p53 interactions. Chemical inhibitors of Mdm2–p53 interactions, e.g., Nutlin [91], which insert

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themselves within the binding pocket of Mdm2 and block p53-interaction, promote p53-mediated activation of gene expression in cellulo without phosphorylation by stress-activated kinases. This finding suggests that high levels of wild-type p53, lacking stress-induced post-translational modifications and no longer blocked or ubiquitylated by Mdm2, may function in tumor suppression. Potential therapeutic agents, which restore p53 activity, are of great interest and several are currently being tested or are targeted for clinical trials.

Subcellular Localization of p53 An important component in regulation of p53-signaling is control of nuclear-cytoplasmic shuttling and the subcellular localization of p53 [62]. In response to stress, the balance is tipped toward nuclear localization and increased p53 protein levels. In the absence of stress, p53 is not only maintained at low levels but also may be held in a “latent” state by multiple mechanisms. In one mechanism, Mdm2 interacts with p53 bound to chromatin at p53RE sites within regulatory regions of specific genes, such as CDKN1A encoding p21, blocks co-­ activator, and/or promotes co-repressor interactions with DNA-bound p53 [92]. Additionally, low levels of Mdm2 may promote exposure of the NES at the C-terminus and “escort” p53 to the nuclear periphery [93]. In all states, whether normal homeostasis or stress-induced activation, p53 likely exists in multiple protein complexes, which can function in gene- and/or cell-specific ways. In response to stress, Mdm2-p53 interaction is disrupted and the NLS, within the TD domain (Fig. 23.1), is subjected to several post-translational modifications. S315 is phosphorylated by CDK2/cyclin A kinase and K320 is acetylated by p300/CBP-associated factor (PCAF); both act to increase DNA binding of p53 [27]. Additionally, post-translational modifications at the C-terminus of p53 allow recruitment of cofactors to mask the NES and abolish p53 nuclear export. Once Mdm2 restores its interactions with p53, to effect feedback-regulated termination of p53-signaling, the NES is exposed to promote p53 nuclear export, mediated by CRM1 and protesomal degradation in the cytoplasm [94]. Other negative regulators of p53-protein stability, described above, share many of the features of Mdm2regulated stability of p53 but are not characterized in similar detail.

W.-W. Tsai and M. C. Barton

Dysfunction of p53, in the absence of mutation, can be promoted by disruption of the nuclear/cytoplasmic balance of p53 localization. The hepatitis B virus (HBV), as well as hepatitis C (HCV), exploits multiple mechanisms to disrupt p53-mediated transcription [95, 96]. There is some evidence that hepatitis virus exacerbates or cooperates with formation of DNA adducts that cause R249 mutations in TP53 when aflatoxin is ingested and metabolized [97]. Several of these rely on the trans-acting factor encoded within the HBV genome, the factor HBx. The HBx protein can interact with p53 and sequester it within the cytoplasm of an infected cell, where p53 is degraded [98, 99]. This sequestering of p53, in addition to HBx-mediated disruption of p53 DNA binding and/or interactions between p53 and trans-acting factors [100], likely plays an important role in the 200-fold disposition toward development of HCC displayed by patients chronically infected with HBV [101].

Stress Response: Arrest of Cell Cycle and DNA Repair A considerable body of literature focuses on p53responses to cellular stress; in this chapter, we pri­ marily discuss pathways involved in downstream, regulatory functions [102]. Following cellular insults that cause DNA damage, such as ionizing radiation (IR), ultra–violet (UV) radiation, oxidative stress, or stresses that do not cause DNA damage, e.g., hypoxia, p53 is stabilized and rapidly accumulates in the nucleus (Fig.  23.3). Nuclear accumulation of p53 promotes arrest of proliferating cells in the G1-phase of the cell cycle, which may facilitate repair of DNA damage prior to continued cell cycling [103]. The induction of p53 triggers several signaling pathways in mediating cell cycle arrest [104], primarily through activation of p21CIP1/WAF1 gene expression [44, 105]. The p21 protein is a cyclin-dependent kinase inhibitor (CDKI) that blocks multiple cyclin/CDK complexes, e.g., cyclinD/ CDK4(6), cyclinE/CDK2, and cyclinA/CDK2, which promote sustained cellular proliferation. During G1 of the cell cycle, retinoblastoma (Rb) protein is hypo-phosphorylated and binds to E2F proteins and HDAC complexes to inhibit E2F-activities in S-phase progression [106]. CyclinD/CDK4(6) phosphorylates Rb protein to release E2F transcription factors

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from inactive Rb/E2F complexes. The unrestrained E2F transcription factors are then free to activate cell cycle genes, essential for S-phase progression [107, 108]. p21, induced by p53, plays a crucial role in a G1mediated stopgap of E2F-functions by inhibiting the activity of CDK/cyclin protein complexes. Gadd45, another p53-activated downstream gene target, interacts with p21 to augment regulation of cyclin/CDK complexes and mediate cell cycle arrest [109, 110]. The essential role of p53 in G1-arrest of cell cycle is supported both in mouse models, MEFs generated from p53-null mice fail to undergo G1-arrest in response to IR [109], and in ­cultured cells derived from human tumors, where ­p53-mediated G1-arrest fails to occur when TP53 is mutated or p53 is otherwise dysfunctional [111, 112]. When cells encounter DNA damage, p53 not only induces G1-arrest but also blocks G2-M transition and prevents cells from entering mitosis (Fig. 23.3) [113]. CDK1(Cdc2) is activated by CDK-activating kinase (CAK)-mediated phosphorylation; it binds cyclinB and, as an activated complex, is a key component in cellular entry into mitosis [114, 115]. Several downstream targets of p53, e.g., p21, Gadd45, and 14-3-3s, regulate cyclinB/CDK1 functions in response to DNA damage or stalling of ongoing replication [116, 117]. The p21 protein binds and inactivates CDK1 directly [118, 119], while Gadd45 causes dissociation of CDK1 from the cyclinB/CDK1 complex [120–122]. Further, 14-3-3s chaperones CDK1 from the nucleus to the cytoplasm to sequester CDK1 from its nuclear activities [123, 124]. To augment arrest of cell cycle, p53 may repress transcription of the genes encoding cyclin B and CDK1 [116, 125, 126], and is a direct repressor of CDC25C expression to disrupt mitotic progression [22]. In the absence of p53, arrest of the G2-M transition is accomplished by p53-independent pathways, which are less efficient in preventing cellular transformation following DNA damage [113]. In addition to mediating cell cycle arrest that allows repair of DNA damage before cell division, p53 may also play a more direct role in repair itself. Normal, wild-type (WT) p53, but not mutant p53, directly interacts with AP-endonuclease (APE) and DNA polymerase beta, key players in base excision repair (BER), and stimulates BER in vitro [127]. The ribonucleotide reductase gene (p53R2) is a direct downstream target of activation by p53. Ribonucleotide reducatse is essential for synthesis of deoxyribonucleotides, required for

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Fig.  23.4  Apoptosis is induced by p53 via transcription-­ dependent and -independent processes. Activated p53 may act in extrinsic, Fas-mediated pathways or intrinsic, cytochrome-Cdriven cell death. Less understood are the direct interactions between p53 and the mitochondria, which can lead to cytochrome-C release. Each of the pathways converges on cleavage of caspase 3 to drive apoptosis

DNA replication and repair [128, 129]. These studies, which suggest that p53 has a direct impact on DNA repair, further cement the role of p53 in maintaining genomic stability.

Stress Response: Apoptosis The protective functions of p53, when confronted by cellular insult and stress, are mediated not only through cell cycle arrest but also by p53-regulated pathways to apoptosis [103, 130]. p53 can regulate two different branches of apoptosis : the intrinsic mitochondrial pathway and the extrinsic death receptor pathway (Fig. 23.4). In addition to transcription-dependent mechanisms invoked through these pathways, p53 has also been shown to act independently of transcription by direct interactions with mitochondria [131, 132]. The decisive

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factors directing p53 toward one or more of these pathways, as well as dictating arrest versus apoptosis, are not fully understood. In the intrinsic mitochondrial pathway, p53 induces transcription of several genes encoding Bcl-2 family proteins [36], e.g., Bax, Puma, and Noxa [133–136], which are pro-apoptotic proteins. Additionally, p53 down-regulates the gene encoding Bcl-2, which opposes apoptosis by binding and inhibiting Bax [137]. Bcl-2-mediated inhibition of Bax is disrupted by Puma and Noxa, which interact with Bcl-2 to oppose this inhibition and promote cell death. As a result of induction by p53 and inhibition of Bcl-2, Bax is translocated to the mitochondrial ­membrane in order to effect release of cytochrome C. After release from the mitochondria, cytochrome C binds to Apaf1, which additionally is a direct target of ­p53-regulated activation of transcription [138]. Asso­ ciation of cytochrome C and Apaf1 activates Caspase 9 to trigger an apoptotic response. Although Puma and Noxa also interact with the mitochondrial outer membrane to effect release of cytochrome C, their roles may be redundant as ablation of Bax and Bax-related protein Bak, in mouse models, completely abolishes apoptotic response in mouse thymocytes [139, 140]. In addition to transcription-dependent and -independent roles for p53 in control of mitochondrial/ intrinsic apoptosis, p53 is a critical regulator of the extrinsic pathway of apoptosis (Fig. 23.4). The extrinsic apoptosis pathway occurs through cellular membrane-bound, CD95/Fas death receptors. In response to apoptotic stimuli, p53 activates the extrinsic apoptosis pathway by upregulating gene expression of both the ligand (FasL) and the receptor (CD95) to induce an apoptotic cascade [141–143]. When FasL ligand binds to the CD95 receptor, the CD95 receptor recruits several adaptor proteins, such as FADD and FAF, to activate Caspase 8 and Caspase 10, and lead to cell death. Both intrinsic and extrinsic pathways of cell death converge on Caspase 3, which is cleaved during induc­ tion of apoptosis. Ectopic expression of WT p53 in M1 mouse myeloid leukemia cells, which lack endogenous p53, triggers both the intrinsic and the extrinsic pathways to induce Caspase 8, 9, and 10 activities [144]. Functions of p53 in transcriptional control of apoptosis are likely augmented by members of the p53-family, p63 and p73, which have roles in tissuespecific differentiation and tumor suppression [6].

W.-W. Tsai and M. C. Barton

Selectivity in p53 Downstream Response Numerous factors contribute to the outcome of p53 induction, whether to arrest the cell cycle or to induce cell death. This is an active area of research and several models have been suggested regarding the determinants of p53 response to cellular stress [102]. The hypothesis that relative degrees of DNA damage or induced stress determine the p53-dependent response, with arrest in response to repairable damage and apoptosis when damage is extensive, is likely too simple to  explain cellular response to activation of p53. An auxiliary model is that the sequence of a p53RE within the regulatory region of p53 target genes dictates the efficiency of p53 binding and thus the type of response. The binding affinity of p53 and DNA depends on the length of the spacer nucleotides between two halfsites of the p53RE. High affinity p53RE’s, which have a spacer length of zero and diverge from consensus with few mismatches, are predominantly found in genes active in cell cycle arrest, such as CDKN1A (p21) and GADD45. In contrast, low affinity p53RE’s are located in regulatory elements of apoptosis-related genes, such as those that encode Bax, PERP, and IGFBP3, and these genes require additional p53 binding sites and/or co-regulatory proteins to augment p53–p53RE interactions [145, 146]. Therefore, the amount of p53 protein present within the nucleus may, in part, dictate a response of cell cycle arrest or apoptosis. Factors that set the levels of p53 within nuclei may include the type, periodicity, and extent of cellular stress, the expression levels of co-regulators of p53 functions or a combination of some or all of these variables in a specific cell type, as well as other means that remain undetermined. Cell-type specificity in p53-response to DNA damage is fundamentally supported by studies of WT mice exposed to 5Gy of IR [147]. In a highly tissuespecific response to damage, p53 accumulates in the nuclei of cells and induces apoptosis in spleen, thymus, bone marrow, intestine, and ependyma. Likewise p53 levels increase in kidney, osteocytes, myocardium, and salivary glands, but no cell death occurs. Finally, in liver, skeletal muscle, and brain tissue, there is no response to IR at the level of p53 stability or apoptosis. Further studies show that cell-type specific responses are not limited to stress induced by DNA damage. Knock-in mouse models, expressing p53 with

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tumor-derived mutations of specific amino acids, exhibit tissue-specific response in stability of the mutant p53 proteins and profiles of tumor development [32, 148]. Interestingly, the liver may be a unique environment in terms of p53-regulation: among the eleven different tissues examined, Mdm2-regulated stabilization of mutant p53 protein in vivo was absent only in the liver [148]. Recent studies of tumor development in p53/p63/p73depleted mice support the likelihood that p53 and p73 exert endoderm-specific functions: 15% of p53+/-;p73+/mice develop HCC and a similar number develop acinar pancreatic carcinoma within 5–7 months of age [20]. This tissue-specific tumor profile is observed only in genetic depletion of p53/p73, among p53-family members, suggesting potentially important roles for p53 and p73 in progenitor cells of pancreatic and hepatic cells, an area that requires further study.

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Summary

›› Tumor suppressor p53 protects the cell from

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Liver-Specific Challenges to p53 Functions In this chapter, we have noted several examples of hepatic-specific influences on p53 regulation and dysfunction. Tumor suppressor p53 acts by promoting cell cycle arrest or apoptosis, and the selection of distinct pathway and outcome is highly influenced in a tissue-specific manner. The determinants specific to the liver are unknown, as are the specific post-translational modifications of p53 that occur in liver tissue. In addition to damage to TP53 directly induced by aflatoxin, there are nonmutational effects of hepatitis infection, which greatly predispose the liver toward development of HCC [97, 100]. How p53 may remain functional and responsive to extrinsic stresses imposed on the liver, when faced with normal, intrinsic challenges such as the polyploidy of aging hepatocytes, is unknown. The factors that determine the radiation insensitivity exhibited by liver tissue likewise remain a mystery. Additionally, how do mature hepatoctyes of an adult liver efficiently re-enter cell cycle to regenerate without irredeemable loss of p53 function? These and other questions regarding tissuespecificity of p53 regulation, how it is disrupted and, most importantly, how it may be restored, require further investigation by in vitro, in cellulo, and in vivo methodologies.

››

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stresses that promote tumor development and is the primary guardian of genomic sta­ bility. Transcription regulation is the principal mechanism employed by p53 to promote either cell cycle arrest or apoptosis. Various stresses induce post-translational modifications of p53, which promote its activation, repression, or destruction. The levels of p53 are exquisitely controlled by Mdm2 and other negative regulators of p53 protein stability. There are numerous ways in which p53 may become dysfunctional without mutation of its encoding gene. Interactions between p53 and specific protein partners dictate its response to stress in the form of arrest versus apoptosis in a cell-specific manner. Geographical distribution of HCC is correlated with regions where there is high exposure to aflatoxin and/or hepatitis infection. Each of these may promote tumor development by interference with p53-functions. It is important to test potential regulatory mechanisms in vivo as cultured cells are generally tumor-derived and may display dysfunction in pathways that impinge on p53. The activities of p53 are likely influenced by the members of the larger p53-family. consisting of p53, p63, and p73 isoforms. Control of p53 subcellular localization is an important regulatory mechanism in controlling p53-signaling.

Multiple Choice Questions 1. How can p53-signaling be dysfunctional when the p53-encoding gene is not mutated? (a) By over expression of negative regulators of p53 (b) By cytoplasmic sequestering of p53 effected by HBx

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(c) By HBx-mediated interaction with p53 to block DNA binding of p53 (d) By binding to the large T antigen of SV40 virus (e) All of the above 2. What structural domain of p53 is most often mutated in tumor-derived cells? (a) The trans-activating domain (b) The DNA-binding domain (c) The PXXP-region of AD2 (d) The negative regulatory domain (e) The tetramerization domain 3. How might exposure to aflatoxin promote development of HCC? (a) By formation of DNA adducts (b) By specific mutation of TP53 within the DBD (c) By exacerbating the effects of chronic hepatitis (d) By activating Mdm2 (e) All of the above 4. Which of the following is not a mechanism by which p53 suppresses tumor development? (a) Arrest of cell cycle at G1/S (b) Activation of the intrinsic pathway of cell death (c) Arrest of cell cycle at G2/M (d) Interaction with HDACs to inhibit Rb (e) Activation of apoptosis independently of transcription 5. Ionizing radiation at levels that induce DNA damage does not cause accumulation of nuclear p53 in which of the following adult tissues? (a) Thymus (b) Intestine (c) Liver (d) Kidney (e) Bone marrow

References   1. Vousden KH, Lane DP (2007) p53 in health and disease. Nat Rev Mol Cell Biol 8(4):275–283   2. Laptenko O, Prives C (2006) Transcriptional regulation by p53: one protein, many possibilities. Cell Death Differ 13(6): 951–961   3. Riley T et al (2008) Transcriptional control of human p53regulated genes. Nat Rev Mol Cell Biol 9(5):402–412   4. Laurent-Puig P, Zucman-Rossi J (2006) Genetics of hepatocellular tumors. Oncogene 25(27):3778–3786   5. Hu W et al (2008) p53: a new player in reproduction. Cell Cycle 7(7):848–852

W.-W. Tsai and M. C. Barton   6. Stiewe T (2007) The p53 family in differentiation and tumorigenesis. Nat Rev Cancer 7(3):165–168   7. Feng Z et al (2008) The tumor suppressor p53: cancer and aging. Cell Cycle 7(7):842–847   8. Donehower LA et al (1992) Mice deficient for p53 are developmentally normal but susceptible to spontaneous tumours. Nature 356(6366):215–221   9. Donehower LA, Bradley A (1993) The tumor suppressor p53 (review). Biochem Biophys Acta 1155(2):181–205 10. Sah VP et al (1995) A subset of p53-deficient embryos exhibit exencephaly. Nat Genet 10:175–180 11. Melino G et al (2003) Functional regulation of p73 and p63: development and cancer. Trends Biochem Sci 28(12):663–670 12. Moll UM, Slade N (2004) p63 and p73: roles in development and tumor formation. Mol Cancer Res 2(7):371–386 13. Courtois S, de Fromentel CC, Hainaut P (2004) p53 protein variants: structural and functional similarities with p63 and p73 isoforms. Oncogene 23(3):631–638 14. Yang A et al (1998) p63, a p53 homolog at 3q27–29, encodes multiple products with transactivating, death-inducing, and dominant-negative activities. Mol Cell 2(3):305–316 15. Yang A et al (1999) p63 is essential for regenerative proliferation in limb, craniofacial and epithelial development. Nature 398(6729):714–718 16. Yang A et al (2000) p73-deficient mice have neurological, pheromonal and inflammatory defects but lack spontaneous tumours. Nature 404(6773):99–103 17. Kaghad M et al (1997) Monoallelically expressed gene related to p53 at 1p36, a region frequently deleted in neuroblastoma and other human cancers. Cell 90(4):809–819 18. Benard J, Douc-Rasy S, Ahomadegbe JC (2003) TP53 family members and human cancers. Hum Mutat 21(3):182–191 19. De Laurenzi V, Melino G (2000) Evolution of functions within the p53/p63/p73 family. Ann N Y Acad Sci 926:90–100 20. Flores ER et al (2005) Tumor predisposition in mice mutant for p63 and p73: evidence for broader tumor suppressor functions for the p53 family. Cancer Cell 7(4):363–373 21. Lee KC, Crowe AJ, Barton MC (1999) p53-mediated repression of alpha-fetoprotein gene expression by specific DNA binding. Mol Cell Biol 19:1279–1288 22. St Clair S et al (2004) DNA damage-induced downregulation of Cdc25C is mediated by p53 via two independent mechanisms: one involves direct binding to the cdc25C promoter. Mol Cell 16(5):725–736 23. Ho JS et al (2005) p53-Dependent transcriptional repression of c-myc is required for G1 cell cycle arrest. Mol Cell Biol 25(17):7423–7431 24. Hoffman WH et al (2002) Transcriptional repression of the anti-apoptotic survivin gene by wild type p53. J Biol Chem 277:3247–3257 25. Koumenis C et al (2001) Regulation of p53 by hypoxia: dissociation of transcriptional repression and apoptosis from p53-dependent transactivation. Feb;(4). Mol Cell Biol 21: 1297–1310 26. Ho J, Benchimol S (2003) Transcriptional repression mediated by the p53 tumour suppressor. Cell Death Differ 10(4): 404–408 27. Bode AM, Dong Z (2004) Post-translational modification of p53 in tumorigenesis. Nat Rev Cancer 4(10):793–805 28. Choi J, Donehower LA (1999) p53 in embryonic development: maintaining a fine balance. Cell Mol Life Sci 55(1): 38–47

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W.-W. Tsai and M. C. Barton   93. Li M et al (2003) Mono- versus polyubiquitination: differential control of p53 fate by Mdm2. Science 302(5652): 1972–1975   94. Gottifredi V, Prives C (2001) Molecular biology. Getting p53 out of the nucleus. Science 292(5523):1851–1852   95. Shirota Y et al (2002) Hepatitis C virus (HCV) NS5A binds RNA-dependent RNA polymerase (RdRP) NS5B and modulates RNA-dependent RNA polymerase activity. J Biol Chem 277(13):11149–11155   96. Murakami S (2001) Hepatitis B virus X protein: a ­multifunctional viral regulator. J Gastroenterol 36(10): 651–660   97. Hussain SP et al (2007) TP53 mutations and hepatocellular carcinoma: insights into the etiology and pathogenesis of liver cancer. Oncogene 26(15):2166–2176   98. Elmore LW et al (1997) Hepatitis B virus X protein and p53 tumor suppressor interactions in the modulation of apoptosis. Proc Natl Acad Sci U S A 94:14707–14712   99. Cheng Z et al (1997) Inhibition of hepatocellular carcinoma development in hepatitis B virus transfected mice by low dietary casein. Hepatology 26:1351–1354 100. Ogden SK, Lee KC, Barton MC (2000) Hepatitis B viral transactivator HBx alleviates p53-mediated repression of  a-fetoprotein gene expression. J Biol Chem 275: 27806–27814 101. Murakami S (1999) Hepatitis B virus X protien: structure, function and biology. Intervirology 42:81–99 102. Murray-Zmijewski F, Slee EA, Lu X (2008) A complex ­barcode underlies the heterogeneous response of p53 to stress. Nat Rev Mol Cell Biol 9(9):702–712 103. Liebermann DA, Hoffman B, Vesely D (2007) p53 induced growth arrest versus apoptosis and its modulation by survival cytokines. Cell Cycle 6(2):166–170 104. Schwartz D, Rotter V (1998) p53-dependent cell cycle control: response to genotoxic stress. Semin Cancer Biol 8(5):325–336 105. Di Leonardo A et al (1994) DNA damage triggers a prolonged p53-dependent G1 arrest and long-term induction of Cip1 in normal human fibroblasts. Genes Dev 8(21): 2540–2551 106. Genovese C et al (2006) Cell cycle control and beyond: emerging roles for the retinoblastoma gene family. Oncogene 25(38):5201–5209 107. Sherr CJ (1998) Tumor surveillance via the ARF-p53 pathway. Genes Dev 12(19):2984–2991 108. Giaccia AJ, Kastan MB (1998) The complexity of p53 modulation: emerging patterns from divergent signals. Genes Dev 12(19):2973–2983 109. Kastan MB et al (1992) A mammalian cell cycle checkpoint pathway utilizing p53 and GADD45 is defective in ataxia-telangiectasia. Cell 71(4):587–597 110. Zhan Q et al (1994) The p53-dependent gamma-ray response of GADD45. Cancer Res 54(10):2755–2760 111. Kuerbitz SJ et al (1992) Wild-type p53 is a cell cycle checkpoint determinant following irradiation. Proc Natl Acad Sci U S A 89(16):7491–7495 112. Kessis TD et al (1993) Human papillomavirus 16 E6 expression disrupts the p53-mediated cellular response to  DNA damage. Proc Natl Acad Sci U S A 90(9): 3988–3992 113. Taylor WR, Stark GR (2001) Regulation of the G2/M transition by p53. Oncogene 20(15):1803–1815

23  p53 114. Nurse P (1990) Universal control mechanism regulating onset of M-phase. Nature 344(6266):503–508 115. Pines J (1995) Cyclins and cyclin-dependent kinases: a biochemical view. Biochem J 308(Pt 3):697–711 116. Taylor WR et al (1999) Mechanisms of G2 arrest in response to overexpression of p53. Mol Biol Cell 10(11): 3607–3622 117. Agarwal ML et al (1995) p53 controls both the G2/M and the G1 cell cycle checkpoints and mediates reversible growth arrest in human fibroblasts. Proc Natl Acad Sci U S A 92(18):8493–8497 118. Bunz F et al (1998) Requirement for p53 and p21 to sustain G2 arrest after DNA damage. Science 282(5393): 1497–1501 119. Boulaire J, Fotedar A, Fotedar R (2000) The functions of the cdk-cyclin kinase inhibitor p21WAF1. Pathol Biol (Paris) 48(3):190–202 120. Vairapandi M et al (2002) GADD45b and GADD45g are cdc2/cyclinB1 kinase inhibitors with a role in S and G2/M cell cycle checkpoints induced by genotoxic stress. J Cell Physiol 192(3):327–338 121. Wang XW et al (1999) GADD45 induction of a G2/M cell cycle checkpoint. Proc Natl Acad Sci U S A 96(7): 3706–3711 122. Zhan Q et al (1999) Association with Cdc2 and inhibition of Cdc2/Cyclin B1 kinase activity by the p53-regulated protein Gadd45. Oncogene 18(18):2892–2900 123. Hermeking H et al (1997) 14–3-3 sigma is a p53-regulated inhibitor of G2/M progression. Mol Cell 1(1):3–11 124. Chan TA et al (1999) 14–3-3Sigma is required to prevent mitotic catastrophe after DNA damage. Nature 401(6753):616–620 125. Passalaris TM et al (1999) The G(2) checkpoint is maintained by redundant pathways. Mol Cell Biol 19(9): 5872–5881 126. Innocente SA et al (1999) p53 regulates a G2 checkpoint through cyclin B1. Proc Natl Acad Sci U S A 96(5): 2147–2152 127. Zhou J et al (2001) A role for p53 in base excision repair. EMBO J 20(4):914–923 128. Tanaka H et al (2000) A ribonucleotide reductase gene involved in a p53-dependent cell-cycle checkpoint for DNA damage. Nature 404(6773):42–49 129. Nakano K et al (2000) A ribonucleotide reductase gene is a transcriptional target of p53 and p73. Oncogene 19(37): 4283–4289 130. Bargonetti J, Manfredi JJ (2002) Multiple roles of the tumor suppressor p53. Curr Opin Oncol 14(1):86–91 131. Palacios G et al (2008) Mitochondrially targeted wild-type p53 induces apoptosis in a solid human tumor xenograft model. Cell Cycle 7(16):2584–2590

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The MYC Network and Cancer

24

Snorri S. Thorgeirsson and Valentina M. Factor

Introduction

MYC Transcriptome Network

Deregulation of c-Myc (referred to as MYC] contributes to the development of the most human tumors [1–3]. In addition to MYC, the Myc gene family contains three related genes, N-Myc, L-Myc, and S-Myc, which are also implicated in the genesis of specific human tumors (for review, see ref [4]). MYC is a nuclear transcription factor, which is first identified as the cellular homologue of the cancer-causing gene in the avian myelocytomatosis retrovirus [5]. MYC functions in a heterodimeric complex with MAX to bind E-Box motifs in DNA, and transcriptionally regulates hundreds to thousands of target genes. The most recent estimates suggest that MYC could regulate as many as 15% of genes in genomes from flies to human [6]. A compilation of MYC-regulated genes and studies on MYC alterations in human cancers is available online at www.myccancergene.org [6]. This database emphasizes both the critical role of MYC in human cancers and the significance of MYC target genes in driving its oncogenic activity. The target genes are involved in diverse programs including cell cycle, cell growth, protein synthesis, cell adhesion and cytoskeleton, metabolism, apoptosis, angiogenesis, DNA repair, and microRNA [6–8]. The diversity of MYC target genes is illustrated in Fig.  24.1. Numerous excellent and comprehensive reviews have been written about MYC [9–12]. Therefore, in this chapter, we will focus mainly on the role of MYC in cancer with the emphasis on the most recent findings.

To define the MYC transcriptome network, it is essential to identify the target genes regulated by MYC. The MYC-responsive genes are either genes directly bound by MYC or genes that require the activities of the direct target genes (indirect targets). Direct targets are defined as genes that are bound by MYC and respond to the changes in MYC levels and/or MYC activity. Most current models used to study Myc target genes rely on responses to the changes in MYC protein levels. This is well illustrated by experiments with serum starvation and restimulation of cells in culture, in which serum stimulation leads to a rapid activation of the MYC as a part of the early response genes program [13, 14]. The target genes are then investigated by measuring the kinetics of mRNA and protein levels induced by the changes in MYC levels. It is important to emphasize that MYC functions may be regulated by post-translational modifications that include phosphorylation and ubiquitylation, as well as interactions with a variety of proteins [15–22]. Various methodologies have been used to identify the direct MYC target genes (for a comprehensive review, see ref 6) but only two will be discussed here. The inducible MycER system, originally developed in Bishop’s laboratory, has proven to be a powerful tool for searching MYC target genes [23]. In MycERtransduced cells, the chimeric MycER protein is constitutively bound to the chaperone HSP90 in the cytoplasm. When the cells are treated with estrogenic compounds (e.g., 4-OH-tamoxifen), the chimeric protein separates from the chaperone via conformational changes, and translocates into the nucleus. Consequently, the MycER protein engages MYC target sites and initiates tran­ scription of the target genes in the absence of newly synthesized proteins, which is a property of a direct MYC target gene as discussed earlier. Thus, in this system

S. S. Thorgeirsson (*) Laboratory of Experimental Carcinogenesis, Center for Cancer Research, National Cancer Institute, National Institutes of Health, 37 Convent Drive, Bethesda, MD 20892, USA e-mail: [email protected]

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_24, © Springer-Verlag Berlin Heidelberg 2010

359

360

S. S. Thorgeirsson and V. M. Factor

Unknown

Cell cycle

Transport Protein phosphorylation

Transcription

RNA localization

Metabolism

Cell growth or maintenance Carbohydrate metabolism

RNA processing Protein dephosphorylation

Signal Transduction

Protein degradation Protein biosynthesis

Cell organization and biogenesis Protein transport

Stress response DNA repair

Fig. 24.1  Distribution of Myc targets by gene ontology (GO). One thousand five hundred and sixty-one Myc targets (small circles) from http://www.myccancergene.org are displayed in concentric rings by the OSPREY software (http://www.biodata.mshri.on.ca/

osprey/servlet/Index) with functional groups colored and labeled. GO groups highlighted in red are statistically over-represented as determined by EASE analysis (http://www.david.niaid.nih.gov/ david/ease.htm). Reproduced from [6] with permission

genes responding to the ligand-stimulated MycER are considered to be the direct MYC target genes. Chromatin immunoprecipitation (ChIP) is an important technique that has advanced our understanding of the association between transcription factors and cognate genomic sites in vivo [24, 25]. At present, ChIP is the only method that provides direct physical evidence of a transcription factor association with a specific target gene. By applying the ChIP method, target genes in sheared chromatin are cross-linked to a specific transcription factor and are subsequently immunoprecipitated with an antibody specific for that transcription factor. The chemical cross-links are then reversed, and the de-proteinized DNA is assayed by PCR or hybridized to microarrays to detect the specific genomic sequences which are precipitated along with the transcription factor in question. Although ChIP provides the most direct physical evidence of the association of a transcription factor with target genomic sites, its sensitivity is somewhat limited by the size distribution of

the sheared DNA fragments (from several hundred base pairs to about 1 kb). In order to obtain more detailed information on the localization of MYC binding within these DNA fragments, it is possible to use the preferential binding of MYC to E-boxes over other non-­ canonical sites [25–27]. Many studies have examined the changes in gene expression associated with the induction of MYC expression in cells [2, 28–33]. In addition, comparative analyzes of gene expression profiles between MYCinduced tumors in mouse models and human tumors in liver and prostate cancers have provided useful information [34, 35]. The analyses showed the similarities in gene expression between the experimental MYCinduced tumors and human tumors. However, these studies did not formally identify MYC-associated gene products responsible for driving the tumorigenesis. To address this issue, Felsher et al. have recently attempted to analyze the gene expression profiles in tumors generated in conditional transgenic mouse models in order to

24  The MYC Network and Cancer

361

MYC Off

MYC Reactivate common MYC associated tumorigenesis: 8 genes

Ptpru Gtf2f2 Bzw2 Time (hour) Dnajc2 Ppih H2afy Sfrs3 Nap1l1 Nola2 Mapk8 Blmh Mnat1 Ddx17 0610042115Rik Glud1 Ranbp3 Rcor1 Hrb Dhcr7 Mrg1 Ercc5 Ube2d2 Epc1 9430080K19Rik Son Ube2b 2810452K22Rik Trip13 Mmp12 9430080K19Rik 2700088M22Rik Jmjd1a Ccng1 Rsn

Lifr AA415817 Bat5 Tcn2 Fabp3 Edg1 Igh-6 Slc35b1

-2.00 -1.33 -0.67 0.00 0.67 1.33 2.00

common MYC associated tomorigenesis: 34 genes

p<0.01 StepMiner Analysis

Time (hour)

MYC Reactivate

0 4 8 12 18 24 36 48 4 8 12 18 24 36 48

MYC Off

MYC osteosarcomas time course data filtered with MYC pancreatic tumor repressed genes list

p<0.01 StepMiner Analysis

MYC osteosarcomas time course data filtered with MYC pancreatic tumor induced genes list

gene expression were associated with the permanent changes in the ability of MYC to bind to the promoter regions. Finally, to validate the role of the gene signatures associated with MYC in human tumorigenesis, the authors examined the expression of the human homologues in 273 published human lymphoma microarray datasets (using a Boolean analysis approach, ref 37). Among these genes, a large functional group comprised the ribosomal structural proteins, but genes involved in diverse cellular functions (i.e., BZW2, H2AFY, SFRS3, NAP1L1, NOLA2, UBE2D2, CCNG1, LIFR, FABP3, and EDG1) were also identified. This work demonstrates the power of using a defined transgenic mouse model of conditional tumorigenesis in which inactivation of MYC in tumor cells results in a permanent loss of neoplastic phenotype, allowing identification of a

0 4 8 12 18 24 36 48 4 8 12 18 24 36 48

identify the gene expression signature specifically associated with the ability of MYC to initiate and maintain tumorigenesis [36]. In this study, mRNA samples from a time-course experiment with MYC inactivated and then reactivated in osteosarcoma were used for mi­croarraybased global gene expression analysis. The StepMiner method [37] was used to analyze the microarray time courses, identify genes undergoing abrupt transitions in expression level, and determine the times at which these transitions occur. Consequently, the StepMiner analysis revealed genes whose expression was most strongly correlated with the capacity of MYC to induce a neoplastic state (Fig.  24.2). In addition, genes displaying permanent expression changes following MYC inactivation were also recognized. By employing ChIP analysis, the authors demonstrated that the stable changes in

color scale

Fig. 24.2  A common gene signature associated with the ability of MYC to induce tumorigenesis in murine conditional tumor models. Microarray data from the time-course experiment in MYC-induced osteosarcoma was filtered with the list of tumor maintenance genes from pancreatic tumors. 34 genes from the

induced gene list and eight genes from the repressed gene list were identified (p < 0.01) as common MYC target genes associated with MYC-induced tumorigenesis in mice. Genes with E-box sequences in their promoter regions (−2,000 to +2,000) are labeled with red. Reproduced from Ref 36 with permission

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gene list specifically correlating with the capacity of MYC to maintain tumorigenesis. These results can be used in combined analysis of gene expression data from human cancers (e.g., human lymphomas) to reveal novel cancer-type specific gene signatures correlating with MYC’s ability to maintain the neoplastic stage.

Biological Output of MYC In vivo One of the many functions of the MYC protein involves engaging and coordinating expression of diverse genes necessary for the efficient and orderly proliferation of somatic cells. In normal cells, the mRNA and protein expression of MYC are low and strictly dependent on mitogen signalling [38–40]. In stark contrast, MYC expression is either deregulated or elevated in most human tumors. These modifications can be caused by alterations in the MYC gene itself which perturb its normal regulation (i.e., chromosomal translocation, retro­ virus integration, or gene amplification), by increased MYC mRNA or MYC protein stability, or by abrogated MYC autorepression (reviewed in [41, 42]). However, in most tumors the MYC gene appears normal. In all likelihood, the upregulation of MYC activity in human tumors is due to persistent induction via multiple upstream oncogenic pathways including RAS/RAF/ MPAK [43], JAK/STAT, and WNT/b-catenin [38, 44, 45] which may result in a significant overlap between the MYC and other oncogene-induced gene expression signatures. The notion that spontaneous tumorigenesis is suppressed to a large extend by a strict coupling of proliferation to the anti-oncogenic programs such as senes­cence and apoptosis is generally accepted [46, 47]. This situation is well illustrated by MYC. Tumorigenic activation of MYC not only induces a strong proliferation of cells but also utilizes the tumor suppressor programs such as ARF/p53 pathway [48–51] and apoptosis [52, 53], thus antagonizing cell expansion and restricting the MYC oncogenic potential . Since MYC can induce proliferation of normal cells, it can be assumed that the activation of ARF/p53 and apoptosis is restricted to the oncogenic function of MYC. How cells distinguish between the mitogenic and oncogenic MYC activities has been recently addressed by Evan and colleagues [54]. In an elegant study, the authors employed an MYC transgenic mouse model in which the latent expression

S. S. Thorgeirsson and V. M. Factor

of the reversibly switchable variant of MYC, MycERT2, was driven by the constitutive and ubiquitously active Rosa26 promoter. Explicit MycERT2 expression was triggered in any target tissue by the hit-and-run action of Cre recombinase. Because the Rosa26 promoter is relatively weak, the MycER levels were similar to the physiological MYC levels after a normal mitogen stimulation. This study demonstrates that the distinct threshold levels of MYC dictate its output in vivo, that is, low levels of deregulated MYC (1.5 to two fold normal MYC level) are competent to drive ectopic proliferation of somatic cells and oncogenesis. However, more than a two fold increase in MYC level is needed to activate the apoptotic and ARF/p53 intrinsic tumor surveillance pathways. Therefore, keeping the activated Myc (and, presumably, other oncogenes) at low levels, which prevents engaging tumor suppressors, may constitute an important selective pressure governing the early stages of tumor microevolution. The work of Evan et al. has many important implications with respect to MYC-induced oncogenesis. In addition, as has been discussed in [55], the work may have serious implications for cancer therapies based on reducing MYC protein levels. Treatment with the kinase inhibitor imatinib can cause a partial inhibition of oncogene function and thereby induce remission of some tumors. However, even though treatment based MYC inhibition may be able to suppress the high MYC levels found in many advanced tumors, the residual MYC levels might be still sufficient to support the proliferation of tumor cells which have also acquired attenuated tumor suppressor and/or apoptotic response.

Conclusions The impact of MYC deregulation on human cancer incidence is enormous given the diverse cellular programs regulated by this transcription factor which include cell cycle, cell growth, protein synthesis, cell adhesion and cytoskeleton, metabolism, apoptosis, angiogenesis, stem cell fate, DNA repair, and selective control of microRNA expression. Here, we highlight two recently discovered properties of the MYC pathway, the gene network maintaining MYC driven oncogenesis and the mechanism by which low-level deregulation of MYC expression can exert a potent oncogenic effect by circumventing tumor

24  The MYC Network and Cancer

suppressor defense mechanisms. Although much has been learned about the oncogenic functions of MYC, there are still numerous questions to answer, such as what determines when MYC functions as a transcriptional activator or repressor, and how MYC is regulated in stem cells and cancer stem cells. Deciphering MYC’s role in cancer remains a challenge.

Summary

›› Myc is an oncogenic transcription factor which ›› ›› ›› ›› ›› ››

functions in a heterodimeric complex with MAX MYC could regulate as many as 15% of genes in the genome MYC functions can be regulated by posttranslational modifications that include phosphorylation and ubiquitylation In normal cells, expression of MYC is low and strictly dependent on mitogen signalling. MYC expression is deregulated or elevated in most human tumors Low levels of deregulated MYC are comptent to drive cell proliferation and oncogenesis More than two fold increase in MYC level is needed to activate the apoptotic and ARF/p53 tumor surveillance pathways.

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(b) Increase in mRNA expression (c) Increase in MYC protein stability (d) 2 & 3 only (e) All of the above 3. MYC regulates expression of (a) 0.1% of all genes (b) 1% of all genes (c) 10% of all genes (d) 15% of all genes (e) 25% of all genes 4. MYC functions include regulation of: (a) Cell proliferation (b) Apoptosis (c) Cell growth (d) Cell differentiation (e) All of the above 5. Outcomes of different levels of deregulated MYC expression: (a) Only high levels of MYC can cause cancer (b) Low levels of MYC are not sufficient to induce proliferation (c) Both high and low levels of MYC expression can drive oncogenesis (d) Suppression of MYC is not sufficient for tumor regression (e) Other members of MYC gene family do not contribute to carcinogenesis

Selected Readings Multiple Choice Questions 1. Role of MYC disregulation in cancer (a) MYC plays a limited role in cancer (b) MYC is essential only for cancer initiation (c) MYC is essential for cancer maintenance (d) Suppression of MYC overexpression is not sufficient for tumor regression (e) MYC is essential both for cancer initiation and maintenance 2. Deregulation of MYC expression is caused by (a) Alterations in MYC gene itself

1. Eisenman RN. Deconstructing myc. Genes Dev. 2001;15:2023-30 (comprehensive analysis of the problem derives from the apparent gap between Myc’s biological role and what is surmised to be its molecular function) 2. Eilers M, Eisenman RN. Myc’s broad reach. Genes Dev. 2008; 22:2755 (two major aspects of MYC - the nature of the genes and pathways that are targeted by Myc, and the role of MYC in stem cell and cancer biology - are reviewed in this article 3. Meyer N, Penn LZ. Reflecting on 25 years with MYC. Nat Rev Cancer. 2008; 8:976-990 (in this article the authors chronicle the major advances in our understanding of MYC biology since the discovery of MYC 25  years ago, and examine the future trends of MYC research)

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S. S. Thorgeirsson and V. M. Factor 24. Eberhardy SR, Farnham PJ (2001) c-Myc mediates activation of the cad promoter via a post-RNApolymerase II recruitment mechanism. J Biol Chem 276:48562–48571 25. Zeller KI, Haggerty TJ, Barrett JF et al (2001) Charac­terization of nucleophosmin (B23) as a Myc target by scanning chromatin immunoprecipitation. J Biol Chem 76: 48285–48291 26. Mao DY, Watson JD, Yan PS et al (2003) Analysis of Myc bound loci identified by CpG island arrays shows that Max is essential for Myc-dependent repression. Curr Biol 13:882–886 27. Haggerty TJ, Zeller KI, Osthus RC et al (2003) A strategy for identifying transcription factor binding sites reveals two classes of genomic c-Myc target sites. Proc Natl Acad Sci U S A 100:5313–5318 28. Coller HA, Grandori C, Tamayo P et al (2000) Expression analysis with oligonucleotide microarrays reveals that MYC regulates genes involved in growth, cell cycle, signaling, and adhesion. Proc Natl Acad Sci U S A 97:3260–3265 29. Guo QM, Malek RL, Kim S et al (2000) Identification of c-myc responsive genes using rat cDNA microarray. Cancer Res 60:5922–5928 30. Schuldiner O, Benvenisty N (2001) A DNA microarray screen for genes involved in c-MYC and N-MYC oncogenesis in human tumors. Oncogene 20:4984–4994 31. Marinkovic D, Marinkovic T, Kokai E et al (2004) Identification of novel Myc target genes with a potential role in lymphomagenesis. Nucleic Acids Res 32:5368–5378 32. Lawlor ER, Soucek L, Brown-Swigart L et al (2006) Reversible kinetic analysis of Myc targets in vivo provides novel insights into Myc-mediated tumorigenesis. Cancer Res 66:4591–4601 33. O’Connell BC, Cheung AF, Simkevich CP et al (2003) A large scale genetic analysis of c-Myc-regulated gene expression patterns. J Biol Chem 278:12563–12573 34. Ellwood-Yen K, Graeber TG, Wongvipat J et al (2003) Mycdriven murine prostate cancer shares molecular features with human prostate tumors. Cancer Cell 4:223–238 35. Lee JS, Chu IS, Mikaelyan A et al (2004) Application of comparative functional genomics to identify best-fit mouse models to study human cancer. Nat Genet 36:1306–1311 36. Wu CH, Sahoo D, Arvanitis C et al (2008) Combined analysis of murine and human microarrays and ChIP analysis reveals genes associated with the ability of MYC to maintain tumorigenesis. PLoS Genet 4:1–16 37. Sahoo D, Dill DL, Tibshirani R et al (2007) Extracting binary signals from microarray time-course data. Nucleic Acids Res 35:3705–3712 38. Liu J, Levens D (2006) Making myc. Curr Top Microbiol Immunol 302:1–32 39. Rabbitts PH, Watson JV, Lamond A et al (1985) Metabolism of c-myc gene products: c-myc mRNA and protein expression in the cell cycle. EMBO J 4:2009–2015 40. Ramsay G, Evan GI, Bishop JM (1984) The protein encoded by the human proto-oncogene c-myc. Proc Natl Acad Sci U S A 81:7742–7746 41. Popescu NC, Zimonjic DB (2002) Chromosome-mediated alterations of the MYC gene in human cancer. J Cell Mol Med 6:151–159 42. Spencer CA, Groudine M (1991) Control of c-myc regulation in normal and neoplastic cells. Adv. Cancer Res 56:1–48 43. Sears R, Leone G, DeGregori J et al (1999) Ras enhances Myc protein stability. Mol Cell 3:169–79 44. He TC, Sparks AB, Rago C et al (1998) Identification of c-MYC as a target of the APC pathway. Science 281:1509–12

24  The MYC Network and Cancer 45. Kolligs FT, Bommer G, Goke B (2002) Wnt/beta-catenin/tcf signaling: a critical pathway in gastrointestinal tumorigenesis. Digestion 66:131–144 46. Evan G, Littlewood T (1998) A matter of life and cell death. Science 281:1317–1322 47. Lowe SW, Cepero E, Evan G (2004) Intrinsic tumour suppression. Nature 432:307–315 48. Eischen CM, Weber JD, Roussel MF et al (1999) Disruption of the ARF-Mdm2–p53 tumor suppressor pathway in Mycinduced lymphomagenesis. Genes Dev 13:2658–2669 49. Kamijo T, Weber JD, Zambetti G et al (1998) Functional and physical interactions of the ARF tumor suppressor with p53 and Mdm2. Proc Natl Acad Sci U S A 95:8292–8297 50. Schmitt CA, McCurrach ME, de Stanchina E et al (1999) INK4a/ARF mutations accelerate lymphomagenesis and

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The WNT/b-Catenin Pathway

25

Satdarshan P. S. Monga

Background Genetic studies in species such as Xenopus, Drosophila, and Caenorhabditis have lent themselves quite well to further our understanding of the molecular basis of human diseases. A classical example is the identification and characterization of the Wnt/b-catenin pathway that is crucial in normal development including embryogenesis, organogenesis, and epithelial-mesenchymal transition and at the same time its deregulation is implicated in disorders such as cancers (reviewed in [1–3]). This pathway has remained conserved through the evolutionary process. In Drosophila, the role of Wnt or Wingless (Wg) was initially identified in normal wing development, however, it was later recognized for multiple functions such as inducing segment polarity and anterior–posterior patterning that were imperative for a viable embryo [4–6]. As the importance of Wnt emerged, several key components of this pathway were identified. The discovery of armadillo (or b-catenin) added a significant player to this orchestra and although circumstantial evidence suggesting such a relationship existed earlier it was a few years later that b-catenin was positively identified as a central component of the canonical Wg pathway [5, 7–9]. These studies led to the emergence of a model system for cell adhesion and signal transduction [10]. This was also the beginning of the understanding of the Wnt/b-catenin pathway and its role in complex cellular processes such as cell– cell adhesion, mitogenesis, motogenesis, and morphogenesis in the vertebrates.

S. P. S. Monga Division of Experimental Pathology, University of Pittsburgh, School of Medicine, Pittsburgh, PA 15261, USA e-mail: [email protected]

Next several years focused on the discovery of various components of this pathway that improved our understanding of the regulation of this complex signaling pathway in normal physiology and disease. Several important players such as the Wnt receptor frizzled (Fz), zeste-white 3 kinase or glycogen synthase kinase 3b  (GSK3b), adenomatous polyposis gene product (APC), axin, Disheveled (Dsh), and T cell factor-1 (TCF1), and their interactions were identified that were directly influenced by the Wnt signaling [11–18]. Many new components and interactions as well as expanding list of target genes are being identified. Also, research is focused on their role in regulation of this pathway in health and disease. Furthermore, crosstalks have now been established between the Wnt pathway and other prominent pathways such as the Jagged/Notch, HGF/Met, EGF, and TGF pathways that could have additional implications [19–25]. Presently, the role of Wnt/b-catenin pathway is well established in vertebrates in embryogenesis and carcinogenesis [3, 26]. b-Catenin knockout yields an embryonic lethal phenotype in mice due to defect in gastrulation [27]. Other studies in vertebrates have also shown its role in anterior–posterior axis specification and mesoderm formation [28, 29]. Availability of conditional knockouts to overcome embryonic lethality has been key to understanding a more ubiquitous role of b-catenin and other Wnt components in the development of many organs such as kidneys, lungs, brain, limbs, muscles, and skin [30–35]. Its role in liver development is beginning to be uncovered and is discussed in Sect.  25.3.1. This pathway is crucial in stem cell biology where it is known to regulate stem cell renewal in multiple tissues including hematopoietic, epidermal, and intestinal compartments [36–40]. It has also been identified as a prominent player in angiogenesis and vasculogenesis and maintenance of endothelial cell

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_25, © Springer-Verlag Berlin Heidelberg 2010

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adhesion [41–44]. Finally, its role in regulating metabolism ranging from carbohydrate, protein, fat, and xenobiotics is also being better understood and might have additional implications.

The WNT/b-Catenin Signal Transduction Pathway The binding of an extracellular secreted glycoprotein Wnt to its cell surface receptor Fz induces specific downstream events consisting of many intricate protein– protein interactions involving meaningful changes in their binding, phosphorylation, and localization [45]. Although the most understood and predictable events are the result of the activation of the canonical Wnt pathway, the signaling can be transduced to at least two other branches. These two pathways are the planar cell polarity pathway and the Wnt/Ca2+ pathway. How is the diversification of these signals being modulated still remains under intense investigation! However, a concept that is gradually vanishing is that of designated canonical and noncanonical Wnts (reviewed in [46]). In fact, Wnts from either of these historical classes can signal through b-catenin-dependent or independent manner. It has becoming apparent that diversification of the signal is highly dependent on the receptors, which in turn can be highly context dependent. In fact, the final outcome of a particular signaling might be highly stage-specific, tissue-specific, and might be dictated by several protein–protein interactions occurring at the time of ongoing signaling, making this process highly complex. For the sake of simplicity and clarity, many recent changes in Wnt signaling nomenclature have been suggested. These are extremely relevant although readers need to be conscious of the fact that this field is rapidly evolving and hence such terminology will have to stand the test of time. From a liver perspective, it will be most relevant to outline the major components and interactions in various Wnt signaling pathways, as elaborated in the forthcoming sections.

The Canonical WNT Pathway or the WNT/b-Catenin Signaling In a normal steady state where excess of b-catenin, a key component the pathway and a powerful “­oncoprotein,”

S. P. S. Monga

is not needed or in the absence of a Wnt signal, the free monomeric form of b-catenin in the cytoplasm is actively targeted for degradation by ubiquitination. This is comparable to the pathway being in “OFF” mode (reviewed in [3]). In this situation, b-catenin is being phosphorylated at serine and threonine residues in its amino terminal region, specifically at serine-45 (Ser45), Serine-33 (Ser33), and threonine-41 (Thr41) by Casein kinase Ia (CKIa and GSK3 [47–50]. CK and GSK3b are part of a larger multiprotein degradation complex that includes axin, which acts as a scaffold to form homodimeric or heterodimeric complexes with axin2/conductin, APC, and diversin and each of these, play a role in b-catenin degradation [12, 51, 52]. Once phosphorylated, this larger complex enables recognition and ubiquitination of b-catenin by b-transducin repeat-containing­protein (bTrCP) and its ensuing proteosomal degradation [53]. Thus free levels of b-catenin are kept low and it is prevented from translocating to the nucleus to induce target gene transcription. These events can also be observed if Wnts are sequestered or prevented from binding to their receptors. Several such modulators have now been identified. Fz related proteins (FRPs) are smaller proteins (30 kDa) with Fz-like cysteine-rich domain that bind and sequester Wnts [54]. Similarly, Wnt inhibitory factors (WIFs) bind Wnts to inactivate the pathway [55]. Cerebrus is a more nonspecific inhibitor that represses Wnt, nodal, and bone morphogenic protein (BMP) signaling [56] (Fig. 25.1). Any of the Wnts (19 members in humans) in the absence of their negative regulators, bind to their seven-transmembrane receptor Fz that further induces a ternary complex formation with LRP5 or LRP6 (or arrow) [11, 57–59]. This complex is crucial in dictating the downstream canonical Wnt/b-catenin signaling. One of the inhibitors, Dickkopf (Dkk), prevents Wnt-induced Fz-LRP complex formation and hence, Wnt signaling [60]. Upon formation of the ternary complex, signal is transduced through multiple ­intermediate proteins to finally induce ­hypophosphorylation of b-catenin at the APC-axin-GSK3b-CK complex. One such interaction is the activation of Dsh that blocks b-catenin degradation by recruiting GSK3b-binding protein (GBP)/Frat-1 that displaces GSK3b from axin, resulting in its inactivation [61, 62]. Also, Dsh can bind to phosphatase PP2C that enables it to dephosphorylate axin [63]. Dsh can also potentiate b-catenin stabilization following its activation by the serine/threonine kinase Par-1 [64]. CK1epsilon and CK2 are yet

25  The WNT/b-Catenin Pathway

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Fig. 25.1  Two state canonical pathway signaling. On the left, in absence of Wnt or presence of its inhibitors, b-catenin is phosphorylated at ser/thr residues to be degraded. The right panel shows activation of the pathway in presence of Wnt that allows b-catenin to be released from its cytoplasmic complex to translocate to the nucleus and bind to TCF family members and induce target genes. Black boxesOncogenes; Gray boxestumor suppressor genes

two another unrelated kinases that interact with Dsh to induce b-catenin activation [65–69]. The end result is the hypophosphorylation of b-catenin at specific serine and threonine residues, its release from the multiprotein complex, cytoplasmic stabilization of its monomeric form, and ensuing nuclear translocation where it binds to an HMG box containing DNA-binding ­protein T cell factor/lymphoid enhancing factor (TCF/LEF) family member [70, 71]. Transcription under the control of b-catenin/TCF4 is a highly complicated event and is reviewed comprehensively elsewhere [72]. Briefly, in the absence of an activating Wnt signal, TCF inside the nucleus acts as a repressor of the target genes and it does so at least in conjunction with a corepressor Groucho and interactions with histone deacetylase Rpd3 [73–75]. In the presence of a Wnt signal, b-catenin can induce the  transcriptional activation capability of TCF and the two important players identified at this level are the legless or Bcl9 and pygopos. Legless promotes recruitment of pygopos to b-catenin in the nucleus and permits it to become transcriptionally active [76, 77]. Other positive regulator to be identified is Brg-1, which is a ­component of mammalian SWI/SNF and Rsc chromatin-remodeling complexes. It has been shown that b-catenin recruits Brg-1 to the TCF target

gene promoters to assist in chromatin remodeling that is necessary for transcriptional activation [78]. CREBbinding protein (CBP), which is a known coactivator for several transcription factors, was shown to repress TCF in Drosophila [79]. However, in vertebrates, CBP and another related acetyltransferase p300 acted as a transcriptional coactivator in b-catenin-TCF transcription machinery [80, 81]. More recently, P300 and CBP have shown to have differential and sometimes, opposite effects on target gene promoters e.g., survivin [82]. Two other homologous proteins – pontin52 and reptin52, bind to b-catenin, and function as its antagonistic regulators [83, 84]. Finally, another protein that deserves a mention is chibby, which functions as a competitive inhibitor of b-catenin-mediated transcriptional activation by competing with LEF-1 [85]. Once the TCF-b-catenin complex is formed in the nucleus, there is transcriptional activation of several target genes that have now been identified (Table 25.1). An emerging concept is the stage and tissue-specificity of the transcriptional targets of this pathway. Apart from the targets listed in Table  25.1, several Wnt components such as AXIN, DKK, dFz7, Fz2, FRP2, WISP, bTrCP, and TCF are themselves targets, suggesting existence of several regulatory loops within this pathway (Fig. 25.2).

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Table 25.1  List of prominent target genes of the Wnt/b-catenin pathway Target genes Model Axin-2 Human colon cancer BMP4 Human colon cancer Xenopus C-Jun Human colon cancer C-Myc Human colon cancer Cdx1 Mouse Wnt3A Cdx4 Zebrafish Connexin-43 Xenopus, Mouse Cyclooxygenase-2 Mouse (Wnt1) 3T3L1preadipocytes Cyclin-D1 Human colon cancer E-cadherin Mouse hair follicles Epidermal growth factor receptor Mouse liver FGF4 Mouse tooth bud Fibronectin Xenopus G-protein-coupled receptor 49 (Gpr49) Hepatocellular cancer Glutamate transporter-1(GLT-1) Mouse liver Glutamine synthetase (GS) Mouse liver IGF-I/IGF-II 3T3L1 preadipocytes Keratin Mouse hair follicle MMP-7 Human colon cancer Ornithine aminotransferase Mouse liver PPAR-d Human colon cancer Survivin Human colon cancer TCF-1 Human colon cancer uPAR Human colon cancer VEGF Human colon cancer

Regulation Upregulated Upregulated Downregulated Upregulated Upregulated Upregulated Upregulated Upregulated Upregulated Upregulated Upregulated Downregulated Upregulated Upregulated Upregulated Upregulated Upregulated Upregulated Upregulated Upregulated Upregulated Upregulated Upregulated Upregulated Upregulated Upregulated Upregulated

Reference [259] [260] [261] [262] [263] [39] [264] [265] [266] [267] [268, 269] [270] [176] [271] [272] [273] [180] [180] [267] [274] [275, 276] [180] [261] [277] [16] [262] [278]

The Alternative WNT Signaling Pathways

The WNT/Ca2+ Pathway

As mentioned earlier, the past classification of Wnts into canonical and noncanonical Wnts is a fading concept. Thus, it is important to emphasize that the canonical Wnts such as Wnt1 have been shown to inhibit b-catenin activity. Also, the traditional noncanonical Wnts such as Wnt5a and Wnt11 have now been shown to be able to induce activation of b-catenin. In fact, while the participation of Fz and Dvl in the planar cell polarity (PCP) or Wnt/Ca2+ pathways is undeniable, the requirement of actual Wnt proteins in these pathways has been challenged. Wnts as ligands for alternate signaling, which has been traditionally divided into the Wnt/Ca2+ or PCP pathway, have now been suggested to be due to the interactions with nonfrizzled receptors such as atypical tyrosine kinase Ryk and single pass receptor tyrosine kinase Ror2, respectively [46]. However, while these interactions are being elucidated, it is relevant to give a brief overview of these alternate Wnt signaling cascades as they stand today.

The first evidence of the existence of such a pathway came from a comparable phenotype that was observed in Xenopus following the overexpressions of Wnt5A and 5HT1c serotonin receptor [86, 87]. At that time serotonin receptor was known to stimulate Ca2+ release in a G-protein-dependent fashion suggesting the possibility that a similar might be occurring in response to Wnt5A [86]. Further analysis identified a rat Fz-2 (rFz2) that induced intracellular Ca2+ release in response to Wnt5A activation via interaction with the phosphatidylinositol pathway in a G-protein-dependent manner [88, 89]. This induced an increase in intracellular Ca2+ that in turn stimulated two major Ca2+-sensitive enzymes-Ca2+/ calmodulin-dependent protein kinase II (CamKII) and protein kinase C (PKC) [90, 91]. CamKII activation was shown by in vitro kinase activity and increased autophosphorylation and the activation of PKC was demonstrated in vitro kinase activity and membrane translocation. These events were shown to occur in a

25  The WNT/b-Catenin Pathway

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Fig. 25.2  All component signaling in the canonical Wnt pathway. This cartoon is a comprehensive schematic that includes most of the components of this pathway demonstrating various protein–protein interactions occurring in the extracellular region, subcellular and cytoplasmic region as well as in the nucleus. This figure has been borrowed with permission from Dr. Roel Nusse’s website at www. stanford.edu/~rnusse/ wntwindow.html

b-catenin-independent manner as shown by the inability of Wnt-8 and Rfz-1 (activate canonical signaling) to activate either CamKII or PKC activation (reviewed in [92]). More functional evidence of this pathway came from the examination of this pathway as a “ventralizing” inducer in Xenopus. It was shown that the elevated levels of intracellular Ca2+ in response to Wnt also activated phosphatase calcineurin that initiated dephosphorylation of the transcription factor NF-AT, allowing its nuclear translocation and activation of target genes [93]. How this pathway is regulated in relation to the canonical Wnt pathway is still unclear but it is suggested that NF-AT or

its targets might influence the canonical pathway downstream of Dsh and upstream of b-catenin to balance the dorsal-ventral axis formation [93] (Fig. 25.3).

The Planar Cell Polarity Pathway Additional studies uncovered yet another pathway that involves Wnt signaling and is distinct from the two pathways described so far (reviewed in [94]). The clues for existence of this branch of the pathway emerged initially from the studies in Drosphila wing. As with

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Fig. 25.3  Left panel represents the Wnt/Ca2+ pathway. Wnt signaling incites intracytoplasmic Ca2+ accumulation through Ins [47, 53, 190] P3 receptor. This in turn activates Ca2+ -calmodulindependent protein kinase II, protein kinase C or calcineurin. Calcineurin induces nuclear translocation of NF-AT to activate target genes. Right panel is the planar polarity pathway where frizzled and disheveled (or axin) via Daam1 activate the JNK or Rho-associated kinase (ROCK) to induce cytoskeletal changes to achieve planar polarity

all epithelial cells, these specialized cells are polarized along their apical-basal axis. In addition, these cells also exhibit planar polarity that arranges the cells within the epithelial plane of the wing in a proximaldistal axis. This involves rearrangement of cellular cytoskeleton along the proximal-distal axis such that actin is polymerized at its distal tip that forms wing hairs that uniformly point distally (reviewed in [95]). Quest for such genes led to the discovery of Fz and Dsh as central players in this rearrangement [96]. However, no role of b-catenin could be identified in planar polarity [97]. This triggered intense research to elucidate bifurcation of the pathway at the level of Fz and disheveled [98]. The summary that emerges shows activation of Jun-N-terminal kinase (JNK) in response to Fz. In wing, Rho and Rho kinase are important intermediates that are downstream of Fz, whereas in eye, another tissue exhibiting planar polarity, a small guanosine triphosphatase Rho and JNK-MAPK are prominent players [99–103]. Also, Dsh and axin have now been shown to directly activate the JNK pathway suggesting that they might function in cooperation [104]. Interestingly, loss of function studies of JNK and JNKK show no compromise in planar polarity and it has been suggested that other MAPK components might have a redundant role in this process [105]. This also led to identification of two additional kinases – TAK1 (MAPKKK homologue) and MAPK family member Nemo, which are proposed to function in parallel to the canonical Wnt pathway to confer planar polarity [106, 107]. As it stands now, this pathway branches of from the canonical pathway at Dsh; and

involves cadherin-related transmembrane protein flamingo (Fmi) or Starry Night; the proteoglycan knypek (Kny); and the PDZ molecule Stbm [108–110]. Dsh is connected to Rho and Rho-associated kinase (ROCK) via Daam [111]. How the signal is detoured towards planar polarity pathway remains obscure. Recently, a product of the Wnt target gene naked (Nkd) has been shown to bind to Dsh and stimulate JNK pathway at the same time blocking b-catenin [112]. What upstream proteins or which Wnts or Fz specifically, if at all, favor one pathway vs. the other, or is it more of a tissue or stage specific decision remains undecided. However, it is now evident that planar polarity might be a function of establishing a gradient of Wnt and Fz signaling along a specific axis within the sheet of epithelial cells. This has led to the discovery of a role of cadherin superfamily of adhesion molecules-Fat (Ft), dachsous (Ds), and four-jointed (Fj) in Drosophila eye [113]. Wnt regulates expression of Ds and Fj that further generates a gradient of Ft activity that in turn establishes Fz activity gradient along the desired axis. Although more analysis will be vital, similar conserved pathways might operate in mammals conferring cell polarity to specialized cells as hepatocytes (Fig. 25.3).

b-Catenin-E-Cadherin Interactions Apart from playing a central role in the canonical Wnt pathway as a transcriptional coactivator, b-catenin performs a yet another crucial function by acting as a

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bridge between the cytoplasmic domain of the cadherins and the actin-containing-cytoskeleton [114–116]. It is interesting that each of these two roles are played by a distinct b-catenin in C. elegans [117]. We have reported a smaller species of b-catenin during liver development, that appears to be located at the membrane and associating to E-cadherin, and a functional characterization is pending [118]. Cadherins consist of an extracellular domain, a transmembrane domain, and a cytoplasmic tail that is the most conserved region among various subtypes. Type I cadherins are the most characterized and consist of E-cadherin and N-cadherin. Structurally, the cytoplasmic tails of cadherins show dimerization and connect to the actin-cytoskeleton via p120, b-catenin and a-catenin (Fig.  25.4). Specific b-catenin-binding sites on the cytoplasmic domain of cadherins have been characterized [119, 120]. The significance of regulation of b-catenin–cadherin interactions is not only important in modulating cell–cell adhesion but has been extended to transcriptional activation function of b-catenin as well. These interactions are regulated by tyrosine phosphorylation and not phosphorylation at serine/threonine residues (reviewed in [121]). There is a large body of literature that has shown significance of such interaction by multiple means. Phosphorylation of b-catenin destabilizes cadherin–b-catenin bond, a-catenin–bcatenin complex, uncouples cadherin from actin cytoskeleton and promoting loss of intracellular adhesion [122,  123]. Dephosphorylating b-catenin at tyrosine residues enhanced E-cadherin, b-catenin, and

a-catenin reassembly [124]. Following tyrosine 654 phosphorylation of b-catenin, its cytosolic pool is greatly increased as is its ability to bind to TATA-box binding protein (TBP) and increased transcriptional activity of b-catenin/TCF complex [125]. Another important ramification of tyrosine phosphorylation of b-catenin and dissociation of b-catenin–E-cadherin complex is that it leaves the cytoplasmic domain of E-cadherin vulnerable to degradation [119]. How is the cadherin–b-catenin complex being regulated? The answer to this question is quite complex and only the most pertinent regulators are mentioned. One level of regulation of catenin–cadherin complex is via the GTP-bound form of Ga subunit of heterotrimeric G proteins and overexpression of Ga12/13 results in dissociation of this complex [126, 127]. Another key regulator of this complex is the protein tyrosine phosphatase 1B (PTP1B) that directly associates to the intracytoplasmic tail of cadherins [128, 129]. There is a partial overlap in the binding domains of PTP1B, b-catenin, and Ga12, thus, adding complexity to the regulation of cadherin function (reviewed in  [121]). Other specific interactions that regulate phosphotyrosine-b-catenin include (a) nonreceptor kinases – src and Fer [130, 131]; (b) transmembrane kinases – EGF receptor (EGFR) and Met (HGF receptor) [21, 132–135]; (c) Protein tyrosine phosphatases including LAR-PTP, the chondroitin sulfate proteoglycan PTPb/z, and members of the Meprin/A5/Mu (MAM) domain containing family [136–140].

Miscelleneous Interactions/Crosstalks Extracellular domain

E-Cadherin

*

Intracytoplasmic tail

*

*

p120

*

�-Catenin

�-Catenin Actin

*

Fig. 25.4  Schematic showing the cadherin–catenin complex at the cell membrane. The key regulation sites are marked by asterisks

There are a few other interactions that are worth mentioning. We reported a novel Met–b-catenin complex at the hepatocyte membrane that appears to be independent of E-cadherin–b-catenin complex and is liver-specific [22]. HGF induced tyrosine phosphorylation-dependent nuclear translocation of b-catenin with an increase in c-myc by interactions involving the Met–b-catenin complex. In a follow up study, tyrosine residues 654 and 670 were identified as targets of HGF-induced b-catenin phosphorylation [141]. Other reports had identified a similar effect of HGF on positively regulating b-catenin/ TCF transactivation, albeit via other mechanisms [20, 24, 142]. These observations are relevant as high levels of HGF and have been observed in patients with liver pathologies that might be influencing b-catenin redistribution and altering the course of the disease [143, 144].

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Similarly, direct interactions of b-catenin with EGFR have been reported as well [21]. In fact, ErbB2 has also been shown to be associated with b-catenin [133]. Although, the fate of b-catenin and nuclear signaling in this context is unclear its effect on cell–cell adhesion is well defined [145]. Another key crosstalk is with the transforming growth factor b (TGFb). There exists a physical interaction between the b-catenin–TCF complex and smad4. Smad4 is a mediator of the TGFb signaling that interacts with smad2-smad3 heterodimers following TGFb signaling. Considering the role of TGFb signaling in liver growth and regeneration and also the phenotypes observed in the “loss of function” studies involving components of this pathway such as embryonic lethality due to compromised liver development and alterations in b-catenin, E-cadherin, and b1-integrin in mice lacking a copy of smad2 and smad3, indicates a great deal of relevance of this crosstalk in liver [146–149].

WNT/b-Catenin Signaling in Liver: Physiological Relevance The importance of the Wnt/b-catenin pathway in liver began to be recognized only in late 1990’s. The earlier studies focused on the altered immunohistochemical expression of b-catenin and E-cadherin in hepatocellular cancer (HCC). Other groups initiated studies to examine the mechanism of such an increase that led to recognition of mutations in the Ctnnb1 (b-catenin gene) as well as in other components of the multiprotein-degradation machinery. This also led to identification of aberrant Wnt/b-catenin signaling in pediatric liver tumors and hepatic adenomas. Studies were also focused in liver growth and development, to better understand the regulation of this pathway in liver physiology and pathology.

Role in Liver Development Loss of b-catenin led to embryonic lethality due to defects in gastrulation [27]. Due to availability of “floxed” b-catenin mice, it can be conditionally knocked out utilizing tissue-specific cre recombinase transgenic mice, which has yielded important information on multiple tissues [30, 150]. In 2003, the first

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study reported identification of the role of Wnt/bcatenin signaling in liver development [151]. Utilizing in vitro organ cultures and a comprehensive ontogenic analysis, the role of b-catenin was demonstrated in early liver development [118, 151]. Extremely tight regulation of Wnt/b-catenin signaling is essential at multiple steps during hepatic development [152]. In Xenopus, the impact of Wnt/b-catenin signaling on liver development can be seen before gastrulation, during the maternal phase. Maternal Wnt/b-catenin, which acts on the dorsal side of the embryo during the preblastula stage, can induce anterior endomesoderm (AE), a subset of endoderm cells fated to form the liver [25]. In Xenopus, b-catenin expression after gastrulation is necessary for intestinal formation in the posterior endoderm, while repression in the anterior endoderm is correlated with liver and pancreas development. Repressing b-catenin in the posterior endoderm causes organ buds expressing liver markers to form [153]. Other studies have found regulated expression of Wnt inhibitors during the early stages of hepatic development. Sfrp5, an antagonist of Wnt, is expressed in the ventral foregut endoderm that gives rise to the liver at mouse E8.5 [154], resembling the expression pattern of Hex. The expression of this inhibitor may function to modulate Wnt activity by delineating borders between organs in the developing gut [155]. Another study in zebrafish also demonstrates modulation of b-catenin expression during early liver development. A conditional mutant of prometheus, a homologue of Wnt2bb, which is expressed in the mesoderm directly adjacent to the developing liver, causes a severe but transient defect in liver formation. Further analysis revealed that expression of genes such as Hex and Prox1, which are essential in hepatoblast formation, is impaired in these mutants [156]. While in this study Wnt/b-catenin axis appears to be positive regulator of hepatic specification of foregut endoderm, it is likely that b-catenin is actually activated in the hepatic induction phase, which might partially overlap with the specification stage, especially in zebrafish due to shorter development period. Thus, while an initial repression of Wnt signaling is necessary for hepatic specification, it might immediately be followed by b-catenin activation. This is in agreement with our earlier studies that clearly demonstrate b-catenin activation during early stages of liver development once liver is specified in mice. b-Catenin gene and protein expression peaks at E1014 in mouse embryonic liver and during this time b-catenin is localized in the nucleus, cytoplasm, and

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membrane in different epithelial cells and coincided well with ongoing cell proliferation [118]. Mouse embryonic livers from E9.5-10 stages cultured in the presence of a b-catenin antisense oligonucleotides, showed decreased proliferation and a simultaneous increase in apoptosis, two processes vital to hepatic morphogenesis that follows hepatic specification and induction [151]. This correlated well with a subsequent study that found overexpression of b-catenin in developing chicken livers leads to a threefold increase in liver size, which is due at least in part to an expanded hepatoblast population [157]. In the same study, blocking b-catenin expression through overexpression of pathway inhibitors resulted in decreased liver size and altered liver shape. The effect on cell proliferation noted in both the cases may be due to cell cycle mediators such as cyclin-D1, which is a known downstream target of b-catenin. Subsequent decreases in b-catenin gene expression and increased protein degradation coincide with a dramatic decrease in total b-catenin protein expression after E16, at which time it is also localized to the membrane of maturing hepatoblasts and hepatocytes, although some nuclear localization is also observed. The later stages represent hepatoblast maturation to hepatocytes that begin to express genes associated with hepatocyte differentiation such as transferrin, cyochrome P450s, coagulation factors, haptoglobin, and many others [158]. All these phenotypes were also visible in b-catenin-conditional null mice utilizing floxed b-catenin and Foxa3-Cre mice [159]. Interestingly, an earlier study employing GSK3b gene knockouts, demonstrated a phenotype of increased liver cell death and liver degeneration that resulted in embryonic lethality. Whether this effect was due to untimely b-catenin stabilization or due to the fact that GSK3b is at the crossroads with several other signaling pathways critical to liver biology, such as PI3 kinase, insulin, and others, remains to be investigated further [160]. The concept of premature b-catenin stabilization on liver growth and survival during development is also supported by a more recent in vivo study, which utilizes APC deletion during liver development. This study shows a dramatic increase in cell death and a counterintuitive decrease in cell proliferation, however, this was associated with untimely differentiation of hepatoblasts into biliary cells [161]. This clearly supports a highly temporal expression, activation, and function of Wnt/b-catenin signaling during the process of normal liver development. Role of b-catenin in bile duct differentiation of hepatoblasts is intriguing. Antisense against b-catenin

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in embryonic liver cultures led to absent bile duct differentiation and the addition of Wnt3a to the embryonic liver cultures induced a biliary phenotype [162]. These observations were supported by in vivo studies that showed lack of biliary differentiation in b-catenin conditional null livers and enhanced biliary differentiation in APC-null (increased b-catenin) livers, during prenatal development [159, 161]. Role of b-catenin in hepatocyte maturation is also important and was also initially observed in antisensemediated b-catenin knockdown in embryonic liver cultures. This phenotype was confirmed by continued expression of stem cell markers in hepatocytes [151]. Using another in vitro model of matrigel induced hepatocyte differentiation, a total increase in b-catenin protein at the hepatocyte membrane was also reported [163]. The strongest evidence of the role of b-catenin in hepatocyte maturation was the in vivo study that clearly demonstrates lack of b-catenin to result in dramatic decreases in nuclear enriched transcription factors such as CEBPa and HNF4a, with significant impact on hepatocyte maturation and fetal viability [159]. Thus, a clear bimodal expression (repression followed by activation) and dual role (proliferation/survival and differentiation) of Wnt signaling is apparent in liver development in more than one species [153, 159, 164]. A major undetermined aspect of Wnt/bcatenin signaling during liver development remains the obscurity of upstream effectors such as Wnt/Fz genes and related proteins that are dictating the temporal expression and activation of b-catenin. Some studies are beginning to explore the upstream effectors and their regulation in liver development. If early activation of b-catenin is indeed observed in earliest phases of hepatic morphogenesis in zebrafish, then Wnt2bb might be one of the earliest upstream effectors [156]. Recently Matsumoto et al. demonstrated Wnt9a expression in endothelial and stellate cells of the embryonic sinusoidal wall in developing liver [165]. They also provided evidence that Wnt9a promotes in vivo stabilization of b-catenin through binding with frizzled 4, 7, and 9. Based on the interaction of HGF/ Met and b-catenin and the role of HGF/Met in liver development, it seems that HGF/Met/b-catenin signaling might have important implications [22, 166–168]. Expression of FGF-10 in the mouse liver correlates with peak b-catenin activation; moreover, the release of FGF-10 from stellate cells stimulates b-catenin expression in hepatoblasts [169]. Our laboratory previously reported that FGF-2, FGF-4, and FGF-8 could

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impact b-catenin activation in embryonic liver cultures [170]. Thus, while b-catenin is undeniably necessary during hepatic morphogenesis, there might be several upstream effectors that might, in a concerted effort, precisely regulate the amount, timing, and extent of b-catenin activation for normal hepatic development.

Role in Liver Growth, Metabolism, and Homeostasis Due to aberrant activation of Wnt/b-catenin signaling in HCC, it was essential to understand the normal expression and regulation of various components of this pathway in the liver. A recent study has identified expression of many Wnt and Fz genes in an adult liver and within various cell types in the liver [171]. The liver continues to grow during neonatal stages. In fact, in mice an early postnatal hepatic growth spurt is known for quite some time. Wnt/b-catenin signaling was recently identified to be active during these stages and correlated well with ongoing hepatocyte proliferation [172]. Several additional models have been employed, which demonstrate a positive role of b-catenin in liver growth. A transgenic mice overexpressing a stable mutant of b-catenin generated under the transcriptional control of calbindin-D9K (CaBP9K) promoter and liver-specific enhancer of the aldolase B gene displayed 3–4 times larger livers due to increased cell proliferation [173]. Interestingly they did not detect any changes in any of the conventional target genes of the pathway such as c-myc and cyclin-D1. Subsequent analysis of transgenic livers and subtractive hybridization led to the identification genes involved in glutamine metabolism as targets of the Wnt/b-catenin pathway. However, no signs of neoplastic transformation were reported in these animals, although, APC-conditional null mice that exhibit b-catenin stabilization leads to development of robust HCCs [174]. Other transgenic mice have shown similar hepatic growth advantage [175, 176] and also enabled identification of new targets such as EGFR [176]. A more useful approach to study the role of b-catenin in liver biology has been the generation and characterization of b-catenin conditional null mice [30]. After the advent of floxed b-catenin mice, at least two independent groups including ours have reported to have utilized these mice [177–179]. These models have

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identified a decreased liver weight/body weight ratio due to compromise chiefly in proliferation, along with mild effects on hepatocyte survival. This model enabled verification of the regulatory role of b-catenin in ammonia metabolism as well as also reaffirmed its role in xenobiotic metabolism through regulation of various cytochrome P450s and GSTs. In fact, three genes involved in glutamine metabolism encoding glutamine synthetase, ornithine aminotransferase, and the glutamate transporter GLT-1 are all the targets of Wnt/bcatenin signaling in transgenic and knockout mice [178–180]. Likewise, overexpression of glutamine synthetase was noted selectively in mouse liver tumors that contained b-catenin mutations [181]. Also several CYP isoenzymes are upregulated in liver tumors harboring b-catenin mutations [182]. Likewise, hepatocyte specific knockout of b-catenin in mice leads to a loss of expression of several CYPs, especially 2E1 and 1A2 [178, 179]. These findings have also been complemented by the overall concept of APC/b-catenin, playing a major role in the zonation within the liver. While it has long been known that the liver is structurally divided into different zones, which express specific proteins and partake in differing metabolic functions, not much is known about its molecular basis. Benhamouche et al. recently reported that b-catenin and APC distribution within various hepatic zones might be critical in the zonation process [183]. They identified high APC expression in periportal region with no b-catenin activation and conversely, identified absence of APC expression in centrizonal area where b-catenin activation is high, which maintains high expression of target genes such as GS and CYP2E1. Thus, b-catenin plays major roles in regulating hepatic homeostasis.

Liver Regeneration Due to the importance of aberrant Wnt/b-catenin signaling in liver cancer, it is imperative to understand the regulation of this pathway in a “regulated” growth environment. One such widely accepted system in the liver is the two-third partial hepatectomy model in mice and rats [184, 185]. Wnt/b-catenin pathway has been comprehensively studied in these models [186]. There was a significant increase in total b-catenin protein within first few minutes of hepatectomy that was mediated by posttranslational mechanisms. Interestingly, while the increase was transient due to activation of

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b-catenin degradation complex including axin and APC, b-catenin persisted in the nuclei of the hepatocytes until around 24 h. Thus, there are crucial modulators of the pathway that come into play to monitor b-catenin levels in a regulated growth milieu. This is not surprising, owing to the abundance of b-catenin, a potent “oncoprotein” at the membrane of normal hepatocyte and it will be only devastating to not have a stringent monitor to limit unnecessary or sustained b-catenin activation. Another inference drawn from this study was that Wnt/b-catenin pathway might be one of the earliest pathways to become activated following hepatectomy that might initiate a cascade of events including but not limited to inducing gene expression of c-myc, cyclin-D1, and uPAR or yet undiscovered targets. Met–b-catenin complex in hepatocytes might also be one of the contributing sources of nuclear b-catenin as elevated tyrosine phosphorylation of Met and activation of HGF is also observed during early liver regeneration [22, 187, 188]. More recent studies with b-catenin conditional-null mice from two groups reinforce the importance of b-catenin in optimal liver regeneration after partial hepatectomy. Following hepatectomy, absence of b-catenin in hepatocytes led to the delay in peak hepatocyte proliferation by 24  h, which coexisted with absent increases in the expression of cyclins D, A and E [177, 179]. While this study demonstrates the role of b-catenin in optimal regeneration, it does highlight the level of redundancy in signal transduction pathways that ensures regeneration [184]. Another study using phospho-morpholino-antisense driven b-catenin knockdown after hepatectomy in rats, led to a more persistent decrease in hepatocyte proliferation and recovery of liver mass [189]. A major difference between studies in rats and mice was a cell-autonomous suppression by b-catenin-antisense injection in rats vs. absence of b-catenin exclusively in hepatocytes in mice, which could possibly explain a more dramatic suppression of regeneration in the rat study.

Stem Cells in Adult Liver or Oval Cells Based on the role of Wnt/b-catenin signaling in development and in stem cell biology in general, the status of Wnt/b-catenin signaling has also been assessed in adult liver stem cells or “oval cells.” These cells, although oblivious in normal adult liver, can be forced to expand

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in animal models through specific protocols. The protocols such as administration of 2-acetylaminofluorine before partial-hepatectomy in rats or administration of diet containing 3,5-diethoxycarbonyl-1,4-dihydrocollidine diet to mice, will lead to appearance and expansion of transient amplifying progenitors or oval cells. Two independent studies demonstrated activation of b-catenin in oval cells secondary to autocrine or paracrine Wnt/ frizzled signaling [190, 191]. This led to proliferation of oval cells in these models. Presence of active b-catenin in expanding oval cells might have two opposing consequences. Because of its relevance to stem cell renewal, Wnt signaling might have positive implications in oval cell biology, regenerative medicine, hepatic tissue engineering, and cell therapeutics. On the other hand based on the building evidence of a subset of HCC might have stem cell origin, inhibition of Wnt signaling might have therapeutic implications as well [192].

WNT/b-Catenin Signaling in Liver Benign Liver Neoplasms The major benign hepatic tumors that have displayed activation of b-catenin consist of focal nodular hyperplasia (FNH) and hepatocellular adenoma. Recently, transcriptome analysis of FNH identified activation of Wnt/b-catenin pathway without any mutations in b-catenin gene [193]. While the significance of these findings is unclear, especially since these tumors are thought to be a result of vascular disturbances, these observations might be a result of alternate mechanisms of b-catenin activation such as growth factor-dependent activation [22, 194]. More recently, immunostaining for GS, a target of Wnt/b-catenin signaling in the liver, has been proposed to identify FNH [195]. Significant subsets of hepatic adenomas display inactivating mutations in HNF1a or TCF1 gene. Biallelic inactivating mutations in TCF1 genes were identified in around 50% of HA. The tumors in this scenario display marked steatosis and excess glycogen accumulation. HA with HNF1a inactivation displays an extremely low risk of malignant transformation. However, another subset of HA’s, display Wnt/bcatenin activation secondary to mutations in CTNNB1 gene. These adenomas can range from 15 to 46%, but the numbers might be closer to the lower percentage

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when only CTNNB1 mutations as a mechanism of b-catenin activation are taken into account [196, 197]. These tumors show frequent cytological abnormalities and pseudo-glandular formation and have been shown to occur at higher frequency in males. Most importantly, these HAs have been shown to possess a higher propensity for malignant transformation [197]. More recently, GS immunohistochemistry has been utilized to identify this group of tumors [198].

Hepatoblastomas This tumor is the most common malignant hepatic tumor during early childhood. These embryonal tumors are frequently sporadic; however, the incidence is the highest in patients suffering from familial adenomatous polyposis coli [199]. This led to the identification of APC mutations as the molecular etiology for hepatoblastomas in familial cases [200]. Increased frequency of diverse APC mutations (57%) were then reported in sporadic form of the disease as well [201]. Since APC regulates b-catenin levels, next set of analyzes focused on and revealed abnormal b-catenin accumulation and associated amino-terminal mutations (exon 3) in around 50% of all sporadic hepatoblastomas [202]. A number of reports that followed illustrated nuclear and cytoplasmic localization of b-catenin in 90–100% of all hepatoblastomas [203–205]. Predominantly, inframe mutations in the b-catenin gene in the form of deletions or missense were observed in 70–90% of such cases [203, 205]. Mutations in AXIN1 were also identified in less than 10% of these tumors [206]. Hepatoblastomas as a component of syndromes such as Beckwith-Wiedemann syndrome have also revealed abnormal Wnt/b-catenin activation [205, 207]. Thus, there is compelling data that shows Wnt/b-catenin aberrations as an obligatory event in the etiopathogenesis of hepatoblastomas. Use of b-catenin nuclear reactivity as a prognostic indicator for the disease was suggested, but is not a widespread practice [208]. Interesting analysis of new members of the Wnt pathway such has as the pathway inhibitor Dkk1 has shown overexpression in hepatoblastomas and is believed to be due to negative feedback related to uncontrolled Wnt signaling [209]. It also appears that not all hepatoblastomas with nuclear b-catenin might have deleterious activation of b-catenin. Based on studies in

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development, it should become apparent in future, if there are additional cofactors that might be essential for target gene specificity and hence dictate if nuclear b-catenin is promoting differentiation or proliferation, the latter being more relevant to tumor behavior and progression [159].

Hepatocellular Cancer A disease of extremely poor prognosis, HCC remains among the leading cause of mortality and morbidity around the globe. The disease bears a strong etiological association with viral hepatitis, hemochromatosis, chemical carcinogens, and toxins (mycotoxins) (reviewed in [210]). Preexisting cirrhosis due to any number of factors such as metabolic disease, inflammation, or infection predisposes to HCC. Inappropriate Wnt/b-catenin activation has been implicated in many cancers and is one of the important aberrant pathways identified in HCC in animal and patients [3, 211, 212]. Abnormal localization of cadherins and catenins in liver cancer was first shown by immunohistochemistry [213]. A more comprehensive study identified anomalous b-catenin expression as well as mutations in the Ctnnb1in around 25% of all HCC cases and up to 50% of all hepatic tumors in transgenic lines such as c-myc or H-ras [214]. Several subsequent studies corroborated these observations, although the mutations in b-catenin gene ranged from 12 to 34%, abnormal b-catenin redistribution has been reported in up to 90% of HCCs suggesting additional mechanisms [215–218]. Table  25.2 shows common mutations in Ctnnb1 in some HCC patients. Additional mechanisms of b-catenin activation have also been described. AXIN1 and AXIN2 mutations were also detected in around 5–10% and 3% of HCCs respectively [206, 219]. Reports analyzing GSK3b studies are conflicting although elevated levels of inactive GSK3b are observed in HCCs harboring b-catenin accumulation [220–222]. Activation of Frizzled-7/ Wnt-3a axis has been reported in significant numbers of HCCs analyzed [223, 224]. Epigenetic inactivation of inhibitors of Wnt signaling such as soluble frizzled related proteins (sFRP) has also been reported in HCC [225]. More recently, fibrolamella HCC showed significant tyrosine-phosphorylated b-catenin suggesting receptor tyrosine kinase activation in these tumors (in

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Table 25.2  List of studies showing spectra of mutations in Ctnnb1 gene Study Cases with mutations Mutation sites (includ. Del*) S33 S37 S45 T41 [221] 15/45 5 [273] 16/38 2 2 3 1 [206] 14/73 1 7 1 [229] 5/62 1 2 [227] 9/22 1 3 2 1 [217] 12/35 1 2 2 [216] 21/119 3 2 8 4 [214] 6/26 2 1 1 [214] 9/32 2 2 1 2

Additional information Others 10 10  5  2  3  7  5  2  2

No GSK3b mutations Multiple mutations in two patients one insertion between S33 and G34 Aflatoxin study Multiple mutations in one patient Multiple mutations in two patients Multiple mutations in a patient One patient had deletion Two patients had mutations at D32

*Deletions usually involved one of the key sites-S33, S37, S45, or T41

press). There have been reports identifying differences in molecular targets of the pathway secondary to modes of b-catenin activation [226]. Aberrant immunohistochemical findings for b-catenin in HCC include nuclear and/or cytoplasmic with or without membranous localization and represent heterogeneity in mechanisms inducing this redistribution. Similarly, variations in frequency of mutations might be reflective of differences in geographical, dietary, and other factors, influencing the molecular pathogenesis of this disease. One study detected an inverse correlation between b-catenin mutations and loss of heterozygosity in the genome suggesting chromosomal instability (involving tumor suppressor genes) and mutations in Ctnnb1 representing alternative modes of tumor progression [115]. Interestingly, a much higher frequency of Ctnnb1 mutations are observed in HCC associated with hepatitis C virus (HCV) infection. More than 40% of HCV associated HCCs demonstrate stabilizing mutations in b-catenin gene (mostly at Ser45) as well as nuclear accumulation of its protein [227]. HBV related HCC had overall lesser frequent b-catenin mutations [228]. Also, although mutations in its gene were infrequent, aflatoxin associated HCC showed increased accumulation of b-catenin in around 45% of tumors [229]. Analysis has also extended to identify distinct molecular signatures of HCC arising in cirrhotic vs. noncirrhotic livers and although preliminary, this analysis suggests unique pathogenetic events in the two subsets. While HCC in noncirrhotic livers demonstrates more frequent Wnt/b-catenin involvement along with other pathways, HCC arising in cirrhosis showed mainly p53 alterations [210]. Along similar lines another study reported more frequent Wnt/b-catenin

aberrations in HCV associated HCCs as compared to alcoholism associated HCC that more frequently involved RB1 and p53 pathways [230]. Prognostic implications of aberrant b-catenin localization have also been addressed in patients, however, the reports are once again conflicting. There are examples of b-catenin activation associated with poorer prognosis [217, 227] and also with noninvasive form of tumor and better prognosis [228, 231]. Another report shows a non-nuclear type of b-catenin overexpression related to poor cell differentiation, larger tumor size and significantly shorter disease-free survival lengths [218]. Another study found a significant relation of nuclear cellular retinol-binding protein-1, nuclear b-catenin, low Ki-67 positivity and favorable prognosis, and 2-year survival [232]. Thus, this issue remains unresolved and would need to be addressed in future studies taking into account, geography, etiology, and patient numbers before a consensus is reached. Wnt/b-catenin pathway in HCC in animal models is also unsettling. The studies involving b-catenin transgenic mice that overexpress truncated b-catenin in liver do not show any evidence of spontaneous carcinogenesis [173]. Another study overexpressing nontruncated human b-catenin gene under transcriptional control of albumin promoter/enhancer also yielded no tumors in the liver [176]. Similar lack of tumorigenesis was also observed following adenoviral-mediated overexpression of dominant stable b-catenin mutant in liver [175]. Interestingly APC-null mice showed significant HCC through activation of b-catenin [174]. Overall, it might be b-catenin mutations might be insufficient on their own and require cooperation with other pathways for hepatocarcinogenesis. Devising newer liver models including transgenic lines as well

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as newer chemical carcinogenesis strategies such as tumors induced by 2-amino-3, 4-dimethylimidazo [4,5f] quinoline, that selectively induce anomalies in the Wnt/b-catenin pathway would be really useful to better understand the role and regulation of this pathway as well as have significant therapeutic implications [233].

Bile Duct Tumors The most common tumor that arises from the biliary tree is the cholangiocarcinoma that can either originate from the intrahepatic portion intrahepatic cholangiocarcinoma (ICC) or the hilum (hilar cholangiocarcinoma) (reviewed in [234]). The molecular pathogenesis is not yet characterized and along with several other oncogenic pathways analyzed there have been reports implicating aberrant Wnt/b-catenin signaling in a subset of tumors. One thing that is worth mentioning here is that there is a definite role of this pathway in biliary development and survival [151, 162]. Also there is a significant crosstalk of this pathway with Notch/jagged pathway that is associated with developmental defects in biliary tree [235–239]. In cholangiocarcinoma, reduced expression of b-catenin and E-cadherin at the membrane is observed as compared to the surrounding noncancerous ducts [240]. More importantly, nuclear localization of b-catenin is seen in a subset of tumors based on histology and location of the tumor (reviewed in [241]). For most ICCs, aberrant nuclear localization is observed in around 15% and a decrease in membranous localization is related to poorer histological differentiation [242]. This study failed to identify any mutations in exon 3 of b-catenin gene although it did not analyze mutations in any other components of the Wnt pathway. A larger study detected exon 3 mutations in 7.5% of biliary tract cancer and in 57% of gall bladder adenomas [243]. Higher frequency of mutations is seen in ampullary and gall bladder carcinomas than the bile duct cancers. A higher correlation of Ctnnb1 mutation and papillary adenocarcinoma is also observed. Intraductal papillary neoplasms also show anomalous nuclear localization of b-catenin in around 25% of patients without any b-catenin gene mutation in the GSK3b-phosphorylation region [244]. Again, other components of the pathway were not analyzed for mutations in this study. Thus, while we can incriminate the Wnt/b-catenin pathway in a subset of biliary tract

S. P. S. Monga

neoplasms, more studies are needed to comprehend the mechanism of its observed deregulation.

Miscelleneous Pathologies The Wnt/b-catenin pathway is also gaining importance in molecular pathogenesis of several nonmalignant conditions. Autoimmune and inflammatory conditions such as osteoarthritis, rheumatoid arthritis, idiopathic pulmonary fibrosis and renal fibrosis are a few examples that show activation of this pathway [245–248]. In liver, increased expression of b-catenin protein as well as a Wnt pathway gene (apoptosis-related protein 3) has been identified in HCV associated cirrhosis and not autoimmune hepatitis cirrhosis [249]. Although we have previously discussed activation of this pathway in HCC and cirrhosis, we should point out that the frequency of activation of Wnt/b-catenin in cirrhosis associated HCCs and non-HCV hepatitis associated HCC is generally lower [210, 230]. We must reiterate that these are recent studies and would require more corroboration. cDNA array analysis was also utilized to examine the alterations in gene expression in primary biliary cirrhosis (PBC) as compared to disease free livers and primary sclerosis cholangitis (PSC) associated cirrhosis [250]. This analysis revealed over expression of numerous genes of the Wnt pathway, prominently Wnt5A, Wnt13, FRITZ, and b-catenin in the PBC samples again implicating the Wnt pathway in pathogenesis of PBC by probably contributing to the accompanying inflammation, fibrosis, and regeneration.

Therapeutic Implications Activation of Wnt/b-catenin pathway: Based on several observations demonstrating role of Wnt/b-catenin signaling in regeneration, development, stem cells, and growth, it might be beneficial to be able to induce or modulate this signaling pathway for intervention in hepatic insufficiency. This might have positive implications in the in vitro systems for applications in stem cell transdifferentiation to hepatocytes; cell transplantation to improve homing and efficiency; and in artificial liver devices for improved and prolonged functionality. Regulated activation of b-catenin might

25  The WNT/b-Catenin Pathway

have positive implications in treatment of acute liver failure, which would clearly need to be investigated extensively in preclinical models. Inhibition of Wnt/b-catenin signaling: Due to aberrant activation of Wnt/b-catenin pathway in multiple cancers including HCC, it is an accepted therapeutic target [251]. The most crucial component of this pathway especially in liver is perhaps b-catenin, and most of the relevant pathologies are effect of either b-catenin loss from the membrane or its cytoplasmic stabilization and nuclear translocation resulting in an increase in target gene expression. Several proof-of-principle

Summary

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embryogenesis (bile duct differentiation), stem cell biology, carcinogenesis, angiogenesis, and now in metabolism as well. In the canonical pathway, in the absence of a Wnt signal, the free monomeric form of b-catenin remains cytoplasmic and is targeted for debradation after phosphorylation and ubiquitination. Upon formation of a ternary complex between one Wnt ligand, Fz receptor, and LRP, Disheveled potentiates b-catenin stabilization and its nuclear translocation to bind to TCF/ LEF transcription factor. Among the many genes targeted by the Wnt/bcatenin pathway, several belongs to the pathway itself and among the others it is worth to remember glutamine synthetase, GRP49, c-myc, cyclin D1, and VEGF. Two noncanonical pathways have been recently recognized: a Wnt/calcium pathway and the planar cell polarity pathway involving JNK. b-Catenin plays also a crucial role as a bridge between the cytoplasmic domain of the cadherins and the cytoskeleton. Normally in the liver, APC is expressed in periportal regions with no b-catenin activation and conversely, b-catenin is active in the centrizonal regions with absence of APC expression. Wnt/b-catenin pathway is important for liver regeneration and has been found to be frequently active in hepatic tumors.

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preclinical studies have clearly demonstrated an important benefit of therapeutic inhibition of b-catenin as a means of treatment for HCC [252]. Various cox-2 inhibitors such as rofecoxib have shown efficacy in decreasing b-catenin levels along with shrinkage of tumors [253]. R-Etodolac, an enantiomer of a cox-2 inhibitor that lacks an inhibitory effect on cox-2 has also shown anti-b-catenin effect [254]. This might be more meaningful in light of and the unwanted side effects associated with cox-2 inhibitors. Imatinib (Gleevec) has shown to decrease tyrosine-phosphorylated b-catenin levels only [255]. Similarly, recently approved Sorafenib for HCC treatment might have more dramatic response in HCC showing receptor tyrosine kinase activation, which also leads to the tyrosine phosphorylation of b-catenin [256]. Another group of agents including Exisulind and analogs that are inhibitors of cyclic GMP phosphodiesterases (PDE) have been shown to activate protein kinase G (PKG) that in turn decrease b-catenin levels via a novel GSK3b-independent processing mechanism [257]. Another important strategy will be to identify novel tissue-­specific targets of the pathway that are contributing to the disease to develop therapies against such molecules. But, successful inhibition of Wnt/b-catenin signaling would be one key therapeutic strategy in treatment or chemoprophylaxis of HCC [258].

Multiple Choice Questions 1. Cross-talks have now been established between the Wnt pathway and others signaling pathways except: (a) Jagged/Notch (b) Toll-like receptors (c) HGF/Met (d) EGF (e) TGF 2. Wnts are (a) Transcription factors (b) Transmembrane receptors (c) Extracellular secreted glycoproteins (d) Scaffold proteins (e) Channels 3. What is the name of the factor preventing the formation of Wnt-induced Fz-LRP complex (a) Disheveled

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(b) TCF (c) Axin (d) APC (e) Dickkopf 4. Regarding Wnt/b-catenin pathway and liver regenration, what is correct: (a) b-catenin protein increases in the minutes following partial hepatectomy (b) b-catenin is redistributed to the plasma membrane (c) b-catenin pathway is important in the late phase of liver regeneration (d) In b-catenin conditional-null mice absence of b-catenin in hepatocytes led to increased expression of cyclins (e) b-catenin is massively ubiquinated 5. Which hepatic tumor has been linked to Wnt/bcatenin pathway: (a) Focal nodular hyperplasia (b) Hepatic adenoma (c) Hepatoblastoma (d) Hepatocellular carcinoma (e) All the above Recommended Other Reading (Or Website): Website – “The Wnt Homepage” at http://www.stanford. edu/~rnusse/wntwindow.html. (This site gives an in depth and updated information about the Wnt pathway. An absolute must for anybody working in the Wnt field.)

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Sonic Hedgehog Pathway

26

Alessia Omenetti and Anna Mae Diehl

Introduction The Hedgehog Pathway The Hedgehog (Hh) Pathway, originally identified in Drosophila, [1–4] is a highly conserved signaling pathway which orchestrates multiple, disparate aspects of embryogenesis, development and tissue remodeling in a wide spectrum of systems [5–8]. This is usually accomplished by autocrine/paracrine signaling and aims to control the size and localization of Hh-responsive cell populations in response to local/long distance signals [7, 9]. Hh pathway activation typically enhances the growth and viability of Hh-responsive cells, whereas abrogating Hh signal transduction usually triggers apoptosis in such cells, unless other locally available differentiating factors expedite cellular differentiation to a more mature phenotype that no longer requires Hh viability signals [5, 9]. Thus, up-/downregulation of the Hh pathway provides a selective growth advantage for cell types that are capable of responding to Hh ligands, when compared to neighboring cells that lack Hh receptors. This leads to ­expansion/ contraction, respectively, of Hh-responsive cells, thereby, orchestrating the cellular composition of several tissues [5–7, 9]. In certain conditions, Hh producing cells (which may or may not be Hh-responsive themselves) release Hh ligands into the extracellular environment. Hh

A. M. Diehl () Department of Medicine, Division of Gastroenterology, Duke University Medical Center, GSRBI, 595 LaSalle Street, Suite 1073, Box 3256, Durham, NC 27710, USA e-mail: [email protected]

ligands (Sonic, Shh; Indian, Ihh; Desert, Dhh) are ­soluble, lipid-modified morphogens [10–14] that may be secreted in two different forms: a short range acting (poorly diffusible) type, and a second form for longrange transport, “packed” in membranous structures [7, 14, 15]. Hh proteins are able to interact with Patched (Ptc: Ptch1 in vertebrates), a membrane-spanning receptor on the surface of Hh-responsive cells [16]. In the absence of Hh ligands (Fig.  26.1), Ptc keeps the ­co-receptor Smoothened (Smo) in its inactive form and silences the Smo-dependent down-stream intracellular signaling [7, 14]. Hence, when Smo-signaling is inhibited by “free”-Ptc, Hh-regulated transcription factors (which typically reside in the cytosol) undergo multiple phosphorylation by glycogen synthase kinase 3 (GSK3), protein kinase A (PKA), and casein kinase (CSK); the phosphorylated (inactive) forms become target for proteasome degradation, and their nuclear translocation is prevented [17, 18]. In contrast, when the extracellular microenvironment is enriched with soluble Hh ligands (Fig.  26.2), ligand-receptor interaction de-represses Smo. Activation of Smo, in turn, inhibits Hh transcription factor phosphorylation, leading to an intracellular signaling cascade that ultimately drives the activation and nuclear translocation of Glioblastoma (Gli) family zinc-finger transcription factors [17, 18]. The latter consist of three distinct Gli proteins (Gli1, Gli2, and Gli3) in vertebrates, [19] while a single mediator (Cubitus interruptus, Ci) is known to mediate Hh signaling in Drosophila [2, 19]. The binding of Gli proteins to their cognate cis-acting elements regulates the expression of Hh target genes. The latter include several components of the Hh pathway itself, such as Ptc, Smo, and Glis. Gli1 and Gli2 are mostly responsible for providing prolonged cellular responses to Hh ligands, while Gli3 primarily acts as signaling repressor [6, 7, 14]. Thus Hh activity is auto-regulated by complex positive

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_26, © Springer-Verlag Berlin Heidelberg 2010

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Cilium

Cilium Active Smo

Inactive Smo

Hedgehog ligand

“free”-Ptc

Inactive Ptc

CSK PKA GSK3

Gli

CSK PKA GSK3

Gli

Gli

p Proteasome

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rGli

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Fig.  26.1  Hedgehog Pathway OFF In absence of Hedgehog ligands, Patched (Ptc), a membrane-spanning receptor on the surface of Hh-responsive cells, keeps the co-receptor Smoothened (Smo) in its inactive form. “Free”-Ptc silences the Smo-dependent down-stream intracellular signaling, and Hh-regulated transcription factors undergo multiple phosphorylation by glycogen synthase kinase 3 (GSK3), protein kinase A (PKA) and casein kinase (CSK). The phosphorylated (inactive) forms become target for proteasome degradation, and their nuclear translocation is prevented

Fig.  26.2  Hedgehog Pathway ON When the extracellular microenviroment becomes enriched with soluble Hedgehog ligands, they bind to the receptor Patched (Ptc) and this interaction de-represses Smoothened (Smo). Activation of Smo, in turn, inhibits Hedgehog transcription factor phosphorylation mediated by glycogen synthase kinase 3 (GSK3), protein kinase A (PKA) and casein kinase (CSK), leading to an intracellular signaling cascade that ultimately drives the activation and nuclear translocation of Glioblastoma (Gli) family zinc-finger transcription factors

and negative feedback mechanisms that are tightly conserved across species [2, 6, 7, 14]. Despite the conservation of the Hh pathway between invertebrates and vertebrates, Hh pathway regulation seems to diverge and differentiate at some point [14, 20]. For example, unlike in Drosophila, Hh-responsive vertebrate cells have a functional active primary cilium, an organelle that protrudes from the cell surface and that acts as Hh-signaling center [21, 22]. Moreover, only vertebrates have an additional trans-membrane protein, Hh-interacting protein (Hhip, also a Hh target gene), that competes with Ptc for binding to Hh soluble ligands [23, 24]. Thus, in vertebrates, when levels of Ptc exceed those of the Hh ligands, or when Hhip sequesters Hh ligands and subtracts activating signals to Ptc, the Hh pathway is turned off.

Hedgehog-Producing and HedgehogResponsive Cell Types in Adult Liver Repair Adult hepatic damage evokes an intricate woundhealing response aimed to reconstitute the normal structure and function of injured livers. As in many other tissues, this complex repair process involves the post-natal reactivation of mechanisms that regulate tissue construction during development, including Hh signaling. Despite the lack of direct evidence for Hh involvement in fetal liver development, several types of cells in adult livers are capable of producing and responding to Hh ligands. Specifically, the resident hepatic cell populations that are most engaged in

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liver remodeling (e.g., liver myofibroblasts, hepatic progenitors, hepatic stellate cells, immature cholangiocytes, endothelial cells, and T lymphocytes) [25–36] activate Hh pathway signaling both in human diseases and in related animal models. The proximity of these cells in damaged livers suggests that Hh ­signaling orchestrates complex cell-to-cell cross talk. This variably engages different cell populations depending on the identity of injured-related factors that are released into the hepatic microenvironment, leading to remodeling and re-construction of the injured liver. During liver injury, metabolic and/or inflammatory stress causes surviving/dying damaged cells and infiltrating immune cells to produce injury-related proinflammatory cytokines (e.g., Interferon (IFN)-g and Tumor Necrosis Factor (TNF)-a) and growth ­factors (e.g., Platelet-Derived Growth Factor-BB (PDGF-BB), Transforming Growth Factor (TGF)-b1, Epidermal Growth Factor (EGF), Hepatocyte Growth Factor (HGF), and Insulin-like Growth Factor (IGF)) [37]. Some of these soluble factors (PDGF-BB, TGF-b1, EGF) have been demonstrated to stimulate Hh pathway activation [29, 30, 32, 35, 36]. For example, each of these factors stimulates cells that are able to produce  Hh ligands, to up-regulate their synthesis and release of Hh proteins. Because mature hepatocytes are not able to respond to Hh ligands, enrichment of the microenvironment with these factors provides a ­selective survival advantage for cell types that are Hh-responsive, leading to the outgrowth of these ­populations as long as injury persists. However, when the insult abates and injury subsides, the Hh pathway turns off, and other factors promote the differentiation of the  progeny of Hh-responsive cells toward one cell population or another. Because PDGF-BB, EGF, and IGF-1 activate AKT-dependent post-translational mechanisms that stabilize Gli transcription factors in Hh-responsive cells [38] (in addition to stimulating production of Hh ligands), Hh signaling may modulate the actions of multiple growth factors, and vice versa. Thus, variations in tissue remodeling during liver injury probably ultimately reflects differences in: (1) local cytokine/growth ­factor accumulation, (2) the dose and duration of Hh ligand exposure, (3) the balance between Hh-responsive/-not responsive cell types, and (4) the presence/absence of poorly understood factors that regulate cell differentiation when these injury-related signals wane.

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Hedgehog Pathway in Non-Malignant Liver Diseases Biliary Fibrosis Biliary injury elicits compensatory repair responses that attempt to reconstruct the damaged bile ducts. This process variably involves replication of surviving mature cholangiocytes, as well as replication/differentiation of immature ductular epithelial cells [39, 40]. Injured and reactive cholangiocytes release soluble factors that act in autocrine/paracrine fashion to induce compensatory ductular proliferation and accumulation of matrix-producing myofibroblasts in portal areas [41–48]. When the biliary insult becomes chronic, the protracted activation of these tissue repair mechanisms causes portal tracts to expand into the lobule. This eventuates in an extensive fibro-ductular reaction that bridges adjacent portal tracts and eventually culminates in biliary cirrhosis [39, 47]. PDGF-BB released from damaged cholangiocytes is one of the key biliary injury-related mediators because PDGF-BB is known to be a potent mitogen for HSC. Thus, cholangiocyte-derived PDGF-BB elicits the accumulation of myofibroblastic stellate cells in portal tracts [43–46]. Interestingly, PDGF-BB also induces production of Shh by myofibroblastic stellate cells [35, 36] and immature cholangiocytes [30, 35]. In both cell types, Shh acts as a viability factor by increasing proliferation and/or decreasing apoptosis [30, 35, 36]. Consistent with these findings, expression of Shh parallels that of PDGF-BB during experimental biliary fibrosis induced by bile duct ligation (BDL) in rodents. In the BDL model, when biliary injury is surgically relieved by biliary decompression via Roux-en-Y biliary-enteric anastomosis, Hh signaling gradually subsides [30]. Thus, during injury and repair of the biliary tree, Hh pathway activation follows the expected kinetics of the remodeling process: namely, Hh-responsive ductular cell populations expand and regress along with up- and down-­regulation of Shh and Hh-target genes in injured livers [30]. Further support for the concept that Hh pathway activation mediates biliary remodeling during cholestatic liver injury is provided by studies of mice with an overly-active Hh pathway (due to partial deficiency of Ptc) [31, 32, 49–51]. Such mice show aberrant responses to BDL, with an  exuberant ductular

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reaction, overly-expanded and fibrotic portal areas, and greater number of lobular necrotic foci [31, 32]. Evaluation of liver specimens from patients with Primary Biliary Cirrhosis (PBC) and control healthy subjects confirmed that Hh pathway activation occurs during human cholangiopathies [28, 31]. Immunemediated destruction of biliary epithelium in PBC drives a repair response that eventually leads to biliary-type cirrhosis and progressive ductopenia [39]. Inter­estingly, portal tracts of PBC livers harbor several cell types that express Hh ligands and/or Hh target genes [28, 31]. Further phenotypic evaluation of these cells demonstrated that myofibroblastic and immature ductular cell populations were particularly enriched with Hh-responsive cells, whereas more matureappearing ductular cells seemed to be the main source of Hh ligands [28]. These findings suggest that Hh signaling might regulate epithelial-mesenchymal cell cross talk that promotes fibroproliferative responses in PBC, [28, 31] as it does in experimental biliary fibrosis [30, 32]. In both types of chronic biliary injury, ­sustained Hh pathway activation seems to divert immature-Hh-responsive populations of ductular cells away from duct replacement, and to promote their differentiation  into myofibroblastic cells [30, 31]. This concept is strongly supported by recent experimental evidence that Hh pathway activation stimulates cultures of imma­ture ductular-type cells to undergo epithelial-to-­mesenchymal transition (EMT), and by immunohistochemical and QRT PCR evidence for cholangiocyte EMT in liver samples from patients with PBC, as well in rodents with experimental biliary fibrosis [31].

Nonalcoholic/Alcoholic Fatty Liver Disease Fatty acids are potent triggers for oxidative-, metabolic-, and endoplasmic reticulum- stress in hepatocytes. In order to limit the replication of defective/ damaged cells, chronically “stressed” cells are­ typically prevented from traversing the cell cycle. Therefore, although increased hepatocyte apoptosis occurs in nonalcoholic/alcoholic fatty liver disease, and this ­provides an ongoing stimulus for liver regeneration, the proliferation of mature hepatocytes is inhibited [52–56]. Consequently, restoration of hepatic

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architecture necessitates mobilization of progenitor populations, and steatotic liver damage is usually accompanied by the so-called “ductular reaction” [40, 57]. The ductular reaction is characterized by periportal accumulation of atypical bile ductules (derived from proliferation of pre-existing ductular cells and expansion of the progenitor compartment) intermingled with matrix-producing myofibroblasts [40, 47, 58]. The intensity of this ductular reaction is tightly associated with progressive fibrosis in human nonalcoholic steatohepatitis (NASH) [57]. It was recently shown that Hh pathway activation closely parallels the severity of the fibroductular response in experimental models of both nonalcoholic [25] and alcoholic fatty liver disease, [29] as well as in humans with alcoholic hepatitis [29]. The  latter finding suggests that Hh pathway activation  might play a role in remodeling/ reconstruction of fatty  liver damage, as it does in chronic biliary-type disease. Animal studies in wild-type mice fed with high-fat diets, [29] and in leptin-deficient (ob/ob) mice exposed to the hepatotoxin ethionine, [25] demonstrated that hepatic progenitor populations accumulate during experimental NASH, and that these populations are enriched with Hh-responsive cells. Furthermore, the relationship between mature hepatocyte injury and the expansion of hepatic progenitors that produce and/or respond to Hh ligands was strictly dose-dependent (i.e., greater expression of Hh ligands was accompanied by more accumulation of Hh-responsive progenitors) [25, 29]. Intriguingly, this process was associated with increased hepatic expression of TGFb1, [25, 29] a factor that is known to promote the death of mature hepatocytes, and to divert hepatic progenitors toward more ductular and/or fibroblastic phenotypes. When alcoholic steatohepatitis was modeled by ethanol intragastric feeding in mice, TGFb1 expression was also induced and numerous Hh-responsive cells accumulated [29]. Similar striking enrichment with Hh-responsive populations was also demonstrated in liver specimens from severely ill patients with alcoholic hepatitis [29]. In vitro treatment of hepatocytes (but not immature cholangiocytes or bipotent hepatic progenitor cells) with TGFb1 caused significant apoptosis [29]. However, the cells that survived were found to be Hh-responsive (unlike mature hepatocytes), and to exhibit features suggestive of immature liver epithelial cells that were undergoing EMT [29]. In liver samples from patients with fibrosing

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alcoholic hepatitis, quantitative immunohistochemical analysis revealed a striking correlation between the numbers of such cells and the Maddrey discriminant function (a reliable predictor of short term mortality) [29]. The latter finding proves that accumulation of Hh-responsive progenitors parallels alcoholic liver disease severity in humans (as it does in experimental models) and suggests that Hh-mediated EMT might promote liver repair at the expense of liver-specific function.

Vascular Remodeling During Cirrhosis Cirrhosis is characterized by a complex tissue remodeling response that eventually results in bridging fibrosis, parenchymal nodularity, and changes in sinusoidal architecture together with extrahepatic vasculature rearrangement [59]. Hh signaling is active in myofibroblasts (derived from transition of quiescent stellate cells and immature ductular cells) that accumulate in fibrotic/cirrhotic livers [31–33, 36]. This supports the concept that Hh pathway activation may orchestrate various aspects of the complex woundhealing process that occurs during cirrhosis. The Hh pathway is a key regulator of vascular remodeling during development; [60] PDGF-BB (which activates Hh signaling in adult liver cell populations) [30, 36] has also been demonstrated to regulate hepatic vascular structure and function [61]. Hence, it was postulated that Hh signaling might regulate cirrhosis-related vascular remodeling. Supporting this concept, biologically active Hh ligands were recently identified in exosomes that were released from myofibroblasts and immature cholangiocytes after these cells were exposed to PDGF-BB [35]. Moreover, treatment of other cells that contained Hh-reporter constructs with exosomes purified from myofibroblast- or cholangiocyte-conditioned medium activated Hh transcriptional activity, proving that Hh-containing exosomes are capable of initiating Hh signaling in distant Hh-target cells [35]. Consistent with the in vitro data, BDL-induced fibrosis/cirrhosis elicited the release of membrane-associated Hh ligands into both plasma and bile [35]. Even more interestingly, when exposed to either plasma- or bile-derived exosomeenriched Hh-containing membrane particles, sinusoidal endothelial cells were stimulated to undergo phenotipic

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changes that are known to occur during the capillarization process that accompanies cirrhosis-related vascular remodeling [35]. These findings identify a potentially novel mechanism for vascular remodeling during cirrhosis, namely, Hh-induced phenotypic changes in endothelial cells.

Hedgehog Pathway in Hepatocarcinogenesis Dysregulation of the Hedgehog pathway has been implicated in the genesis of malignancies derived from various tissues [5, 8]. Hh pathway activity has been reported in some cholangiocarcinoma and hepatocellular carcinoma cell lines, [5] and subgroups of patients with hepatocellular carcinoma [62]. Hh pathway inhibitors were also shown to block the growth of hepatoblastoma cells in culture, [62] raising the possibility that inappropriate activation of Hh signaling plays a role in hepatocarcinogenesis. Indeed, a novel Smo mutation has been reported in at least one HCC patient [62]. A pro-carcinogenic role for the Hh pathway is further supported by other evidence that HCC typically arise in the context of cirrhosis, a condition that favors the outgrowth of Hh-responsive cells.

Cholangiocarcinoma As discussed previously, there is abundant evidence that bile ductular cells are capable of both producing and responding to Hh ligands. Indeed, cholangiocarcinoma was the first type of liver cancer that was demonstrated to exhibit Hh activity [5, 8, 63]. Cholangiocarcinomas typically arise in the context of chronic fibrosing biliary and/or hepatic parenchymal injury and such conditions are characterized by striking induction of Hh signaling in bile ductular cells. These findings suggest that Hh pathway activation may play a permissive role in the pathogenesis of cholangiocarcinoma. Recent insight into mechanisms by which this might occur were provided by a careful analysis of apoptotic signaling in cultured cholangiocarcinoma cells. That work demonstrated that Hh signaling induced expression of survival factors and inhibited propagation of death receptor-initiated

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signaling that would have otherwise resulted in caspase activation. (Kurita et al., AASLD 2008 Abstract 1489, Hepatology, Volume 48, Number 4, Suppl.) [64].

Hepatoblastoma The Hh pathway is very active in liver progenitor cells and falls exponentially during cellular differentiation. For example, expression of Hh target genes was thousands of fold greater in pluripotent embryonic stem cells than in more differentiated hepatic progenitors, and pathway activity in the latter types of cells was thousands of fold greater than in healthy, mature hepatocytes [34]. Hepatic progenitors isolated from human fetal livers exhibit Hh pathway activity [34] Hepatic Hh pathway activity also increases dramatically in various types of adult liver injury that induce hepatic accumulation of oval cells (i.e., bipotent hepatic progenitors that are capable of differentiating into either hepatocytes or cholangiocytes). Consistent with this observation, cultured bipotent epithelial progenitors and immature ductular-type cells have been shown to produce and respond to Hh ligands ([31, 32, 34] and Diehl, unpublished data). Given these data, it is not surprising that several groups have demonstrated Hh signaling in Hep3B cells, a line of malignant liver cells that were derived from a child with hepatoblastoma. In Hep3B cells, pharmacologic or genetic inhibition of Hh signaling results in dramatic growth inhibition, suggesting that the Hh pathway plays a role in the pathogenesis of hepatoblastoma [65, 66].

Hepatocarcinoma Hepatocellular carcinoma (HCC) is one of the leading causes of cancer related mortality world-wide and the incidence of this tumor is rising in the United States. HCC typically develops in the context of cirrhosis, and cirrhosis is characterized by sustained activation of the Hh pathway, raising the possibility that induction of Hh signaling in liver might play a general role in the pathogenesis of HCC. This concept is supported by recent evidence that the Hh pathway inhibitor, Hhip,

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was hypermethylated and silenced in almost two-thirds of HCC in one study [67]. Several other studies have demonstrated increased expression of Hh-target genes in substantial subsets of HCC samples [68, 69]. Evidence for Hh pathway activation in HCC has been demonstrated predominately at the level of gene expression. Thus, the types of cells that are Hh-responsive within these tumors remain unclear. The possibility that stromal elements (including tumor fibroblasts) comprise of at least some of the Hh-responsive cells seems reasonable, but has not been examined directly. Interestingly, evidence for Hh signaling in hepatocellular carcinoma cells themselves has been variable, with some, but not other, groups reporting effects of silencing Hh signaling on HepG2 cell growth.

Conclusions Regardless of the type of injury, remodeling responses that aim to restore anatomical integrity and liver-­specific functions are triggered. When liver repair is forced to occur in the context of ongoing injury, remodeling res­ ponses include hepatic activation of a normally-­dormant developmental pathway, dubbed Hedgehog (Hh). Virtually all of the types of cells that participate in reparative remodeling of chronically injured livers are capable of producing and responding to Hh ligands, including liver progenitors, immature ductular cells, stellate cells, myofibroblasts, endothelial cells and ­certain types of lymphocytes. Hh signaling amongst these cells occurs via a complex network of autocrine/­ paracrine signals, and it helps to orchestrate various aspects of the wound-healing process, including expansion of progenitor populations, hepatic accumulation of ­myofibroblasts and inflammatory cells, and vascular remodeling. Unfortunately, sustained or dysregulated Hh pathway activation also compromises hepatocytespecific ­function and contributes to liver fibrogenesis, and ­carcinogenesis. Thus, activation of Hh signaling in Hh-responsive adult liver cells appears to represent a “double-edged sword,” necessary for repair of damaged livers, but potentially responsible for cirrhosis and liver cancer. Given that either insufficient or excessive Hh activity may be problematic, more research is needed to  delin­eate the mechanisms that control hepatic Hh ­signaling in adults.

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Summary

›› Hh ligands (Sonic, Shh; Indian, Ihh; Desert,

››

››

››

››

››

›› ›› ›› ››

Dhh) are soluble, lipid-modified morphogens that may be secreted in two different forms: a short range acting (poorly diffusible) type, and a second form for long-range transport, “packed” in membranous structures. Hh proteins are able to interact with Patched (Ptc: Ptch1 in vertebrates), a membrane-spanning receptor on the surface of Hh-responsive cells. In the absence of Hh ligands, Ptc keeps the co-receptor Smoothened (Smo) in its inactive form, and silences the Smo-dependent down-stream intracellular signaling. Hh ligand-receptor interaction de-represses Smo. Activation of Smo, in turn, inhibits Hh transcription factor phosphorylation, leading to an intracellular signaling cascade that ultimately drives the activation and nuclear translocation of Glioblastoma (Gli) family zinc-finger transcription factors. Since mature hepatocytes are insensitive to Hh ligands, enrichment of the microenvironment during hepatic injury with Hh ligands provides a selective survival advantage for cell types that are Hh-responsive, leading to the outgrowth of these cell populations. During cholangiopathies, PDGF-BB released from damaged cholangiocytes induces production of Shh by myofibroblastic stellate cells and immature cholangiocytes. BDL-induced fibrosis elicits the release of membrane-associated Hh ligands into both bile and plasma leading to a sinusoidal endothelial cell phenotypic change that is known to occur during the capillarization process. In alcoholic liver disease, accumulation of Hh-responsive progenitor cells parallels the severity of the disease. Cholangiocarcinoma exhibits Hh activity. The Hh pathway is very active in liver progenitor cells. There is an increased expression of Hh-target genes in a substantial subset of HCCs. Moreover, the Hh pathway inhibitor, Hhip, is hypermethylated and silenced in more than 50% of HCCs.

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Multiple Choice Questions 1. Which of the following statements is wrong? (a) The Hedgehog signaling pathway is normally very active in healthy adult livers (b) The Hh pathway affects the expression of several of its components (c) Gli1 and Gli2 are mostly responsible for providing prolonged cellular response (d) There is no direct evidence for hh involvement in fetal liver development (e) When levels of Ptc exceed those of the Hh ligands, the Hh pathway is turned off 2. Which of the following types of liver cells are NOT Hh-responsive? (a) Mature hepatocytes (b) Immature ductular cells (c) Myofibroblastic stellate cells (d) Sinusoidal endothelial cells (e) Hepatic progenitor cells 3. Which of the following growth factors have been shown to elicit production of Hh ligands by epithelial cells? (a) Hepatocyte growth factor (b) Interferon-g (c) Platelet-derived growth factor-BB (d) Insulin-like growth factor (e) TNF-a 4. Which of the following statement is true? (a) Epithelial-to-mesenchymal transitions induced by transforming growth factor beta are likely to involve Hh signaling (b) Phosphorylation of Gli by GSK3 promotes its nuclear translocation (c) Smo is activated by phosphorylation by casein kinase (d) Mice with overly-active Hh pathway have no duc­ tular reaction in response to bile duct ligation (e) Hh ligands are insoluble lipid-modified mor­ phogens 5. In the pathogenesis of which types of liver cancers does Excessive Hh signaling probably not play a role? (a) Cholangiocarcinoma (b) Hepatoblastoma (c) Hepatocellular carcinoma (d) Hemangioma (e) TUMORS derived from Hep3B cells

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401 55. Yang S, Koteish A, Lin H et al (2004) Oval cells compensate for damage and replicative senescence of mature hepatocytes in mice with fatty liver disease. Hepatology 39:403–411 56. Yang SQ, Lin HZ, Mandal AK et al (2001) Disrupted signaling and inhibited regeneration in obese mice with fatty livers: implications for nonalcoholic fatty liver disease pathophysiology. Hepatology 34:694–706 57. Richardson MM, Jonsson JR, Powell EE et al (2007) Progressive fibrosis in nonalcoholic steatohepatitis: association with altered regeneration and a ductular reaction. Gastroenterology 133:80–90 58. Popper H, Kent G, Stein R (1957) Ductular cell reaction in the liver in hepatic injury. J Mt Sinai Hosp N Y 24:551–556 59. Friedman SL (2008) Mechanisms of hepatic fibrogenesis. Gastroenterology 134:1655–1669 60. Vokes SA, Yatskievych TA, Heimark RL et al (2004) Hedgehog signaling is essential for endothelial tube formation during vasculogenesis. Development 131:4371–4380 61. Semela D, Das A, Langer D et al (2008) Platelet-derived growth factor signaling through ephrin-b2 regulates hepatic vascular structure and function. Gastroenterology 135: 671–679 62. Sicklick JK, Li YX, Jayaraman A et al (2006) Dysregulation of the Hedgehog pathway in human hepatocarcinogenesis. Carcinogenesis 27:748–757 63. Jinawath A, Akiyama Y, Sripa B, Yuasa Y (2007) Dual blockade of the Hedgehog and ERK1/2 pathways coordinately decreases proliferation and survival of cholangiocarcinoma cells. J Cancer Res Clin Oncol 133:271–278 64. Kurita S, Mott JL, Cazanave S, et al (2008) Inhibition of Hedgehog signalling sensitizes human cholangiocarcinoma cells to TRAIL-mediated apoptosis by TRAIL-RI/DR4 up-regulation. Hepatology 48 (Suppl. 1): 969A 65. Huang S, He J, Zhang X, Bian Y et al (2006) Activation of the hedgehog pathway in human hepatocellular carcinomas. Carcinogenesis 27:1334–1340 66. Wang Q, Huang S, Yang L et al (2008) Down-regulation of Sonic hedgehog signaling pathway activity is involved in 5-fluorouracil-induced apoptosis and motility inhibition in Hep3B cells. Acta Biochim Biophys Sin (Shanghai) 40:819–829 67. Tada M, Kanai F, Tanaka Y et al (2008) Down-regulation of hedgehog-interacting protein through genetic and epigenetic alterations in human hepatocellular carcinoma. Clin Cancer Res 14:3768–3776 68. Kim Y, Yoon JW, Xiao X et al (2007) Marcusson EG. Selective down-regulation of glioma-associated oncogene 2 inhibits the proliferation of hepatocellular carcinoma cells. Cancer Res 67:3583–3593 69. Patil MA, Zhang J, Ho C et al (2006) Hedgehog signaling in  human hepatocellular carcinoma. Cancer Biol Ther 5:111–117

Hypoxia-Inducible Factor-1 Signaling System

27

Deborah Stroka and Daniel Candinas

Introduction Oxygen (O2) homeostasis is regulated by balancing the supply of O2 from the vasculature and O2 consumed by mitochondrial oxidative phosphorylation. If the balance of O2 is disturbed, cells are exposed to a state of either oxidative stress or oxidative depression. Hypoxia is a state of oxidative depression and occurs when there is a reduction in blood flow or O2 content of the blood. In the liver, as within other tissues, hypoxic cells initiate adaptive responses to help ensure their survival. These adaptive responses are aimed at increasing O2 supply and compensate for loss of energy via physiological, metabolic, and molecular mechanisms. Studies are continuing to define the molecular pathways of these mechanisms and reveal that they occur at every regulatory level, including gene transcription, protein translation, and posttranslational modifications.

Identification of HIF-1 One of the most illustrative molecular adaptations to hypoxia is activation of the HIF signaling pathway. This pathway was initially identified by studies aimed at ­isolat the factors responsible for the production of erythropoietin (EPO). Tissue hypoxia is the primary physio­ logical stimulus of increased EPO gene transcription [1]. Although the kidney is the main site of EPO

production, other sites have been identified. In the liver, it was demonstrated using a transgenic mouse expressing the human EPO gene, that hepatocytes surrounding central veins synthesized large amounts of human EPO mRNA when the mouse was bled from a hematocrit of 55 to 10% [2]. This study importantly demonstrated the impact of O2 availability on controlling EPO gene expression and showed that EPO is transcriptionally controlled via an O2-regulated mechanism that senses and responds to reduced O2 availability. In 1991, the O2sensitive DNA control element that conferred responsiveness of the EPO gene to hypoxia was identified in its 3′ flanking region [3–5]. Further investigations identified a 120-kDa nuclear factor binding protein and it was named hypoxia-inducible factor-1 (HIF-1) [6, 7]. In extended studies investigating the O2-dependent activity of the EPO 3′-enhancer in a wide variety of non EPOproducing cells, it became clear that HIF was not exclusively an inducer of EPO transcription, but also operated in an O2-sensing system that is wide-spread in mammalian cells [7–9]. Subsequently, HIF-1 protein was cloned from the human hepatoma cell line, Hep3B, and determined to be a heterodimeric transcription factor consisting of two subunits, HIF-1a and HIF-1b, the latter being also known as aryl hydrocarbon receptor nuclear translocator (ARNT). Both HIF-1a and ARNT belong to the basic-helix-loop-helix (bHLH), Per/ARNT/ SIM (PAS) family of transcription factors [10, 11].

Structure of HIF-1 D. Stroka (*) Visceral Surgery Research Laboratory, Department of Clinical Research, University of Bern, Murtenstrasse 35, 3010 Bern, Switzerland e-mail: [email protected]

The a-subunit (120 kDa) of HIF-1 is a class II bHLHPAS protein and is the regulatory subunit that specifically mediates responses to hypoxia [12]. The bHLH domain contains the basic DNA binding region and

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_27, © Springer-Verlag Berlin Heidelberg 2010

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404 Fig 27.1  Sites targeted by post translational modifications that influence HIF-1a stability and transcriptional activity

D. Stroka and D. Candinas

HIF-1a P402

1

bHLH

PAS

ODD

K532 P564 S641 S643 NTAD

K391 K477

N803 CTAD

826

C800

HIF-1b/ARNT bHLH

1

HIF-1a

CTAD

PAS

P402, P564 N803 K532 S641, S643 K391, K477 C800

bHLH primary dimerization interface. The adjacent PAS domain mediates protein–protein interactions and acts as a second dimerization interface in conjunction with the HLH motif [13]. Two transactivation domains, N-TAD (N-terminal transactivation domain) and C-TAD (C-terminal transactivation domain) activate transcription of target genes [14]. The C-TAD interacts with the coactivators p300/CBP to activate transcription. The N-TAD is located within the O2-dependent degradation (ODD) domain and is a target of several posttranslational modifications, which influence HIF1a stability and transcriptional activity (Fig. 27.1). To date two other HIF a-subunits were cloned from human, rat and mouse sources, HIF-2a [15,16] and HIF3a [16,17]. All three HIF a-subunits have one of the same heterodimerization partners, ARNT1, 2, and 3. HIF-2a was also referred to as endothelial PAS domain protein 1 (EPAS1) [18], HIF1a-like factor (HLF) [19], and HIF-related factor (HRF) [20], and is also a member of the PAS superfamily 2 (MOP2) [15]. HIF-1a and HIF-2a subunits are structurally similar in their DNA binding and dimerization domains but differ in their transactivation domains, implying they may have unique target genes. The functional role and expression pattern of HIF-3a still need to be elucidated; however, it appears to be involved in negative regulation of hypoxic responses, through an alternately spliced transcript termed inhibitory PAS domain protein (IPAS) [21]. The HIF-1b-subunit (91–94 kDa) is a class I bHLHPAS protein and is also referred to as ARNT. Other

789

PHD Hydroxylation FIH-1 Hydroxylation ARD1 Acetylation MAPK Phosphorylation SUMOylation S-nitrosation

family members include ARNT2, ARNT3 (also known as BMAL1/MOP3), and MOP9. ARNT is a constitutively expressed nuclear protein that functions in a variety of transcriptional systems with alternative dimerization partners [22]; for example, ARNT is known to heterodimerize with AhR (aryl hydrocarbon receptor) following activation by xenobiotic ligands such as dioxin or other aryl hydrocarbons. This complex binds to the xenobiotic response element that controls expression of genes involved in xenobiotic metabolism including cytochrome P-450 and glutathione S-transferase Ya. Activity of ARNT is generally not affected by hypoxia; however, nuclear stabilization of the heterodimer may result in a perceived increase in total cellular ARNT levels in some cell lines [23]. The universal importance of the HIF system is demonstrated by the ubiquitous expression of HIF-1a and HIF-1b (ARNT) mRNA in most if not all adult and embryonic mouse and human tissues; furthermore, HIF-1a, HIF-2a, and ARNT subunits are all absolutely required for normal embryonic development. Murine embryonic lethal phenotypes were found in gene targeting experiments at both the HIF-1a and HIF-2a loci. HIF-1a knockout embryos die around mid-gestation, showing abnormal vascular development and open neural tube defects [24–26], indicating that HIF-1a is required for mesenchymal cell survival. Targeted inactivation of HIF-2a resulted in differing phenotypes, with defects in vascular remodeling [27], a defect in fetal ­catecholamine production [28], or defects in lung

27  Hypoxia-Inducible Factor-1 Signaling System

maturation involving surfactant deficiency [29]. Mice containing only one mutant HIF-1a allele develop normally but show impaired physiological responses to chronic hypoxia such as reduced polycythemia, right ventricular hypertrophy, pulmonary hypertension, pulmonary vascular remodeling, and electrophysiological responses [30, 31].

405

modifications. Numerous studies revealed the complexity of its regulation showing that its protein ­stability and transcriptional activity can be influenced by posttranslational modifications such as, hydroxylation, acetylation, phosphorylation, S-nitrosylation, and SUMOylation (Fig. 27.1). Although each of these modifications affects HIF-1a protein, the major and most clearly defined mechanism is its O2-dependent hydroxylation.

HIF Target Genes The HIF-dependent O2-sensing system has a widespread distribution and mediates adaptive responses to hypoxia in both physiological and pathological conditions. More than 100 HIF-1 target genes have been identified so far. HIF target genes encode proteins that play key roles in both immediate and prolonged adaptations to O2 deficiency. Among these genes are those involved in angiogenesis, vascular tone and remodeling, glucose uptake and metabolism, cell proliferation and survival, and iron homeostasis (Table 27.1). HIF target genes are defined by the presence of a functional HIF-1 binding site containing the core recognition sequence 5'-R/CGTG-3' in regulatory regions of its responsive genes. The presence of a HIF binding site is necessary but not sufficient to direct gene expression in response to hypoxia. Binding of additional transcription factors in the vicinity of HIF-1, such as activator protein-1 (AP-1) for VEGF transcription, the presence of adjacent HIF binding sites, present in the iron transport protein transferrin, or the glycolytic enzyme phosphoglyerate kinase-1 (PGK1), are generally required. Other proteins that interact with HIF-1a include various transcriptional coactivators, thereby suggesting that HIF target genes expression can be induced in a cell-type specific manner depending on the cellular composition of accessory coactivating proteins (Table 27.2).

Regulation of HIF-1 The transcriptional activity of HIF-1 is predominately determined by environmental factors that influence the expression of its a subunit. While HIF-1a mRNA levels generally remain constant, its protein expression is tightly controlled by various posttranslational

Hydroxlation When O2 is present, HIF-1a is hydroxylated by 2-oxoglutarate-dependent oxygenases at two specific prolyl residues (P402 and P564) located in the ODD [32– 35]. This enzymatic reaction is inherently O2-dependent as the oxygen of the hydroxyl group is derived from molecular O2. In addition to molecular O2, prolyl hydroxy­lation requires 2-oxoglutarate and iron as cofactors, thereby accounting for the well known “hypoxia-mimic” effects of iron antagonists such as desferoxamine and cobalt chloride. The hydroxylated prolyl residues of HIF-1a serve as a recognition site for the tumor suppressor protein VHL [36–38]. pVHL is a recognition component of the E3 ubiquitin ligase complex that contains elongin B, elongin C, cullin2, and Ring box protein 1 (Rbx1), as well as an E2 ubiquitin conjugating enzyme (E2). Ubiquitination of HIF-1a by this multiprotein complex leads to HIF-1a degradation by the 26S proteasome. The O2-dependent prolyl hydroxylation of HIF-1a is carried out by three orthologs of Caenorhabditis elegans (C. elegans) Egl-9 (called EGLN1-3), which in mammalian cells are referred to as prolyl hydroxylase (PHD) 1–3, or HIF-1 prolyl hydroxylases (HPH) 1–3 [39–42]. It was demonstrated that specific gene silencing of PHD2 by siRNA was sufficient to stabilize HIF1a under normoxic conditions. However, silencing of PHD1 and PHD3 had no effect on HIF-1a stability [43]. Consistent with these findings, it was shown that PHD2 has the highest specific activity for HIF-1a hydroxylation [44]. Interestingly, it was demonstrated that PHD2 and PHD3 mRNAs, as well as PHD2 protein, were induced by hypoxia [40, 45]. These findings provided evidence of an autoregulatory mechanism that helps to explain why the more severe the hypoxic stress (leading to a strong induction of PHD2 protein) the faster HIF-1a is degraded upon reoxygenation [46].

406 Table 27.1  HIF-1 target genes Categories of HIF target Genes O2 transport Erythropoietin Iron metabolism Ceruloplasmin (iron oxidation) Transferrin (iron transport) Transferrin receptor (iron uptake) Ferrochelatase Angiogenesis, vascular tone, and extracellular matrix metabolism Vascular endothelial growth factor (VEGF) VEGF receptor Flt-1 EG-VEGF a-1B-andrenergic receptor Heme-oxygenase-1 (CO production) Inducible nitric oxide synthase (NO production) Endothelin-1 Atrial natriuretic protein Plasminogen activator inhibitor-1 (PAI-1) Adrenomedullin LDL-receptor-related protein 1 Transforming growth factor-b3 Leptin Endoglin Connective tissue growth factor Prolyl-4-hydroxylase a 1 Collagen type V (a1) Fibronectin Matrix metalloproteinase 2 Urokinase plasminogen activator receptor Cathepsin D Glucose metabolism and uptake Glucose transporters 1 and 3 Adenylate kinase-3 Aldolase A and C Enolase 1 Phosphoglyerate kinase-1 Phosphofructokinase L and C Lactate dehydrogenase A Glyceraldehyde-3-phosphate-dehydrogenase Glucose-6-phosphate Isomerase Hexokinase 1,2 6-Phosphofructo-2-kinase/Fructose-2,6biphos-4-3 and -4 Triosephosphate isomerase Glucokinase Phosphoenolpyruvate carboxykinase pH regulation Carbonic anhydrase-9 Na+/H+ exchanger isoform 1

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References [6] [113] [114] [115, 116] [117]

[118–120] [121] [122] [123] [124] [125] [126] [127] [128] [129, 130] [131] [132] [133, 134] [135] [136] [137] [131] [138] [138] [138] [138] [118, 139] [140] [141] [141, 142] [142–144] [142] [140, 143] [145] [146] [24] [147, 148] [149] [73, 150] [151]

[152] [153]

Categories of HIF target Genes Cell proliferation and apoptosis Insulin-like growth factor binding protein-1-3 p21 (cell proliferation) Transforming growth factor-a Cyclin G2 Human telomerase reverse transcriptase Nucleophosmin Nip3 (proapoptotic) Bid, Bax BNIP3, NIX RTP801 NOXA HGTD-P Myeloid cell factor Survivin Inflammation COX-2 b2-integrin CYP4B1 Adrenomedullin Adenosine A2B receptor Macrophage migration inhibitory factor Retrotransposon VL30 Intestinal trefoil factor Regulation of PHDs HIF-prolyl hydroxylase 2 HIF-prolyl hydroxylase 3 Cancer-related N-myc downstream regulated 1 Stromal cell-derived factor-1 c-Met Wilms’ tumor suppressor CXCR4 Breast cancer resistance protein Transcriptional regulation Differentiated embryo-chondrocyte gene 1, 2 ETS-1 NUR77 CITED (p35srj; CBP/p300 antagonist) PP1 nuclear targeting subunit Miscellaneous Glucose-regulated protein 94 (ER-stress) b-glycoprotein (MDR1, drug resistance) Furin (proprotein conversion) Cytochrome P450 2C11 (drug metabolism) Inhibitor of differentiation (ID2; development) Adenylate kinase 3 (nucleotide metabolism) Ecto-5′-nucleotidase (nucleotide metabolism) Leptin (metabolism) Visfatin (metabolism)

References [154, 155] [156] [138] [131] [157] [158] [159] [160] [161] [162] [163] [164] [165, 166] [167] [168] [169] [170] [171] [172] [173] [174] [175] [176] [177] [178] [179] [88] [180] [89, 181] [182] [183, 184] [185] [186] [187] [188] [189] [90] [190] [191] [192] [140] [193] [133, 134] [194, 195]

27  Hypoxia-Inducible Factor-1 Signaling System Table 27.2  HIF-1a interacting proteins Transcriptional Function cofactors CBP/p300 Histone acetyltransferase SRC-1 Histone acetyltransferase TIF2 Histone acetyltransferase Ref-1 Redox factor HNF4 Gene-specificity Smad3 Gene-specificity FIH-1 Hydroxylase Nucleolar sequestration p14ARF

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References [196, 197] [198] [198] [198] [199] [200] [201] [202]

In addition to the PHDs, HIF-1 activity is regulated by another 2-oxoglutarate-dependent dioxygenase,

Normoxia Asparaginyl hydroxylase (FIH)

OH

Hypoxia

HIF-1α

O2, Fe2+ 2-oxoglutarate

N803

termed factor inhibiting HIF-1 (FIH). The C-TAD of HIF-1a interacts with the transcriptional coactivator p300/CBP to enhance transcription of HIF-1 target genes. Hydroxy­lation of a specific asparagine residue (N803) in the C-TAD by FIH interrupts the interaction between C-TAD and p300/CBP, thereby inhibiting HIF-1 transcriptional activity [47]. In summary, the mammalian O2-sensing pathway involves prolyl and asparaginyl hydroxylation of the HIF-1a subunits, which in oxygenated cells inactivates HIF by proteolytic destruction and inhibition of coactivator recruitment. Under hypoxic conditions, these mechanisms are blocked by the lack of molecular O2, allowing HIF-1a stabilization, nuclear translocation, binding to target genes, and coactivator recruitment (Fig. 27.2).

P402

HIF-1α

OH Ac OH

Acetyltransferase

Stabilization of Hif-1α protein

P564

Prolyl hydroxylases

p300/CBP

HIF-1α

OH Ac OH

VHL EL B

Cul2

EL C

RBX1

Ubiquitination

Nucleus p300/CBP

HIF-1α

mRNA EPO, VEGF, etc. HIF-1α

HIF-1β Proteosomal Destruction of HIF-1α

Fig 27.2  Post translational regulation of HIF-1a

HRE

DNA

408

Acetylation Acetylation is another posttranslational modification that results in the instability of HIF-1a protein. Acetylation is controlled by two opposing groups of enzymes, histone acetyl-transferases (HATs) and histone deacetylases (HDACs). These enzymes are best known by their ability to modify the acetylation level of histone proteins. More recently, HATs and HDACs have been shown to be modified nonhistone proteins, among which are several key transcription factors (e.g., p53, E2F1, NF-kB, MyoD) and transcriptional coactivators (e.g., p300, PGC-1a). The acetylation of transcription factors influences their stability, transcriptional activity, and interaction with transcriptional coactivators, as well as their DNA binding affinity. In the case of HIF-1a, conflicting data concerning its acetylation by an Naacetyltransferases, termed arrest-defective-1 (ARD1) protein, have been published. It was demonstrated in a yeast two-hybrid assay that a mouse ARD1 acetylates a specific lysine residue (K532) located in the ODD domain of HIF-1a under normoxic conditions. Acetylated HIF1a showed enhanced binding to VHL, thereby promoting HIF-1a degradation [48]. However, human ARD1 associates with the ODD domain of human HIF-1a, but does not acetylate and destabilize HIF-1a [49,50]. Although the above mentioned publications provide evidence that hARD1 is unable to acetylate HIF-1a, one report suggested an antiangiogenic role of connective tissue growth factor (CTGF) by accelerating HIF-1a degradation through ARD1-dependent acetylation [51]. Whereas only one relevant acetyl-transferase has been described thus far, several HDACs have been implicated in the regulation of HIF-1a. HIF-1a has been shown to interact with HDAC1 and 3 (class I) [52], HDAC4 [53] and HDAC7 (class IIa) [54], and HDAC6 (class IIb) [53]. So far, no data have been published concerning the interaction between class III HDACs (sirtuins) and HIF-1a. Inhibition of the class II HDACs-4 and -6 results in HIF1a degradation in a VHL-independent manner [53]. Inhibition of HDAC6 was suggested to decrease HIF-1a stability by interfering with chaperone protein HSP90. HSP90 is responsible for the correct folding and maturation of several proteins including HIF-1a. Inhibition of HDAC6 increases the acetylation level of HSP90, thereby impairing its chaperone function and concomitantly

D. Stroka and D. Candinas

degrading HIF-1a levels in a VHL-independent, proteasome-mediated pathway [55]. HDACs are generally involved in transcriptional silencing. Therefore, it is interesting that in the case of HIF-1, HDACs are needed for transcriptional activation by stabilizing HIF-1a protein. In hepatocellular carcinoma (HCC), HDAC1 and metastasis-associated protein 1 (MTA1) are induced by hepatitis B virus X protein (HBx) [56]. MTA1/HDAC1 complex stabilizes HIF-1a by deacetylation, thus potentially playing a critical role in angiogenesis and metastasis of HCC [57]. In the past years, a number of small-molecule inhibitors of HDACs (HDACi) have been identified to possess antitumoral activity. Various mechanisms of the HDACimediated antitumoral effects have been proposed, including directly altering HIF-1a stability and/or transcriptional activity and thereby targeting HIF-mediated tumor angiogenesis (reviewed in [58]).

Phosphorylation Phosphorylation of HIF-1a is another posttranslational modification that increases its transcriptional activity. Direct phosphorylation of HIF-1a occurs after stabilization of the protein under normoxic or hypoxic conditions [59,60]. Two specific serine residues (S641 and Ser643) have been identified as phosphorylation targets of p42/p44 mitogen-activated protein kinases (MAPK) [61]. It was suggested that phosphorylation of the two serine residues promotes nuclear accumulation and transcriptional activity of HIF-1 by inhibiting the interaction between the nuclear export signal (NES) of HIF-1a and the nuclear export protein, CRM1 [62]. Inhibition of phophorylation by site-directed mutagenesis of the two serine residues or by MAPK pathway inhibitors showed much lower transcriptional activity [61]. The phosphorylation status of HIF-1a can also determine whether cells undergo apoptosis or survive. Whereas phosphorylated HIF-1a binds to HIF-1b, dephosphorylated HIF-1a preferentially interacts with p53, thus promoting apoptosis [63]. Other family members of MAPK such as p38 MAPK and c-Jun N-terminal kinase (JNK) do not phosphorylate HIF-1a [60] unless activated by specific viral oncogenes [59]. Unlike phosphorylation by MAPK that results in HIF-1a stabilization, phosphorylation by glycogen

27  Hypoxia-Inducible Factor-1 Signaling System

synthase kinase 3 (GSK3) results in its degradation. Inhibition of GSK-3 or mutations of the GSK-3 phosphorylation sites (S551, T555, and Ser589) in the ODD enhanced HIF-1a protein levels [64].

S-Nitrosylation S-nitrosylated proteins form when a cysteine thiol reacts with nitric oxide (NO) to form an S–NO bond. S-nitrosylation of a cysteine residue (C800) within the C-TAD of HIF-1a allows the interaction between HIF1a and the transcriptional coactivator p300, thereby promoting HIF-1 transcriptional activity. This effect was not observed when the cysteine residue was ­substituted by alanine (C800A) [65]. A recent study reported that normoxic HIF-1 activity increased upon NO-mediated S-nitrosylation and stabilization of HIF1a. In murine tumors, ionizing radiation stimulated the production of NO, thereby promoting S-nitrosylation of  the only cysteine residue (C533) within the ODD of  murine HIF-1a [66]. The mechanism, by which NO-mediated S-nitrosylation was suggested to enhance HIF-1a stability, is interruption of VHL binding to the S-nitrosylated ODD of HIF-1a. These findings were further confirmed by demonstrating that mutation of the cysteine residue in the ODD (C533S) did not decrease binding with VHL in the presence of NO, thereby leading to its continuous degradation [66].

SUMOylation SUMOylation has recently been discovered as an important and dynamic posttranslational modification of proteins. A general increase of protein SUMOylation occurs under certain stress conditions such as hypoxia [67]. It was shown that HIF-1a was targeted for SUMOylation; however, what influence SUMOylation has in the stabilization and transcriptional activation of HIF-1a is still controversial. On one hand, a study demonstrated increased HIF-1a stability and transcriptional activity by SUMOylation of two lysine residues (K391 and K477) within its ODD [68]. Consistent with these results, another group described a role for RSUME (RWD-

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containing SUMOylation enhancer), a protein that is induced by hypoxia and that enhances HIF-1a SUMOy­ lation, thus promoting its stabilization and transcriptional activity [69]. On the other hand, there are data demonstrating reduction of HIF-1 transcriptional activity by SUMOylation [70]. A recent study revealed a critical role for SUMO-specific protease 1 (SENP1) in the regulation of HIF-1a stability in hypoxia. SENP1−/− mice embryos show severe fetal anemia because of deficient EPO production [71]. SENP1 controls EPO by the regulation of HIF-1a stability during hypoxia. Hypoxia induces HIF-1a SUMOylation and promotes VHL binding and consecutive ubiquitination and degradation. SENP1 reverses HIF-1a SUMOylation.

HIF Expression in the Liver In the liver an O2 gradient is formed within the parenchyma as a result of the unidirectional blood flow from the portal vein and hepatic artery (periportal) to the central vein (perivenous). This O2 gradient is further delineated by the O2-consuming metabolic processes of the parenchymal cells along the hepatic plate in which the O2 tension drops from 60–65 mmHg in the periportal area to 30–35 mmHg in the perivenous area [72]. The O2 gradient is an important factor in the regulation of genes encoding various enzymes of the carbohydrate metabolism [72]. For example, glycolytic enzymes like glucokinase or pyruvate kinase show an enhanced expression in the less aerobic, perivenous zone of the liver, whereas gluconeogenic enzymes like phosphoenolpyruvate carboxykinase or glucose-6phosphatase are expressed predominantly in the aerobic, periportal zone [72]. The transcription factors mediating the zonated expression of glycolytic enzymes are not fully known at this time; however, recent advances have demonstrated that a regulatory transcriptional complex consisting of HIF-1, HNF-4, and p300 appears to be involved in insulin-dependent gluocokinase gene activation [73]. All three HIFa-subunits are expressed in the liver; in situ hybridization showed that HIF-1a, HIF2a, and HIF-3a mRNA were found predominantly in the perivenous zone of the liver. However, expression of HIFa-proteins was not zonated and the

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proteins were found evenly distributed in the cytosol of hepatocytes at low levels and in the nuclei at higher levels in both periportal and perivenous regions [16]. As they are more susceptible to hypoxic damage, the increased amount of observed HIFamRNAs in hepatocytes around the perivenous region may facilitate a quick de novo protein synthesis enabling a quick response to initiate adaptation mechanisms [16]. A homogeneous nuclear expression of HIF-1a protein was also observed in hepatocytes of normal mouse liver tissue, and as expected its expression was further increased in an O2-dependent manner [74]. Using a model of systemic hypoxia, HIF-1a protein expression was increased in liver tissue of mice exposed to decreasing pO2 in a time- and O2-dependent manner. There was no zonal protein expression in the parenchyma or no increase of HIF-1a mRNA in response to hypoxia [74]. Using a similar model, rats exposed to 0.1% CO, which induced a functional anemia, were used to localize the expression of HIF-2a. Strong staining of hepatocytes was seen in the vicinity of the central vein, in virtually every cell the nucleus was positive, whereas weak or no staining was found around the portal triads; these suggest a zonal expression of HIF-2a protein [75]. Interestingly the kinetics of HIF-1a expression in mice differed from HIF-2a expression observed in rats. Under a continuous hypoxic stimulus, HIF-1a was induced in the liver within 1  h of exposure, peaked after 2 h, and returned to undetectable levels at 3 h. In contrast, the response of HIF-2a is delayed for up to 3 h and then shows a rather marked increase that was prolonged for more than 6 h. Although this difference may be a result of experimental conditions, including the species variation, these findings suggest that there may be a coordinated response to hypoxia that is not a redundancy of function between the family members; it suggests that a differential function could be achieved in cells that activate both HIF-a isoforms at the protein level. In comparative studies of hypoxia-induced genes, it was demonstrated that HIF-1a and HIF-2a have unique targets. HIF-1a, but not HIF-2a, stimulates glycolytic gene expression in various cell types [76]. Additionally, it was shown that hepatic EPO production is preferentially regulated by HIF-2a, while other HIF target genes in the liver such as PGK-1 are preferentially regulated by HIF-1a [77].

D. Stroka and D. Candinas

Importance of Hypoxia in Liver Pathologies Although the liver receives approximately 30% of its blood supply from the hepatic artery and the remaining blood is supplied from the portal vein, the hepatic artery accounts for more than 50% of the O2 delivered to the liver. O2 is the main energy source for ATP synthesis via oxidative phosphorylation; interruptions in O2 supply can abolish the cell’s main energy supply and further compromise membrane function, reduce protein synthesis, and create alterations in hepatocellular iron homeostasis [78]. The high-energy consumption of hepatocytes renders them vulnerable to reductions in O2 availability; therefore, hypoxia can lead to hepatocellular damage and can be a factor in several secondary and primary liver diseases. Listed below are examples of various liver pathologies in which hypoxia may be an augmenting stimulus; further elucidation is required to determine if the effects of tissue hypoxia are HIF-mediated responses.

Ischemia-Reperfusion Injury and Ischemic Preconditioning Ischemia-reperfusion (I/R) injury occurs in the liver when cellular damage during restrictions of blood flow is accentuated following the restorations of O2 delivery. I/R injury is a complex problem of hepatic resections and liver transplantation and remains a serious complication in clinical practice. During the ischemic state, lack of O2 causes mitochondrial de-energization, ATP-depletion, alterations of H+, Na+, and Ca+ homeostasis that impair cell volume regulation and microcirculation once blood flow is restored. The severity of damage depends on the duration of the ischemic stress, whereby for short periods the cell damage can be repaired, whereas long periods result in acute and chronic liver dysfunction. Interestingly, the ability of liver cells to withstand short periods of ischemia was shown to have a beneficial effect. Ischemic preconditioning, which is a brief period of ischemia followed by a short interval of reperfusion before a prolonged ischemic stress, protects against I/R injury. The cellular mechanism behind the protective effect of ischemic preconditioning is complex involving the interaction between various cell types and activation of many

27  Hypoxia-Inducible Factor-1 Signaling System

signaling pathways. One mechanism responsible for early preconditioning is the release of adenosine by hepatocytes, which subsequently activates A2A-receptors and various intracellular signaling pathways. Recently, stimulation of adenosine A2A-receptors was shown to induce HIF-1, leading to the proposed involvement of HIF-1 in preconditioning of the liver [79]. HIF-1 activation by preconditioning was also suggested to prevent hepatocyte apoptosis induced by Fas-ligand, mainly through the activation of cytoprotective genes [80]. These studies advocate HIF-1 as a potential target for the pharmacological preconditioning of the liver; however, additional studies are still needed to determine if its prolonged activation has adversative effects.

Liver Cirrhosis Cirrhosis is a chronic disease of the liver in which diffuse destruction and regeneration of hepatic parenchymal cells result in an increase of connective tissue, creating a disorganization of the lobular and vascular architecture. The disorganization of the vascular architecture creates local hypoxic areas subjecting the parenchymal cells to reduced O2 availability. Results from an experimental biliary cirrhosis model in rats concluded that there is a sequential induction of two major angiogenic factors, VEGF and FGF-2, during biliary type of liver fibrogenesis. This suggests that hypoxia might be a major factor in the induction of VEGF and in the marked angiogenesis occurring at an early stage before the onset of cirrhotic lesions [81]. In additional studies, it is suggested that hepatocellular hypoxia causes inhibition of HGF and of c-Met-mediated proliferation, and thereby might contribute to failure in liver regeneration in the cirrhotic liver [82]; hepatocellular hypoxia after a liver injury directly contributes to the progression of liver fibrosis [83]. The role of the HIF-1 signaling pathway in these responses is not determined at this time.

Liver Cell Carcinomas The oxygenation state of tumors is used as a prognostic indicator of potential therapeutic outcome, as hypoxic tumors are more resistant to chemo- and radiation therapy, and have a more aggressive phenotype [84]. As a

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tumor increases in its cell mass and glycolytic rate, the O2 tension within its microenvironment drops as it is no longer adequately supplied with O2 from the existing blood vessels [85–87]. Tumor cells turn the lack of O2 to their advantage, using it as a powerful stimulus for tumor progression by selecting cells with enhanced glycolytic activity, as well as promoting tumor angiogenesis through the upregulation of angiogenic factors, i.e., VEGF, and increasing its metastatic potential [88,89]. Furthermore, the multidrug resistance (MDR1) gene is hypoxia-responsive and implicates hypoxia-induced P-glycoprotein expression as a pathway for resistance of some tumors to chemotherapeutics [90]. It has been suggested in HCC that shortage of blood supply due to portal hypertension (an effect of liver cirrhosis) and the rapid proliferation of tumor cells lead to local hypoxia, which in turn stimulates the synthesis of angiogenic factors. The majority of HCC tissue samples exhibit strong expression of angiogenic agents, such as VEGF, FGF-2, and IGF-1, and are implicated as important factors in the neo-vascularization of HCC. With more clarification on the importance of the cyto­kine networks and of tumor angiogenesis in HCC, agents that affect these pathways will be of great interest [91]. Interestingly, these factors, as well as many other tumor adaptive responses, involve the molecular adaptation to hypoxia in part through HIF-1. The importance of HIF-1 in tumor biology is well established. A recent study of metastatic liver cancers demonstrated that glycolysis induced by HIF-1 is the predominant energy source in the hypoxic environment and, at least in some transcatheter arterial embolization-pretreated HCC cases, cancer cells obtained energy for growth by switching the metabolic profile to glycolysis through HIF-1 [92,93]. Finally, in addition to tumor hypoxia, HIF-1a is over expressed in human cancers as a result of genetic alterations, such as over expression of the v-src oncogene [94] or inactivation of the tumor suppressor genes p53 [95], PTEN [96], or pVHL [36]. The most marked effect observed thus far is in tumors that have lost VHL function [36]. The importance of VHL function in the liver was demonstrated using conditional targeted disruption of hepatic VHL. Mice heterozygous for the 1-lox allele develop cavernous hemangiomas of the liver that histologically displayed hepatocellular steatosis and focal proliferations of small vessels. The mRNA encoding HIF targets genes, such as VEGF, GLUT-1, and EPO, were upregulated in these tumors [97]. Further experiments performed demonstrated the

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development of hemangiomas with the targeted deletion of HIF-1a, thereby suggesting the vital role of HIF-2a in the development of hepatic VHL-associated vascular tumors [98,99].

D. Stroka and D. Candinas

Summary

›› The importance of O2 tension in liver patholo-

Liver Regeneration Currently the role of HIF-1 in liver regeneration has not been fully established. However, it is attractive to speculate that it does have a regulatory function. At the molecular level, the entry of a hepatocyte into the cell cycle is stimulated by various cytokines and growth factors, many of which have been shown to either be influenced by hypoxia or effect HIF-1a protein stabilization. These factors include IL-1, IL-6, HGF, EGF, TNF-a, and insulin, all of which also result in the activation of other transcription factors including NF-kB, STAT3, activator protein 1 (AP-1), and CCAAT/ enhancer-binding protein (C/EBP)b (reviewed in [100]). Interestingly, using a murine 70% partial hepatectomy (PH) model together with a high density oligonucleotide microarray, HIF-1a mRNA was shown to increase 2.7-fold 4 h post PH, suggesting that HIF-1 may play a role in the beginning of tissue remodeling [101]. Recently, the product of the HIF target gene, EPO, was suggested to increase the regenerative capacity of the liver after major hepatectomy [102].

››

›› ›› ››

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gies has long been appreciated. In addition to the diverse signaling pathways activated by an abundance of O2 in the form of free radicals, pathways that sense and respond to the absence of O2 are emerging as dynamic and versatile signaling mechanisms. Tissue hypoxia can provide a strong physiological stimulus that is an important modulator in the progression of liver disease and tumor development. The HIF signaling system provides a molecular mechanism describing how cells and tissues adapt to hypoxic environments. Targeted intervention of HIF activity may be beneficial both in the promotion and inhibition of hypoxia driven responses. Therapeutic over-expression of HIF-1 may help the revascularization of ischemic tissues; on the contrary, inactivation of HIF-1 activity may be advantageous in inhibiting cancer progression as this would help starve growing tumors of O2 and nutrient supply. Further characterization of the HIF system, namely of the recently described class of O2dependent regulating enzymes, has the potential to offer new and exciting therapeutic approaches for the treatment of liver cancers and disease.

Other Signaling Pathways Influenced by Hypoxia In addition to the HIF-1 signaling pathway, the regulation and activity of many other cellular processes and proteins are influenced by O2 availability. The activity of other protein kinases has been shown to be regulated by hypoxia; these include p44/p42MAPK [103], p38 MAPK [104,105], and diacylglycerol kinase [106]. Likewise, hypoxia induces the activity of other transcription factors such as AP-1 [107], NF-kB [108], and Egr-1 [109]. Finally, hypoxia has been shown to increase the stability of messenger RNA, such as for VEGF [110–112], as well as influence the splicing of specific alternative mRNA transcripts [21].

Multiple Choice Questions 1. HIF-1a is targeted by the 2-oxoglutarate-dependent oxygenase called factor-inhibiting HIF-1 (FIH) at the following residue(s) (a) P402 (b) P564 (c) P402 and P564 (d) N803 (e) P402, P564 and N803 2. Which of the following post-translational modifications is most likely to increase the transcription activity of HIF-1?

27  Hypoxia-Inducible Factor-1 Signaling System

(a) Hydroxylation (b) Acetylation (c) Phosphorylation by glycogen synthase kinase 3 (d) S-nitrosylation (e) None of the above 3. An autoregulatory mechanism of HIF-1a can be induced through the hypoxic upregulation of which of the following enzymes? (a) HIF-1 prolyl hydroxylase-1 (b) HIF-1 prolyl hydroxylase-2 (c) HIF-1 prolyl hydroxylase-1 and -2 (d) HIF-1 asparaginyl hydroxylase (e) HIF-1 asparaginyl hydroxylase and HIF-1 ­prolyl hydroxylase-1 4. Which are the following cellular functions that can be regulated by hypoxic stress? (a) Production of carbon dioxide (b) Drug metabolism (c) Regulation of intracellular pH (d) Apoptosis (e) All of the above 5. In the liver, HIF-1 is the predominant family member responsible for hypoxia-induced… (a) Erythropoietin production (b) Hepatic steatosis (c) Phosphoglyerate kinase-1 expression (d) VHL-associated vascular tumors (e) All of the above

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27  Hypoxia-Inducible Factor-1 Signaling System 181. Schioppa T et al (2003) Regulation of the chemokine receptor CXCR4 by hypoxia. J Exp Med 198:1391–1402 182. Krishnamurthy P et al (2004) The stem cell marker Bcrp/ ABCG2 enhances hypoxic cell survival through interactions with heme. J Biol Chem 279:24218–24225 183. Ivanova AV, Ivanov SV, Danilkovitch-Miagkova A, Lerman  MI (2001) Regulation of STRA13 by the von Hippel-Lindau tumor suppressor protein, hypoxia, and the UBC9/ubiquitin proteasome degradation pathway. J Biol Chem 276:15306–15315 184. Miyazaki K, Kawamoto T, Tanimoto K, Nishiyama M, Honda H, Kato Y (2002) Identification of functional hypoxia response elements in the promoter region of the DEC1 and DEC2 genes. J Biol Chem 277:47014–47021 185. Oikawa M, Abe M, Kurosawa H, Hida W, Shirato K, Sato Y (2001) Hypoxia induces transcription factor ETS-1 via the activity of hypoxia-inducible factor-1. Biochem Biophys Res Commun 289:39–43 186. Choi JW, Park SC, Kang GH, Liu JO, Youn HD (2004) Nur77 activated by hypoxia-inducible factor-1alpha overproduces proopiomelanocortin in von Hippel-Lindaumutated renal cell carcinoma. Cancer Res 64:35–39 187. Bhattacharya S, Michels CL, Leung MK, Arany ZP, Kung AL, Livingston DM (1999) Functional role of p35srj, a novel p300/CBP binding protein, during transactivation by HIF-1. Genes Dev 13:64–75 188. Yoshida D, Kim K, Noha M, Teramoto A (2006) Hypoxia inducible factor 1-alpha regulates of platelet derived growth factor-B in human glioblastoma cells. J Neurooncol 76: 13–21 189. Paris S, Denis H, Delaive E, Dieu M, Dumont V, Ninane N, Raes M, Michiels C (2005) Up-regulation of 94-kDa glucose-regulated protein by hypoxia-inducible factor-1 in human endothelial cells in response to hypoxia. FEBS Lett 579:105–114 190. McMahon S, Grondin F, McDonald PP, Richard DE, Dubois CM (2005) Hypoxia-enhanced expression of the proprotein convertase furin is mediated by hypoxia-inducible factor-1: impact on the bioactivation of proproteins. J Biol Chem 280:6561–6569 191. Liu M, Alkayed NJ (2005) Hypoxic preconditioning and tolerance via hypoxia inducible factor (HIF) 1alpha-linked induction of P450 2C11 epoxygenase in astrocytes. J Cereb Blood Flow Metab 25:939–948 192. Lofstedt T, Jogi A, Sigvardsson M, Gradin K, Poellinger L, Pahlman S, Axelson H (2004) Induction of ID2 expression

419 by hypoxia-inducible factor-1: a role in dedifferentiation of hypoxic neuroblastoma cells. J Biol Chem 279: 39223–39231 193. Synnestvedt K et al (2002) Ecto-5’-nucleotidase (CD73) regulation by hypoxia-inducible factor-1 mediates permeability changes in intestinal epithelia. J Clin Invest 110: 993–1002 194. Segawa K et al (2006) Visfatin in adipocytes is upregulated by hypoxia through HIF1alpha-dependent mechanism. Biochem Biophys Res Commun 349:875–882 195. Bae SK et al (2006) Hypoxic induction of human visfatin gene is directly mediated by hypoxia-inducible factor-1. FEBS Lett 580:4105–4113 196. Arany Z, Huang LE, Eckner R, Bhattacharya S, Jiang C, Goldberg MA, Bunn HF, Livingston DM (1996) An essential role for p300/CBP in the cellular response to hypoxia. Proc Natl Acad Sci U S A 93:12969–12973 197. Ema M, Hirota K, Mimura J, Abe H, Yodoi J, Sogawa K, Poellinger L, Fujii-Kuriyama Y (1999) Molecular mechanisms of transcription activation by HLF and HIF1alpha in response to hypoxia: their stabilization and redox signal-induced interaction with CBP/p300. Embo J 18:1905–1914 198. Carrero P, Okamoto K, Coumailleau P, O’Brien S, Tanaka H, Poellinger L (2000) Redox-regulated recruitment of the transcriptional coactivators CREB-binding protein and SRC-1 to hypoxia-inducible factor 1alpha. Mol Cell Bio 20:402–415 199. Zhang W, Tsuchiya T, Yasukochi Y (1999) Transitional change in interaction between HIF-1 and HNF-4 in response to hypoxia. J Hum Genet 44:293–299 200. Sanchez-Elsner T, Botella LM, Velasco B, Corbi A, Attisano L, Bernabeu C (2001) Synergistic cooperation between hypoxia and transforming growth factor-beta pathways on human vascular endothelial growth factor gene expression. J Biol Chem 276:38527–38535 201. Mahon PC, Hirota K, Semenza GL (2001) FIH-1: a novel protein that interacts with HIF-1alpha and VHL to mediate repression of HIF-1 transcriptional activity. Genes Dev 15:2675–2686 202. Fatyol K, Szalay AA (2001) The p14ARF tumor sup­ pressor protein facilitates nucleolar sequestration of hypoxia-inducible factor-1alpha (HIF-1alpha ) and inhibits HIF-1-mediated transcription. J Biol Chem 276: 28421–28429

VEGF Signaling

28

David Semela and Jean-François Dufour

Introduction Vascular endothelial growth factor (VEGF) is the main growth factor for angiogenesis and vasculogenesis. Identified as a vascular endothelial cell mitogen and survival factor, it has been sequenced and cloned by Ferrara and Connolly in 1989 [1, 2]. Intense research over the past years has deciphered the gene, molecular pathways, receptors, and functions of this angiogenic factor [3]. VEGF plays a key role in liver regeneration, hepatic fibrogenesis, portal hypertension, hepatocarcinogenesis, and malignant ascites formation.

VEGF Biological Functions of VEGF VEGF is the key angiogenic factor of developmental, physiological, and pathological angiogenesis, which is defined as formation of new microvessels from a preexisting vascular bed. VEGF is a glycoprotein which can be produced and secreted by most cells in mammals. The main target of VEGF are vascular endothelial cells. VEGF has been shown to promote proliferation [4] and survival of vascular endothelial cells in vivo and in vitro, and acts as an endothelial survival factor by inducing expression of the antiapoptotic proteins Bcl-2 and A1 in vascular endothelial cells [5–8].

J.-F. Dufour (*) Institute of Clinical Pharmacology and Visceral Research, University of Bern, Murtenstrasse 35, 3010 Bern, Switzerland e-mail: [email protected]

Furthermore, VEGF induces the expression of proteases like collagenase [9], matrix metalloproteinases [10], urokinase- and tissue-type plasminogen activators [11], which enable endothelial cells to breakdown the surrounding basal membrane and extracellular matrix in order to migrate and form new blood vessels. Initially described as vascular permeability factor (VPF) [12], VEGF increases the permeability of blood vessels up to 50,000 more than histamine [13]. Transmission and scanning electron microscopy studies showed that VEGF regulates hepatic sinusoidal permeability by inducing fenestration in hepatic sinusoidal endothelial cells possibly through caveolin-1 protein [14]. During embryonic development liver organogenesis and vasculogenesis that is defined as the de novo formation of blood vessels from hemangioblasts, are regulated through the VEGF signaling system [15–17]. The importance of VEGF signaling during embryonic development is highlighted by the fact that lack of a single VEGF gene allele results in abnormal blood vessel development and embryonic lethality in mice [15, 16]. Knockout of the genes for the VEGF receptors VEGFR-1 or -2 also results both in embryonic lethality [18, 19]. Kidney development, skeletal growth, enchondral bone formation, wound healing, and ovarian angiogenesis are the further fundamental physiological processes regulated by VEGF [20]. Elevated levels of systemically circulating VEGF recruit hematopoietic stem cells and endothelial progenitor cells from bone marrow to sites of neovascularization (reviewed in [21]). These cells home to sites of neovascularization like tumor microvasculature and the regenerating liver, where they are incorporated into the microcirculation and contribute to angiogenesis [22–25]. The level of circulating VEGF in healthy individuals is low [26]. Significant amount of VEGF are

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_28, © Springer-Verlag Berlin Heidelberg 2010

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bound to plasma proteins such as a2-microglobulin and are stored in platelets, which secrete VEGF from their a granules upon activation during blood clotting [27]. The complex biology of VEGF, its release from platelets, and the different isoforms (see below) have to be considered while choosing a VEGF detection test (free vs. total VEGF, plasma vs. serum) [26–28].

VEGF Gene and Splice Variants The human VEGF gene is localized on chromosome 6p21.3 [29] and contains eight exons (Fig. 28.1) [30]. VEGF is highly conserved across species with a homology of approximately 85% between human and rat VEGF [3]. Alternative exon splicing of the human VEGF pre-mRNA produces six VEGF isoforms containing 121, 145, 165, 183, 189, or 206 amino acids [30]. All VEGF isoforms are secreted as covalently linked homodimers but display differences in the basic amino rich domains encoded by exons 6 and 7 [30, 31]. These domains are important sites for molecular interaction and mediate binding of VEGF to heparin,

heparan sulfate proteoglycans, and to elements of the extracellular matrix, which results in sequestration of certain VEGF isoforms in the extracellular matrix or at the cell surface: VEGF121, an isoform which lacks the domains encoded by exons 6 and 7, does not bind to heparin and therefore, is freely diffusible after secretion [32]. In contrast, the highly basic isoforms, VEGF189 and VEGF206 containing exons 6 and 7, remain almost entirely bound to the extracellular matrix and cell surface [32]. VEGF165 (exon 6 absent), the most abundant isoform (46  kDa as homodimer), displays an intermediate behavior with 50–70% sequestration [32]. Sequestered VEGF can be released by heparin, heparan sulfate, and heparinase [32] and proteases like plasmin or urokinase-type plasminogen activator [33].

VEGF Protein Family VEGF is the member of a gene family of growth factors consisting of VEGF itself (also named VEGFA), VEGFB, VEGFC, VEGFD, VEGFE, and placenta growth factor (PlGF, also known as PGF, consisting of

VEGF121

VEGF145

VEGF165

VEGF183

VEGF189

VEGF206 Exons 1-5

Exon 6

Exon 7

Exon 8

(141 amino acids)

(41 aa)

(44 aa)

(6 aa)

Fig. 28.1  The six different splice variants of the VEGF (VEGFA) gene ordered by length of the amino acid sequence (single asterisk truncation in exon 6, double asterisk additional sequence encoded by exon 6)

28  VEGF Signaling Fig. 28.2  VEGF receptors (VEGFR) -1, -2, -3 and the neuropilin (NRP) receptors 1 and 2 with their function in VEGF signaling

423 VEGF A, C, D, E VEGF A, B, E VEGF A, B

VEGF C, D

PIGF

VEGF A

PIGF

VEGFR-1

VEGFR-2

HGF, IL-6 release cross talk VEGFT-2 Decoy for VEGF Induction MMP-9

VEGFR-3

Lymphangiogenesis

NRP1

NRP2

Enhance binding of VEGF to VEGFR-2

Angiogenesis

three isoforms PlGF-1, -2, and -3). They share significant sequence homology and are ligands to the same receptors as VEGF (see below and Fig. 28.2) [31]. PlGF expression is not only restricted to the placenta [34] but can be upregulated in different cell populations such as endothelial, smooth muscle, inflammatory, and malignant cells [35]. VEGFB is mainly expressed in heart and skeletal muscle [36]. VEGFC, also referred to as VEGFrelated protein [37, 38], and VEGFD [39] are involved in lymphangiogenesis [40] and fetal lung development (VEGFD) [41]. Overex­pression of VEGFC induces selective proliferation of lymphatic, but not vascular, endothelial cells and lymphatic vessel enlargement [42]. However, recent evidence suggests that VEGFC might play a role in angiogenesis by inducing fenestrations in vascular endothelial cells and by inducing blood capillaries under certain circumstances [43]. VEGFE has been found in the genome of orf virus, which affects sheep, goats, and humans [44]. It is believed that the gene has been originally acquired from a mammalian host [44]. Interestingly, skin lesions induced by orf virus show extensive dermal vascular endothelial cell proliferation and vasodilatation [44].

VEGF Gene Expression The regulatory factors for VEGF expression can be divided into two main categories: hypoxia through hypoxia-inducible factor (HIF)-1 and a group of cytokines, growth factors, and transcription factors other than HIF-1. Additionally, mutations in tumor suppressor genes and oncogenes like p53, ras, raf, VHL, myc, c-fos and others can induce and upregulate VEGF expression [45–49]. Focal gains of VEGFA (i.e., VEGFA amplification, VEGFA high level gains, and chromosme 6 gains) have been shown to contribute to the increased VEGF expression in hepatocellular carcinoma (HCC) [50]. Hypoxia Hypoxia is a potent stimulator of VEGF expression. HIF-1 is the key transcription factor in hypoxic tissues and induces the expression of several hypoxia-response genes such as VEGF and VEGFR-1 [51, 52]. The detailed molecular mechanisms of HIF-1 and the regulation of VEGF expression by HIF-1 are reviewed in Chap. 27.

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Hypoxia not only induces and upregulates the expression of VEGF but also stabilizes the intrinsically labile VEGF mRNA, which contains destabilizing elements in its 3’ and 5’ untranslated and coding regions [53–56]. The hypoxia-induced RNA-binding protein HuR binds with high affinity and specificity to regulatory elements of VEGF mRNA and prevents its degradation [57].

Cytokines and Growth Factors Many cytokines like IL-1a, IL-1b, IL-6, nitric oxide (NO), and growth factors such as fibroblast growth factor (FGF), TGF-a, TGF-b, TNF-a, platelet-derived growth factor (PDGF), epidermal growth factor (EGF), hepatocyte growth factor (HGF), and insulin-like growth factor-1 upregulate VEGF highlighting the complex regulation and redundancy of the angiogenesis network [20, 31, 49].

VEGF Receptors The effects of VEGF are mediated mainly through two cell surface receptor tyrosine kinases, namely VEGFR-1 and VEGFR-2. Neuropilin-1 (NRP1) and neuropilin-2 (NRP2) are the recently discovered VEGF receptors belonging to the Semaphorin subfamily. VEGFR-3 is a receptor for VEGFC and VEGFD but not for VEGFA. VEGFR-1, -2, and -3 belong to the flt subfamily of receptor tyrosine kinases [31] and consist of seven extracellular immunoglobulin-like domains, one transmembrane region, and a conserved tyrosine kinase domain intracellularly, which is interrupted by a kinase insert domain [58–60]. The functions of the different VEGF receptors and their ligands are summarized in Fig. 28.2.

VEGFR-1 VEGFR-1 (also known as fms-like-tyrosine kinase (Flt)-1 or FLT-1 in humans [61]) is a 180-kDa glycoprotein binding VEGF, VEGFB, and PlGF [31, 62]. In contrast to VEGFR-2, VEGFR-1 is upregulated by hypoxia through HIF-1 (see Chap. 27) [52]. It is highly

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expressed on the surface of the different endothelial cell populations (quiescent and cycling) such as endothelial cells of liver sinusoids and hepatic arterioles [63–65], but also on hepatocytes during liver regeneration [63, 66] and on few other cells such as hepatic stellate cells [67, 68], monocyte-macrophages including Kupffer cells [69, 70], pericytes [71], smooth muscle cells [72], nerve cells [73], and hematopoietic cells [20]. The exact role of VEGFR-1 is still under debate and depends on the biological situation (i.e., physiological vs. pathological angiogenesis). Although, there is a knockout of the VEGFR-1 gene resulting in the formation of abnormal vascular channels, excessive angioblast proliferation and embryonic lethality by day E8.5 in mice [18, 74], VEGFR-1 lacking only the tyrosine kinase domain is sufficient for normal development and angiogenesis [75]. In contrast, VEGFR-1 tyrosine kinase-deficient mice showed impaired angiogenesis during carcinoma growth [76]. No proliferative or migratory response in endothelial cells or hepatocytes has been attributed to VEGFR-1 [63, 77, 78]. Signaling through VEGFR-1 was shown to contribute to the regulation of endothelial cell permeability [79]. Activation of VEGFR-1 by the VEGF homolog PlGF enhances VEGF-driven angiogenesis through VEGFR-2 [35, 80] (see below). Further, VEGFR-1 is involved in the recruitment of endothelial progenitor cells [22] and bone marrow-derived myeloid progenitors [81], and promotes the survival of hematopoietic stem cells [82]. Matrix metalloproteinase 9 (MMP-9) has been shown to be induced specifically via VEGFR-1 in endothelial cells of the lung promoting pulmonary metastasis [83]. Recent evidence suggests an angiogenesis-independent endothelial protection of hepatocytes through VEGFR-1 [64]: Selective activation of liver sinusoidal endothelial cells via VEGFR-1 induces paracrine secretion of the potent mitogens HGF and interleukin 6 (IL-6, see Chap. 1), which promote hepatocyte proliferation and reduce liver damage in mice exposed to the hepatotoxin CCl4. Besides a membrane-bound form of VEGFR1, a soluble form of VEGFR1 (sVEGFR1, sFlt-1), which is produced by endothelial cells and monocytes by alternative splicing has been identified [84, 85]. sVEGFR-1 has been detected in human serum and plasma of normal male and female blood donors [85]. By binding VEGF, sVEGFR1 is a naturally occurring VEGF antagonist. The function of sVEGFR1 in

28  VEGF Signaling

physiologic angiogenesis and malignant neovascularization is unclear.

VEGFR-2 VEGFR-2 (also known as mouse fetal liver kinase (Flk)-1 or kinase domain region KDR in humans [86]) is a 200–230-kDa VEGF receptor, which shares 85% of sequence identity with VEGFR-1 [60] but binds VEGFA, VEGFC, VEGFD, and VEGFE instead of VEGFB or PlGF [31]. In contrast to the nonmitogenic and nonmotogenic VEGFR-1, VEGFR-2 induces endothelial cell proliferation, migration, and survival [4, 6, 87]. VEGFR-2 is involved in the mechanotransduction of blood flow, shear stress to the vascular endothelium by nuclear translocation of VEGFR-2 and consecutive binding to the cytoskeleton together with VE-cadherin and b-catenin [88]. VEGFR-2 is highly expressed on adult and embryonic endothelial cells, embryonic angioblasts, and hematopoietic stem cells [20, 31]. In resting liver, VEGFR-2 expression is limited to endothelial cells of the larger hepatic vessels [63], although, earlier reports described VEGFR-2 mRNA expression also in liver sinusoidal endothelial cells [65]. During liver regeneration, VEGFR-2 expression predominantly increases on endothelial cells of large vessels (portal venules, arterioles, central venules) and to a lesser extent on sinusoidal endothelial cells often in close proximity to large vessels [63]. Additionally, hepatic stellate cells have been found to express VEGFR-2 in vitro [68]. Homozygous loss of VEGFR-2 results in lack of endothelial cells, impaired liver organogenesis [17], and embryonic lethality at day E9.5–E10.5 [19], whereas heterozygous mice are normal [19]. VEGFR-2 is phosphorylated in resting liver, but has been shown to increase in activation during liver regeneration [63].

VEGFR-3 VEGFR-3 (or fms-like-tyrosine kinase (Flt)-4) is a receptor for VEGFC and VEGFD and does not bind VEGF [89, 90]. VEGFR-3 is mainly expressed on

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lymphatic endothelium and involved in lymphangiogenesis [42, 91]. VEGFR-3 could not be detected on endothelial cells in resting or regenerating liver [63].

Neuropilin-1 and -2 NRP1 and NRP2 are additional VEGF receptors unrelated to VEGFR-1, -2, and -3 [92]. They are isoformspecific receptors binding VEGF165, but not VEGF121 and are involved in neuronal guidance [93]. Coexpres­ sion of VEGFR-2 and NRP1 enhances binding of VEGF to VEGFR-2 [94]. Neuropilins can form complexes with VEGFR-1 and probably with VEGFR-2 [94]. No intrinsic receptor signaling has been shown after binding of VEGF to NRP1 or NRP2 [94]. To date, nothing is known about the presence or function of NRP1 and NRP2 in liver biology.

VEGF Receptor Signaling After binding, soluble VEGF dimers induce VEGF receptor dimerization leading to homo- or heterodimers [80]. The juxtapose cytoplasmic tyrosine kinase domains of the VEGFR molecules, transphosphorylate several tyrosine residues in the neighbor molecule [62, 80, 95, 96]. Activated receptors in turn activate proteins of different signaling pathways by phosphorylation (reviewed in [96]): phospholipase C (PLC), phosphatidylinositol 3’-kinase (PI-3 kinase)/Akt, Ras GTPaseactivating protein (GAP), Src family kinases and Raf [6, 97–99]. A recent study showed that although both VEGF and PlGF bind VEGFR-1, they activate this receptor differently leading to a distinct gene expression profile, where only PlGF was capable of switching on downstream target genes [80]. Many of these PlGFregulated genes have a role in cell cycle (Ets2, Map4k4, Fst, Jak2, Egr1), angiogenesis (Flt-1, NRP2, Angptl4, Dcn), and apoptosis (Birc2) [80]. The same authors were able to show that there is an intra- and intermolecular cross talk between VEGFR-1 and VEGFR-2: activation of VEGFR-1 by PlGF resulted in intermolecular transphosphorylation of VEGFR-2, thereby amplifying VEGF-driven angiogenesis through VEGFR-2 [80]. Further, VEGF/PlGF heterodimers

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activated intramolecular VEGF receptor cross talk through formation of VEGFR-1/VEGFR-2 heterodimers [80].

VEGF Signaling in Specific Liver Conditions Liver Organogenesis During embryonic development of the liver, hepatic cells are induced within the endoderm by day E8.5 of mouse gestation [100]. Interaction between these cells and surrounding endothelial cells or angioblasts induces outgrowth of the liver bud into the mesenchyma [17]. This stage is then followed by the formation of a de novo local vascular network (vasculogenesis) and the recruitment of hematopoietic cells [17]. Mice with homozygous deficiency in VEGFR-2 lacking mature endothelial cells and blood vessels show normal thickening of the hepatic endoderm but lack liver bud emergence [17]. Vasculogenic endothelial cells with intact signaling through VEGFR-2 are therefore critical already in the earliest stages of liver organogenesis, even prior to blood vessel function [17]. In addition to the formation of the liver sinusoidal vascular network, VEGF signaling is also fundamental for the development of the peribiliary vascular plexus. It has been shown that the developing bile ducts drive arterial development in the liver [101]. Thereby, hepatic artery branches are formed in close proximity with the ductal plates, which are the precursors of the intrahepatic bile duct. Similar to hepatocytes, which secrete VEGF for liver sinusoidal endothelial cells, cholangiocytes secrete VEGF for peribiliary endothelial cells [101].

Liver Regeneration Liver regeneration is angiogenesis-dependent [102]; inhibition of angiogenesis by antiangiogenic substances such as angiostatin or TNP-470 impairs liver regeneration [102, 103]. In the course of liver regeneration after hepatectomy, there is an initial wave of  hepatocyte proliferation with the formation of

D. Semela and J.-F. Dufour

avascular hepatocellular islands [63]. This stage is followed by endothelial cell proliferation [104] and consecutive penetration of these avascular islands by endothelial cells with the formation of new sinusoids [105]. Recent evidence suggests that endothelial progenitor cells are mobilized from bone marrow and participate in this neovascularization by committing to sinusoidal endothelial cells [25]. HIF-1a and VEGF as key regulators have been shown to be upregulated during this process [105, 106]: hepatocellular production of VEGF peaks 48–72  h after hepatectomy and is detected mainly in periportal hepatocytes [105, 107]. VEGF production is accompanied by an increase in the expression of VEGFR-1 on hepatocytes and of VEGFR-1 and VEGFR-2 on sinusoidal endothelial cells [63, 66, 104, 106]. Administration of VEGF in hepatectomized rodents increases hepatocyte and sinusoidal endothelial cell proliferation [107, 108], accelerates gain in liver mass [64], and improves functional hepatic recovery [106]. Neutralizing antibodies against VEGF inhibit hepatocyte and endothelial cell proliferation after partial hepatectomy [107]. LeCouter and coworkers showed that this effect is likely due to a VEGFinduced release of HGF by sinusoidal endothelial cells [64, 65]. Additional effects of VEGF on functional hepatic recovery could be a stimulatory effect on the formation of new blood vessels and/or a direct effect on the hepatocytes, which express VEGF receptors after partial hepatectomy [63, 106]. Transduction of VEGF before hepatic resection also hastens functional hepatic recovery in mice with fatty liver, which is known for its impaired regenerative capacity [106].

Liver Fibrosis and Cirrhosis Accumulating evidence suggests that VEGF and its receptors are also involved in fibrogenesis and cirrhotic remodeling. The concept of an interplay between fibrosis and angiogenesis probably mediated through the stimulus of hypoxia is supported by two facts: First, the deposited matrix proteins – a hallmark of fibrosis – contain and sequester different angiogenic factors such as VEGF, which are liberated during remodeling of the connective tissue framework by proteolytic enzymes such as matrix metalloproteinases

28  VEGF Signaling

[109, 110]. Second, in the fibrotic tissue around regenerative nodules develops an abundant vascular network [111, 112] leading to a remodeled and abnormal hepatic microcirculation in cirrhotic liver [113, 114]. Experimental liver fibrogenesis after common bile duct ligation in rats induces VEGF expression in hepatocytes and angiogenesis [115]. The authors of this study showed that this process is hypoxia-driven and that the percentage of VEGF expressing hepatocytes increases from 3% at the time of ligation to >95% 2  weeks after ligation. The upregulation of VEGF is followed by vascular endothelial cell proliferation and angiogenesis in fibrotic areas by week 3 [115]. A similar animal study with diethylnitrosamineinduced cirrhosis confirmed the concept that hepatocellular hypoxia and angiogenesis progress together with fibrogenesis after liver injury [116]. Using neutralizing monoclonal antibodies of VEGFR-1 and VEGFR-2 in murine CCl4-induced liver fibrosis both significantly attenuated the development of fibrosis and suppressed neovascularization in the liver [117]. Fibrosis markers (hepatic hydroxyproline, serum hyaluronic acid, and procollagen III-N-peptide), the number of smooth muscle actin positive cells and procollagen mRNA expression were also suppressed by this treatment [117]. The inhibitory effect of the anti-VEGFR-2 antibody was more potent than that of anti-VEGFR-1 and combination treatment with both almost completely attenuated fibrosis development [117]. Hypoxia directly contributes to the progression of liver fibrosis by inducing the expression of VEGF, VEGFR-1, and type I collagen in activated hepatic stellate cells, the key player in the pathogenesis of hepatic fibrosis [116, 118, 119]. VEGF increases a1(I)-procollagen mRNA expression and stimulates proliferation of activated hepatic stellate cells [117]. VEGF expression in hypoxic hepatic stellate cells has been shown to be mediated by cyclooxygenase-2 (COX-2) protein and COX-2 inhibitors significantly blocked VEGF production via the HIF-1a pathway [118]. VEGF signaling through VEGFR-1 was shown to inhibit hepatic stellate cell contraction probably through attenuation of smooth muscle a-actin expression [68]. In vitro studies have shown that a basal expression of VEGF is essential for VEGF-stimulated NO by liver sinusoidal endothelial cells to prevent hepatic stellate cell activation or promote reversion of activated hepatic stellate cells to quiescence [120].

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Portal Hypertension A major complication of chronic liver diseases is portal hypertension, which is a consequence of hyperdynamic circulation and increased vascular resistance in the liver. The development of portal-systemic collateral vessels in portal hypertension is classically explained as a mechanical consequence of increased portal pressure with subsequent opening of collateral vessels [121]. Recent evidence suggests that active, VEGFdependent angiogenesis is also involved in this process: VEGF, VEGFR-2, and CD31 protein levels in splanchnic organs increased after partial portal vein ligation in mice in a time-dependent fashion during the evolution of portal hypertension [122]. A monoclonal antibody against VEGFR-2 given to these animals after ligation decreased the expression of VEGFR-2 and CD31 significantly, and inhibited the formation of portal-systemic collateral vessels measured with labeled microspheres [122]. Experiments using partial portal vein-ligated rats and a VEGFR-2-specific tyrosine kinase inhibitor (SU5416) confirmed that the formation of portal-systemic collateral vessels is an angiogenesis-dependent process which can be inhibited by antagonization of the VEGF/VEGFR-2 signaling pathway [122]. Similar results were found using different animal models of portal hypertension and different anti-VEGF strategies: the multikinase inhibitor sunitinib decreased portal pressure in rats with CCl4induced cirrhosis [123] as well as the mTOR inhibitor sirolimus (rapamycin) after partial portal vein ligation in rats [124].

Viral Hepatitis Hepatitis B virus X protein (HBx) is a hepatitis B virusencoded transcriptional activator, which is involved in hepatocarcinogenesis and hypoxia-induced angiogenesis [125]. Recent studies have shown that HBx protein stabilizes HIF-1a and enhances transcriptional activity of HIF-1a through activation of mitogen-activated protein kinase (MAPK) pathway under normoxic and hypoxic conditions [126, 127]. The expression of HIF1a and VEGF was increased in the liver of HBxtransgenic mice [126] and in HBx-transfected HCC cell lines [125, 128]. Immunohistochemical staining for

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VEGF in chronic hepatits B correlated with the degree of injury (grade) and amount of fibrosis (stage) [129]. Patients with chronic hepatitis C infection were shown to have elevated levels of serum VEGF [130]. Antiviral treatment with ribavirin and pegylated interferon a-2b decreased VEGF levels. Furthermore, in biopsies from patients with chronic hepatitis C, an increase in VEGF expression and angiogenesis in portal tracts has been described [131]. In vitro studies showed that infection of human Huh-7 cells with hepatitis C virus (HCV) stimulates the expression of VEGF by stabilization of HIF-1a [132]. HCV core protein has been found to activate expression of VEGF in HepG[2] cells [133].

Hepatocellular Carcinoma Tumor growth beyond the size of 1-2 mm3 requires the formation of new blood vessels in order to supply the malignant tissue with nutrients and oxygen [134, 135]. Central hypoxia is thought to be the main driving force of tumor angiogenesis and upregulates proangiogenic growth factors like VEGF (Fig. 28.3) [51]. Therefore, it is not surprising that VEGF is upregulated in most human tumors and that direct correlation with intratumoral microvessel density exists [4]. The angiogenic switch, which describes the acquisition of the capacity to stimulate angiogenesis by shifting the balance between stimulatory and inhibitory factors of angiogenesis towards proangiogenic factors, is a rate-limiting step in tumoral development [136]. VEGF and other growth factors promote survival, proliferation, and migration of endothelial cells, which will finally result in the formation of new tumoral blood vessels enhancing further tumor growth. These growth factors are secreted by neoplastic cells, adjacent stroma, hepatocytes, stellate cells, and tumor-infiltrating inflammatory cells [136–139]. Besides hypoxia, mutations in tumor suppressor genes and oncogenes and certain viral proteins are also involved in the upregulation of VEGF during hepatocarcinogenesis (see above). In vitro studies have shown that different inflammatory cytokines (interleukin-1b, interferon-a, interferon-g, tumor necrosis factor-a) and growth factors (epidermal growth factor, platelet-derived growth factor, basic fibroblast growth factor, transforming growth factor-a) increase the secretion of VEGF in HCC cell lines [140].

Fig. 28.3  VEGF signaling and its effect on tumoral endothelial cells and HCC growth

HCC is a hypervascular tumor [140–142] and arterial hypervascularization on imaging studies is one of the diagnostic criteria for HCC [143]. Several studies report on the overexpression of VEGF in HCC [127, 140, 144–151]. FISH analyzes in tissue microarrays of the chromosomal locus 6p21 of VEGFA showed increased VEGF expression by focal gains in a third of 210 human HCCs [50]. Grafting HCC tissue onto chick embryo chorioallantoic membrane, which is a classical angiogenesis assay, stimulates neovascularization [152]. It has been shown that during hepatocarcinogenesis expression of VEGF increases gradually from lowgrade dysplastic nodules to high-grade dysplastic nodules to early HCC [139]. The degree of VEGF expression during development of HCC correlates with the density of vessels, unpaired arteries (i.e., arteries not accompanied by bile ducts, indicative of tumor angiogenesis), CD34 staining (as a marker of sinusoidal capillarization), and the proliferation of hepatocytes assessed by

28  VEGF Signaling

staining with PCNA [139]. Tumor expression of VEGF (mRNA and protein expression) significantly correlated with serum VEGF level per platelet in patients with HCC providing the basis for using circulating VEGF as a prognostic marker [153]. Small HCCs showed a higher status of neoangiogenesis and cell proliferation activity than advanced HCCs [139]. Circulating concentration of VEGF increases with the stage of HCC, the highest levels being in patients with metastasis [154]. A prospective study of 100 patients suffering from HCC found that high serum levels of VEGF significantly correlated with the absence of tumor capsule, presence of intrahepatic metastasis, microscopic venous invasion, advanced stage, and postoperative recurrence [155]. Similar results have been found using serum VEGF per platelet count in 52 HCC patients [156]. In a recent study, preoperative serum VEGF in 98 patients with resectable HCC was a significant and independent predictor of tumor recurrence, disease-free survival and overall survival [150]. In 80 patients with inoperable HCC undergoing transarterial chemoembolization (TACE), Poon and colleagues evaluated the prognostic significance of serum VEGF levels prospectively [157]: pretreatment serum VEGF levels were significantly higher in patients with progressive disease than those with stable or responsive disease. Patients with serum VEGF >240 pg/ml had significantly worse survival than those with serum VEGF <240  pg/ml (median survival 6.8 vs. 19.2  months, p = 0.007). In a Cox multivariate analysis, serum VEGF >240  pg/ml was an independent prognostic factor of survival [157]. In a phase III trial, the multityrosine kinase inhibitor sorafenib (which inhibits VEGFR-1, -2, -3 among others) has been shown to prolong survival in patients with advanced HCC [158]. This study showed for the first time that systemic therapy is effective in case of advanced HCC and that patients can benefit from antiangiogenic treatment. Several other tyrosine kinase inhibitors with antiangiogenic profile are currently being tested in patients suffering from HCC [159]. Expression of VEGF in patients and animals with HCC increased significantly after transcatheter arterial chemoembolization (TACE) [160–162]. VEGF antisense oligodeoxynucleotides mixed with lipiodol inhibited HCC growth in rats significantly more than arterial embolization with lipiodol alone [163]. These studies suggest that VEGF signaling pathway

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plays an important role in tumor response to hypoxia after arterial embolization treatment and that antiVEGF strategies such as tyrosine kinase inhibitors might enhance the efficacy of arterial embolization in HCC [164].

Ascites Formation In 1983 Senger and coworkers reported that HCC cells secrete a vascular permeability factor (VPF, later named VEGF) which promotes accumulation of ascites fluid [12]. Ascites, which is defined as accumulation of excess fluid within the peritoneal cavity is encountered in many patients with cirrhosis, other forms of liver disease and malignancies. The pathogenesis of ascites depends on the underlying disease. An important component in the formation of ascites in patients with malignancies of the liver and abdominal cavity is microvascular hyperpermeability of tumor vessels due to tumor-secreted VEGF with consecutive extravasation of plasma and plasma proteins. In fact, levels of biologically active VEGF in patients with malignant ascites are higher in comparison to patients with ascites due to nonmalignant or cirrhotic causes [165–168]. Four pathways for macromolecular extravasation have been described: endothelial fenestrae [169, 170], interendothelial cell gaps [171], transendothelial cell pores [172], and vesiculo-vacuolar organelles [173, 174]. VEGF has been shown to induce fenestration and increase permeability in normal and tumoral microvascular endothelium [14, 43, 169, 170, 175]: within 10  min of VEGF application fenestrations appear even in vascular beds which do not have fenestrated endothelium under physiological circumstances [169]. Liver sinusoidal endothelial cells incubated with 100 ng VEGF/ml increased the number of fenestrations and cell pores in vitro [14, 176]. Neutralizing antibodies against VEGF or against VEGFR-2 significantly suppressed the volume of ascites, the number of tumor cells in ascites and the peritoneal capillary permeability and prolonged the survival of ascites-bearing mice suffering from HCC [177]. Soluble VEGFR-1 and VEGF-trap (both soluble decoy receptors for endogenous VEGF), monoclonal antibody against VEGF or VEGFR-2, and the VEGF receptor tyrosine kinase inhibitor valatinib prevented the formation of

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malignant ascites in animal tumor models [178–182]. Similarly, inhibition of VEGFR-2 by the tyrosine kinase inhibitor sunitinib reduced development of ascites in cirrhotic rats [183]. These results suggest that the VEGF/VEGFR-2 interaction is a major regulator of ascites formation. Interestingly, ascites VEGF levels are higher in cirrhotic patients with spontaneous bacterial peritonitis, which is a frequent complication of cirrhotic patients with ascites, than in noninfected cirrhotic patients [184]. VEGF is thereby produced in peritoneal macrophages of cirrhotic patients and is markedly upregulated by bacterial lipopolysaccharide and cytokines like interleukin-1 (IL-1) [184].

Liver Transplantation Hepatocyte and endothelial cell damage due to ischemia/reperfusion injury in liver transplantation after cold preservation is an important determinant of graft function. Activation of sinusoidal endothelial cells by cold ischemia alters expression of different adhesion molecules and sequesters leukocytes and platelets during reperfusion leading to microcirculatory disturbance and liver injury. VEGF expression is upregulated in hepatocytes of rat livers preserved in University of Wisconsin (UW) solution for orthotopic liver transplantation probably due to hypoxic stress [185]. VEGF is expressed and released in a biphasic pattern by Kupffer cells and hepatocytes during the early postoperative period after transplantation in a syngeneic rat orthotopic liver transplantation model [186]. Anti-VEGF antibody treatment, administered during reperfusion, decreased the degree of damage (measured as liver function tests, lipid peroxidation, and metalloproteinase activity), suggesting that VEGF may have a role in ischemia/reperfusion injury to liver grafts [186]. The anti-VEGF antibody bevacizumab was reported to induce regression of hepatic vascular malformation in a patient suffering from hereditary hemorrhagic telangiectasia, which prevented the need for liver transplantation [187]. Sirolimus (also known as rapamycin) is a potent immunosuppressive drug used after liver transplantation. Sirolimus has been shown to downregulate VEGF expression and is often used in patients with HCC recurrence after liver transplantation to prevent tumor progression [188–190].

D. Semela and J.-F. Dufour

Summary

›› Alternative splicing of the human VEGF pre-

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›› ›› ››

mRNA produces six VEGF isoforms, which differ in their sequestration in the extracellular matrix or at the cell surface. VEGF expression is stimulated not only by hypoxia-inducibe factor, but also by many cytokines and growth factors. Besides a membrane-bound form of VEGR1, a soluble form of VEGFR1, which is produced by endothelial cells and monocytes, acts as a naturally occurring VEGF antagonist. In contrast to the non-mitogenic and nonmotogenic VEGFR-1, VEGFR-2 induces endothelial cell proliferation, migration, and survival. VEGF signaling is necessary for liver organogenesis, liver regeneration, and development of portal hypertension. VEGF expression levels may be useful as prognostic marker in HCC patients and as predictor of tumor response to treatment. Recent approval of angiogenesis inhibitor drugs such as sorafenib and bevacizumab have opened a wide range of clinical applications for VEGF-based strategies in liver diseases (fibrosis, cancer).

Multiple Choice Questions 1. VEGFR-1 is expressed on except: (a) Quiescent endothelial cells (b) Quiescent hepatocytes (c) Hepatic stellate cells (d) Kupffer cells (e) Smooth muscle cells 2. Regarding the role of VEGF during liver regeneration, which statement is wrong: (a) VEGF peaks in the first 6 h following hepatectomy (b) VEGF is expressed mainly in periportal hepato­ cytes (c) The expression of VEGFR-1 on hepatocytes increases

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(d) Administration of VEGF after hepatectomy hastens gain in liver mass (e) Inhibition of angiogenesis impairs liver regenera­ tion 3. During hepatic fibrosis, VEGF (a) Is degraded by metalloproteinases (b) Is liberated from the extracellular matrix during remodeling (c) Is promoting fibrosis (d) Is not expressed in stellate cells (e) Is increased by COX2 inhibitors 4. VEGF plays an important role in portal hypertension; which statement is wrong: (a) Formation of collaterals can be prevented by antagonizing the VEGF/VEGFR-2 signaling pathway (b) The multikinase inhibitor sunitinib decreases portal pressure in experimental cirrhosis (c) Inhibition of mTOR decreases portal pressure in experimental cirrhosis (d) The antibody against VEGF, bevacizumab has been associated to variceal bleeding in patients with portal hypertension (e) The expression of VEGFR-2 is decreased in splanchnic organs during the development of portal hypertension 5. It has been found that VEGF in patients with HCC: (a) Has an expression which correlates with micro­ vessel density (b) Has an tumor expression which correlates with serum VEGF (c) Circulating levels of VEGF have a prognostic significance (d) Circulating levels of VEGF are increasing after trans-arterial chemoembolisation (e) All the above statements are correct

Selected Reading 1. Ferrara N, Gerber HP, LeCouter J (2003) The biology of VEGF and its receptors. Nat Med 9:669–676 (this review provides a detailed and comprehensive description of the different VEGF isoforms and receptors and discusses their role under physiological and pathological conditions [20]) 2. Shibuya M, Claesson-Welsh L (2006) Signal transduction  by  VEGF receptors in regulation of angiogenesis and ­lymphangiogenesis. Exp Cell Res 312:549–560 (this review

431 o­ utlines the current knowledge about the signal transduction properties of the different VEGF receptors [191]) 3. Ellis LM, Hicklin DJ (2008) VEGF-targeted therapy: mechanisms of antitumor activity. Nat Rev Cancer 8:579–591 (this article reviews the recent advances and mechanisms of VEGF-targeted therapies in tumors [192]) 4. LeCouter J, Moritz DR, Li B et al (2003) Angiogenesisindependent endothelial protection of liver: role of VEGFR1. Science 299:890–893 (describes the paracrine cross talk between hepatocytes and liver sinusoidal endothelial cells during hepatocyte growth [64]) 5. http://www.nature.com/focus/angiogenesis/ (this joint web focus on angiogenesis with a special section on VEGF signaling is a project between the journals Nature Medicine and Nature Reviews Cancer. The web site provides review articles and a selection of “classic” papers nominated by experts in the field of angiogenesis)

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D. Semela and J.-F. Dufour 152. Marzullo A, Vacca A, Roncali L, Pollice L, Ribatti D (1998) Angiogenesis in hepatocellular carcinoma: an experimental study in the chick embryo chorioallantoic membrane. Int J Oncol 13:17–21 153. Poon RT-P, Lau CP-Y, Cheung S-T, Yu W-C, Fan S-T (2003) Quantitative correlation of serum levels and tumor expression of vascular endothelial growth factor in patients with hepatocellular carcinoma. Cancer Res 63:3121–3126 154. Jinno K, Tanimizu M, Hyodo I, Nishikawa Y, Hosokawa Y, Doi T, Endo H et al (1998) Circulating vascular endothelial growth factor (VEGF) is a possible tumor marker for metastasis in human hepatocellular carcinoma. J Gastroenterol 33:376–382 155. Poon RT, Ng IO, Lau C, Zhu LX, Yu WC, Lo CM, Fan ST et al (2001) Serum vascular endothelial growth factor predicts venous invasion in hepatocellular carcinoma: a prospective study. Ann Surg 233:227–235 156. Kim SJ, Choi IK, Park KH, Yoon SY, Oh SC, Seo JH, Choi CW et al (2004) Serum vascular endothelial growth factor per platelet count in hepatocellular carcinoma: correlations with clinical parameters and survival. Jpn J Clin Oncol 34:184–190 157. Poon R, Lau C, Yu W, Fan S, Wong J (2004) High serum levels of vascular endothelial growth factor predict poor response to transarterial chemoembolization in hepatocellular carcinoma: a prospective study. Oncol Rep 11: 1077–1084 158. Llovet JM, Ricci S, Mazzaferro V, Hilgard P, Gane E, Blanc JF, de Oliveira AC et al (2008) Sorafenib in advanced hepatocellular carcinoma. N Engl J Med 359:378–390 159. Llovet JM, Bruix J (2008) Molecular targeted therapies in hepatocellular carcinoma. Hepatology 48:1312–1327 160. Liao X, Yi J, Li X, Yang Z, Deng W, Tian G (2003) Expression of angiogenic factors in hepatocellular carcinoma after transcatheter arterial chemoembolization. J Huazhong Univ Sci Technolog Med Sci 23:280–282 161. Suzuki H, Mori M, Kawaguchi C, Adachi M, Miura S, Ishii H (1999) Serum vascular endothelial growth factor in the course of transcatheter arterial embolization of hepatocellular carcinoma. Int J Oncol 14:1087–1090 162. Li X, Feng G, Zheng C, Zhuo C, Liu X (2003) Influence of transarterial chemoembolization on angiogenesis and expression of vascular endothelial growth factor and basic fibroblast growth factor in rat with Walker-256 transplanted hepatoma: An experimental study. World J Gastroenterol 9:2445–2449 163. Wu H, Feng G, Liang H, Zheng C, Li X (2004) Vascular endothelial growth factor antisense oligodeoxynucleotides with lipiodol in arterial embolization of liver cancer in rats. World J Gastroenterol 10:813–818 164. Strebel BM, Dufour JF (2008) Combined approach to hepatocellular carcinoma: a new treatment concept for nonresectable disease. Expert Rev Anticancer Ther 8:1743–1749 165. Kraft A, Weindel K, Ochs A, Marth C, Zmija J, Schumacher P, Unger C et al (1999) Vascular endothelial growth factor in the sera and effusions of patients with malignant and nonmalignant disease. Cancer 85:178–187 166. Zebrowski BK, Liu W, Ramirez K, Akagi Y, Mills GB, Ellis LM (1999) Markedly elevated levels of vascular endothelial growth factor in malignant ascites. Ann Surg Oncol 6:373–378

28  VEGF Signaling 167. Verheul HMW, Hoekman K, Jorna AS, Smit EF, Pinedo HM (2000) Targeting vascular endothelial growth factor blockade: ascites and pleural effusion formation. Oncologist 5:45–50 168. Dong W, Sun X, Yu B, Luo H, Yu J (2003) Role of VEGF and CD44v6 in differentiating benign from malignant ascites. World J Gastroenterol 9:2596–2600 169. Roberts WG, Palade GE (1995) Increased microvascular permeability and endothelial fenestration induced by vascular endothelial growth factor. J Cell Sci 108(pt 6): 2369–2379 170. Roberts WG, Palade GE (1997) Neovasculature induced by vascular endothelial growth factor is fenestrated. Cancer Res 57:765–772 171. Hirata A, Baluk P, Fujiwara T, McDonald DM (1995) Location of focal silver staining at endothelial gaps in inflamed venules examined by scanning electron microscopy. Am J Physiol 269:L403–L418 172. Neal CR, Michel CC (1995) Transcellular gaps in microvascular walls of frog and rat when permeability is increased by perfusion with the ionophore A23187. J Physiol 488(Pt 2):427–437 173. Kohn S, Nagy JA, Dvorak HF, Dvorak AM (1992) Pathways of macromolecular tracer transport across venules and small veins. Structural basis for the hyperpermeability of tumor blood vessels. Lab Invest 67:596–607 174. Feng D, Nagy J, Dvorak A, Dvorak H (2000) Different pathways of macromolecule extravasation from hyperpermeable tumor vessels. Microvasc Res 59:24–37 175. Grunstein J, Roberts WG, Mathieu-Costello O, Hanahan D, Johnson RS (1999) Tumor-derived expression of vascular endothelial growth factor is a critical factor in tumor expansion and vascular function. Cancer Res 59:1592–1598 176. Funyu J, Mochida S, Inao M, Matsui A, Fujiwara K (2001) VEGF can act as vascular permeability factor in the hepatic sinusoids through upregulation of porosity of endothelial cells. Biochem Biophys Res Commun 280:481–485 177. Yoshiji H, Kuriyama S, Hicklin D, Huber J, Yoshii J, Ikenaka Y, Noguchi R et al (2001) The vascular endothelial growth factor receptor KDR/Flk-1 is a major regulator of malignant ascites formation in the mouse hepatocellular carcinoma model. Hepatology 33:841–847 178. Stoelcker B, Echtenacher B, Weich H, Sztajer H, Hicklin D, Mannel D (2000) VEGF/Flk-1 interaction, a requirement for malignant ascites recurrence. J Interferon Cytokine Res 20:511–517 179. Shibuya M, Luo J, Toyoda M, Yamaguchi S (1999) Involvement of VEGF and its receptors in ascites tumor formation. Cancer Chemother Pharmacol 43 Suppl:S72–S77 180. Mesiano S, Ferrara N, Jaffe RB (1998) Role of vascular endothelial growth factor in ovarian cancer: inhibition of ascites formation by immunoneutralization. Am J Pathol 153:1249–1256

437 181. Byrne AT, Ross L, Holash J, Nakanishi M, Hu L, Hofmann JI, Yancopoulos GD et al (2003) Vascular endothelial growth factor-trap decreases tumor burden, inhibits ascites, and causes dramatic vascular remodeling in an ovarian cancer model. Clin Cancer Res 9:5721–5728 182. Xu L, Yoneda J, Herrera C, Wood J, Killion J, Fidler I (2000) Inhibition of malignant ascites and growth of human ovarian carcinoma by oral administration of a potent inhibitor of the vascular endothelial growth factor receptor tyrosine kinases. Int J Oncol 16:445–454 183. Melgar-Lesmes P, Tugues S, Ros J, Fernandez-Varo G, Morales-Ruiz M, Rodes J, Jimenez W (2009) Vascular endothelial growth factor and angiopoietin-2 play a major role in the pathogenesis of vascular leakage in cirrhotic rats. Gut 58:285–292 184. Perez-Ruiz M, Ros J, Morales-Ruiz M, Navasa M, Colmenero  J, Ruiz-del-Arbol L, Cejudo P et al (1999) Vascular endothelial growth factor production in peritoneal macrophages of cirrhotic patients: regulation by cytokines and bacterial lipopolysaccharide. Hepatology 29: 1057–1063 185. Arai S, Mochida S, Ohno A, Ishikawa K, Matsui A, Arai M, Shibuya M et al (1999) Decreased expression of receptors for vascular endothelial growth factor and sinusoidal endothelial cell damage in cold-preserved rat livers. Transplant Proc 31:2668–2672 186. Boros P, Tarcsafalvi A, Wang L, Megyesi J, Liu J, Miller CM (2001) Intrahepatic expression and release of vascular endothelial growth factor following orthotopic liver transplantation in the rat. Transplantation 72:805–811 187. Mitchell A, Adams LA, MacQuillan G, Tibballs J, Vanden Driesen R, Delriviere L (2008) Bevacizumab reverses need for liver transplantation in hereditary hemorrhagic telangiectasia. Liver Transpl 14:210–213 188. Cho ML, Cho CS, Min SY, Kim SH, Lee SS, Kim WU, Min DJ et al (2002) Cyclosporine inhibition of vascular endothelial growth factor production in rheumatoid synovial fibroblasts. Arthritis Rheum 46:1202–1209 189. Guba M, von Breitenbuch P, Steinbauer M, Koehl G, Flegel  S, Hornung M, Bruns CJ et al (2002) Rapamycin inhibits primary and metastatic tumor growth by antiangiogenesis: involvement of vascular endothelial growth factor. Nat Med 8:128–135 190. Luan FL, Ding R, Sharma VK, Chon WJ, Lagman M, Suthanthiran M (2003) Rapamycin is an effective inhibitor of human renal cancer metastasis. Kidney Int 63: 917–926 191. Shibuya M, Claesson-Welsh L (2006) Signal transduction by VEGF receptors in regulation of angiogenesis and lymphangiogenesis. Exp Cell Res 312:549–560 192. Ellis LM, Hicklin DJ (2008) VEGF-targeted therapy: mechanisms of anti-tumour activity. Nat Rev Cancer 8: 579–591

Apoptosis and Mitochondria

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Jose C. Fernández-Checa and Carmen Garcia-Ruiz

Introduction Among the various recognized forms of cell death that include necrosis and autophagy, apoptosis or programmed cell death is evolutionarily conserved, highly organized, and characterized by unique nuclear changes, chromatin shrinkage, DNA fragmentation, membrane blebbing, and formation of apoptotic bodies that contain components of the dying cell. Apoptosis is a crucial component of life that eliminates unwanted cells and is vital for embryonic development, homeostasis, and immune defense. Dysregulation of apoptosis underlies many pathophysiological states and diseases. The key mediators of apoptotic cell death are cysteine proteases, called caspases, that work in a coordinated cascade to cleave key substrates and dismantle the cell [1]. The caspase cascade involves “initiator” caspases and “executioner” caspases that can be activated in different ways by different apoptotic stimuli. While changes in nuclei are characteristic in apoptotic cell death, other subcellular organelles are also involved such as endoplasmic reticulum, lysosomes, and, particularly, mitochondria. Moreover, although caspases are crucial in apoptosis, similar morphologic changes can be produced in a caspase-independent fashion. In vertebrates, caspase-dependent apoptosis occurs through two main pathways, the extrinsic pathway and the intrinsic pathway (Fig. 29.1). The extrinsic

J. C. Fernández-Checa (*) Liver Unit and Centro de Investigaciones Biomédicas Esther Koplowitz, IMDiM, Hospital Clínic i Provincial and CIBEREHD, IDIBAPS, C/Villarroel 170, 08036-Barcelona, Spain e-mail: [email protected]

pathway is initiated upon the binding of an extracellular ligand to transmembrane death receptors of the TNF superfamily (see below), which leads to the assembly of the death-inducing signaling complex (DISC). The DISC then activates an initiator caspase, which triggers the enzymatic cascade that leads to apoptotic death. The intrinsic pathway, also known as the mitochondrial pathway, is activated by stimuli that lead to the permeabilization of the outer mitochondrial membrane (OMM) and the subsequent release of proteins from the mitochondrial intermembrane space (IMS), which initiate or regulate caspase activation, such as cytochrome c. Cytochrome c normally resides within the cristae of the inner mitochondrial membrane (IMM) and is effectively sequestered by narrow cristae junctions. Within the IMM, cytochrome c participates in the mitochondrial electron-transport chain, using its heme group as a redox intermediate to shuttle electrons between complex III and complex IV. However, when the cell detects an apoptotic stimulus, such as DNA damage, or metabolic stress, the intrinsic apoptotic pathway is triggered and mitochondrial cytochrome c is released into the cytosol. This process is thought to occur in two phases, first the mobilization of cytochrome c and then its translocation through permeabilized OMM. In addition to cytochrome c, other IMS proteins are mobilized and released into the cytosol where they are engaged in a strategic battle to promote or counteract caspase activation and hence cell death. In this chapter, we will examine signaling pathways involved in the release of deadly proteins from mitochondria and the relevance of this process to liver diseases. Moreover, although proteins interacting with mitochondria have been a major focus in the field and are considered as the major regulators of OMM permeabilization, we will present evidence for a role of lipids as death effectors and as regulators of

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_29, © Springer-Verlag Berlin Heidelberg 2010

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440 Extrinsic (receptor-mediated) Death ligand FADD TRADD

CD95L TRAIL TNF-a

Intrinsic (mitochondria-mediated)

Death receptor

Adaptor

DISC Procaspase-8

TRADD tBid

Caspase-8

Bax

Bcl-2

Bak

BcI-xL

Bid Caspase-9

Procaspase-9

Cytochrome c Smac/DIABLO

Caspase-3 Apaf- 1 Death substrates

IAP

Cell death

Fig. 29.1  Extrinsic and intrinsic death pathways. Extrinsic pathway of cell death is executed mainly by the binding of specific ligands to death receptors, which leads to the recruitment of adaptor proteins resulting in the activation of caspases. Both

pathways cross-talk by the activation and translocation of BH3only Bid which facilitates the activation of Bax to cause OMM permeabilization and release of proteins secured in the intermembrane space of mitochondria

mitochondrial-dependent apoptosis and highlight their potential involvement in acute and chronic liver diseases.

Fas and TNF are recognized to mediate several forms of liver injury and diseases, such as alcohol-induced liver damage, non-alcoholic steatohepatitis, or ischemia/reperfusion (I/R) liver injury. Indeed, Fas was shown to play a minor role in I/R liver injury as opposed to TNF [4]. Fas, however, is expressed in many liver cell types including hepatocytes, cholangiocytes, activated stellate cells, and Kupffer cells and it is central to liver pathobiology [3]. The TNF superfamily of cytokines comprises 19 members and the corresponding TNF receptors (TNFR) superfamily includes 23 related receptors. A subgroup of this family includes the death receptors, TNFR1, CD95, the TNF-related apoptosis-inducing ligand (TRAIL) receptors (TRAIL R1, also known as DR4 and TRAIL R2 also known as DR5), DR3, DR6, and p75NTR. These receptors share the “death domain,” a conserved 80-amino-acid sequence in the cytoplasmic tail that is necessary for the direct activation of the apoptotic programme by some of these receptors (TNFR1, CD95, TRAIL R1, and TRAIL

Death Receptor Mediated Apoptosis Death receptors are transmembrane cytokine receptors that belong to the tumor necrosis factor/nerve growth factor superfamily. The signaling through the death receptors is triggered by the binding of specific ligands, which initiates a cascade of events that ultimately results in the survival of the cell or in its death due to the activation of caspases. Recent findings have revealed a role for receptor internalization and endosomal trafficking, which selectively transmit the signals towards cell death or survival [2]. Some of these ligands (e.g., Fas and TNF) are of great relevance to liver diseases, as their expression and levels increase in many forms of liver diseases and mediate hepatocyte apoptosis [3].

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R2). TNF-induced apoptosis is mediated by the recruitment of the adaptor proteins, TNFR-associated ­death-domain (TRADD) protein, FAS-associated death-domain protein (FADD), and caspase-8 to the cytoplasmic death domain of the receptor [5]. CD95 and TRAIL receptors do not require TRADD for the recruitment of FADD and caspase-8 [6, 7]. TRADD diverges TNFR1 signaling from the death domain so that interaction of TRADD with receptor-interacting protein (RIP) and TNFR-associated factor-2 (TRAF2) leads to the activation of the survival transcription factor nuclear factor-kB (NF-kB) and to the induction of the c-Jun N-terminal kinase (JNK) cascade [8, 9]. Previous reports described that the mediating TNFinduced apoptosis follows a different mechanism than that activated by the CD95 ligand or TRAIL [10, 11]. In a seminal report, Micheau and Tschopp proposed a model in which TNFR1 signaling involves the assembly of two molecularly and spatially distinct signaling complexes that sequentially activate NF-kB and caspases [11]. Within a few minutes of TNF binding, TNFR1 recruits TRADD, RIP1, and TRAF2 to form a signaling complex at the cell surface (called “complex I”) that activates NF-kB. This model implies that, at later time points and after TNFR1 internalization, RIP1, TRAF2, and TRADD become modified and dissociate from the receptor. TRADD and/or RIP then bind to FADD, which then recruits caspase-8 to a secondary signaling complex within the cytosol (called “complex II”). This complex subsequently mediates apoptosis. However, in type II cells, such as hepatocytes, the activation of caspase-8 by complex II is weak for sustained activation of downstream caspases, needing an amplification loop through mitochondria, which are recruited through caspase-8-mediated Bid cleavage [12]. The resulting truncated Bid fragment (tBid) translocates to mitochondria where it activates other proapoptotic Bcl-2 proteins, Bax and Bak (see below). Whereas Bak resides in OMM, Bax translocates to mitochondria and inserts itself in OMM. Bax and Bak then uncover N-terminal epitopes and oligomerize, which is considered a central step in OMM permeabilization and cell death because it allows the release of cytochrome c into the cytosol to assemble the apoptosome (see below). Thus, although TNFR1 gives out simultaneously death and survival signals through caspase-8 and NF-kB activation, respectively, the mechanisms governing this balance that ultimately controls cells’ fate are poorly understood.

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Tnfr1 Internalization and Endosomal Trafficking Binding of TNF initiates the rapid clustering of TNFR1, followed by internalization of the ligand-receptor complex via clathrin-coated pit formation [2, 13]. TNF activates two types of sphingomyelinases, an endolysosomal acid sphingomyelinase (ASMase) and a membranebound neutral sphingomyelinase (NSMase). The lipid second messenger ceramide, which is generated by sphingomyelinases, is a potent pro-apoptotic mediator [14]. Conflicting data on the role of ceramide and sphingomyelinases (SMases) in signaling have been published [15–17]. NSMase is activated via FAN (factor associated with NSMase) and leads to the accumulation of ceramide at the plasma membrane although its role in TNF signaling remains unclear [18]. Activation of ASMase is dependent on TNFR1 internalization and is mediated via the death domain of TNFR1 by the recruitment of the adaptor proteins TRADD and FADD [2, 13]. A role for ASMase in transmitting apoptotic signals of death receptors has been reported for TNF (see below) [19, 20], FasL [21, 22], and TRAIL [23]. Blocking the formation of clathrin-coated pits inhibits the activation of the endolysosomal ASMase and JNK, as well as TNF-induced cell death. In contrast, inhibition of TNFR1 internalization did not affect the interaction of the adaptor molecules FAN and TRADD with TNFR1 at the cell surface, the activation of plasmamembrane-associated NSMase, and the stimulation of proline-directed protein kinases. The Asp-protease cathepsin D (ctsD) is a direct downstream target of ceramide within the same endolysosomal compartment [24]. The pro-apoptotic protein Bid colocalizes with ctsD-positive vesicles and, following TNF stimulation, both ctsD and Bid are located in Rab5-positive early endosomes, indicating that Bid is located at the subcellular site of ctsD activation. After TNF-induced, ceramide-mediated translocation through the endosomal membrane, ctsD cleaves Bid. This leads to the activation of caspase-9 and caspase-3. In addition, studies by Guicciardi et al. showed that cathepsin B (ctsB), a lysosomal enzyme, was required for TNF-mediated hepatocellular cell death and for hepatic injury and fibrosis during cholestasis [25, 26]. Using hepatocytes lacking cathepsin B, these authors showed that caspase-mediated release of cathepsin B from lysosomes enhanced mitochondrial release of cytochrome c and subsequent

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caspase activation in TNF-treated hepatocytes. Thus, as with caspases, endolysosomal proteases such as ctsD or ctsB can be activated in a cascade-like manner downstream of ASMase activation. Once released to the cytosol, these proteases might execute caspase-­ independent apoptosis or might individually participate in different apoptotic or cell-death-signaling cascades by connecting the endosomal compartment to the classic apoptotic signaling pathways. In addition, recent data have provided evidence for a role for ASMase in liver fibrogenesis by controlling ctsD and ctsB activation in hepatic stellate cells [27]. While these findings highlight the relevance of ceramide generation in TNFmediated hepatocellular cell death, they also illustrate the pathological significance of ceramide generation in liver diseases. Hence, if Fas and TNF overexpression are of relevance to liver diseases, ASMase-mediated ceramide generation emerges as a key signaling entity in TNF or Fas, and may be a novel therapeutic target for liver diseases.

Mitochondrial Membrane Permeabilization and Release of Proapoptotic Proteins As mentioned above, cytochrome c release from IMS has been the major focus of mitochondrial-confined proteins that are released upon apoptosis signaling. However, other proteins are also released from the intermembrane space provided the breakage of OMM occurs. For instance, the release of Smac/Diablo into the cytosol ensures the efficiency of caspase 3 in proteolyzing target proteins through inhibition of inhibitor of apoptosis proteins (IAPs) [28, 29]. Furthermore, the mitochondrial protein Omi/HtrA2 promotes cell death in a dual fashion. Besides its IAP activity Omi/HtrA2 also functions as a serine protease, thereby contributing to both caspase-dependent and caspase-independent cell death [30, 31]. Moreover, other specialized mitochondria-residing proteins, such as the apoptosis inducing factor (AIF) [32] and endonuclease G [33], are translocated to the nuclei following their release from mitochondria and promote peripheral chromatin condensation and high molecular weight DNA fragmentation. The intricacy of this pathway highlights the central role of mitochondria in controlling cell death, regardless of the phenotype of death (caspase-

J. C. Fernández-Checa and C. Garcia-Riuz

dependent apoptosis, caspase-­independent apoptosis, or necrosis). Hence, understanding the mechanisms leading to the release of mitochondrial pro-apoptotic factors constitutes an important advance in designing therapies aimed to regulate cell death and for the treatment human pathologies, including liver diseases. Because these potentially toxic proteins are normally secured in the IMS, mitochondrial membrane permeabilization that culminates in the rupture of the physical barrier (OMM) limiting their release into the cytosol constitutes a point-of-no-return in cell death. This subject has been an intense field of investigation with evidence for two possible mechanisms leading to the breakage of OMM: the mitochondrial permeability transition (MPT), and the permeabilization of OMM without disruption of the inner membrane. The former is a process characterized by mitochondrial swelling, IMM permeabilization, and OMM rupture as a secondary event [34]. On the other hand, there is strong evidence indicating the selective permeabilization of OMM in the absence of disrupted inner membrane [35]. The relative prevalence of these pathways in the regulation of cell death is not definitively established. One important feature of mitochondrial permeabilization is the obvious loss of function resulting in the inability of mitochondria to synthesize ATP through the oxidative phosphorylation. However, while the final outcome of mitochondrial dysfunction is cell death, the phenotype of death, apoptosis, and/or necrosis will depend on the level of cellular ATP as ATP is required for the efficient assembly of the apoptosome.

Cytochrome c Mobilization In addition to critical understanding of the mechanisms leading to OMM permeabilization to tailor strategies to control cell death, other levels of uncertainty are the mechanisms underlying the mobilization of cytochrome c from IMS. It has been proposed that during mobilization cytochrome c detaches from the IMM and dissociates from the membrane phospholipid cardiolipin [36]. A significant proportion of the cytochrome c in the mitochondria seems to be associated with cardiolipin, involving two major mechanisms. At physiological pH, cytochrome c has +8 net charges, establishing an electrostatic bond with the anionic cardiolipin. In addition, cytochrome c has a hydrophobic

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channel through which one of the four acyl chains of cardiolipin inserts. The other chains of cardiolipin remain in the membrane, thereby anchoring cytochrome c to the IMM [37]. One mechanism involved in cytochrome c detachment from IMM involves cardiolipin oxidation because oxidized cardiolipin has a much lower affinity for cytochrome c than the unoxidized form [38, 39]. Cardiolipin can be oxidized by phospholipase A2, by reactive oxygen species (ROS) or by the cardiolipin–cytochrome c complex [36, 40]. Detachment of cytochrome c from cardiolipin might also be triggered by increased cytosolic calcium, which weakens the electrostatic interaction between cytochrome c and cardiolipin. In addition, as described below, oxidized cardiolipin modulates the biophysical properties of OMM to allow oligomerized Bax to insert itself and permeabilize OMM [39]. In contrast to this view, other evidence neglects the mobilization of cytochrome c from IMM as a requirement for its release into the cytosol. In isolated mitochondria, exposure to 50–80 mM K+, a concentration of intracellular K+ that is well within physiological limits, is sufficient to release cytochrome c from the IMM [41]. In addition, live-cell imaging has revealed that there are no differences in the kinetics of the release of cytochrome c compared with that of other IMS proteins in cells undergoing apoptosis [42]. These results imply that cytochrome c mobilization might not require an additional step in its release from the mitochondria. In addition, mobilization of cytochrome c might also involve its removal from narrow cristae junctions. About 85% of the total cytochrome c resides within mitochondrial cristae, which are connected to the peripheral portion of the IMS by relatively narrow crista junctions. It has been suggested that remodeling of cristae is a required step in the release of this interior pool of cytochrome c [43], and recent studies correlated the disassembly of opa1 oligomers, a structural determinant of cristae morphology, with the remodeling of cristae [44]. Hence to account for the rapid and extensive release of cytochrome c during apoptosis, it has been suggested that cristae are remodeled such that cytochrome c is redistributed in the mitochondria prior to translocation through the OMM. However, using fluorescence microscopy followed by three-dimensional electron microscope tomography, Sun et al. found that cristae remodeling is not required for efficient release of cytochrome c [45]. Moreover, Yamaguchi et al. reported that BH3-only proteins Bid and Bim induced

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full cytochrome c release but only a subtle alteration in crista junctions, which involved the disassembly of opa1 complexes [46]. Furthermore, mitochondria fission or fragmentation is observed during apoptosis, and this change in mitochondrial morphology has been thought to mediate or contribute to cytochrome c release. In this regard, Sheridan et al., studying mutants overexpressing genes involved in the regulation of mitochondrial fusion/fission, reported that Bax/Bakdependent mitochondrial fragmentation coincides with apoptosis-associated cytochrome c release, but it is not necessarily required in this process, thus uncoupling both events in apoptotic cell death [47].

Omm Permeabilization: Mpt Vs The Bcl-2 Network Once cytochrome c detaches from IMM it is available for its release in the cytosol to engage the apoptosome provided that the OMM becomes permeabilized, as this particular membrane is not normally permeable to proteins. Two major mechanisms have been described in OMM permeabilization, which are differentiated on the basis of IMM permeabilization or not. The usually impermeable IMM prevents unrestrained influx of lowmolecular-weight solutes into the mitochondria. The MPT features mitochondrial swelling, uncoupling, and IMM permeabilization to small solutes, which results in a colloidal osmotic pressure that leads primarily to massive swelling of the mitochondrial matrix [34]. Because of of MPT the OMM ruptures and cytochrome c and other IMS proteins are released into the cytosol. MTP is most likely assembled at the contact sites of the OMM and the IMM, and the actual components of the MTP remain ill defined. One model proposes that an aggregation of misfolded integral membrane proteins present at high density can form MTP and are regulated by chaperone-like proteins [48]. A current accepted model proposes that VDAC (voltage-dependent anion channel), the adenine nucleotide translocator (ANT), and cyclophilin D (CypD) are the main components of MTP [34]. Mitochondrial phosphate carrier, another member of the mitochondrial carrier-protein family, has also been implicated as a MTP component because its depletion results in delayed cytochrome c mobilization and apoptosis [49]. However, the role of MPT in OMM permeabilization is unclear. For instance, some

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studies suggest that the MTP is an initiator of OMM permeabilization, whereas others indicate that it is a consequence of it. Recent experiments have provided a major role for MPT in ischemia-reperfusion injury but not in other forms of cell death such as developmental apoptosis. Mitochondria from CypD-deficient mice showed a general defect in the Ca2+-induced permeability transition, but had no developmental defects associated with a lack of apoptosis [50]. Furthermore, OMM permeabilization that was induced by the pro-apoptotic protein Bid/ Bax or by apoptotic stimuli was intact in these knockout animals. In contrast, cell death induced by ischaemia–reperfusion injury in the heart or brain was defective in CypD-deficient mice. Although VDAC has been proposed as an integral component of MPT and it participates in OMM permeabilization, recent data have discarded a role for VDAC in apoptosis. Murine cells deficient in all three VDAC isoforms successfully undergo intrinsic apoptosis [51]. Moreover, an isoform of VDAC (VDAC2) has been described to be antiapoptotic [52]. Similarly, mitochondria from murine cells lacking ANT1 and ANT2 can still undergo Ca2+-induced swelling and MPT, although at a higher threshold, which has been interpreted as evidence against a role for ANT in MPT and hence in OMM permeabilization [53]. However, the ability of ANT1/ANT2-deficient cells to undergo MPT might be due to the functional compensation by a novel ANT isoform identified recently [54], or by other mitochondrial carriers able to form pores in the inner membrane such as the ornithine/citrulline transporters or the phosphate carrier [49]. Alternatively to MPT, in the control of OMM permeabilization a major role for Bcl-2 family members has been put forward. Bcl2-family death agonists induce OMM permeabilization, thereby promoting cytochrome c release, whereas Bcl2-family death antagonists prevent it. Thus, Bcl2-family proteins control mitochondrial integrity, regulate cytochrome c release and intrinsic apoptosis [55]. Under non-­ apoptotic conditions, Bax is inactive and present in the cytosol as a monomer. Following an apoptotic stimulus, Bax is activated and translocates to the mitochondria, where it undergoes a conformational change and inserts itself into the OMM. Bax oligomerization is associated with the formation of openings in the OMM to allow the release of cytochrome c and other IMS proteins into the cytosol, and hence Bax oligomerization is considered a critical regulatory point in cell

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death [55]. In contrast to Bax, Bak resides on the OMM rather than in the cytosol but, like Bax, it undergoes a conformational change in response to apoptotic stimuli. This change allows Bak to oligomerize, contributing to OMM permeabilization. One potential mechanism involved in the permeabilizing activities of Bax/Bak oligomers is the formation of pores in OMM. However, attempts to visualize these oligomers have shown that large clusters of Bax are localized near, but not on, the OMM. An alternative model suggests that the insertion of activated, oligomerized Bax and/or Bak into the OMM creates a positive curvature stress on the membrane, leading to supramolecular pores that include lipids (lipidic pores) in the OMM [56, 57]. Clearly, understanding the mechanisms underlying OMM permeabilization may provide novel strategies to regulate cytochrome c and hence apoptosis.

Mitochondrial Ros and Gsh Excessive reactive oxygen species (ROS) generation leads to apoptotic and necrotic cell death, and mitochondria are the primary source of ROS, which are generated by leakiness of the electron transport chain (ETC) [58]. The basal stimulation of ROS from ETC is low, with estimates of 2–4% of electrons leaking from ETC to molecular oxygen to form superoxide anion. Recent data described the onset of superoxide flushes originating from MPT that occur randomly in space and time with all-or-none of the properties and the enhanced frequency of flushes contributes to hypoxia/reoxygenation injury [59]. Mitochondrialderived ROS can contribute to OMM permeabilization, cytochrome c release, and cell death, and hence the regulation of this burst of ROS from mitochondria may be of vital importance in preserving cells’ survival. The first line of defence against superoxide anion is the presence in mitochondria of superoxide dismutase (Mn-SOD), which dismutates superoxide anion into hydrogen peroxide. Although hydrogen peroxide is not strictly a free radical it is a potent oxidant, which can be transformed into hydroxyl radical in the presence of transition metals if it is not efficiently reduced into water. Peroxiredoxin-III (Prx-III) and thioredoxin-2 (Trx-2) regulate hydrogen peroxide metabolism and hence apoptotic signaling [60, 61]. In addition, the GSH redox cycle ensures

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efficient hydrogen peroxide catabolism. Mitochondrial GSH (mGSH) limitation has been shown to control hepatocyte survival in response to stimuli that trigger mitochondrial ROS such as TNF, Fas, or hypoxia despite Prx-III or Trx-2 defense [39]. In examining the mechanisms involved in the mGSH-dependent hepatocellular susceptibility to TNF, the functions of complexes I and II, assembled upon the binding of TNF to TNFR1 and causing the activation of NF-kB and caspase-8, respectively, were analyzed under selective mGSH depletion. Interestingly, TNF activated caspase-8, resulting in Bid cleavage, mitochondrial Bax translocation, and oligomerization in OMM despite the lack of inactivation of NF-kB. Interestingly, the predicted consequences of these events on OMM permeabilization, cytochrome c release, caspase-3 activation, and hepatocellular death occurred only upon mGSH depletion. These events were preceded by stimulated mitochondrial ROS that predominantly oxidized cardiolipin (CLOOH). Indeed, CLOOH potentiated oligomerized Bax-induced OMM-like liposomes permeabilization by restructuring the lipid bilayer, without effect on membrane Bax insertion/ oligomerization. These findings are one of the rare examples of enhanced susceptibility to TNF despite NF-kB activation, the master regulator of TNF susceptibility, as its inactivation sensitizes hepatocytes to TNF. In addition these findings show that OMMlocalized oligomeric Bax is not sufficient for TNFinduced OMM permeabilization and hepatocellular death. Moreover, mGSH controls the susceptibility of hepatocytes to TNF by controlling the efficiency of Bax to cause OMM permeabilization via mitochondrial membrane remodeling through CLOOH generation. Although the model of Micheau and Tschopp [11] established that NF-kB protects against TNF by blocking caspase-8 activation via FLIP, our data indicate that TNF activates a default level of hepatic caspase-8 independently of NF-kB inactivation. Although the level of caspase-8 activation may seem modest compared to when NF-kB is first inactivated, it is sufficient for OMM permeabilization if mGSH levels are low. In addition, these data indicate a dual role for CL in cell death regulation, not only by modulating the level of unbound cytochrome c available for release but also by controlling membrane stability via bilayer-to-hexagonal lipid phase transition, which facilitates the pore-forming activity of Bax in OMM.

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Cholesterol and Sphingolipids in Cell Death Cholesterol and sphingolipids (SLs) are integral components of biological membranes that play a major structural role. However, in addition to this view, recent evidence has shown that both lipids are more than just components of structural membranes, and actively participate and regulate signaling pathways and cell death [14, 62]. Cholesterol and SLs are not randomly distributed within membranes, but are concentrated in specific domains called lipid rafts where specific signaling pathways occur [63, 64]. Moreover, these lipids are unevenly distributed among cell ­membranes with the plasma membrane being highly enriched and mitochondria being relatively poor in endoplasmic reticulum [62].

Cholesterol and Hepatocyte Apoptosis Although cholesterol plays a vital role in regulating physical properties of membranes [65], its accumulation in cells is toxic and causes fatal diseases. For instance, Niemann Pick type C (NPC) disease is a fatal neurodegenerative disease characterized by lysosomal storage of cholesterol and glycosphingolipids. Although most patients exhibit neurologic symptoms, cholesterol accumulation affects the liver and pancreas, and some patients die early from liver failure before manifestations of the neurological symptoms. Moreover, NPC disease is the second most common cause of neonatal cholestasis. Ten percent of NPC infants presenting with neonatal cholestasis die from liver failure before they reach 6 months of age, and patients who survive often live with persistent liver disease accompanied with fibrosis, and in some rare cases, cirrhosis. Recent studies in NPC deficient mice showed that hepatocyte apoptosis is a primary cause of liver dysfunction and liver failure [66]. In addition, recent data have demonstrated a key role for TNF in NPC-mediated hepatocyte apoptosis [67]. These data suggest that cholesterol accumulation in NPC sensitizes to TNF-induced hepatocellular cell death. While the molecular mechanisms underlying these observations are not completely understood, recent findings have shown that free cholesterol in NPC hepatocytes accumulates in mitochondria but not in

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endoplasmic reticulum, and that this results in selective mGSH depletion [68]. Indeed, although mitochondrial cholesterol fulfils an important physiological function such as bile acids synthesis, its overaccumulation in mitochondria impairs vital functions of the membrane including transport of GSH from the cytosol into mitochondria, resulting in mGSH depletion [62, 69]. Interestingly, free cholesterol modulates membrane fluidity and its enrichment in mitochondria has been shown to impair MPT and the permeabilizing ability of Bax, hinting an antiapoptotic function of cholesterol as described in hepatocellular carcinoma [70, 71]. Intriguingly, unlike untransformed hepatocytes, hepatoma cell lines are able to maintain physiological levels of mGSH despite enhanced mitochondrial cholesterol levels [71]. Therefore, due to the emerging role of mitochondrial cholesterol in hepatocellular sensitization to cell death, a better understanding of the mechanisms of cholesterol trafficking to mitochondria may be of relevance to disease pathogenesis, including steatohepatitis or hepatocarcinogenesis (Fig. 29.2) (see below).

Sphingolipids and Cell Death SLs are more than mere structural components of biological membranes. Many different stimuli generate or stimulate the upregulation of SLs, particularly,

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ceramide which, upon interaction with downstream targets, mediate specific cell responses that range from proliferation to growth arrest, apoptosis, differentiation, or recognition. Intriguingly, some SLs such as ceramide and sphingosine 1-phosphate (S1P) exert opposing functions in the regulation of cell death and survival, and hence the relative balance between ceramide and S1P determines the fate of cells in response to specific stimuli [14]. Among SLs, ceramide has attracted considerable attention because of its recognized role as a key intermediate in inducing many stimuli, particularly in stress and death ligands (e.g. TNF/Fas-induced cell death). Ceramide can be synthesized by two general mechanisms, one involving its rapid and transient formation from sphingomyelin hydrolysis upon activation of SMases, of which the ASMase and NSMase isoforms are of major relevance in cell signaling. As mentioned above, while NSMase in TNF/Fas-mediated hepatocyte apoptosis is controversial, the role of ASMase has been shown to involve two distinct mechanisms, involving the recruitment of mitochondria through ganglioside GD3 generation [72], and the downregulation of liver-specific methionine adenosyltransferase-1A (MAT1A), the ratelimiting enzyme responsible for the synthesis of S-adenosyl-l-methionine (SAM) [73]. As a key intermediate of TNF/Fas, ASMase therefore contributes to hepatocyte apoptosis, thus emerging as a novel therapeutic target for liver diseases (see below). In addition to recruiting mitochondria stimulating OMM permeabilization and cytochrome c release [74], ganglioside GD3 has been shown to prevent the nuclear translocation of DNA binding competent members of NF-kB, thereby disabling survival factors and to sensitize hepatocytes to TNF-mediated cell death [75]. In addition to these targets ceramide is a very resourceful molecule and recruits many different intermediates to exert its multiple effects including the activation of cell death pathways [14].

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Fig. 29.2  Paradoxical role of mitochondrial cholesterol in cell death. Although mitochondrial cholesterol plays a physiological role, its accumulation perturbs mitochondrial membranes modulating their susceptibility to be permeabilized by multiple stimuli. Mitochondrial cholesterol accumulation in diseases such as ASH/NASH results in mGSH depletion sensitizing hepatocytes to TNF/Fas-mediated cell death, while in hepatocellular carcinoma, mitochondrial cholesterol loading contributes to chemotherapy resistance

Apoptosis and Liver Diseases Steatohepatitis Steatohepatitis (SH) represents an advanced stage in the spectrum of fatty liver diseases that encompasses alcoholic (ASH) and non-alcoholic steatohepatitis

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(NASH), two of the most common forms of liver disease worldwide. Although the primary etiologies of ASH and NASH are different, these two diseases show almost identical histology features characterized by steatosis, mixed lobular inflammation with scattered leukocytes and mononuclear cells, and hepatocellular cell death due to sensitivity to oxidative stress [76]. Despite significant progress in recent years, the pathogenesis of SH is still incompletely understood, although the two-hits hypothesis is the most prevalent view to explain the progression from steatosis to SH. In this hypothesis, the accumulation of fat within hepatocytes constitutes the first hit, which somehow sensitizes the liver to an upcoming stress, (e.g. oxidative stress, inflammation) with inflammatory cytokines TNF/Fas playing a key role in the disease [77]. Testing the hypothesis that the type rather than the amount of fat in hepatocytes is a crucial determinant of the susceptibility to TNF/Fas and using nutritional and genetic models of hepatic steatosis, it has been described that cholesterol accumulation in hepatocytes, as opposed to that of triglycerides or free fatty acids sensitizes to TNF/Fas-mediated SH [68]. Cholesterol accumulation in the ER or the plasma membrane did not cause ER stress or alter TNF signaling. Rather, the trafficking of cholesterol to mitochondria accounted for the hepatocellular susceptibility to TNF because of mitochondrial GSH (mGSH) depletion. In addition, hepatocytes from NPC1 knockout mice exhibit accumulation of cholesterol into mitochondria resulting in mGSH depletion and susceptibility to TNF [68]. Boosting the pool of mGSH or preventing its depletion by blocking cholesterol synthesis with atorvastatin, the susceptibility of obese ob/ob mice to LPS-mediated liver injury was blunted, highlighting the relevance of mitochondrial-mediated oxidative stress in the susceptibility to TNF-induced liver injury and NASH. Similar to these findings with nutritional or genetic models of NASH, alcohol-induced liver injury is characterized by the susceptibility to TNF-mediated cell death [78–80]. Although the mechanisms for this transition from resistance to susceptibility to TNF upon alcohol intake may be multifactorial [81], it has been reported that alcohol feeding causes mGSH depletion due to alcohol-stimulated cholesterol synthesis [69, 82]. Cholesterol accumulation in mitochondrial membranes impairs the mitochondrial transport of GSH from the cytosol, resulting in its depletion [83, 84]. Similar findings in the liver have been observed in the

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intragastric model of alcohol administration in mice [85] and in alveolar type-II cells [86]. mGSH has been associated with increased susceptibility to ethanolinduced liver injury and lethality in mice deficient in Nrf2, a transcription factor that regulates GSH biosynthesis and homeostasis [87]. In order to unambiguously test whether mGSH determines the hepatocellular susceptibility to TNF/Fas, selective mGSH depletion was accomplished with 3-hydroxy-4-pentenoate without effect on the cytosol pool of GSH [39]. In this paradigm, mGSH limitation sensitized to TNF/Fas by enhancing the mitochondrial generation of ROS via ASMase-induced ceramide generation, causing the peroxidation of cardiolipin, which facilitates the permeabilizing activity of oligomerized BAX in the OMM. In summary, cholesterol, particularly mitochondrial cholesterol, emerges as a key factor that contributes to the progression from steatosis to SH by sensitizing hepatocytes to TNF/Fas via mGSH depletion. Recent findings have confirmed the critical role of cholesterol in SH [88, 89], strongly suggesting that therapy aimed at lowering cholesterol synthesis and/or expanding the mitochondrial GSH defence may be of relevance in SH.

Ischemia/Reperfusion Liver Injury Hepatic ischemia/reperfusion (I/R) damage can occur in diverse settings including liver transplantation, trauma, hemorrhagic shock, or liver surgery and is a serious clinical complication that may compromise liver function because of extensive hepatocellular loss. Despite intense research in this area, the molecular mechanisms responsible for hepatic I/R injury are not well understood [90]. Multiple cellular and molecular mechanisms are ultimately involved in hepatic I/R damage. In addition to recruitment and activation of inflammatory cells, as well as Kupffer cells, platelet adhesion in the sinusoidal lining has been involved in sinusoidal endothelial cell death and hepatic I/R injury. Molecular events include NF-kB activation, although its ultimate role in hepatic I/R injury is controversial, JNK activation, MPT, and ROS generation. Interestingly, TNF has been identified as a key player in hepatic I/R damage [4]. Consistent with these findings, ischemic preconditioning has been shown to reduce ischemic injury in the liver via activation of oxidative stress that induces a

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cytoprotective response. Thus, it has been shown that preconditioning with TNF and Fas protects the liver against ischemic injury [91]. Moreover, cardiotrophin-1 knockout mice are more susceptible to hepatic I/R injury, and cardiotrophin-1 preconditioning has been shown to be protective [92]. In line with its role in TNF/ Fas signaling, it has been shown that ASMase plays a role in hepatic I/R injury [93]. Indeed, ASMase-induced ceramide generation targeted mitochondria via JNK and BimL activation. Thus, in addition to the previous mechanisms described and consistent with the role of ceramide in recruiting mitochondria pathway of cell death, ASMase emerges as a potential target for intervention against hepatic I/R.

Hepatocarcinogenesis One of the striking features of tumorigenesis is the deregulation of cholesterol metabolism [94]. This is exemplified by the desensitization of hydroxymethylglutaryl-CoA reductase (HMG-CoAR) to inhibition by sterols, and by the continued cholesterol synthesis in growing solid tumors. Cholesterol synthesis is oxygen dependent [95], and hypoxia is a prominent feature of solid tumor development and considered a major driving force for tumor progression, invasiveness, and survival [96, 97]. In addition, increased cholesterol levels in mitochondria have been observed in heterotopic Morris hepatoma xenografts in Buffalo rats, compared to the content found in mitochondria from host liver [98]. As cholesterol enrichment can adversely affect mitochondrial functions [68, 70], it is conceivable that the accumulation of cholesterol within mitochondrial membranes may actually account for or contribute to the known mitochondrial dysfunction of cancer cells, underlying the Warburg effect and dependence on glucolysis [99]. Indeed, using human and rat hepatocellular carcinoma cell lines or mitochondrial fraction isolated from human HCC patients, a dramatic increase in the levels of mitochondrial cholesterol compared to untransformed cells was observed, which translated into increased membrane order parameter [71]. This outcome was accompanied by increased resistance to chemotherapy selectively acting via mitochondria, which was reversed either by cholesterol extraction or by fluidization of mitochondrial membranes. Moreover, considering the role of steroidogenic acute regulatory

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protein in regulation of mitochondrial cholesterol homeostasis [62], its downregulation in hepatocellular carcinoma cell lines by siRNA reduced the net levels of mitochondrial cholesterol, increasing their susceptibility to mitochondria-targeted chemotherapy. In line with these data, Lucken-Ardjomande have recently shown that treatment of HeLa cells with U18666A, which caused mitochondrial cholesterol upregulation, showed a delay in the release of Smac/Diablo and cytochrome c, as well as in Bax oligomerization and partial protection against stress-induced apoptosis [100]. Moreover, the inhibitory effect of cholesterol on mitochondrial Bax activation was demonstrated in liposomes, and this effect was exerted by a dual mechanism involving changes in membrane order parameter and in the decrease of Bax penetration into the membrane [71]. Thus, by inhibiting Bax-driven OMM permeabilization, cholesterol modulates cell death susceptibility. Furthermore, the potentiation of hepatocellular carcinoma chemotherapy by squalene synthase inhibition by YM-53601, which reduces cholesterol levels including that in mitochondria, without perturbing isoprenoid metabolism, validates the specificity of cholesterol in chemotherapy resistance, and revitalizes the potential benefit of cholesterol downregulation in cancer therapy.

Closing and Future Remarks Although apoptosis is essential for life, especially during early stages of development, its deregulation is known to mediate a number of pathologies and diseases. In particular, hepatocellular apoptosis is a prominent characteristic feature of many forms of liver disease, some of which have been briefly described in this chapter, and is triggered by factors that induce the expression of death receptors and/or their ligands [101]. Unlike in other cell types, mitochondria play a key role in hepatocellular death, and hence factors that modulate mitochondrial membrane permeabilization will have a profound impact on hepatocyte cell and its survival. In this regard, cholesterol and, more specifically, mitochondrial cholesterol emerges as a key factor in modulating the mitochondrial or intrinsic pathway of cell death (Fig. 29.2), establishing the mitochondrial cholesterol paradox in cell death. While in ASH/NASH, mitochondrial cholesterol enrichment plays a proapoptotic

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role determined by the depletion of mGSH, the mitochondrial cholesterol loading observed in hepatocellular carcinoma is antiapoptotic. Paradoxically this paradigm is accompanied by undepleted mGSH state despite cholesterol accumulation in mitochondria by uncharacterized mechanisms. Despite this contrast, mitochondrial cholesterol depletion would be expected to halt hepatocyte death in ASH/NASH and increase chemotherapeutic efficiency of hepatocellular carcinoma, which remains to be established awaiting well-designed clinical trials in both cases.

Summary

›› Caspase-dependent apoptosis occurs through ››

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two main pathways, the extrinsic pathway and the intrinsic pathway. The extrinsic pathway is initiated upon the binding of an extracellular ligand to transmembrane death receptors of the TNF superfamily, which leads to the assembly of the deathinducing signaling complex (DISC). The DISC then activates an initiator caspase, which triggers the enzymatic cascade that leads to apoptotic death. Selected reading 3(104) The intrinsic pathway, also known as the mitochondrial pathway, is activated by stimuli that lead to the permeabilization of the outer mitochondrial membrane (OMM) and the subsequent release of proteins from the mitochondrial intermembrane space (IMS), which initiate or regulate caspase activation, such as cytochrome. Selected reading 1, 2(102, 103) Cytochrome c normally resides within the cristae of the inner mitochondrial membrane (IMM) and is effectively sequestered by narrow cristae junctions. Cytochrome c participates in the mitochondrial electron-transport chain, using its heme group as a redox intermediate to shuttle electrons between complex III and complex IV. Although mitochondrial cholesterol fulfils an important physiological function with bile acid synthesis, its overaccumulation in mitochondria impairs vital membrane functions including import of GSH resulting in mGSH depletion and increased sensitivity to TNF and Fas.

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Multiple Choice Questions 1. Which is one of the following factors is not released by the mitochondria? (a) cytochrome c (b) Smac/Diablo (c) Caspase 3 (d) Endonuclease G (e) Apoptosis inducing factor 2. Which of the following statements regarding the interaction cytochrome c/cardiolipin is wrong? (a) at physiological pH there is an electrostatic bond between the two (b) detachment of cytochrome c from cardiolipin involves oxidation of cardiolipin (c) cardiolipin anchors cytochrome c to the OMM (d) calcium weakens the interaction between cytochrome c and cardiolipin (e) Oxidized cardiolipin modulates the biophysical properties of OMM 3. Which is not a component of the MTP? (a) cytochrome c (b) VDAC (voltage dependent anion channel) (c) ANT (adenine nucleotide translocator) (d) Cyclophilin D (e) Mitochondrial phosphate carrier 4. Regarding Bax, which statement is correct? (a) Inactive Bax is located in the cytosol (b) When activated, it translocates to the mito­ chondria (c) It undergoes conformational change and inserts into OMM (d) It oligomerizes (e) All the above statements are correct 5. Which statement is correct? (a) Ceramide is a potent anti-apoptotic factor (b) Ceramide activates sphingomyelinase (c) TNF activates only the membrane-bound neutral sphingomyelinase (d) The endolysosomal acid sphingomyelinase is activated by cathepsins (e) Activation of endolysosomal acid sphingomyelinase is dependent on TNFR1 internalization Acknowledgments  This work was supported in part by the Research Center for Liver and Pancreatic Diseases Grant P50 AA 11999 funded by the US National Institute on Alcohol Abuse

450 and Alcoholism, Plan Nacional de I + D Grants: SAF200503923, SAF2005-03943, SAF2006-06780, and FIS06/0395 and by the Centro de Investigacion Biomedica en Red de Enfermedades Hepaticas y Digestivas (CIBEREHD) supported by the Instituto de Salud Carlos III.

Selected Reading 102. Liu X, Kim CN, Yang J, Jemmerson R, Wang X (1996) Induction of apoptotic program in cell-free extracts: requirement for dATP and cytochrome c. Cell 86: 147–157. (This study is hallmark in the field of cell death regulation as it described for the first time the identification of cytochrome c released from mitochondria as a trigger for caspase activation, thereby hinting at the existence of the mitochondrial pathway of apoptosis.) 103. Kuwana T et al. (2002) Bid, Bax, and lipids cooperate to form supramolecular openings in the outer mitochondrial membrane. Cell 111: 331–342. (This study showed that Bax permeabilizes lipid membranes sufficiently to allow large molecules to pass through, and that this effect requires an activation signal that is provided by Bid. It also showed that specific lipids, such as cardiolipin, are implicated in the function of Bax.) 104. Schneider-Brachert, W. et al. (2004) Compartmen­talization of TNF receptor 1 signaling: internalized TNF receptosomes as death signaling vesicles. Immunity 21: 415–428. (This study highlights the relevance of TNFR1 internalization to recruit and activate downstream intermediates that signal cell death in endosomes and provides the mechanisms whereby TNFR1 activates SMases in acidic com­partments.)

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452 61. Hansen JM, Zhang H, Jones DP (2006) Mitochondrial thioredoxin-2 has a key role in determining tumor necrosis factor-a induced reactive oxygen species generation, NF-kB activation and apoptosis. Toxicol Sci 91:643–650 62. Garcia-Ruiz C, Mari M, Colell A et al (2009) Mitochondrial cholesterol in health and disease. Histol Histopathol 24(1): 117–132 63. Ikonen E (2008) Cellular cholesterol trafficking and compartmentalization. Nat Rev Mol Cell Biol 9:125–138 64. Van Meer G, Voelker DR, Feigenson GW (2008) Membrane lipids: where they are and how they behave. Nat Rev Mol Cell Biol 9:112–124 65. Maxfield FR, Tabas I (2005) Role of cholesterol and lipid organization in disease. Nature 438:612–621 66. Beltroy EP, Richardson JA, Horton JD et al (2005) Choles­ terol accumulation and liver cell death in mice with Niemann Pick type C disease. Hepatology 42:886–893 67. Rimkunas VM, Graham MJ, Crooke RM et al (2009) TNF plays a role in hepatocyte apoptosis in Niemann Pick type C liver disease. J Lip Res 50: 327–333 68. Mari M, Caballero F, Colell A et al (2006) Mitochondrial free cholesterol loading sensitizes to TNF- and Fas-mediated steatohepatitis. Cell Metab 4:185–198 69. Fernández-Checa JC, Kaplowitz N (2005) Hepatic mitochondrial glutathione: transport and role in disease and toxicity. Toxicol Appl Pharmacol 204:263–273 70. Colell A, Garcia-Ruiz C, Lluis JM et al (2003) Cholesterol impairs the adenine nucleotide translocator-mediated mitochondrial permeability transition through altered membrane fluidity. J Biol Chem 278:33928–33935 71. Montero J, Morales A, Llacuna L et al (2008) Mitochondrial cholesterol contributes to chemotherapy resistance in hepatocellular carcinoma. Cancer Res 68:5246–5256 72. Garcia-Ruiz C, Colell A, Morales A et al (2002) Trafficking of ganglioside GD3 to mitochondria by tumor necrosis factor-alpha. J Biol Chem 277:36443–36448 73. Mari M, Colell A, Morales A et al (2004) Acidic sphingomyelinase downregulates the liver-specific methionine adenosyltransferase 1A, contributing to tumor necrosis factor-induced lethal hepatitis. J Clin Invest 113:895–904 74. Garcia-Ruiz C, Colell A, Paris R et al (2000) Direct interaction of GD3 ganglioside with mitochondria generates reactive oxygen species followed by mitochondrial permeability transition, cytochrome c release, and caspase activation. FASEB J 14:847–58 75. Colell A, Garcia-Ruiz C, Roman J et al (2001) Ganglioside GD3 enhances apoptosis by suppressing the nuclear ­factor-kappa B-dependent survival pathway. FASEB J 15: 1068–1070 76. Angulo P (2002) Nonalcoholic fatty liver disease. N Engl J Med 346:1221–1231 77. Tomita K, Tamiya G, Ando S et al (2006) Tumour necrosis factor a signaling through activation of Kupffer cells plays an essential role in liver fibrosis of non-alcoholic steatohepatitis in mice. Gut 55:415–424 78. Colell A, Garcia-Ruiz C, Miranda M et al (1998) Selective glutathione depletion of mitochondria by ethanol sensitizes

J. C. Fernández-Checa and C. Garcia-Riuz hepatocytes to tumor necrosis factor. Gastroenterology 115: 1541–1551 79. Minagawa M, Deng Q, Liu ZX et al (2004) Activated natural killer T cells induce liver injury by Fas and tumor necrosis factor-alpha during alcohol consumption. Gastroenterology 126:1327–1399 80. Pastorino JG, Hoek JW (2000) Ethanol potentiates tumor necrosis factor cytotoxicity in hepatoma cells and primary rat hepatocytes by promoting induction of the mitochondrial permeability transition. Hepatology 31:1141–1152 81. Shulga N, Hoek JB, Pastorino JG (2005) Elevated PTEN levels account for the increased sensitivity of ethanolexposed cells to tumor necrosis factor-induced cytotoxicity. J Biol Chem 280:9416–9424 82. Lluis JM, Colell A, García-Ruiz C et al (2003) Acetaldehyde impairs the mitochondrial glutathione transport in HepG2 cells through endoplasmic reticulum stress. Gastroenter­ ology 124:708–724   83. Colell A, Garcia-Ruiz C, Morales A et al (1997) Transport of reduced glutathione in hepatic mitochondria and mitoplasts from ethanol-treated rats: effect of membrane physical properties and S-adenosyl-L-methionine. Hepatology 26:699–708   84. Zhao P, Kahorn TF, Slattery JT (2002) Selective mitochondrial glutathione depletion by ethanol enhances acetaminophen toxicity in rat liver. Hepatology 36:326–335   85. Nakagami M, Wheeler MD, Bradford BU et al (2001) Overexpression of manganese superoxide dismutase prevents alcohol-induced liver injury in the rat. J Biol Chem 276:36664–36672   86. Velasquez A, Bechara RI, Lewis JF et al (2002) Glutathione replacement preserves the functional surfactant phospholipid pool size and decreases sepsis-mediated lung dysfunction in ethanol-fed rats. Alcohol Clin Exp Res 26:1245–1251   87. Lamlé J, Marhenke S, Borlak J et al (2008) Nuclear factoreythroid 2-related factor 2 prevents alcohol-induced fulminant liver injury. Gastroenterology 134:1159–1168   88. Matsuzawa N, Takamura T, Kurita S et al (2007) Lipidinduced oxidative stress causes steatohepatitis in mice fed an atherogenic diet. Hepatology 46:1392–1403   89. Zheng S, Hoos L, Cook J et al (2008) Ezetimibe improves high fat and cholesterol diet-induced non-alcoholic fatty liver disease in mice. Eur J Pharmacol 584:118–124   90. Selzner N, Rudiger H, Ralf R et al (2003) Protective strategies against ischemic injury of the liver. Gastroenterology 125:917–936   91. Jang JH, Moritz W, Graf R et al (2008) Preconditioning with death ligands FasL and TNF-a protecs the cirrhotic mouse liver against ischaemic injury. Gut 57:492–499   92. Iñiguez M, Berasain C, Martinez-Anso E et al (2006) Cardiotrophin-1 defends the liver against ischemia-reperfusion injury and mediates the protective effect of ischemic preconditioning. J Exp Med 203:2809–2815   93. Llacuna L, Mari M, Garcia-Ruiz C et al (2006) Criticial role of acidic sphingomyelinase in murine hepatic ischemia-reperfusion injury. Hepatology 44:561–572

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Calcium Signaling

30

Thierry Tordjmann

Introduction In normal eukaryotic cells, the concentration of free Ca2+ in the cytosol ([Ca2+]i) is actively kept much lower (100–200 nM) than extracellular (1–2 mM) and endoplasmic reticulum (ER) (0.5 mM) Ca2+ concentrations [1]. The cytosol, with its very low concentration of free calcium, is located at the interface of these two highly calcium-rich environments. This results in the cytosol, being a site of major and rapid variations in [Ca2+]i in response to the transfer of small quantities of Ca2+ from the extracellular medium or intracellular storage compartments [2]. These variations in [Ca2+]i (“calcium signals”), induced by agonists such as hormones and neurotransmitters, constitute a kind of language which is translated into physiological responses by the cells. Such calcium signals, highly organized in space and time, orchestrate a wide array of physiological processes from the subcellular to the whole tissue ­levels [2–4]. It was first shown by Woods et al. [5] that the Ca2+ signals in response to hormonal stimuli consist of a series of spikes in [Ca2+]i (oscillations) with a period of a few seconds to a few minutes. It is important that in this landmark study, the experimental cell model was the isolated rat hepatocyte. Many papers have been published since this one, concerning calcium signals in hepatocytes, and it appeared later that each Ca2+ spike was also organized spatially: the Ca2+ concentration first increased locally, then it propagated in the whole

T. Tordjmann INSERM U757, Université Paris sud, Bât. 443, 91405, Orsay, France e-mail: [email protected]

cell as a wave, traveling at a speed of 10–20 mm s−1 [6]. Also, at the level of the intact liver, calcium signals have been studied and well characterized [7–9]. In the agonist-perfused tissue, Ca2+ waves propagate from cell to cell along hepatocyte plates. Although this phenomenon of intercellular Ca2+ wave propagation has been observed in other tissues, the mechanisms involved are liver-specific, involving both coordination of Ca2+ signals and directional programming through gradual expression of signaling molecules along hepatocyte plates [10]. However, although a precise picture of the hepatocyte calcium signals has been built at the subcellular, cellular, and intercellular levels, their precise impact on liver physiology and pathology remains poorly understood and studied. Also, the liver is not made only of hepatocytes. Cholangiocytes have been well studied with respect to calcium signaling [11], but only a few studies are available on hepatic stellate cells [12], and other cell types (endothelial, Kupffer cells) which have been almost neglected in terms of their own calcium signaling and the regulatory role they may have on hepatocyte calcium signaling. An integrated full picture of the “liver calcium signaling” is obviously not yet available.

Hepatocyte Ca2+ Oscillations Preliminary. Hepatocytes have long been considered by biochemists and molecular biologists as an “easyto-get material” (several hundreds of millions cells per rat) on which numerous studies were performed, without any relevance to the organ from which the cells were isolated. That has been also the case for calcium

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signaling studies. As already mentioned, the first calcium oscillations were reported in hepatocytes [5], and numerous studies, both experimental and theoretical, have been conducted after that to decipher, in the hepatocyte, the machinery by which agonists generated cytosolic calcium signals. However, hepatocyte calcium signaling remains far less understood in terms of functional aspects. In hepatocytes, as in most nonexcitable cells, Ca2+ oscillations originate from the periodic opening of Ca2+ channels located in the membrane of the ER, ­following the activation of the phosphoinositide cascade. The binding of an agonist to a membrane-bound receptor activates the Ga-subunit of a G-protein complex coupled to the receptor. This activated G protein in turn stimulates phospholipase C (PLC) activity. The latter enzyme catalyzes the hydrolysis of the ­m embrane-bound phosphatidyl-inositol bisphosphate (PIP2) into diacyl-glycerol and inositol trisphosphate (InsP3). Ca2+ release from the internal stores is ensured by the InsP3R, an homotetramer that can bind up to four InsP3 molecules, forming a Ca2+ channel which equilibrium opening probability presents a bell-shaped dependence on cytosolic Ca2+ [13]. The decrease of [Ca2+]i in the cytosol is due to the activity of the Ca2+ ATPases (SERCA pumps), which actively transport Ca2+ from the cytosol into the ER. Ca2+-regulated InsP3Rs and Ca2+ ATPases are together sufficient to generate Ca2+ oscillations [3]. In most cases, hormone-induced Ca2+ oscillations in hepatocytes take the form of repetitive, sharp spikes sometimes preceded by a slower, pacemaker-like elevation in the cytosolic Ca2+ concentration. These periodic increases in the level of free Ca2+ in the cytosol from about 0.1 mM up to 1 mM have been observed in hepatocytes in response to stimulation by a large number of agonists such as noradrenaline, vasopressin, phenylephrine, angiotensin II, adenosine triphosphate (ATP), histamine, thrombin, etc., the shape of the oscillations being ­agonist-dependent [14]. The oscillation frequency increases with the level of stimulation, i.e., the concentration of external agonist, a phenomenon known as “frequency-encoding.” The level of extracellular Ca2+ – and thus the rate of 2+ Ca influx – affects the frequency of Ca2+ oscillations. Moreover, a basal level of external Ca2+ is required to avoid a progressive damping of the oscillations. As in many cell types, the decrease of Ca2+ concentration in the Ca2+ stores appears as the driving force for Ca2+

T. Tordjmann

entry from the external medium into the cytosol. The molecular basis of this mechanism, known as “store operated Ca2+ entry”, has been recently clarified [15]. Two major players, STIM1 (“stromal interaction molecule”) and “Orai,” have been discovered. STIM1, as the sensor of Ca2+ depletion in the ER, redistributes to the plasma membrane and signals to Orai which constitutes pore-forming subunits of SOCs (store operated Ca2+ channels). There is recent evidence that this machinery – discovered in mast and T cells – is also effective in hepatocytes or liver cell lines [16–18]. There are also some data suggesting that a TRP (transient receptor potential) protein, TRPC1 or TRPC6, may constitute the pore of SOCs in hepatoma cell lines [17, 19]. Finally, Jones et al. [18] recently reported the role of STIM1 and Orai1 on the maintenance of Ca2+ oscillations in primary rat hepatocytes, and emphasized that a non-SOCs component of Ca2+ entry also exists in these cells and needs to be explored.

Intracellular Ca2+ Waves Since early studies by Allbritton et al. [20], it appeared clearly that Ca2+ waves did not result from the simple diffusion of Ca2+ throughout the cytosol, due to abundant intracellular buffers. Instead, InsP3 which is more soluble in the cytosol, can diffuse freely along a 25-µm long distance from its production site(s), and mobilize Ca2+ from the storage compartments throughout the cell, creating the intracellular calcium wave. Initially, intrahepatocyte Ca2+ waves had been reported to start at a specific locus [21] and to propagate with constant amplitude and rate, whatever the agonist concentration [4]. Since these earlier studies, molecular basis for directional waves were dissected. Especially, it has been shown that expression and distribution of the InsP3R isoforms may vary among cells and tissues, the hepatocyte expressing only the types I (20%) and II (80%) [22]. Type II InsP3R, in contrast to types I and III, particularly supports sustained repetitive Ca2+ oscillations [23], and was shown to exhibit the highest affinity for InsP3 [24]. In addition to this isoform specificity, it has been shown in hepatocytes that the type I was evenly distributed in the cytosol and the type II was concentrated at the canalicular domain [25]. This spatial pattern of distribution was reported to support the direction of intracellular Ca2+ waves, starting from the canalicular region containing

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the most abundant and affine InsP3R isoform, and spreading towards the other regions of the cytosol, less sensitive to InsP3 [26]. The same authors further demonstrated that lipid rafts were responsible for the pericanalicular localization of InsP3R and for the polarized Ca2+ waves associated with this distribution [27].

Intercellular Ca2+ Waves in Hepatocytes As in many other cell types, intracellular movements of Ca2+ in hepatocytes, induced by hormones and neurotransmitters, may be propagated from cell to cell. In multicellular rat hepatocyte systems – i.e., doublets or triplets of hepatocytes connected by gap junctions [10, 28–30] – agonists such as vasopressin or noradrenaline, induce tightly coordinated intracellular Ca2+ increases. A striking feature of these responses is their sequential pattern that always follows the same order for a given agonist, resulting in apparent unidirectional

intercellular Ca2+ waves (Fig. 30.1). Such coordinated and sequential signals were also observed in the intact perfused liver in which vasopressin elicits waves of [Ca2+]i increases running along hepatocyte plates across the lobules, at a dose-dependent speed of 20–120 mm s−1 [7–9]. Although these waves propagate towards only one direction, the starting area of vasopressin-induced waves in the liver lobule (periportal or perivenous region) remains a matter of controversy [7–9]. We demonstrated that unidirectional Ca2+ waves resulted from a gradually decreasing cellular sensitivity to hormonal stimuli from the first to the last responding cell. It is well known that hepatocytes contribute unequally to various liver functions according to their position in the liver cell plate [31]. Hepatocytes ­isolated from periportal and perivenous areas, exhibited significantly different cellular sensitivities to the ­agonists. Periportal hepatocytes were more sensitive than perivenous hepatocytes to ATP; in contrast, the ­opposite was true for vasopressin, noradrenaline, and angiotensin II [10, 32]. We found that this heterogeneity in

b

a 3 0s

72 s

2 Time (s) 1 VASOPRESSIN (0.1nM) 78 s

81 s

c 200 nM 1 min

3 2 1

Fig. 30.1  Calcium signals in rat hepatocyte multiplets. Spatiotemporal development of Ca2+ increases in the three cells of a rat hepatocyte triplet. Isolated rat hepatocytes were loaded with fura2 and [Ca2+]i imaging was performed with an intensified charged-coupled camera. The 340 nm and 380 nm frames were captured at a high frequency to provide 1 image/110  ms. (a) Imaging of [Ca2+]i in the three connected cells. Vasopressin (1 nM) was added at time 46 s. The agonist generates an increase in [Ca2+]i in a first cell and then sequentially in a second and a

third cell. (b) Time course of the sequential increase in [Ca2+]i in the three cells of the triplet shown in A. Each point in the three curves is the average of the total [Ca2+]i captured every 110  ms throughout cells 1, 2, and 3. (c) Long-lasting image acquisition with smaller time resolution (1 image/s) shows that [Ca2+]i in the three cells oscillate and that spikes appear in a sequential manner, with the same ordered response at each oscillation train, giving the appearance of repetitive intercellular Ca2+ waves

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T. Tordjmann AGONIST

InsP3 Ca2+

InsP3 Ca2+

InsP3 Ca2+

InsP3 Ca2+

RECEPTOR ORIENTED INTERCELLULAR CALCIUM WAVE

Fig. 30.2  Receptor-oriented intercellular Ca2+ waves in hepatocytes. Receptor-oriented intercellular Ca2+ waves result from a gradient in the number of surface hormone receptors along the hepatocyte plates. Agonist-induced Ca2+ responses initiate in the cell with the most hormone receptors. Successive firing of cells is spatially oriented by the gradient in agonist-binding sites. Coordination between Ca2+ responses of adjacent cells is due to gap junctional transfer of InsP3 and allows repetitive Ca2+ waves to occur. Intercellular unidirectional Ca2+ waves are thought to trigger waves of canalicular contraction

hormonal sensitivity was due to the differences in number of the hormonal receptors [10]. Moreover, InsP3 has been shown to flow through gap junctions and thereby coordinate Ca2+ spiking among adjacent hepatocytes [33]. Such a configuration in which the most responsive hepatocytes drive the response of the less sensitive cells is similar to the cell to cell triggering of cardiac pacemaker cells [10, 34, 35] (Fig. 30.2).

Physiological Significance of Ca2+ Oscillations and Waves in Hepatocytes In general terms, Ca2+ oscillations in hepatocytes optimize the effect of hormonal stimulation. The sensitivity of cellular responses to the frequency of Ca2+ spikes may involve a Ca2+-calmodulin activated protein kinase (CaMKII), which acts as a widespread mediator between the Ca2+ spikes and the physiological response [36]. CaMKII can decode the frequency of Ca2+ oscillations thanks to its complex mode of regulation by Ca2+, in the form of autophosphorylation and CaM trapping [37, 38]. Calcium oscillations have been reported to be crucial for the phosphorylation–dephosphorylation cascade that governs the switch between glycogen synthesis and degradation promoted in hepatocytes by agonists like noradrenaline and vasopressin [39]. Also Dolmetsch et al. [40] showed that the temporal pattern of calcium signals

was of major impact as to the expression of transcription factors in lymphocytes, but this aspect has never been investigated in hepatocytes. The mitochondrial metabolic Ca2+ output has also been shown to be optimized by an oscillatory level of Ca2+ [41, 42]. Some liver functions have long been known to be controlled by intracellular calcium movements. For example, the production of glucose by the liver is mediated by hormone-induced Ca2+ increases [43, 44]. Also, intramitochondrial ATP synthesis can be coordinated with cellular energy demand, by Ca2+ oscillations in hepatocytes, thereby maintaining cell homeostasis and viability [42]. Many events related to bile secretion are also regulated by cytosolic Ca2+, such as vesicular trafficking and canalicular exocytosis of bile acid transporters [45, 46], permeability of tight junctions [47], or canalicular contraction [48, 49]. Intracellular calcium waves, as described above, starting from the canaliculus to the basolateral poles may have physiological impact on secretion, as it has been shown in pancreatic acinar cells [50], although direct evidence in hepatocytes is lacking. We recently reported dual effects of vasopressin-induced calcium signals in polarized rat hepatocyte multiplets: microvilli formation, occurring at the sinusoidal pole, requiring mainly extracellular calcium entry, and canalicular contraction, needing principally internal calcium stores mobilization [49]. These data fit with the pericanalicular location of the type II InsP3 receptor, an area where calcium release appeared to be functionally important for bile canaliculus contraction. Moreover, interhepatocyte calcium waves have been reported to support canalicular peristaltism and thereby to regulate bile flow in the normal and regenerating rat liver [51, 52].

Calcium in the Hepatocyte Nucleus Schematically two views have been proposed, one stating that calcium signals in the nucleus originate from the passive diffusion of cytosolic calcium and the second claiming that an autonomous machinery exists in the nucleus that can actively generate calcium signals. Although these two views may coexist in the same cells according to circumstances, in hepatocytes (hepatoma cell lines) recent literature strikingly favors the second view [53–56]. It has been shown in SkHep cells that an

30  Calcium Signaling

InsP3-sensitive intranuclear calcium compartment (i.e., the “nucleoplasmic reticulum”) exists [53]. PLCb, PIP2, and InsP3R have been found in the nucleus, allowing a local InsP3 production and providing the machinery necessary to generate autonomous Ca2+ signals [57]. Gomes et al. have recently shown that the HGF receptor (c-met) can translocate (upon agonist stimulation) from the plasma membrane to the nucleus, and generate an InsP3 production and calcium elevation in the nucleus, independently of cytosolic calcium, in a hepatoma cell line [56]. Very similarly, it was recently shown by the same group that insulin can induce nuclear calcium signals through a translocation of its receptor to the nucleus, in primary rat hepatocytes [58]. Impor­tantly, the nucleoplasmic reticulum as an intranuclear calcium compartment has not been shown in primary hepatocytes, and some authors claimed that it was not essential for calcium signaling [59]. Important cellular functions are thought to be regulated by nuclear calcium signals, including nuclear pore permeability, transcription factor activity thereby controlling gene expression, and protein kinase translocation [57]. Concerning the liver, a clear picture of functional impact of hepatocyte nuclear calcium has not yet emerged, although one may expect important effect at least on physiological and pathological liver growth [55].

Calcium and Hepatocyte Proliferation: Liver Regeneration and Carcinogenesis It is well established that intracellular Ca2+ is crucial for tissue homeostasis through regulation of cell cycle and apoptosis [2]. Pioneer studies have shown that extracellular calcium was crucial for liver regeneration [60]. Also, modifications of intracellular calcium homeostasis during liver regeneration have been reported, concerning Ca2+-binding proteins [61], membrane Ca2+-ATPases [62], or the InsP3 receptor [63]. Zhang et al. [64] also suggested that the alteration of the InsP3 and Ca2+ mobilization pathway could alter liver regeneration in the rat. More recently, it has been shown in non hepatocyte cell lines that the spatiotemporal organization of Ca2+ signals was determining for the activation of transcription factors like CREB, NF-kB, or NF-AT, and for immediate early genes like c-fos or c-jun [40, 65, 66]. It is also well established

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that the activation of the RAS pathway is controlled by [Ca2+]i oscillations [67]. Finally, it has been demonstrated that hepatocyte calcium signaling was deeply remodeled during liver regeneration in the rat, contributing to the regulation of bile flow and cell proliferation [52, 68]. After partial destruction of the liver (in experimental or pathologic context), the initial mass of the organ is restored through compensatory growth of the remnant liver. A complex network of cellular interactions (direct and indirect) orchestrates the regulation of regeneration. In the first hours after partial hepatectomy, plasma concentrations of some calcium mobilizing agonists (in particular HGF and noradrenaline) strongly rise. Our group has recently demonstrated that there was a vasopressin hypersecretion in 1 h after partial hepatectomy in the rat [52] and that this hormone contributed to “push” quiescent hepatocytes to reenter the DNA synthesis S phase. We observed a deep remodeling of the vasopressin receptor hepatic expression and of the related calcium signaling after partial hepatectomy, contributing to the effects of this agonist on liver regeneration [52, 68]. Among the multiple agonists involved in these interactions, some are known to mobilize intracellular calcium, including noradrenaline [69], vasopressin [52], adenosine triphosphate (ATP) [70], EGF, and HGF [71], although the precise impact of calcium mobilization is far from being understood. Sites of the regulatory action of Ca2+ on cell cycle, yet incompletely known, are thought to be essential for G0 to G1, G1 to S phase transitions of the cell cycle, and during mitosis [72, 73]. It has been shown recently in liver cell lines that Ca2+ influx through the TRPC6 channel was of importance for neoplastic cell proliferation [17]. Also, in an already emphasized study, nuclear calcium signaling has been shown to regulate tumor cell proliferation in vivo [55].

Calcium and Cell Death in the Liver It is well known that calcium is a central regulator of cell death, in particular because alteration of calcium homeostatic control, i.e., the perturbation of calcium ions distribution in the different cell compartments, can interfere with the machinery of apoptosis [74]. In

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this machinery, mitochondria take a central position, partly because if they sequester too much calcium ions, they can swell and rupture, releasing different proapoptotic factors. Calcium ions may orchestrate apoptosis through different mechanisms involving the InsP3R [75]. First, cytochrome c has been shown to bind InsP3R and to elicit an amplification process leading to a massive cytochrome c release and apoptosis [76]. Second, several lines of evidence suggest that the balance between pro- and antiapoptotic Bcl-2/Bax family members regulate the ER calcium content, and that the transfer of calcium from the ER  to the mitochondria triggers cell death [77]. Furthermore, the antiapototic protein Bcl-2 can directly interact with the InsP3R, thereby inhibiting calcium release and apoptosis under certain stimulation [78]. Third, the InsP3Rs can be cleaved by caspases and form constitutively active channels that continuously leak Ca2+, although the involvement of this cleavage in the early stages of apoptosis is not entirely confirmed [79]. Potential impact of these different pathways has been emphasized in T lymphocytes and neurones, but has not been investigated in hepatocytes or other liver cells.

Calcium and Ischemia-reperfusion Injury It has been stated for a long time that intracellular calcium overload is involved in liver lesions occurring in the context of liver transplantation, due to reperfusion of the organ after ischemia [80]. Several studies pointed calcium homeostasis alteration as an important parameter in the pathophysiology of these lesions, and thus as a good candidate for therapeutic targeting. In isolated perfused liver experiments, an increase in hepatocyte calcium content has been reported to occur immediately after reperfusion [81]. At the hepatocyte level, it has been reported after cold preservation and warm reoxygenation in the rat, that steady state levels of Ca2+ were elevated, and that calcium responses elicited by ATP were amplified due to a related increase in the InsP3-sensitive calcium pool [82]. The involvement of an increased Ca2+ influx through yet unknown membrane channels has also been hypothesized to explain these alterations of Ca2+ homeostasis in the reperfusion

T. Tordjmann

phase. In the endothelial cell, a particularly targeted cell during reoxygenation of the liver, calcium responses to agonists were found to be attenuated, with unknown impact on cell viability and adhesion molecule expression [83]. It has also been shown that an elevation of cytosolic calcium was mandatory for acquisition of hepatocyte protection during liver preconditioning [84]. Finally, early Kupffer cell activation after reperfusion, which is crucial for liver damage, can be prevented by calcium channel inhibitors, with protective effects on ischemia reperfusion injury [85].

Calcium and Viral Hepatitis Hepatitis viruses (HBV and HCV) may have effects on calcium signaling, while viral infection, and its consequences may be dependant on intracellular calcium [86]. It has been shown that HBV DNA (HBX gene) can integrate the SERCA1 gene and code for a chimeric protein, in which the truncated SERCA1 cannot pump calcium from the cytosol to the ER [87]. Such truncated SERCA proteins, also found in normal livers, reduce the steady state level of calcium in the ER, increase the Ca2+ leakage from this compartment, and have been proposed to have a role on calcium transfer between the ER and the mitochondria, and thereby, create a potential impact on the proliferation/apoptosis balance [88]. Multiple levels of interaction between the hepatitis B X protein and calcium signaling have been reported. First, the effect of HBX protein on HBV DNA replication has been shown to depend on the elevation of cytosolic calcium [89]. Although the mechanisms are far from being known, this cytosolic Ca2+ increase depends on: mitochondrial HBX expression interfering with calcium release from these organelles [90]; HBX driven caspase 3 dependent PMCA proteolysis leading to an impaired Ca2+ extrusion out of the cell [91]; and on Ca2+ release from the ER. Second, HBV core assembly has been reported to depend on an increase in cytosolic Ca2+. Finally, a number of transcription factors and kinases activated by HBX protein depend also on calcium signaling [86]. HCV core and NS5A proteins have also been reported to interact with calcium signaling in several ways, inducing ER calcium depletion and ER stress, and leading to apoptosis [92].

30  Calcium Signaling

Summary

›› Intracellular calcium orchestrates a very large

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array of biological processes, thanks to modulation of signals in terms of speed, amplitude, and spatiotemporal patterning. Calcium signals induced by agonists such as hormones and neurotransmitters constitute a kind of language which is translated into physiological responses by the cells. Because the hepatocyte has long been a model cell for the study of calcium oscillations and waves, the large body of accumulated data allows drawing a detailed picture of hepatocyte calcium signaling, from subcellular to lobular level. Calcium signals in hepatocytes are highly organized, in the form of oscillations and waves, both at the intracellular and at the intercellular level. Directional calcium waves are, at least in part, generated thanks to the heterogeneous distribution of either the different InsP3 receptor isoforms (at the subcellular level), or the agonist receptors (at the lobular level). However a lot remains to be discovered concerning the functions that these complex signals govern in the liver. Calcium signals, including those generated in the nucleus by growth factors may have important impact on physiological and pathological hepatocyte proliferation. Conversely, hepatocyte apoptosis may be regulated by mitochondrial calcium movements. Finally, intracellular calcium movements may be involved in the pathophysiology of ischemia reperfusion injury and viral hepatitis, with yet unknown pathophysiological consequences on liver diseases. In fact, since classical calciumdependant liver functions have been reported (glucose metabolism, bile secretion…) a broader picture of physiological and pathophysiological impact of intracellular calcium in the liver is only emerging.

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Multiple Choice Questions 1. Intracellular calcium oscillations: (a) Have been first discovered in Kupffer cells (liver resident macrophages) (b) Always originate from the periodic opening of Ca2+ channels located in the plasma membrane in nonexcitable cells (c) Can be transmitted from cell to cell in multicellular systems of coupled hepatocytes (d) Are specifically generated in hepatocytes by the agonist arginine vasopressin, but not by other hormones or growth factors (e) Have not been observed in the isolated perfused liver 2. In eukaryotic cells, intracellular calcium: (a) Is uniformly distributed in the different subcellular compartments (b) Is kept at low (nanomolar) concentrations in the ER (c) Is passively equilibrated between extra and intracellular spaces (d) Easily diffuses across the cytosol (e) Cannot diffuse along large distances due to abundant intracellular buffers 3. SERCA pumps actively transport calcium: (a) From the ER to the cytosol (b) From the cytosol to the ER (c) From the extracellular space to the cytosol (d) From the cytosol to the extracellular space (e) From the mitochondria to the cytosol 4. Calcium signals in hepatocytes can be elicited in response to: (a) Hormones (b) Paracrine agonists like ATP (c) Cytokines (d) Growth factors (e) All the above 5. Which statement is wrong: (a) The temporal pattern of calcium signals is of major impact as to the expression of transcription factors (b) The mitochondrial metabolism can be optimized by calcium oscillations

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(c) The calcium oscillation frequency increases with the concentration of external agonist (d) STIM1 is a sensor of Ca2+ depletion in the ER (e) InsP3 is less soluble than calcium in the cytosol, and from its production site it cannot diffuse freely

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T. Tordjmann 1 inositol trisphosphate receptor accelerates apoptotic cell death and induces inositol trisphosphate-independent calcium release during apoptosis. J Biol Chem 279(41): 43227–43236 80. Nieuwenhuijs VB, De Bruijn MT, Padbury RT, Barritt GJ (2006) Hepatic ischemia-reperfusion injury: roles of Ca2+ and other intracellular mediators of impaired bile flow and hepatocyte damage. Dig Dis Sci 51(6):1087–1102 81. Isozaki H, Fujii K, Nomura E, Hara H (2000) Calcium concentration in hepatocytes during liver ischaemia-reperfusion injury and the effects of diltiazem and citrate on perfused rat liver. Eur J Gastroenterol Hepatol 12(3):291–297 82. Elimadi A, Haddad PS (2001) Cold preservation-warm reoxygenation increases hepatocyte steady-state Ca(2+) and response to Ca(2+)-mobilizing agonist. Am J Physiol Gastrointest Liver Physiol 281(3):G809–G815 83. Auger S, Vallerand D, Haddad PS (2003) Cold preservationwarm reperfusion perturbs cytosolic calcium ion homeostasis in rat liver sinusoidal endothelial cells. Liver Transpl 9(2): 150–159 84. Carini R, Castino R, De Cesaris MG, Splendore R, Démoz M, Albano E, Isidoro C (2004) Preconditioning-induced cytoprotection in hepatocytes requires Ca(2+)-dependent exocytosis of lysosomes. J Cell Sci 117(Pt 7):1065–1077 85. Jiang N, Zhang ZM, Liu L, Zhang C, Zhang YL, Zhang ZC (2006) Effects of Ca2+ channel blockers on store-operated Ca2+ channel currents of Kupffer cells after hepatic ischemia/reperfusion injury in rats. World J Gastroenterol 12(29):4694–4698 86. Chami M, Oulès B, Paterlini-Bréchot P (2006) Cytobiological consequences of calcium-signaling alterations induced by human viral proteins. Biochim Biophys Acta 1763(11): 1344–1362 87. Chami M, Gozuacik D, Saigo K, Capiod T, Falson P, Lecoeur H, Urashima T, Beckmann J, Gougeon ML, Claret M, le Maire M, Bréchot C, Paterlini-Bréchot P (2000) Hepatitis B virus-related insertional mutagenesis implicates SERCA1 gene in the control of apoptosis. Oncogene 19(25):2877–2886 88. Chami M, Gozuacik D, Lagorce D, Brini M, Falson P, Peaucellier G, Pinton P, Lecoeur H, Gougeon ML, le Maire M, Rizzuto R, Bréchot C, Paterlini-Bréchot P (2001) SERCA1 truncated proteins unable to pump calcium reduce the endoplasmic reticulum calcium concentration and induce apoptosis. J Cell Biol 153(6):1301–1314 89. Bouchard MJ, Wang LH, Schneider RJ (2001) Calcium signaling by HBx protein in hepatitis B virus DNA replication. Science. 294:2376–2378 90. McClain SL, Clippinger AJ, Lizzano R, Bouchard MJ (2007) Hepatitis B virus replication is associated with an HBxdependent mitochondrion-regulated increase in cytosolic calcium levels. J Virol. 81:12061–12065 91. Chami M, Ferrari D, Nicotera P, Paterlini-Bréchot P, Rizzuto  R (2003) Caspase-dependent alterations of Ca2+ signaling in the induction of apoptosis by hepatitis B virus X protein. J Biol Chem 278(34):31745–31755 92. Benali-Furet NL, Chami M, Houel L, De Giorgi F, Vernejoul F, Lagorce D, Buscail L, Bartenschlager R, Ichas F, Rizzuto R, Paterlini-Bréchot P (2005) Hepatitis C virus core triggers apoptosis in liver cells by inducing ER stress  and ER calcium depletion. Oncogene 21;24(31): 4921–4933

31

HBV Signaling Massimo Levrero and Laura Belloni

The Hepatitis B Virus Infection with hepatitis B virus (HBV) continues to be a major health problem with about 400 million people chronically infected worldwide who are at high risk of developing liver cirrhosis and hepatocellular carcinoma (HCC) [1]. HBV is a member of the Hepadnaviridae family which includes small enveloped DNA viruses infecting primates, rodents, and birds [2, 3]. One common characteristic of these viruses is their high species and cell-type specificity, as well as a unique genomic organization and replication mechanism. The genome of all hepadnaviruses is extremely compact consisting of four overlapping open reading frames (ORF). The S, Core and Pol ORFs encode viral proteins that are essential structural components of viral replication and  assembly (envelope proteins (SHBs, MHBs and LHBs), core (HBc) and reverse transcriptase (RT)/ polymerase (Pol)). The HBeAg, which is generated by the intracellular processing of the preC/Core protein at the endoplasmic reticulum (ER) levels as well as by intracellular and extracellular proteolysis of free HBc proteins, is thought to play an important role in HBV pathogenesis by influencing the host immune system . The X ORF encodes for the regulatory X protein (hepatitis B virus X protein (HBx)) which is an essential factor for viral replication and it is considered to be one of the most important determinants of HBVinduced hepatocarcinogenesis [4]. Whereas many

aspects of viral replication have been elucidated, the initial phases of hepadnaviral infection (attachment of mature virions onto host cell membranes and viral entry) are still less understood, and the search for putative cellular receptors and coreceptors is still very active. An additional important feature of hepadnaviruses replication is the relative low fidelity of the ­enzimatic machinery that leads to high genomic heterogeneity and variability [2, 3]. HBV infection may occur in HBsAg-negative individuals with or without serologic markers of previous infection (antibody to HBsAg (anti-HBs) and antibody to hepatitis B core antigen (anti-HBc)). This condition, known as “occult” HBV infection, is characterized by a strong suppression of virus ­replication and gene expression, and is usually identified only by highly sensitive molecular biology techniques [5, 6]. In this short review, we will discuss the interactions between HBV viral factors and host proteins that are involved in HBV life cycle and pathogenesis. Chapter 2 will describe the role of cellular proteins in regulating HBV transcription and replication, including the formation of the HBV episomal covalently closed circular DNA (cccDNA) and its epigenetic regulation. Chapter 3 will analyze the role of the HBx protein in modifying cellular transcription and signaling. In Chaps. 4 and 5, the impact of HBV and its proteins on liver physiopathology will be discussed with a focus on hepatocytes viability, metabolism and transformation.

M. Levrero (*) Dipartimento di Medicina Interna, Sapienza Universita’ di Roma, Policlinico Umberto I, Viale del Policlinico 155, 0061 Rome, Italy e-mail: [email protected] J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_31, © Springer-Verlag Berlin Heidelberg 2010

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HBV Life Cycle and the Role of Cellular Proteins HBV Life Cycle After viral particles attachment to an unknown receptor on the host cell and internalization, the virion relaxed circular (RC) DNA is delivered to the nucleus, where it is repaired, by yet unknown cellular factors, to form a covalently closed circular (CCC) DNA [7] (Fig. 31.1). The episomal cccDNA serves as the template for the transcription of the pregenomic RNA (pgRNA) and other viral mRNAs by the host RNA polymerase II. The transcripts are then exported to the cytoplasm, where translation of the viral proteins occurs. RT binds to pgRNA and triggers assembly of the core proteins into immature, RNA-containing nucleocapsids [8, 9].

The immature nucleocapsids undergo a process of maturation whereby pgRNA is reverse transcribed by RT to make the mature RC DNA. A unique feature of hepadnavirus reverse transcription is the RT primed initiation of minus-strand DNA synthesis, which leads to the covalent linkage of RT to the 5’ end of the minus-strand DNA [10–13]. The mature RC DNA-containing nucleocapsids are then enveloped by the viral surface proteins in the ER and secreted as virions (secretion pathway) or, alternatively, are recycled back to the nucleus to further amplify the pool of CCC DNA (recycling pathway) [7, 14].

Cellular Proteins and HBV Replication A number of cellular proteins play important roles in crucial steps of HBV life cycle including the initiation

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Fig. 31.1  Virus–cell interactions in the hepadnaviral life cycle. HBV binds to an unknown receptor on hepatocytes, internalized, uncoated, and delivers the RC DNA genome to the nucleus. RC DNA is repaired to form the CCC DNA that serves as the transcriptional template. Viral RNAs, including the pgRNA, are exported to the cytoplasm, where translation of the viral proteins occurs. The pgRNA is packaged together with RT into immature nucleocapsids composed of core protein and reverse transcribed

within the nucleocapsids into the mature RC DNA form. Mature nucleocapsids are then enveloped by the viral envelope proteins and secreted extracellularly as virions or recycled to the nucleus to amplify the cccDNA pool. Cellular cofactors of viral replication are highlighted. RC DNA relaxed circular DNA; cccDNA covalently closed circular DNA; pgRNA pregenomic RNA; RT reverse transcriptase; P Kinase; PPase Phosphatase; Hsp 90 heat-shock protein 90

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of HBV reverse transcription, nucleocapsid maturation, formation of the HBV episomal DNA and its epigenetic regulation (Fig.  31.1). Additional cellular proteins, which will not be further discussed, are involved in viral entry and virion secretion.

enveloped and secreted [20, 21, 24]. Identification of the host kinase(s) and phosphatase(s), responsible for HBc modifications and nucleocapsid maturation, would provide new molecular targets for antiviral therapies.

Chaperones and RT–pgRNA Interaction

cccDNA Formation

The cellular Hsp90 complex, which includes the Hsp90, Hsp70, Hsp40, Hop, and p23 chaperone proteins, is required to stabilize a transient conformation that permits the formation of the ribonucleoprotein (RNP) complex between the RT and the e region of the pgRNA during viral assembly and reverse transcription [15, 16].

The HBV cccDNA exists as an episome in the nucleus with an estimated copy number of 5–50 molecules per infected cell [14, 29, 30]. The cccDNA associates with cellular histone and non histone proteins and is organized as a minichromosome [31] regulated by epigenetic changes [32]. Although the establishment and maintenance of a pool of viral cccDNA is a critical step in HBV infection, very little is known about the viral and cellular factors involved in the cccDNA formation. Since the conversion of RC-DNA to cccDNA can be thought of as a DNA damage response, it has been speculated that host cell DNA repair enzymes might be involved in this step. The observation that the liver of HBV-transgenic mice fails to accumulate detectable levels of cccDNA, despite the presence of high levels of RC DNA precursor, suggests the need for species-specific host factors [33]. However, no specific host factors participating in cccDNA formation have been identified so far and it is possible that the viral RT may be the only activity required for this conversion step.

Nucleocapsid Maturation The phosphorylation state of hepadnaviruses nucleocapsids, which are composed of 180–240 HBc molecules, correlates to the maturation stage of the virus. The C-terminal functional domain (CTD) of HBc is arginine-rich and contains several serine/threonine phosphorylation sites [17–20]. Immature nucleocapsids that contain RNA are phosphorylated at the six CTD sites, while mature, DNA-containing nucleocapsids inside the cells or in extracellular virions are completely dephosphorylated [20–22]. HBc phosphorylation affects HBV pgRNA packaging and DNA synthesis [22, 23], and is required for DHBV minusstrand DNA synthesis [19, 24]. Purified HBV particles demonstrate in vitro kinase activity [25] and since the viral genome does not code for any proteins with kinase activity, nucleocapsids are predicted to package a kinase of cellular origin during the assembly process [25]. The same kinase may play a role in the uncoating process following viral entry by destabilizing the capsid through the phosphorylation of the core subunits [26]. Despite several candidate kinases have been reported to associate with nucleocapsids and phosphorylate HBc in vitro (a 46-kDa kinase, a cdc2-like kinase, SRPK1 and SRPK2 [26–28]), their direct role in viral replication in vivo remains elusive. A cellular phosphatase has also been predicted to remove the phosphate groups during the late stage of reverse transcription, before the viral particles are

cccDNA Transcription The HBV cccDNA is the sole intranuclear transcriptional template for the synthesis of all viral RNAs, including the pgRNA. Transcription of the HBV genes is controlled in cis by four viral promoters and two enhancer elements. In vitro studies have demonstrated that transcription of the HBV genome is regulated by a variety of transcription factors, such as the liverenriched transcription factors C/EBP, retinoid X receptor alpha (RXRa), peroxisome proliferator-activated receptor alpha (PPARa), hepatocyte nuclear factor (HNF) 3, HNF4 and the ubiquitous transcription factors, Sp1, NFkB and p53 (reviewed in [3]). Members of the nuclear receptor family (i.e., HNF4) may be

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HBV Core H3/H4 Histones Acetylated histones (AcH3/AcH4) Methylated H3/H4 and DNA

Fig. 31.2  The cccDNA chromatin immunoprecipitation assay. The HBV cccDNA ChIP assay is a chromatin immunoprecipitation-based quantitative technique that allows study of the recruitment in vivo of cellular and viral proteins onto the HBV minichromosome. The assay combines a cccDNA ChIP step

with a sensitive and specific real-time PCR protocol for cccDNA quantification. (Right) Schematic representation of cccDNAbound histones acetylation status and the recruitment of chromatin-modifying enzymes onto the viral minichromosome in relation to viral replication and infection phase

involved in the downregulation of some viral promoters [34]. Recently, using a new chromatin immunoprecipitation-based assay (Fig.  31.2), we have reported that HBV replication is regulated by the acetylation status of histones H3 and H4 bound to CCC DNA minichromosome [32]. Furthermore, cellular acetyltransferases (PCAF, p300 and CBP) and cellular deacetylases (HDAC1) are recruited in vitro and in vivo onto the HBV cccDNA. Treatment with histone deacetylase inhibitors (HDACi) increase HBV transcripts, HBV replicative intermediates in the cytoplasm and secreted HBV viral particles [34]. Thus, HBV transcription and replication can be modulated by epigenetic changes to the cccDNA. and host transcription factors regulate transcription of cccDNA by modifying the cccDNA-bound histones via the recruitment of acetyltransferases and deacetylases to the viral promoters [35].

HBX, Transcription and Cell Signaling HBx and the Virus HBx is encoded by the smallest HBV ORF and is 154 amino acids in size, with a molecular weight of about 17.5 kDa. It has been suggested to affect viral, as well as host cell functions, by modulating a wide variety of cellular processes, including transcription, cell cycle progression, DNA damage repair, and apoptosis [4] (Fig. 31.3). HBx is essential for viral infection in vivo [36–38], and potentiates viral replication in cell culture [34, 38]. Although it is not well defined, the subcellular localization of HBx seems to be mainly cytoplasmic, with a small fraction in the nucleus [39]. Thus, it is believed that HBx may have a dual role in transcriptional regulation; cytoplasmic HBx could affect the

31  HBV Signaling Fig. 31.3  Multiple cellular targets of the regulatory protein HBx. The regulatory protein HBx, in addition to be required for viral replication, contributes to hepatocytes transformation by multiple mechanisms mediated by interaction with a large number of cellular targets

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regulation of signal transduction pathways while nuclear HBx may function at the promoter level [40]. By coupling cccDNA quantification [41] and the cccDNA ChIP assay [32], we have shown that the HBx produced in HBV-replicating cells is recruited onto the cccDNA minichromosome [34] HBV mutants that does not express HBx are impaired in their replication and that exogenously expressed HBx transcomplements the replication defects [34]. The kinetics of HBx recruitment on the cccDNA parallels HBV replication and is similar to that of the PCAF/GCN5 acetyltransferase [34]. Despite this observation and the physical interaction between the two proteins, we could not find any significant change in the recruitment of PCAF on the cccDNA in cells replicating the HBx-defective virus. Instead, we found that cccDNA-bound histones are more rapidly hypoacetylated in cells replicating the HBx mutant and the recruitment of the p300 acetyltransferase is severely impaired whereas the recruitment of the histone deacetylases, hSirtl and HDAC1, is increased and occurs earlier [34] (Fig. 31.2). Accordingly, we show that in cells replicating the HBx mutant the pool of cccDNA is not reduced but the HBx mutant cccDNA transcribes significantly less pgRNA [34].

HBx and Cellular Transcription HBx has been shown to transactivate a variety of viral and cellular promoters [3, 39]. Since HBx does not directly bind DNA, its ability to activate transcription of host genes is thought to occur indirectly by

interaction with nuclear transcription factors or by the activation of different signal transduction pathways. In support of its role in transcriptional regulation, HBx was reported to associate with several components of the basal transcriptional machinery, including TFIIB, TFIIH, and RBP5, a subunit of mammalian RNA polymerase [42] (Table 31.1). It has also been demonstrated that it binds to transcription factors including CREB, ATF-2, and AP2 to modify their activities [43,  44] (Table  31.1). It was recently demonstrated that HBx could interact and cooperate with CREB-binding protein (CBP)/p300 to synergistically enhance CREB activity [45]. By combining in silico analysis and anti-HBx

Table 31.1  HBx-interacting proteins Apoptosis p53 Mitochondrial-associated HVDAC3 proteins Heat shock proteins Hsp60, Hsp70 Signal transduction pathways 14-3-3 Proteasome complex PSMA7, PSMC1 DNA repair UV-DDB1 Chromatin remodeling PCAF, CBP, p300, HDAC1, DNMT3a Basal transcription RPB5, TFIIB, TBP, TFIIH (ERCC2, ERCC3) Transcription factors RXR, RMP, Egr-1, NFkB bZip transactivators CREB, ICER IIg, Gadd143/ Chop10, ATF2, ATF3, NF-IL6 Peptidyl-prolyl isomerase Pin1 Unknown HBXIP, PA28 HBx physically interacts with several cellular proteins that modulate cell proliferation, cell death, transcription, and DNA repair

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chromatin immunoprecipitation (ChIP), we could demonstrate that HBx is recruited on several CREB target genes, including PCNA, IL8, FAS, and IGFBP3, and the SREBP1 transcription factor, which are actively transcribed in normal and HBV-infected livers and in HCC samples (Levrero and Guerrieri, unpublished observations) (Fig. 31.4). Interestingly, binding sites for SREBP1 are frequently found together with CRE-sites on HBx direct target genes, suggesting the existence of a cooperative HBx/SREBP1/CREB transcriptional network.

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Fig. 31.4  HBx and gene transcription. (a) HBx, by interacting with DNA-bound transcription factors, may help recruitment of histone acetyltransferases and thus exhibit its transactivation function. HBx interacts with CBP to activate a number of CREB direct target genes (46 and personal observations). (b) HBx interferes with the binding of potential repressor(s) on target genepromoters thus allowing hypomethylation, histone acetylation and expression of the target genes (IGFBP3 and CDH6 in [131]). (c) HBx favors the recruitment of DNMT3A or HDAC1, resulting in hypermethylation and heterochromatinization of the promoter region and gene silencing (MT1F and IL4R genes in [131])

HBx and Signal Transduction Aside from its transactivating capabilities, a number of different cytoplasmic signal transduction cascades appear to be affected by HBx, including NFkB, RasRaf-mitogen-activated protein kinase Ras-Raf-MAPK, extracellular signal-regulated kinase (ERK), stressactivated protein kinases/NH2-terminal-Jun kinase (SAPK/JNK), protein kinase B (PKB/Akt), and Janus kinase/STAT (JAK/STAT) [39, 46–48] (Fig. 31.3 and Table 31.1). HBx also stimulate cellular calcium-signaling pathways, resulting in the release of Ca2+ ions into the cytosol [49, 50]. This leads to the activation of focal adhesion kinase (FAK) and proline-rich tyrosine kinase 2 (Pyk2), and the subsequent activation of Src tyrosine kinases and downstream signaling pathways such as Ras-Raf-MAPK pathway [51, 52]. The consequences of the HBx-mediated modifications of Ca2+signaling are an increase in core particles assembly and HBV replication, activation of AP1 and NFATdependent transcription, and induction of apoptosis (reviewed in [54]). The ability of HBx to mobilize intracellular Ca2+ deposits may provide an unified mechanism by which HBx exerts many of its pleiotropic activities, including transcription, cell cycle control, and apoptosis.

HBV and Liver Pathophysiology HBV and Apoptosis Although HBV is considered as a non cytopathic virus [53] and liver damage has been always regarded as

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immunomediated [54] hepadnavirus-induced apoptosis, cytopathic effects have been described in several experimental model systems. Indeed, the induction of apoptosis is a hallmark of many viruses infecting human cells and different HBV proteins have been described to impact on cell viability. Intracellular retention of the HBV large surface protein has been shown to induce cellular vacuolization and apoptosis in hepatoma cell lines [55, 56] and chronic liver damage eventually followed by the development of liver tumors in transgenic mice [57]. Interestingly, a drug-resistance mutation on the Pol gene selected by the nucleot(s)ide analogs, lamividine and adefovir, results in changes in the overlapping S ORF leading to the expression of envelope proteins with secretory defects that may potentially contribute to HBV pathogenesis in addition to affecting the efficacy of the antiviral therapy [58]. A duck hepatitis B variant containing a single amino acid change in the large surface antigen resulting in accumulation of cccDNA resulted in a strong cytopathic effect in hepatocytes in vitro and in vivo [59, 60]. In this system, the level of viral replication and cccDNA formation correlated with cytopathic effects in infected hepatocytes [59, 60]. It has to be noted that the corresponding mutation has not been found in vivo in HBV infection and it does not recapitulate the duck phenotype in in vitro HBV replication models. A viral variant, associated with fulminant hepatitis and containing two mutations in the basal core promoter, has been shown to induce apoptosis in primary Tupaia hepatocytes [61]. Induction of apoptosis in this model was independent of viral replication suggesting that viral protein synthesis was sufficient for the virus-induced hepatocyte cell death and, since the two core promoter mutations resulted in two amino acid changes in the X ORF, HBx has been indicated as a potential candidate [61]. Indeed, HBx protein has been widely implicated in the induction of apoptosis in both a p53-dependent and p53-independent manner [62–64]. A loss of the proapoptotic function of HBx has been associated to hepatocytes transformation. Apoptosis-defective naturally occurring mutants of HBx have been demonstrated in HCC samples but not in the peritumoral tissues of HBV-related HCC patients [65] The relationship between HBx and apoptosis is nevertheless complex and HBx has also been described to inhibit p53-dependent apoptosis by interacting with and sequestering p53 in the cytoplasm [66]. The molecular mechanism underlying the ability of HBx to modulate apoptosis has long

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remained unclear. Recently, HBx has been shown to interact with c-FLIP and inhibit its recruitment to the death-inducing signaling complex, resulting in hyperactivation of caspase-8 and caspase-3 by death signals [63]. In the context of HBV chronically infected livers, apoptosis of hepatocytes is mediated by several molecular pathways, which involve at least three members of the TNF superfamily (TNF, Fas ligand (FasL) and TRAIL). Different from TNF and FasL, TRAIL preferentially induces apoptosis of tumor cells and virusinfected cells but not normal cells [67–69]; blocking the TRAIL pathway using soluble death receptor 5 (DR5) significantly ameliorates liver inflammation in a mouse model of hepatitis [70]. Both HBx [69] and truncated middle hepatitis B surface protein (MHBs(t)) [71] have been described to sensitize hepatocytes to TRAIL-induced apoptosis. In the case of HBx, TRAILinduced apoptosis was found to be enhanced through Bax upregulation [69]. The real impact of HBV-induced apoptosis in viral pathogenesis and liver disease progression in vivo remains, however, to be further clarified. Whereas direct induction of apoptosis by viral proteins or sensitization to DNA damage or death receptorsinduced apoptosis may contribute to liver damage and HBV pathogenicity, blocking hepatocytes apoptosis may contribute to both viral persistence and hepatocyte transformations as well as to chemoresistance in transformed cells. Recently, HBV core protein (HBc) has been found to be a potent inhibitor of TRAILinduced apoptosis in hepatoma cells [72]. Knocking down HBc expression in hepatoma cells transfected with the whole HBV genome enhanced TRAILinduced apoptosis and, interstingly, when present in the same cell HBc blocked the proapoptotic effect of the HBx. HBc-related resistance to TRAIL-induced apoptosis was associated with a significant reduction in death receptor 5 (DR5) expression and HBc was found to significantly repress the promoter activity of the human DR5 gene [72]. On the basis of the presence of nucleic acid-binding motifs, nuclear localization signals, phosphorylation sites, and its localization in the nucleus, HBc may act as a gene regulatory protein. Indeed, HBc has been reported to inhibit the expression of the b-interferon gene, p53 and HBx acting at the promoter level [73–75]. Dr5 promoter activity is regulated by both p53 and NFkB but microarray analysis indicates that HBc represses P53 but not NFkB expression in hepatocytes [72]. Importantly, HBc

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inhibited hepatocyte death in a mouse model of HBVinduced hepatitis and in patients with chronic hepatitis DR5 expression in the liver was significantly reduced [72]. Thus, different HBV proteins have different, even opposite, effects on TRAIL-induced hepatocyte apoptosis, the predominant death receptor pathway mediating apoptosis of virus infected cells in the liver. If the proapoptotic proteins, such as HBx, are dominant, HBV-infected hepatocytes may die and contribute to liver damage. By contrast, if the antiapoptotic viral proteins, such as HBc, prevail infected hepatocytes may not undergo apoptosis favoring the establishment of chronic HBV infection and persistence of damaged hepatocytes in the early stages and cell transformation and chemo-resistance in the later phases.

HBV and Lipid Metabolism Chronic infection with both HBV and HCV are frequently associated with hepatic steatosis. Indeed, although steatosis is commonly considered a hallmark of HCV chronic infection a variable degree of fat accumulation in hepatocytes is described in 27–51% of HBV-infected patients [76–78]. In experimental animals, high-level HCV replication during acute infection is associated with the modulation of multiple genes involved in lipid metabolism [79]. In addition, drugs that control cholesterol and fatty acid biosynthesis regulate the replication of the subgenomic HCV replicon [79]. HCV core protein has been shown to regulate genes related to fatty acid biosynthesis, including liver X receptor-a (LXRa) and sterol regulatory element binding protein-1c (SREBP-1c) [80]. Much less is known on how HBV infection may modulate steatogenic pathways at the molecular level. cDNA microarrays expression profiling analysis of HBVtransgenic mouse livers has shown an upregulation of genes involved in the biosynthesis of lipids, such as fatty acid synthase (FAS) and SREBP-2 [81]. Moreover, HBx has been shown to cause lipid accumulation in hepatic cells through the activation of SREBP-1 and the peroxisome proliferator-activated receptor g (PPARg) [82]. More recently, LXRa and LXRb have been identified as the primary targets of HBx in HBV-associated hepatic steatosis [83]. LXRa and HBx, colocalized in the nucleus, are physically associated and HBx potentiates the transcriptional activity of LXRa by

M. Levrero and L. Belloni

recruiting CREB binding protein to the promoter of LXRs target genes. As a confirmation in vivo that HBx-induced lipogenesis is LXR-dependent, the expression of LXR was found to be increased in the livers of ­HBx-transgenic mice and there was a significant increase in the expression of LXRb, SREBP-1c, FAS, and stearoyl-coenyzme A desaturase-1 in HCC in comparison with adjacent nontumorous nodules in human HBV-associated HCC specimens [83].

HBV and HCC Viral Epidemiology HCC is one of the most frequent solid tumors worldwide and represents the third cause of mortality among deaths from cancer [84]. HCC frequency is particularly high in Asia and Africa because of the high frequency of viral hepatitis infections and exposure to Aflatoxin B1 (AFB1). Although the increased incidence of HCC in the Unites States and UK over the last 10 years [85, 86] is thought to reflect the increase of viral hepatitis C infections, recent estimates attribute to HBV >50% of HCC cases worldwide. Additional etiological factors that often represent cofactors of an underlying HBV(or HCV)-related chronic liver disease include toxins and drugs (e.g., alcohol, aflatoxins, microcystin, anabolic steroids, and vinyl chloride), metabolic liver diseases (e.g., hereditary haemochromatosis, a1-antitrypsin deficiency), steatosis, nonalcoholic fatty liver diseases and diabetes [87, 88]. Lifetime risk of developing HCC is estimated to be 10–25-fold greater for chronic HBV carriers, as compared with noninfected populations, and the risk of HCC has been shown to be increased even in patients with occult HBV infection and after hepatitis B surface antigen (HBsAg) clearance [85, 89, 90]. Persisting high HBV replication associates with the risk of developing HCC [91]. A number of studies have suggested that HBV genotypes or variants might harbor different oncogenic potential. A recent study has shown that HBV genotype C infection (compared to genotype B) is an independent risk factor for HCC development, with an adjusted relative risk of 2.8%, while the relative risk associated with liver cirrhosis is 10.2% [92]. Other studies have shown that the prevalence of T 1762/A

31  HBV Signaling

1764 mutation in the basal core promoter increases with the progression of liver disease, and that this mutation is significantly associated with the development of HCC, in both genotypes B and C [93]. Furthermore, the T1762/A1764 mutation can be detected in plasma up to 8 years before HCC diagnosis; this mutation might therefore be considered as a strong predictive biomarker [94].

Genomic and Transcriptomic Analysis of HBV-Related HCCs Extensive evidence indicates that HCC is an extremely complex tumor at the molecular and genetic levels [95]. Abnormal hepatocyte proliferation and survival result from a complex network of interacting signaling pathways that include increased growth factor/ growth factor receptor signaling (e.g., EGFR and IGFR), costitutive mitogenic intracellular signaling (e.g., Raf/MEK/ERK, PI3K/AKT and Wnt/-catenin pathways), inactivation of proapoptotic and antiproliferative tumor suppressors (e.g., p53 and pRb), increased antiapoptotic signaling (e.g., PTEN/AKT and NFkB). The initial steps of carcinogenesis are followed by tumor angiogenesis, mediated by tumor cell/stromal cell interactions and the activation of multiple angiogenic factors [95]. In HCCs, p53 gene is mutated in about 20% of cases, with important variations in the mutational rate between tumors of different geographical locations [96]. A hotspot mutation affecting p53 at codon 249 was originally described in HCCs from regions with high prevalence of HBV infection and high levels of dietary aflatoxins, and it is considered as a hallmark of aflatoxin B1 [97]. Genome-wide analysis of genetic alterations in HCC has showed that genetic alterations are not randomly distributed in tumors but are closely associated in clusters. These studies have identified two main mechanisms of hepatocarcinogenesis and allowed the classification of HCCs in subsets of genetically and molecularly homogeneous tumors. In the first mechanism, HBV infection is closely related with a higher chromosome instability together with TP53 and AXIN1 mutations [98]. The second mechanism, defined by the presence of b-catenin mutations and chromosome 8p deletion in a context of lower chromosome instability, is associated with the

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absence of HBV infection [98]. A genome-wide transcriptomic analysis of well-annotated HCCs [99] has identified six subgroups of HCC (termed G1 to G6) associated with specific clinical and genetic characteristics (Fig.  31.5). G1 and G2 tumors were both related to HBV infection and displayed frequent activation of the PI3K/AKT pathway but differed for the overexpression of genes expressed in fetal liver and controlled by parental imprinting (G1) and the frequent mutation of the PIK3CA and TP53 genes (G2). In addition to genetic mutations, epigenetic mechanisms, such as hypermethylation of promoters containing CpG islands, have been shown to frequently modify gene expression patterns in HCC. A number of tumor suppressor genes, including pl6INK4A, SOCS-1, APC, RASSF1A, GSTP1, and E-Cadherin, are silenced by DNA methylation in a large proportion of liver tumors, and this process often starts at preneoplastic (cirrhotic) stages [100, 101]. A higher rate of promoter methylation for specific genes, such as pl6INK4A and E-Cadherin, has been observed in HBV-related tumors compared to nonviral tumors [102–104].

Mechanisms of HBV Oncogenesis The exponential relation between HCC incidence and age suggests that, as in other human cancers, multiple steps are required, probably involving independent genetic lesions. The long latency period between HBV infection and HCC has been often used to support the concept of an indirect action of these viruses. It is generally considered that longterm liver damage due to the immune response against infected hepatocytes triggers chronic inflammation, oxidative DNA damage, continuous cell death, and consequent cell proliferation, and potentiates the action of exogenous carcinogenic factors such as aflatoxins and alcohol. Increasing experimental evidence suggests, however, that HBV contributes to HCC by directly modulating pathways that promote the malignant transformation of hepatocytes [87, 88, 105] (Fig. 31.6). HBV infection directly promotes carcinogenesis by at least four different mechanisms [105].

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M. Levrero and L. Belloni Genetic and epigenetic alterations

Clinical features

IGF1R AKT

Axin

HBV (High viremia)

AKT

p53, Axin

HBV (Low viremia)

Transcriptome Signaling Pathways

G1

G2

Mitosis and cell cycle

LOH 16q 4q 16p 4q 16p 13q 17p

G3

p53

4q 16p 17p 5q 21q 22q

G4

TCF-1

No LOH

ß-catenin

No LOH

Low stress immune response

G5 Wnt pathway

G6

Chromosomal stability

No LOH

ß-catenin E-cadherin

Fig. 31.5  Genome-wide studies and transcriptomic signatures in HCC. The combination of genome-wide assessment of genetic and epigenetic alterations, together with transcriptome and systematic pathway analyses, has confirmed the molecular diversity of HCCs and also allowed the molecular classification of HCCs.

Chromosomal instability

The genome-wide transcriptomic analysis performed by Zucman-Rossi and coworkers [99] identifies six homogeneous subgroups of HCCs (termed G1 to G6) associated with specific clinical and genetic characteristics. HBV-related HCCs belong to the G1 and G2 subgroups

HBV

(overt or occult)

Fig. 31.6  Mechanisms of HBV oncogenesis. HBV contributes to HCC by both directly modulating pathways that promote the malignant transformation of hepatocytes and indirectly by promoting long-term liver damage, chronic inflammation, cell death, regeneration, and oxidative DNA damage

Integration of HBV DNA Into host chromosomes : Insertional mutagenesis of cellular genes

Genetic instability

Prolonged expression of viral genes HBx, LHBs

Modifications of the epigenome

Host immune responses Inflammation Oxydative stress

Cell proliferation Apoptosis

31  HBV Signaling

Genomic Instability The majority of HCC cells display a high incidence of chromosome instability that is already evident in cirrhotic liver tissues, and has been found to increase during the hepatocarcinogenesis process [105]. Chromosome instability measured by the fractional allelic loss has been shown to be an independent prognostic marker of prognosis and recurrence in resected HCC [106]. HBV contributes to chromosome instability by both integration of the viral DNA into the host genome [107] and the ability of HBx to directly induce chromosomal instability by affecting the mitotic checkpoints [108]. HBV DNA integration in host chromosomes, although dispensable for viral replication, is detected in about 80% of HCCs. Although HCC tissues associated with HCV infection show significant losses of heterozygosity (LOH) [109] and HCV NS5a encoded proteins also affects mitotic checkpoints [110], the rate of LOH is much higher in HBV-associated tissues [109] and HBVrelated tumors generally harbor a higher rate of chromosomal abnormalities than tumors linked to other risk factors [98, 111].

Insertional Mutatagenesis Classic retrovirus-like insertional mutagenesis can occur with HBV integration at specific sites providing a growth advantage to a clonal cell population in which additional mutations accumulate. Evidence was first provided in two independent HCCs, with retinoic acid receptors (RARs) and cyclin A as target genes [112]. More recently, 15 new genes were found to be targeted by HBV integration in tumors, including recurrent HBV DNA integration into the hTERT gene encoding the catalytic subunit of telomerase, suggesting that viral integration in the vicinity of genes controlling cell proliferation, viability and differentiation is frequently involved in HBV hepatocarcinogenesis [113–116].

Senescence and Telomerase Reactivation Hepatocytes in cirrhotic livers display decreased proliferation rates with a dominant replicative senescence phenotype characterized by critically shortened

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telomeres and permanent cell cycle arrest [117]. Indeed, the length of telomeres progressively shortens from normal liver to chronic liver disease, and reaches the shortest levels in HCC [118, 119]. The low or absent telomerase activity in cirrhotic liver suggests that the repeated proliferation cycles of hepatocytes in the precirrhotic stages of liver disease leads to the progressive loss of telomere sequences and senescence arrest. The emergence of malignant hepatocytes in the context of senescent cirrhotic tissue requires that transformed cells bypass senescence and can survive despite critically shortened telomeres. Many studies have showed that 80–90% of HCCs display a high telomerase activity [120]. The integration of HBV DNA sequences into TERT gene provides evidence for a virus-induced deregulation of TERT expression [113, 116]. HBx and PreS2 proteins both upregulate TERT expression [121]. How TERT expression is reactivated in HCC cells is, however, only partially clarified. The TERT gene promoter displays binding sites for many transcription factors, including the estrogen receptor, Sp1, Myc, and ER81 acting positively, and vitamin D receptor, MZF-2, WT1, Mad, E2F1 and SMAD interacting protein-1 (SIP1, also called ZEB-2 or ZFHX1B) acting negatively [122]. Despite TERT activation, telomers remain very short in HCC cells and this may predispose to occasional telomere instability leading to the increased rate of chromosomal instability and polyploidy that is quite frequent in these tumors [121].

Long-term Expression of Viral Proteins The predominant mechanism of HBV carcinogenesis linked to HBV infection is based on the ability of viral protein, in particular HBx, to modulate cell proliferation and cell viability, and to sensitize liver cells to mutagens. In transgenic mouse models, unregulated expression of the HBV X and S proteins are associated with hepatocarcinogenesis [57, 123]. The inappropriate expression of the large envelope protein has the potential to be directly cytotoxic to the hepatocyte and initiate a cascade of events that ultimately progress to malignant transformation [57]. In human patients, ground-glass hepatocytes are detected in the late, nonreplicative stages of HBV infection. These histological abnormalities reflect accumulation of envelope proteins in ER membranes, and it has been shown recently that some Pre-S mutants might be

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associated with ER stress and participate in hepatocarcinogenesis [124]. Rearrangement of integrated HBV sequences in HCC may also lead to abnormal expression of the S gene proteins. Specific activation of c-Raf-1/Erk2 signaling by the truncated MHBs proteins results in an increased proliferation rate of hepatocytes [125]. The HBx protein (Fig. 31.2) behaves as a promiscuous transactivator of cellular genes such as oncogenes, growth factors and cytokines; it binds and inactivates p53, and interacts with the DNA repair protein DDB1, which may affect repair functions and allow the accumulation of genetic changes (reviewed in [107]). The ability of HBx to activate calciumdependent signaling events accounts for both the induction of apoptosis and increased cell proliferation (reviewed in [107]). HBx also interacts with the peptidyl-prolyl cis/trans isomerase Pin1 and this interaction leads to HBx stabilization, enhanced HBx-mediated transactivation of target genes and increased cellular proliferation [126]. HBx has been shown to modulate chromatin dynamics in HCC cells and tissues. As mentioned above, HBx binds several nuclear proteins involved in the regulation of transcription including component of the basal transcriptional machinery (RPB5, TFIIB, TBP, TFIIH), coactivators (CBP, p300 and PCAF) and transcription factors (ATF/CREB, ATF3, c/EBP, NF-IL-6, Ets, Egr, SMAD4, Oct1, RXR receptor, p53) (reviewed in [3, 36]). HBx is bound in vivo to the promoters of a number of CREB-regulated genes and favors transcription by increasing the amount of CBP and p300 recruited on the same promoters [45].  In addition to stimulate transcription, HBx can also repress gene expression HBx by upregulating DNMT1, DNMT3A1 and DNMT3A2 levels, increasing total DNA methyltransferase (DNMT) activity and selectively facilitating the regional hypermethylation of the promoters of certain tumor suppressor genes [127, 128]. More recently, it has been shown, by ChIP and sequential-ChIP experiments, that HBx recruits DNMT3A on promoters of the repressed genes, MT1F and IL4R, leading to promoter hypermethylation and suppression of gene expression [129]. Interestingly, in the presence of HBx DNMT3A dissociates from the promoter region of the upregulated genes IGFBP3 and CDH6, thus resulting in hypomethylation and transcriptional activation [129].

M. Levrero and L. Belloni

Summary

›› Infection with HBV continues to be a major

››

››

››

››

››

››

health problem with about 400 million people chronically infected worldwide who are at high risk of developing liver cirrhosis and HCC. The epidemiological evidence linking HBV infection to HCC is very strong and, despite the mechanisms underlying HBV-associated carcinogenesis remain to be fully defined, a growing number of studies support a direct role of HBV in the process through the integration of the viral DNA into the host genome, the induction of genomic instability and the expression of viral proteins that affect cell homeostasis. Many cellular proteins play important roles in crucial steps of HBV life cycle including the initiation of HBV reverse transcription, nucleocapsid maturation, formation of the HBV episomal DNA and its epigenetic regulation. The cellular Hsp90 complex is required for complex formation between the RT and the e region of the pgRNA during viral assembly and reverse transcription. HBc phosphorylation affects HBV pgRNA packaging and DNA synthesis. The cellular kinase that is predicted to be packaged into the viral nucleocapsids during the assembly process remains to be identified. The HBV replicative intermediate cccDNA (covalently closed circular HBV DNA), which serves as a template for the transcription of all viral transcripts including the pgRNA, is organized into a minichromosome in the nuclei of infected hepatocytes by histones and non histone proteins of viral and cellular origin. HBV transcription and replication are modulated by epigenetic changes of the cccDNA. Host transcription factors regulate cccDNA transcription by modifying cccDNA-bound histones via the recruitment of acetyltransferases and deacetylases. The regulatory protein HBx is essential for viral infection in vivo. It potentiates viral replication in cell culture and is thought to contribute to HBV oncogenicity.

31  HBV Signaling

›› Studies

››

››

in transfected cells have shown that HBx expression affects several cellular functions such as cytoplasmic calcium regulation, cell signaling, transcription, cell proliferation, DNA repair and apoptosis. To perform its multiple functions, HBx interacts with many cellular partners that include the tumor suppressor p53, the UV-stimulated E3-ubiquitin ligase DDB1, the nuclear export protein CRM1, the peptidyl-prolyl cis/trans isomerase Pin1, nuclear proteins involved in the regulation of transcription (i.e., the RPB5 subunit of Pol II, TFIIB, TFIIH, and TBP), several members of the basic domain-leucine zipper (bZIP) family of transcription factors (i.e., ATF2, CHOP, and CREB); chromatin modifying enzymes (i.e., CBP, p300, and PCAF). HBx activate transcription both indirectly by modulating cytoplasmic signaling pathways which in turn control the activity of nuclear transcription factors (i.e., MAPKs to activate AP1, b-catenin; PI3K/Akt to activate NFAT, NFkB), or directly by interacting with nuclear transcription factors and chromatin modifying (i.e., CBP/p300) enzymes and favoring their recruitment onto the promoter regions of target genes, and by binding and relocating DNA and histones methylating enzymes (i.e., DNMT3a).

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(a) True (b) False 3. Which of the following HBx properties are true (indicate one or more) (a) Blocking apoptosis by decreasing cytochrome c release from mitocohondria (b) Binding and activating the EGFR (c) Interacting with the bZip nuclear transcription factor ATF2 (d) Potentiating the recruitment of the CBP acetyltransferase onto the promoter of CREB target genes (e) Potentiate HBV replication 4. The activation of cellular genes transcription is mediated by nuclear HBV, whereas cytoplasmic HBx is involved in hepatocytes transformation (a) True (b) False 5. The modulation of steatogenic pathways by HBV are mediated by: (indicate one) (a) HBc-mediated activation of the Fatty Acid Synthase gene (b) The retention of LHBs proteins in the endoplasmic reticulum (c) The accumulation of excess immature core particles in the cytoplasm (d) HBx-mediated activation of the nuclear receptors LXRa and PPARg

Multiple Choice Questions

References

1. Which of these proteins and elements are not part of circulating HBV virions (indicate one or more) (a) HBeAg (b) RT-Pol (c) HBcAg (d) HBxAg (e) LHBs (f) SHBs (g) HBV cccDNA

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2. Multiple copies of the viral cccDNA are integrated into the host chromatin and their transcription is modulated by acetylation and deacetylation of the cccDNA-bound histones

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Hepatitis C Virus and Insulin Signaling

32

Francesco Negro and Sophie Clément

Introduction Hepatitis C virus (HCV) infection is a major cause of liver disease in humans. It is estimated that about 150– 200 million people have been in contact with HCV worldwide. HCV is a positive-strand RNA virus classified in the Hepacivirus genus within the Flaviviridae family. HCV RNA genome comprises a 5′-noncoding region, an open reading frame encoding for both structural and nonstructural proteins, and a 3′-noncoding region. The structural proteins, which are assembled into the mature virion, include the core protein and the envelope glycoproteins E1 and E2. The nonstructural (NS) proteins include the p7 (functioning as an ion channel), the NS2-3 protease, the NS3 serine protease (which possesses also an RNA helicase activity), the NS4A, NS4B and NS5A proteins, and the NS5B RNAdependent RNA polymerase [1]. The lack of proofreading activity of the latter accounts for the elevated sequence heterogeneity of HCV sequences, which have been classified into genotypes and subtypes [1]. Thus, there are six genotypes, diverging from one another in their sequence by about 30–35%, and ­several subtypes. Furthermore, in any given individual there is an array of variants in equilibrium among themselves that are collectively referred to as quasispecies [1]. Approximately, 85% of persons who have been in contact with HCV have persistent infection, mostly associated with a relentlessly progressing chronic hepatitis,

F. Negro () Departments of Internal Medicine and Pathology and Immunology, University of Geneva Medical Center, 1 Rue Michel-Servet, 1205 Geneva, Switzerland e-mail: [email protected]

ultimately leading to the development of cirrhosis and hepatocellular carcinoma. The rate of progression toward the cirrhotic stage varies widely, depending on several host-related cofactors, such as age, gender, level of alcohol consumption, overweight, immune status, and coinfections [2]. One of these cofactors is type 2 diabetes (T2D), which has been recognized to modify the course of hepatitis C even at the early stage of insulin resistance (IR) [3, 4]. Moreover, T2D and IR decrease the rate of response to antiviral therapy [5]. Although T2D is a common complication of all liver diseases, especially at the advanced stage and independently of the etiology, a wealth of clinical data support a direct role of HCV in the derangement of glucose metabolism. The scope of this article is to discuss the evidence suggesting a causal relationship between HCV and T2D/IR and review the experimental data that may explain the mechanisms accounting for such an association. Finally, we will briefly discuss its clinical impact and some directions for management.

Clinical Evidence Supporting an Association Between HCV and T2D Several cross-sectional studies [6–13] have reported that patients infected with HCV may present with T2D more often than patients with liver disease of other etiology. Further evidence has come from case-control studies, where cases were either HCV-infected persons [14–19] or diabetic patients [8, 20–22]. It has been suggested that patients with T2D may be at risk of blood borne infections via repeated use of finger stick devices. A single study from France, evaluating the prevalence of  HCV antibodies in 259 patients with T2D, seen ­during 1998 at a diabetic unit, failed to confirm this

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_32, © Springer-Verlag Berlin Heidelberg 2010

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hypothesis [23]. One cannot exclude that iatrogenic transmission of HCV among diabetic patients may however have been significant in the previous decades. The potential ascertainment bias that may occur in clinic-based studies targeting a specific disease group was overcome in a vast study conducted in the general population, the Third National Health and Nutrition Examination Survey [24]. This study, which included 9,841 subjects aged 20 years and more, showed that persons who were anti-HCV-positive and 40 years or older had an odds ratio of 3.77 (95% CI, 1.80–7.87), after adjusting for sex, body mass index (BMI) and ­ethnicity, of having T2D compared to anti-HCV-negative individuals. Thus, both clinic-based studies and the general ­population-based NANHES-III study came to similar conclusions, reinforcing the hypothesis of a causal association between HCV infection and T2D. Because of the cross-sectional nature of all these surveys, however, a temporal relationship between HCV infection and T2D could not be established. This issue, that is, did the HCV infection came before the occurrence of T2D or vice versa, has been addressed at by longitudinal studies. A prospective, case-cohort study analyzed whether persons who developed T2D were more likely to have had precedent HCV infection when they were enrolled in a community-based cohort of 1,084 persons aged between 44 and 65 [25]. The prevalence of HCV in this population was 0.8%. A total of 548 subjects developed de novo T2D after 9 years of followup. Prior to entry, subjects had been categorized as low-risk or high-risk for T2D, based on age and BMI. Among those at high risk for T2D, persons with HCV infection were more than 11 times as likely as those without HCV infection to develop T2D (relative hazard, 11.58; 95% CI, 1.39–96.6). Among those at low risk, the incidence of T2D was not increased among HCV-infected subjects. The conclusion of this survey was that preexisting HCV infection may increase the incidence of T2D in persons with known risk factors. A second study from Taiwan [26] confirmed the above observations. Thus, there exists an excess T2D risk in HCV-infected persons compared to HCV-negative controls, suggesting a direct role of HCV in inducing the derangement of glucose metabolism. A recent, large meta-analysis, the first of this kind, strongly supports this conclusion [27]. Additional evidence in favor of an association between HCV and T2D comes from longitudinal studies performed in patients having

F. Negro and S. Clément

received a liver or kidney transplantation. T2D is a common complication of liver transplantation, and there is accumulating evidence that HCV is a strong predictor of newly-onset T2D in this setting [28–34]. A similarly increased risk of T2D has been reported after kidney transplantation, and a recent meta-analysis on 10 studies evaluated the pooled relative risk for T2D after kidney transplantation at 2.73 (95% CI 1.94–3.83) [35]. Thus, HCV and T2D are associated more than just by chance, suggesting that HCV may alter glucose homeostasis by its direct action, or via indirect mechanisms such as through cytokine stimulation. The associa­ tion between HCV infection and glucose abnor­malities holds true also if one considers prediabetes conditions, like impaired glucose tolerance (IGT) or IR. The latter is defined as a condition in which higher than normal insulin concentrations are needed to achieve normal metabolic responses or, alternatively, normal insulin concentrations are unable to achieve normal metabolic responses [36]. It has to be said, however, that it is unclear whether HCV-associated IR may invariably evolve towards T2D, especially in patients without other risk factors of T2D. Longitudinal studies are needed to clarify this issue. Hui et al. [4] have compared fasting levels of serum insulin, C-peptide and IR (measured as homeostasis assessment [HOMA] score) in 121 HCV patients with stage 0 or 1 liver fibrosis and 137 healthy volunteers matched for sex, BMI, and waist-to-hip ratio. Results showed that such HCV-infected patients, even at an early stage of liver disease, had higher levels of insulin, C-peptide, and HOMA scores compared with controls. This study suggested also that genotype 3 may have significantly lower HOMA scores than other genotype. Thus, HCV may induce IR irrespective of the stage of the underlying liver disease, an effect that seems genotype specific. In a more recent paper, Moucari et al. [37] analyzed 600 consecutive patients (500 with chronic hepatitis C and 100 controls with chronic hepatitis B). IR was less frequent in chronic hepatitis B than in matched chronic hepatitis C patients (5 vs. 35%, respectively, P < 0.001), irrespective of the presence or absence of cirrhosis. Furthermore, IR was associated with genotypes 1 and 4 and elevated serum HCV RNA, suggesting a trend, even among patients without the typical features of the metabolic syndrome, between HCV replication level and HOMA score. These data further strengthened the hypothesis that

32  Hepatitis C Virus and Insulin Signaling

HCV may be directly implicated in the impairment of insulin signaling. A correlation between HCV replication levels and HOMA score has also been reported by others [38–40]. However, further work is warranted. In particular, there is the need of a careful functional analysis of HCV sequences potentially responsible for the IR. It is impossible, at present, to establish whether HCV replication is responsible for an increased IR or whether HCV replication is favored by hyperinsulinemia, as suggested by some in vitro data [41], and/or by the increased serum levels of free fatty acids [42] typically observed in IR and T2D, two features of the metabolic syndrome that may merely create a favorable environment for HCV replication. Finally, evidence that HCV may interfere with glucose metabolism in high-risk individuals comes from clinical trials of treatment of chronic hepatitis C. These observations are based on the premises that curing HCV should result in an amelioration of the HOMA score and in a decreased incidence of T2D after the end of therapy. Kawaguchi et al. [43] showed that eradication of HCV improved the HOMA score and the intrahepatic expression level of the insulin receptor substrates (IRS) 1 and 2, two cellular transducers of the insulin signal. Romero-Gómez et al. [44] showed that a sustained virological response (SVR) reduces by half the incidence of T2D and/or impaired fasting glucose during a post-therapy follow-up of 27 ± 17 months, and similar data have been reported by a group in Barcelona [45]. However, in an Italian cohort of 202 patients [46] the benefit of SVR was lost after a longer follow-up (8.0 years, range 5–16), even after adjustment for several baseline risk factors of T2D. In conclusion, HCV seems to increase the risk of incident T2D in predisposed individuals. As a result, the association between HCV and T2D is more evident among patients who are older and have higher BMI. When measuring IR before T2D has occurred, some HCV-infected patients are clearly less insulin sensitive than controls, matched for risk factors of T2D and the stage of liver disease. This effect is probably associated with specific HCV sequences, and shows some dose-dependence, that is, may be correlated with HCV replication level. Curing HCV seems to have beneficial effects on the level of insulin sensitivity, although this may not be the rule. We will now analyze the potential mechanisms of interference with the insulin signaling brought about by HCV.

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HCV Interference with the Insulin Signaling: Potential Mechanisms A potential, direct interference of HCV with the insulin signaling cascade was suggested by a study where liver specimens obtained from 42 nonobese, nondiabetic HCV-infected individuals and 10 non-HCV-infected subjects matched for age and BMI were exposed ex vivo to insulin and examined for the contents and phosphorylation/activation status of some insulin signaling molecules [47]. Insulin-stimulated IRS-1 tyrosine phosphorylation was decreased by twofold in HCVinfected patients compared to non-HCV-infected ones, and this was accompanied by significant reductions in IRS-1/p85 phosphatidylinositol 3 (PI3)-kinase association, IRS-1-associated PI3-kinase ­enzymatic activity and insulin-stimulated Akt phosphorylation [47]. The  authors concluded that, in patients with chronic hepatitis C, direct interactions between HCV and insulin signaling components occur that may result into IR,  which in turn may progress to T2D in at risk individuals. In the transgenic mouse [48], however, the IR associated with the expression of the core-encoding region of HCV could be reversed by treatment with antitumor necrosis factor (TNF)-a antibodies, suggesting an increased level of serine phosphorylation of IRS-1 as induced by TNF-a (Fig.  32.1). Thus, in this animal model, the core protein may induce IR indirectly via an increased secretion of TNF-a. It has to be said that in vitro models have largely hinted at a direct – rather than indirect – interaction of the core protein with the insulin signaling pathway. An increased proteasomal degradation of the IRS-1 and -2 via the activation of the suppressor of cytokine signaling (SOCS)-3 has been reported after transient expression of the core protein in hepatoma cells [49]. Direct but genotype-specific mechanisms have been advocated in another in vitro study [50], where a downregulation of peroxisome proliferator-activated receptor (PPAR)-g and an upregulation of SOCS-7 was observed in Huh-7 hepatoma cells transfected with the core protein of genotype 3, whereas the core protein of genotype 1b activated the mammalian target of rapamycin, findings that were confirmed by using agonists for PPAR-g (rosiglitazone) or short interfering RNAs for SOCS-7 [50]. Among the indirect mechanisms, an increased endo­ plasmic reticulum stress has also been reported [51]

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TNF-

Insulin

TNFR1

pi (3,4) P2

pi (3,4,5) P3

(47)

IRS1 P

P

?

p85

p110

(48,53)

IKK

Jnk (53)

(54)

mTOR (50)

Core

ROS

SOCS (49,50)

PKD1/2

AMPK

ER stress (51)

Akt PP2A (52)

Anti-apoptosis

Fatty acid synthesis

Glycogen synthesis Protein synthesis

Glucose uptake

Fig. 32.1  Schematic representation of some of the effects brought about by HCV on insulin signaling in hepatocytes. Numbers refer to the bibliographic references. For the abbreviations, please refer to the text

that may lead to activation of the protein phosphatase 2A (PP2A), an inhibitor of Akt (Fig. 32.1). Activation of PP2A may also dephosphorylate the AMP-activated kinase (AMPK), a key regulator of gluconeogenesis [52]. Thus, PP2A may lead to IR via a dual mechanism, that is, inactivation of the two pivotal kinases Akt and AMPK. A potential role of a stress kinase, the c-Jun N-terminal kinase (JNK) has been emphasized in a recent work [53]. The HCV core protein-mediated Ser (312) phosphorylation of IRS-1 was inhibited by a JNK inhibitor in an in vitro infection assay using cellculture grown HCV [53]. The activation of JNK by the HCV core may be direct, but indirect, proinflammatory cytokine-mediated mechanisms (via an autocrine loop) have not been entirely ruled out.

Studies on chronically infected patients suggest that increased oxidative stress and intrahepatic inflammation may play an important role in inducing an IR state. Mitsuyoshi et al. [54] evaluated 203 chronic hepatitis C patients with HCV genotypes 1 or 2 infection. HOMA and serum levels of thioredoxin, a marker of oxidative stress, were significantly correlated with each other, even after adjustment for BMI. Besides the direct role of HCV proteins in inducing oxidative stress, the indirect role of inflammatory mediators, like TNF-a, seems very likely to occur in chronic hepatitis C patients, in whom an increased intrahepatic TNF-a production has been described [55, 56]. Further work is necessary in this field, and the availability of genotype-specific replicon assays may pave the way to more in-depth mechanistic analyses.

32  Hepatitis C Virus and Insulin Signaling

Clinical Consequences of IR/T2D in Chronic Hepatitis C The clinical consequences of IR and T2D on chronic hepatitis C are dual: accelerated fibrogenesis and reduced response to IFN-a-based therapy. Since one of the most frequent consequences of IR/T2D on the liver is steatosis, many data can be inferred looking at studies where the impact of nonvirally- and nonalcohol-induced steatosis on fibrosis was assessed [2]. In these patients, the most likely cause of fatty liver is IR, and this, rather than steatosis, seems to predict severity and progression of fibrosis [4]. In general, accelerated liver fibrogenesis should be considered in the setting of the consequences of the metabolic syndrome on the liver. Thus, several mechanisms other than IR, such as oxidative stress, increased secretion of proinflammatory adipokines and cytokines, and the peculiar susceptibility to apoptosis that has been associated with steatosis should be taken into account. The relative contribution of adipokines on liver fibrosis in chronic hepatitis C is starting to be unraveled, but it is far from being fully appreciated. In nonalcoholic steatohepatitis, hyperglycemia/hyperinsulinemia may be directly stimulating hepatic stellate cells to produce connective tissue growth factor (CTGF), leading in turn to increased collagen fiber deposition [57]. Increased intrahepatic levels of CTGF have been reported to occur in chronic hepatitis C [58]. The reduction of IR consequent to body weight reduction and increased physical activity may lead to reduced fibrosis score over time and decreased activation of stellate cells [58]. Several proinflammatory cytokines and adipokines may be involved in the pathogenesis of liver injury in chronic hepatitis C, but their relative contribution, if any, is unclear. A study has evaluated the role of TNF-a, interleukin 6, leptin, and adiponectin in the pathogenesis of HCV-associated liver injury [59]. Only TNF-a levels seemed to correlate with severity of portal and periportal inflammation, but none of these cytokines correlated with fibrosis. The role of leptin [60, 61] and adiponectin is controversial [59], as well as that of resistin, where the only positive report awaits independent confirmation [62]. Finally, an increased liver cell apoptosis has been reported to correlate with steatosis [63]. In the presence of steatosis (whether IR-induced or not), apoptosis correlated with activation of stellate cells and fibrosis, consistent with the hypothesis that a steatotic liver is more vulnerable to liver injury.

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Increasing levels of IR are associated with reduced rates of early [64–67] and sustained response in chronic hepatitis C patients treated with pegylated IFN-a and ribavirin [68–73]. The molecular link between IR and poor response to IFN-a seems to be the increased levels of SOCS-3 in liver, in agreement with in vitro data [71, 74]. Interestingly, SOCS-3, as said above, is not only promoting the proteasomal degradation of IRS-1, leading to impaired insulin signaling and IR [49], but, together with other members of the SOCS family, is also a negative regulator in the transduction of the IFN-a signaling [75]. Thus, it is not too unlikely that HCV may have developed, at the evolutionary level, the ability to activate SOCS-3 or other members [50] of the SOCS family as a mechanism to inhibit the IFN-a signaling, one of the main arms of the host’s innate immune response, simultaneously impairing the insulin signaling. This view seems corroborated by the finding that HCV activates PP2A, with the dual effect, again, of interfering with the insulin [52] and the IFN-a [51] signaling pathways. Whether these mechanisms may be exploited pharmacologically, that is, with drugs aimed at reducing the IR while improving the responsiveness to IFN-a, remains to be fully explored.

Perspectives for Clinical Management The treatment of IR and T2D in chronic hepatitis C patients has two goals, as far as the underlying liver disease is concerned: to reduce the fibrogenesis (hence the liver disease progression) and to increase the response to IFN-a-based therapy. The therapy should be aimed at correcting IR, based on the underlying molecular mechanisms, which may differ according to the viral genotype and on the presence or not of a metabolic syndrome. At present, however, the approach that is being followed is rather empirical. Insulin sensitizing agents have been tested with the specific aim of improving the rate of response to IFNa-based therapy. It was suggested that IR should be corrected in patients with chronic hepatitis C not responding to IFN-a-based treatment, in order to improve response upon retreatment. The modalities of this intervention, however, have not been established, nor has the optimal HOMA score been identified to be reached prior to antiviral therapy. Preliminary data from four independent studies [76–79] are not encouraging. Further data are warranted before insulin

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sensitizers can be added to the panoply of drugs to treat hepatitis C. Furthermore, the effects of PPAR agonists on serum lipids and their potential consequences on the HCV life cycle should be investigated in more detail. It is also unclear whether the treatment with the insulin sensitizer should be started at the same time as the antiviral retreatment or precede it, in order to start the pegylated IFN-a/ribavirin combination only when the HOMA score has decreased to a level predictive of an increased SVR. It is not clear whether the best approach would be using a PPAR agonist (and at what dose) or an AMPK activator such as metformin. The final results of the TRIC-1 study [79] show that adding metformin to the pegylated IFN-a/ribavirin combination afforded a marginal, insignificant gain as to the SVR rate, despite an increased rapid virological response after 4 weeks of triple therapy. Thus, further clinical trials aiming at reducing the IR in chronic hepatitis C via different pharmacological interventions are warranted.

Conclusions Thus, as far as the relationship between HCV and insulin signaling is concerned, current clinical and experimental evidence supports the following conclusions: 1. HCV and IR/T2D are associated to an extent that cannot be merely explained by chance; 2. This suggests that HCV interferes directly (through one or more of its proteins) and/or indirectly (by modulating the production of specific cytokines, like TNF-a) with glucose metabolism; 3. Independently of the mechanism, IR and T2D have important effects on the hepatitis C progression, since chronic hepatitis C patients with IR/T2D are at increased risk of developing cirrhosis and hepatocellular carcinoma; 4. IR and T2D reduce the response to IFN-a-based therapy in chronic hepatitis C patients, possibly due to the virally-induced disregulation of intracellular factors that operate promiscuously along insulin and IFN-a signaling pathways; 5. Current clinical management of HCV-infected persons warrants identification and correction of derange­ ments of glucose metabolism; 6. Although lifestyle interventions (reduction of body weight and increased physical activity) are certainly indicated in patients with chronic hepatitis C and the

F. Negro and S. Clément

metabolic syndrome, in order to reduce the cardiovascular morbidity and mortality, it remains to be fully explored whether these measures will impact also on the underlying liver disease; 7. Insulin sensitizers are currently being evaluated in clinical trials, but available data do not warrant their use in all chronic hepatitis C with IR with the specific aim of increasing response to antivirals, at least outside the clinical trials.

Summary

›› This review article focuses on how the hepati-

tis C virus (HCV) infection interferes with insulin signaling. Both cross-sectional and longitudinal studies have shown that chronic HCV infection is associated with an increased risk of developing insulin resistance (IR) and type 2 diabetes (T2D). The direct effect of HCV on the insulin signaling has been analyzed in experimental models. HCV seems to affect glucose metabolism via direct and indirect (i.e. cytokine-mediated) mechanisms. Interestingly, many of these mechanisms appear to be promiscuous with cellular pathways implicated in the host innate immune response. The knowledge of the molecular mechanisms underlying this interaction is paramount to proper clinical management: in fact, IR and T2D not only accelerate the clinical progression of chronic hepatitis C, but also reduce the virological response to interferon-alpha-based treatment.

Multiple Choice Questions 1. HCV is associated with type 2 diabetes: (a) Independently of the patient’s age (b) Especially among individuals who are less than 40 years old (c) Especially among individuals who are more than 40 years old (d) HCV is not associated with type 2 diabetes (e) Type 2 diabetes occurs in HCV-infected individuals only after successful eradication of HCV with antivirals

32  Hepatitis C Virus and Insulin Signaling

2. HCV induces insulin resistance: (a) Independently of the viral genotype (b) Only in obese patients (c) Only in patients with genotype 3 and severe steatosis (d) Only in patients without liver fibrosis (e) In none of the above 3. Insulin resistance reduces the rate of response to antivirals in chronic hepatitis C: (a) Independently of the viral genotype (b) Only in patients with genotype 1 (c) Only in patients with genotypes 2 and 3 (d) Never, insulin resistance does not affect the response to antivirals in hepatitis C (e) Only in patients with cirrhosis 4. Experimental data support an interference with insulin signaling brought about HCV proteins. Which of the following is true?: (a) The mechanism is exclusively direct, that is, interaction between viral proteins and insulin signaling factors (b) The mechanism is exclusively indirect, that is, mediated by proinflammatory cytokines (c) Data are compatible with a dual mechanism, that is, direct and indirect (d) HCV has never been shown to interfere with insulin signaling (e) HCV interferes with insulin signaling by stimulating the secretion of IFN-a 5. The following interventions may reduce the level of insulin resistance in patients with hepatitis C: (a) Moderate alcohol drinking (b) Complete rest (c) Diet poor in sodium chloride (d) Increased physical activity (e) None of the above Acknowledgments  The authors’ experimental work cited in the present manuscript is supported by the Swiss National Science Foundation grant number 320000-116544.

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32  Hepatitis C Virus and Insulin Signaling 53. Banerjee S, Saito K, Ait-Goughoulte M et al (2008) Hepatitis C virus core protein upregulates serine phosphorylation substrate-1 and impairs the downstream Akt/protein kinase B signaling pathway for insulin resistance. J Virol 82:2606–2612 54. Mitsuyoshi H, Itoh Y, Sumida Y et al (2008) Evidence of oxidative stress as a factor in the development of insulin resistance in patients with chronic hepatitis C. Hepatol Res 38:348–353 55. Knobler H, Zhornicky T, Sandler A et al (2003) Tumor necrosis factor-alpha-induced insulin resistance may mediate the hepatitis C virus-diabetes association. Am J Gastroenterol 98:2751–2756 56. Knobler H, Schattner A (2005) TNF-a, chronic hepatitis C and diabetes: a novel triad. QJM 98:1–6 57. Paradis V, Perlemuter G, Bonvoust F et al (2001) High glucose and hyperinsulinemia stimulate connective tissue growth factor expression: a potential mechanism involved in progression to fibrosis in nonalcoholic steatohepatitis. Hepatology 34:738–744 58. Hora C, Negro F, Leandro G et al (2008) Connective tissue growth factor, steatosis and fibrosis in patients with chronic hepatitis C. Liver Int 28:370–376 59. Cua IH, Hui JM, Bandara P et al (2007) Insulin resistance and liver injury in hepatitis C is not associated with virusspecific changes in adipocytokines. Hepatology 46:66–73 60. Muzzi A, Rubbia-Brandt L, Leandro G et al (2005) Insulin resistance is associated with liver fibrosis in non-diabetic chronic hepatitis C patients. J Hepatol 42:39–44 61. Lo Iacono O, Venezia G, Petta S et al (2007) The impact of insulin resistance, serum adipocytokines and visceral obesity on steatosis and fibrosis in patients with chronic hepatitis C. Aliment Pharmacol Ther 25:1181–1191 62. Tsochatzis E, Papatheodoridis GV, Hadziyannis E et al (2008) Serum adipokine levels in chronic liver diseases: association of resistin levels with fibrosis severity. Scand J Gastroenterol 43:1128–1136 63. Walsh MJ, Vanags DM, Clouston AD et al (2004) Steatosis and liver cell apoptosis in chronic hepatitis C: a mechanism for increased liver injury. Hepatology 39:1230–1238 64. Huang JF, Yu ML, Dai CY et al (2007) Pretreatment insulin sensitivity contributes to the treatment response to peginterferon plus ribavirin combination therapy for patients with chronic hepatitis C. Hepatology 46(suppl 1):349A 65. Bortoletto G, Realdon S, Dal Pero F et al (2007) Insulin resistance (IR) defined by the homeostasis model of assessment insulin resistance (HOMA-IR) index has a direct effect on early viral kinetics during pegylated-interferon therapy for chronic hepatitis C. Hepatology 46(Suppl 1):361A 66. Bongiovanni M, Ranieri R, Casana M et al (2008) Insulin resistance affects early virologic response in HIV-infected subjects treated for hepatitis C infection. J Acquir Immune Defic Syndr 47:258–259

491 67. Nasta P, Gatti F, Puoti M et al (2008) Insulin resistance impairs rapid virologic response in HIV/hepatitis C virus coinfected patients on peginterferon-alfa-2a. AIDS 22:857–861 68. Romero-Gómez M, Del Mar Viloria M, Andrade RJ et al (2005) Insulin resistance impairs sustained response rate to peginterferon plus ribavirin in chronic hepatitis C patients. Gastroenterology 128:636–641 69. D’Souza R, Sabin CA, Foster GR (2005) Insulin resistance plays a significant role in liver fibrosis in chronic hepatitis C and in the response to antiviral therapy. Am J Gastroenterol 100:1509–1515 70. Conjeevaram HS, Kleiner DE, Everhart JE et al (2007) Race, insulin resistance and hepatic steatosis in chronic hepatitis C. Hepatology 45:80–87 71. Persico M, Capasso M, Persico E et al (2007) Suppressor of cytokine signaling 3 (SOCS3) expression and hepatitis C virus-related chronic hepatitis: insulin resistance and response to antiviral therapy. Hepatology 46:1009–1015 72. Poustchi H, Negro F, Hui J et al (2008) Insulin resistance and response to therapy in patients infected with chronic hepatitis C virus genotypes 2 and 3. J Hepatol 48:28–34 73. Chu CJ, Lee SD, Hung TH et al (2009) Insulin resistance is a major determinant of sustained virological response in genotype 1 chronic hepatitis C patients receiving peginterferon alpha-2b plus ribavirin. Aliment Pharmacol Ther 29(1):46–54 74. Walsh MJ, Jonsson JR, Richardson MM et al (2006) Nonresponse to antiviral therapy is associated with obesity and increased hepatic expression of suppressor of cytokine signalling 3 (SOCS-3) in patients with chronic hepatitis C, viral genotype 1. Gut 55:529–535 75. Gadina M, Hilton D, Johnston JA et al (2001) Signaling by type I and II cytokine receptors: ten years after. Curr Opin Immunol 13:363–373 76. Overbeck K, Genné D, Golay A, Negro F (2008) Pioglitazone in chronic hepatitis C not responding to pegylated interferonalpha and ribavirin. J Hepatol 49:295–298 77. Elgouhari HM, Cesario KB, Lopez R, Zein NN (2008) Pioglitazone improves early virologic kinetic response to PEG IFN/RBV combination therapy in hepatitis C genotype 1 naïve patients. Hepatology 48(Suppl):383A 78. Conjeevaram H, Burant CF, McKenna B et al (2008) A randomized, double-blind, placebo-controlled study of PPARgamma agonist pioglitazone given in combination with peginterferon and ribavirin in patients with genotype-1 chronic hepatitis C. Hepatology 48(Suppl):384A 79. Romero-Gomez M, Diago M, Andrade RJ et al (2008) Metformin with peginterferon alfa-2a and ribavirin in the treatment of naïve genotype 1 chronic hepatitis C patients with insulin resistance (TRIC-1): final results of a randomized and double-blinded trial. Hepatology 48(Suppl):95A 80. Hickman IJ, Clouston AD, Macdonald GA et al (2002) Effect of weight reduction on liver histology and biochemistry in patients with chronic hepatitis C. Gut 51:89–94

33

MicroRNAs Onpan Cheung and Arun J. Sanyal

History of MicroRNA

Types of miRNA and Its Biogenesis

MicroRNAs (miRNA) are small, naturally occurring single-stranded RNA of about 21–23 nucleotide in length. They are generated from endogenous transcripts that are encoded in the genomes of humans, animals, viruses, and plants. The first short noncoding miRNA, lin-4 that regulates gene expression in nematode C. elegans was identified by Victor Ambros et al. in 1993 [1]. The miRNA world did not take off until the discovery of let-7, a second miRNA discovered by Ruvkun and Horvitz in 2000 [2], and the rise in interest in another class of short RNA, silencing RNA (siRNA) [3, 4]. The highly conserved nature of let-7 also attracted a great deal of attention to miRNA research. Since its discovery, more miRNAs in various organisms, from protozoans to humans have been identified. Currently, a total of 873 miRNAs have been reported in human (miRBase 11.0, April 2008), and many of them are encoded in polycistronic transcripts. The expression of miRNA, in general, is both organ-specific and dependent on the stage of development [5, 6]. miRNAs have diverse functions including regulation of important cellular processes e.g., cancer, cell metabolism, immune function, cell proliferation, apoptosis, tissue development, and differentiation [7–11].

As shown in Fig.  33.1, miRNAs in animals are first transcribed as long primary transcripts (pri-miRNAs) by RNA polymerase II enzyme [12]. They are then cropped into hairpin-shaped pre-miRNA by the nuclear RNase III enzyme Drosha, that forms a microprocessor complex with double-stranded RNA-binding protein DGCR8/Pasha to form the approximately 70 base premiRNA processing intermediates, which are exported out of the nucleus by exportin-5 in the presence of GTP as a cofactor, and are subsequently cleaved by the cytoplasmic RNase III Dicer into ~22-nucleotides miRNA duplexes [13–16]. Next, miRNA duplexes are unwound. The miRNA strand that has its 5¢ terminus at this end is the future mature miRNA, which is incorporated into a ribonucleoprotein complex, miRNP that is similar to the argonaute containing RNA-induced silencing complex (RISC) [17]. In RISC, it provides the specific determinants that direct as yet an unidentified protein nuclease to cleave mRNAs complementary to the miRNA, in cases where there is a near-perfect complement between the miRNA and the mRNA [17–19]. The RISC–miRNA combination can also mediate downregulation of the target mRNA activity by translation inhibition, in cases where miRNA only partially complements its corresponding mRNA [1, 2]. The distribution of miRNA genes within genomes is not random. The majority of known mammalian miRNA genes are within introns of host genes [20]. In fact, many of these miRNAs are coexpressed with their host genes, and approximately 40% of human miRNA genes are in genomic clusters [21]. miRNA genes are frequently located at fragile sites and genomic regions involved in cancers, emphasizing that these miRNAs are involved in cell growth, cell division, and differentiation [22].

A. J. Sanyal () Department of Internal Medicine, Division of Gastroenterology, Hepatology and Nutrition, Virginia Commonwealth University Medical Center, Richmond, VA 23298, USA e-mail: [email protected]

J.-F. Dufour, P.-A. Clavien (eds.), Signaling Pathways in Liver Diseases, DOI: 10.1007/978-3-642-00150-5_33, © Springer-Verlag Berlin Heidelberg 2010

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O. Cheung and A. J. Sanyal

Fig. 33.1  A model for the microRNA biogenesis pathway. See text for details

General Mechanisms of Action The mechanism of action of miRNAs remains controversial. To date, only small numbers of in vivo functions of miRNAs have been demonstrated in many organisms. miRNA functions via either translation inhibition or mRNA degradation. In animals, miRNAs are thought to function through the inhibition of mRNA translation of  target genes through imperfect base-pairing with the  3¢-untranslated region (3¢ UTR) of target mRNAs [23, 24]. Alternative mechanisms of action also include direct mRNA cleavage [25–27], as well as mRNA stability [28, 29]. In vivo miRNA targets are largely unknown, but estimates range from one to hundreds of target genes for a given miRNA, based on miRNA target predictions using a variety of bioinformatics approaches [30–34].

How is miRNA Function Identified and Assessed

can be obtained by several approaches: (1) Generation of stable gain- and loss-of-function phenotypes for individual miRNAs and subsequent identification and isolation of modified cells. This approach utilizes miRNA knockout or knockdown and miRNA overexpression studies. Stable gain-of-function can be achieved by overexpressing individual miRNAs using retro- or lentiviral transfer of suitable miRNA-expression cassettes as previously described [35, 36]. In contrast, no method to induce stable loss-of-function phenotypes for individual miRNAs has yet been reported. So far, chemically modified antisense oligonucleotides complementary to specific miRNAs, i.e., antagomirs and polymorpholinos (PMO), have been shown to transiently interfere with miRNA function in cell culture reporter assays and in mice [7, 37–39]. (2) Micro-array and in situ analysis have revealed specific pattern of miRNA expression, giving hints to the functions of specific miRNAs [38, 40, 41]. (3) Most miRNA target genes come from computational predictions. Only a handful of these predicted targets have been experimentally validated and the majority remains to be verified [42].

Identification of miRNA Function The functions and the targets regulated by individual miRNAs, in particular of those encoded on polycistronic transcripts, are diverse and largely unknown (examples are shown in Table 33.1). Analysis of miRNA function

Luciferase Reporter Assay miRNA function can be assessed using an assay that  employs lentiviral indicator vectors carrying two

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Table 33.1  MicroRNA gene targets Functional class Gene target Transcription factors

Protein synthesis and processing Oxidative stress response

Cell growth and cell cycle

Cell differentiation Anti-apoptosis Pro-apoptosis Anti-inflammatory

Examples of microRNA

Homeobox protein (HOX) Sex-determining region Y-box (SOX) Hepatocyte nuclear factors (HNF) Eukaryotic translation initiation factor (eIF) Ubiquitin conjugating enzymes (UBE) X-box binding protein 1 (XBP1), EDEM Cytochrome C-oxidase Biliverdin reductase Interferon regulatory factor (IRF) Cyclin dependent kinases(CDK) Phosphatidylinositol kinases (PIK) FAD104, AEBP2, MMD BCL2L (1, 2) BCL2L (7, 11) PDCD4, GAS1 Suppressor of cytokine signaling (SOCS)

Pro-inflammatory

Mitogen activated protein kinases (MAPK) C-jun kinase Interleukin precursors

Cholesterol metabolism

Fatty acid metabolism

Oxysterol binding proteins HMGCR, ACAT Oxysterol receptor LXR PPAR, ACSL, ACC, DGAT, SREBP, ACLY, FAS

Carbohydrate metabolism

AMPK, SCD, FABP, LDLR, VLDLR, ELOV ALDOA, GYS, glycogenin, PRKC, PYGL

Insulin signaling

IRS1, myotrophin, Islet1

146b, 199a*, 455, 128b, 128a, 145, 92b, 122 224, 34a, 200a, 199a*, , 27b, 23a, 23b, 125b 146b, 214, 23b, 617, 375, 92b, 26b 23a, 23b, 146b, 128a, 145, 139, 125b 23b, 146b, 27b, 199a*, 16, 128a, 128b, 122 34a, 125b, 200a, 214 99b, 423, 27b, 127, 128a, 128b, 601, 198, 361 99b, 100, 221 16, 24, 23a, 23b, 125b, 128a, 128b, 214, 145 34a, 199a*, 99b, 26, 122 23b, 27b, 26b, 145 23b, 125b, 181a, 122, 146b, 23b, 146b, 26b 16, 24, 214, 200a, 145, 92b 27b, 125b, 16, 24, 181, 199a*, 224, 24, 26b 16, 21, 199a*, 200a, 34a, 122, 145 16, 199a*, 214, 23a, 23b, 27b, 125b, 139, 188 23b, 125b, 199a*, 214, 27b, 128a, 128b, 145 27b, 128a, 128b, 214 125, 214, 16, 128a, 128b, 23b, 122, 188, 191* 128b, 146b, 214, 181b, 23a, 23b, 199a*, 145 125b, 224, 122, 145, 188, 375 423, 145, 191*, 375, 574, 92b, 139 128a, 128b, 146, 122, 125b, 214, 23b, 16, 188 30d, 199a*, 23a, 181b, 200a, 203 23b, 27b, 34a, 221, 21, 125b, 128b, 181b, 122 23a, 16, 27b, 128a, 128b, 375

Gene abbreviations were adopted from Ensembl and NCBI

perfectly complementary target sites for each given miRNA in the 3¢ untranslated region of the Renilla luciferase gene. This assay, as previously described, allows demonstration of the activity of each viral miRNA upon cotransduction of cells with the Renilla luciferase indicator vector together with a firefly luciferase control vector [43–45]. In these studies, the reduced expression of Renilla luciferase in infected cells allowed rapid functional analysis of miRNA translational control of target gene expression.

Translation Translational control is thought to occur at the level of either initiation or elongation [45–47]. Recent studies

have looked at miRNAs that are localized to sites of active translation on polyribosomes, either due to direct interaction with ribosomes or with target mRNAs, by dissociating cultured cells that are fractionated and resolved on sucrose density gradients with subsequent measurements of total mRNA and/or miRNA in the gradient fractions. When initiation of translation is blocked, polysomes are reduced and monosomes (not actively translating) are increased greatly. Along with the reduction in heavy polysomes, the density of mRNAs also shifts to lighter polysomes. A recent study has shown that the sedimentation of miRNA mirrored that of mRNAs when translation is blocked chemically by pactamycin, suggesting miRNAs are associated with mRNA, not ribosome per se [46]. These results reinforced the conclusion that miRNAs are associated with mRNAs undergoing translation.

496

Role of P Bodies in mRNA Degradation Studies have shown that nascent polypeptides are rapidly degraded when mRNAs are under miRNA regulation [47]. Specific cytoplasmic foci, also known as the processing bodies (P bodies), contain untranslated mRNAs and can serve as sites for mRNA degradation [48, 49]. These foci also act as storage sites for mRNAs inhibited by miRNAs [50]. The derepression of mRNA is accompanied by its release from cytoplasmic P bodies and its recruitment to polysomes. Studies have shown that translation repression by RISC (the presence of argonaute proteins within P bodies) led to miRNAmediated repression of protein synthesis by delivering mRNAs to P bodies [51]. Reporter mRNAs that are targeted for translational repression by endogenous or exogenous miRNAs become concentrated in P bodies foci in a miRNA-dependent manner [51]. These results suggest that translation repression by RISC delivers mRNAs to these cytoplasmic foci, either as a cause or as a consequence of inhibiting protein synthesis.

Specific Examples in Liver Disease Role of miRNA miR-122 miR-122 is specifically expressed and is most abundant in the human liver. It accounts for almost 70% of all miRNAs in the liver [52], and has been shown to play an important role in lipid and cholesterol metabolism, and adipocyte differentiation [7, 8, 37], which are at the core of fatty liver disease. Several lines of evidence have suggested the significant impact of this specific miRNA in several liver diseases i.e., nonalcoholic fatty liver disease, hepatitis C virus (HCV) infection, and liver cancers.

Nonalcoholic Fatty Liver Disease It has recently been reported that nonalcoholic steatohepatitis (NASH) is associated with specific differentially expressed miRNAs [38]. Specifically, miRNA miR-122 was found to be underexpressed in subjects with NASH when compared to matched control. The potential targets of these differentially expressed

O. Cheung and A. J. Sanyal

miRNAs are known to play a role in lipid metabolism, cell growth and differentiation, apoptosis, and inflammation; all these are key processes involved in the development and progression of NASH [53]. The potential consequences of these changes can affect insulin signaling, lipid metabolism, cellular responses to stress and apoptosis, inflammation, and response to tissue injury.

Hepatitis C Virus Infection The liver-specific miRNA, miR-122 has been shown to be involved in the replication of HCV in the hepatoma cell line Huh7 [54–56]. It interacts directly with the 5¢ noncoding region of the viral mRNA to enhance its replication [55]. A study by Jopling et al. demonstrated for the first time the link between endogenous expression of this specific miRNA and a major infectious disease, and further suggested that the identification of miR-122 as a critical host factor for HCV viral replication implicates the discovery and the development of a novel therapeutic strategy against HCV infection [55]. In this study, inhibition of miR-122 function using an antisense oligonucleotide indeed led to a significant reduction in the expression of HCV viral RNA. A recent study has also reported that IFN b treatment is associated with a significant reduction in the expression of miR-122. This finding supports the notion that mammalian organisms, through the interferon system, use cellular miRNAs to mount an anti-viral response [57], but this result has been since then challenged [58].

Liver Cancers More than half of the human miRNA genes are located at sites known to be involved in cancers, i.e., fragile sites, minimal regions of loss of heterozygosity, minimal regions of amplification, or common breakpoint regions. These locations suggest that some miRNAs are involved in tumorigenesis [22]. A recent study suggested the role of miRNAs in hepatocellular carcinoma (HCC), i.e., let-7a, miR-21, miR-23, miR-130, miR190, and miR-17-92 family of genes were upregulated, while miR-122 was downregulated in animals with HCC [59]. A recent study has reported differential expression of specific miRNAs in hepatitis C associated

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HCC. Interestingly, miR-122 is again implicated in this disease process, and is found to be overexpressed in these hepatic tumors compared to normal liver parenchyma, along with two other miRNAs, miR-100 and 10a that were also found to be overexpressed in HCV associated HCC [60].

Conclusion The field of miRNA is rapidly growing. miRNAs are predicted to target several thousands of target mRNAs, and approximately 30% of all protein-coding genes [61]. This number is expected to rise as more miRNAs are identified and their targets predicted and validated. miRNAs are involved in diverse biological processes. While the regulation of miRNA function and its mechanism of actions on translational control of target mRNA expression remain unknown, advances in miRNA research allow identification and biochemical characterization of events that limit protein expression. It is crucial to consider the different signaling pathways, as well as different cellular structures, i.e., P bodies in the control of miRNA function. It is hoped that by having a complete and accurate understanding of the functions of miRNAs in various forms of human diseases and their subsequent development, potential therapeutics, and preventive measures can be developed to modify and alter these disease processes.

Summary

›› miRNAs are small, naturally occurring singlestrand RNA of 21–23 nucleotides

›› miRNAs are generated from endogenous tran›› ›› ›› ›› ››

scripts that are encoded by the genome Expression of miRNA is organ and stage of development specific miRNAs are processed from long polycistronic transcripts by RNases (Drosha, Dicer) miRNA functions via either translation inhibition or mRNA degradation miRNAs are predicted to regulate 30% of all protein-coding genes miRNAs have been already implicated in NASH, viral hepatitis, and hepatocellular carcinoma.

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Multiple Choice Questions 1. The first miRNA was discovered in which of the following organism? (a) Caenorhabditis elegans (b) Yeast (c) Chlamydomonas reinhardtii (d) Homo sapiens (e) Mus musculus 2. Which of the following two enzymes are involved in miRNA processing? (a) Drosha, RNA polymerase I (b) Dicer, Drosha (c) Exportin-5, RNase II (d) Pasha, DGCR8 (e) RNase III, Dicer 3. P bodies are associated with which of the following function? (a) mRNA degradation (b) Translation initiation (c) Protein transport (d) miRNA processing 4. Which of the following miRNA is most abundantly expressed in the liver? (a) miR-100 (b) miR-122 (c) miR-145 (d) let-7a (e) miR-21 5. miR-122 has been shown to increase viral replication in: (a) Hepatitis A (b) Hepatitis B (c) Hepatitis C (d) Both Hepatitis B and C (e) Hepatitis D

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33  MicroRNAs 43. Gottwein E, Cai X, Cullen BR (2006) A novel assay for viral microRNA function identifies a single nucleotide polymorphism that affects Drosha processing. J Virol 80: 5321–5326 44. Georgantas RW 3rd, Hildreth R, Morisot S et al (2007) CD34+ hematopoietic stem-progenitor cell microRNA expression and function: a circuit diagram of differentiation control. Proc Natl Acad Sci U S A 104:2750–2755 45. Thermann R, Hentze MW (2007) Drosophila miR2 induces pseudo-polysomes and inhibits translation initiation. Nature 447:875–878 46. Maroney PA, Yu Y, Fisher J et al (2006) Evidence that microRNAs are associated with translating messenger RNAs in human cells. Nat Struct Mol Biol 13:1102–1107 47. Nottrott S, Simard MJ, Richter JD (2006) Human let-7a miRNA blocks protein production on actively translating polyribosomes. Nat Struct Mol Biol 13:1108–1114 48. Pontes O, Pikaard CS (2008) siRNA and miRNA processing: new functions for Cajal bodies. Curr Opin Genet Dev 18:197–203 49. Anderson P, Kedersha N (2008) Stress granules: the Tao of RNA triage. Trends Biochem Sci 33:141–150 50. Bhattacharyya SN, Habermacher R, Martine U et al (2006) Stress-induced reversal of microRNA repression and mRNA P-body localization in human cells. Cold Spring Harb Symp Quant Biol 71:513–521 51. Liu J, Valencia-Sanchez MA, Hannon GJ et al (2005) MicroRNA-dependent localization of targeted mRNAs to mammalian P-bodies. Nat Cell Biol 7:719–723 52. Chang J, Nicolas E, Marks D et al (2004) miR-122, a mammalian liver-specific microRNA, is processed from hcr

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34

Hepatic Clocks Ueli Schibler, Gad Asher, Camille Saini, Jörg Morf, and Hans Reinke

Introduction Most physiological processes are subject to daily ­oscillations that are driven by an endogenous circadian clock. These include rest–activity cycles, cardiovascular functions such as heart rate and blood pressure, the production and secretion of hormones, renal plasma flow and urine production, in addition to metabolic functions of organs associated with the gastrointestinal tract (for review and references, see [1–4]). Since most metabolic functions oscillate in a daily manner, the liver is an organ for which circadian timing is particularly obvious. Thus, genome-wide transcriptome profiling studies have revealed that depending on the stringency of algorithms used for the extraction of oscillating transcripts between 2 and 10% of all liver mRNAs accumulate in a rhythmic fashion [5–10]. The majority of these transcripts encode enzymes and regulators involved in the metabolism of fatty acids, cholesterol, bile acids, carbohydrates, and xenobiotics. Several signaling pathways relevant for hepatic clock outputs (e.g., signaling through PPARs, CAR, LXR, and FXR) are elaborated in previous chapters of this issue. In this chapter, we shall thus focus on putative signaling pathways related to input pathways into the liver clock. Specifically, we will discuss current views and hypothesis on how the master pacemaker in the brain’s suprachiasmatic nucleus (SCN) synchronizes peripheral clocks, in particular those operative in liver. We will also present some findings made with cultured

U. Schibler () Department of Molecular Biology and National Center of Competence in Research “Frontiers in Genetics”, Sciences III, University of Geneva, Quai Ernest Ansermet 30, 1211, Geneva-4, Switzerland e-mail: [email protected]

fibroblasts, since these cells have served as a model system in most in vitro studies. Some of the signaling routes outlined below remain speculative, and their detailed analysis requires additional investigations.

A Circadian Oscillator in Every Single Cell Circadian timing system is actually a more appropriate term than “clock.” In fact, virtually all cells of the body harbor self-sustained and cell-autonomous oscillators of a similar molecular makeup [2, 4, 11, 12]. This begs the question of how phase coherence is established among billions of cellular timekeepers. The answer lies in the hierarchical architecture of the mammalian timing system. Two small aggregates of neurons located in the ventral hypothalamus, known as the suprachiasmatic nuclei, serve as the master pacemakers [13, 14]. For simplicity, the term “SCN” (singular) is generally used when one refers to the master clock, and we shall adhere to this unwritten convention. In the absence of external timing cues, for example in constant darkness, the ­singularfree-runs, that is, it generates rhythms with a period length of approximately – but not exactly – 24 h. Hence the name “circadian,” which is derived from the Latin words “circa diem” (about a day). Therefore, circadian clocks must be daily synchronized in order to stay in resonance with geophysical time. Changes in light intensities are the major Zeitgebers for the SCN pacemaker, but temperature cycles and social cues can also contribute to the phase entrainment [15, 16]. The phase-shifting capacity of the SCN is largely sufficient to adjust circadian time to geophysical time under normal conditions. However, when time zones are suddenly changed by many hours after west- or ­east-bound

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transatlantic flights, we need several days to be in resonance with the photoperiod imposed by the new time zone. This sluggishness in adaptation is experienced as “jet lag,” which ­manifests itself in sleep disturbances and, for some ­individuals, in indigestion and other physiological disturbances [17, 18]. Peripheral timekeepers are just as robust as those of SCN neurons, but they rapidly desynchronize in the absence of an SCN master pacemaker. Hence, the SCN must establish phase coherence in the body by synchronizing billions of individual cell clocks every day [2, 19]. In spite of considerable efforts, we are just starting to have a glimpse at the molecular nature of the involved signaling pathways. The difficulty of studying the signaling routes through which the SCN phase entrains peripheral clocks is not due to the lack of candidates, but due to their overabundance (see below).

The Molecular Oscillator Model The currently held molecular model for circadian rhythm generation is based on interlocked negative feedback loops in gene expression (Fig. 34.1) [4, 19]). The major loop consists of two period genes, Per1 and Per2, and two cryptochrome genes, Cry1 and Cry2. These genes are activated by heterodimers of the PAS domain helix-loop-helix transcription factors BMAL1 and CLOCK or its closely related paralog NPAS2. PER and CRY proteins form heteropolymeric complexes containing CRY and PER isoforms and additional proteins, such as casein kinases 1 epsilon (CK1e) and 1 delta (CK1d), nucleic acid-binding protein NONO, and histone methyl transferase adaptor protein WDR5 [20]. Once these large protein complexes have reached a critical concentration or activity, they inhibit Per and Cry gene expression by attenuating the activity of BMAL1-CLOCK/NPAS2 heterodimers. As a consequence, the levels and activities of PER and CRY mRNA and proteins decrease until they can no longer interfere with BMAL1-CLOCK/NPAS2 transactivators, and a new cycle of CRY and PER expression can ensue. This feedback circuitry also affects the expression of BMAL1 and, to a lesser extent that of CLOCK, through an accessory loop involving transcriptional activators of the retinoic acid-related orphan receptors (RORa, RORb, and RORg) and repressors of the REVERB orphan receptor family (REV-ERBa and REVERBb). RORs activate the expression of BMAL1 and

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compete with REV-ERBa/b for ROR-binding elements (ROREs) within the Bmal1 promoter [21–23]. The transcription of Rev-erba is regulated according to the mechanisms described above for Per and Cry genes: it is activated by BMAL1-CLOCK/NPAS2 and repressed by CRY-PER complexes. The cyclic expression of REV-ERBa/b engenders the rhythmic transcription of Bmal1. As the half-life of BMAL1 is relatively long, the high amplitude of Bmal1 mRNA expression is not accompanied by a high amplitude in BMAL1 protein accumulation. Although the circadian transcription of BMAL1 is not essential for rhythm generation, the coupling between the feedback loops within the negative limb (CRY1/2, PER1/2) and the positive limb (BMAL1/CLOCK) may serve as a rheostat to keep the concentrations of positively and negatively acting clock components within certain boundaries. Posttranslational modifications of core clock proteins play crucial roles in the circadian clockwork circuitry. Thus, phosphorylation of PER proteins by CK1 and other kinases tune the period length of the oscillations by affecting PER protein stability (and probably activity) [24, 25]. For example, the mutation of serine 662 to a glycine in PER2 prevents the cooperative phosphorylation of adjacent S/Ts by CK1e/d, and this leads to a dramatic period shortening [26–28]. As a consequence, the clock of human subjects who carry this dominant mutation runs ahead of time each day, causing a familial advanced sleep phase syndrome (FASPS). Other core clock proteins are also subjected to phosphorylation. Thus, CLOCK, BMAL1, CRY, and REV-ERBa are all phosphoproteins [29–32]. Phosphorylations are not the only posttranslational modifications found on core clock proteins. PER2 and BMAL1 have recently been shown to be acetylated [33, 34]. Moreover, the degradation of PER and CRY proteins is regulated by specific ubiquitin ligase complexes. Finally, CLOCK has itself protein acetyltransferase activity and has been proposed to acetylate BMAL1 [34]. Conceivably, it may also be responsible for the acetylation of PER2. BMAL1 has also shown to be sumoylated [35].

Communicative and Autistic Clocks The same clockwork circuitry appears to be operative in SCN neurons and peripheral cell types. Moreover, circadian gene expression persists indefinitely in dissociated

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Fig. 34.1  Circadian oscillator model. The transcription of Per and Cry genes is activated by heterodimers of BMAL1 (B) and either of the two related proteins, Clock (C) or NPAS2 (N). The polycomb protein EZH2 interacts with these heterodimers and thereby facilitates their action. The accumulation and activity of PER and CRY proteins are also influenced by phosphorylation by protein kinases (CK1d,e), ubiquitination via a complex containing the F-box protein FBXL3 (specific for CRYs), SIRT1dependent decateylation, the histone methyl transferase binding

protein WDR5, and NONO, an RNA and DNA binding protein. DEC1 and DEC2 (D1,2) compete with BMAL1-CLOCK/ NPAS2 heterodimers for E-box binding and thereby reduce E-box-mediated transactivation. A accessory feedback loop, employing the nuclear orphan receptors RORa, RORb, and RORg (RORa,b,g) as activators, and REV-ERBa and REVERBb (REV-ERBa,b) as repressors, regulates the circadian transcription of Bmal1. See text for further explanations. Adapted from [19]

SCN neurons and cultured peripheral cells [36–38]. Hence, circadian oscillators function in self-sustained and cell-autonomous manner. There is, however, a significant difference on how these cellular timekeepers interact with each other in the SCN and in peripheral organs. SCN neurons are strongly coupled via synaptic and paracrine signals [2, 14, 39], while peripheral oscillators (e.g., those in liver) do not communicate with each other [36, 40].

the photoperiod via complex photic signaling through the retinohypothalamic tract [2, 16], must daily synchronize peripheral clocks. Experiments with cultured cells suggest that peripheral oscillators are exquisitely sensitive to phase-shifting agents. Depending on the phase, their phase angle can be reset by up to 180° [37]. Such a phase-resetting behavior manifests itself in a “type zero phase-shifting curve,” since if the new phase is plotted against the old phase after administration of a phase-shifting agent, the slope of the resulting curve is zero. Accordingly, a population of desynchronized cells can be synchronized by a single pulse of a strong chemical timing cue. Surprisingly, a puzzling variety of substances acting as ligands of nuclear and membrane receptors, or as activators of various protein kinases, can synchronize circadian oscillators of cultured cells. These include glucocorticoid hormones [43–46], retinoic acids [47, 48], FGF [49], endothelin [50], TGF-b

Signaling to Peripheral Oscillators The surgical ablation of the SCN leads to immediate arrhythmicity of behavior and physiology [40–42]. Although individual cellular oscillators keep ticking in such SCN-lesioned animals, they rapidly loose phase coherence. Hence, the SCN, which is synchronized to

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[51], prostaglandins [52–54], forskolin [44, 55], tumor promoters [44, 49], Ca++ ionophors [44, 56], and glucose [57]. As serum contains many hormones, growth factors, and cytokines, it efficiently resets the phases of circadian oscillators in cultured cells. The interaction of membrane receptors with their cognate ligands usually results in the activation of protein kinases, which then phosphorylate and activate downstream immediate early transcription factors. These in vitro studies indicate that CREB plays an important role as an immediate early transcription factors in the synchronization of peripheral clocks [44, 55, 58, 59]. Similar to what has been observed for the timekeepers of SCN neurons, activated CREB induces the sudden transcription of Per1 and Per2 genes and thereby either advances or delays the phase of peripheral clocks, depending on the time point of activation. The signaling through nuclear receptors also triggers the stimulation of Per1 and/or Per2 gene expression. For example, the murine Per1 gene harbors two almost perfect GREs in its promoter and first intron, respectively [45]. Some chemicals also reset the phase of cultured fibroblasts by repressing Per gene expression. Hirota and coworkers discovered that acute glucose administration to fibroblasts strongly downregulates Per1 and Per2 expression [57]. These authors found that the glucose-mediated repression of Per transcription requires ongoing transcription, and they speculated that the upregulation of the transcriptional regulators transforming growth factor b (TGFb)-inducible early gene 1 (TIEG1) and vitamin D3 up-regulated protein 1 (VDUP1) may participate in this process. TIEG1 is a corepressor of the transcription factor SP1, and it may thus attenuate Per1 transcription via the SP1 binding sites present in the Per1 and Per2 promoters. As insinuated by its name, TIEG1 is induced by TGFb, a growth factor that also phase shifts circadian clocks in cultured cells [51]. However, these authors proposed an additional pathway for the action on TGFb on the phase of circadian clocks. TGFb activates the activin receptorlike kinase (ALK), which leads to the phosphorylation of SMAD3. This transcription factor then stimulates the transcription of Dec1 as a heterodimer with SMAD4. Dec1 specifies an E-box-binding helixloop-helix transcription factor that activates the expression of Cry1 and represses the transcription of Dbp, Rev-erba, and Per1. Nevertheless, it is ­conceivable that glucose and TGFb synchronize peripheral oscillators via similar – or at least overlapping – mechanisms. Similar to glucose and TGFb, prostaglandin J2 (PGJ2),

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which also resets the phase of circadian clocks in ­fibroblasts, downregulates Per1, Dbp, and Rev-erba [52]. In contrast, other prostaglandins, such as PGE2 (signaling through transmembrane EP receptors) [54] and 15d-PGJ2 (a natural ligand of the peroxisome proliferator-activated receptor gamma (PPARg)) [53], cause phase shifts by augmenting Per1 transcription. Recently, Loudon and colleagues have shown that synthetic ligands of the orphan nuclear receptor REVERBa can also elicit phase shifts in cultured cells [60]. The oscillators of cultured cells can also be phase entrained by physical cues. For example, ­low-amplitude temperature rhythms that resemble body temperature oscillations can efficiently synchronize circadian clocks of cultured fibroblasts [61], and as suggested below, body temperature cycles may also participate in the phase resetting of hepatocyte clocks. Moreover, DNA damage caused through ionizing radiation also acts as a strong phase shifting cue in cultured cells and animals [62].

Synchronization of Liver Clocks Signaling Through Feeding–Fasting Cycles and Redox Sensing Some candidate signaling pathways that may be employed by the SCN to synchronize hepatic clocks are schematically presented in Fig.  34.2. We would like to reemphasize, however, that the involvement of most of these signaling routes is still speculative, since it has not yet been scrutinized by genetic lossof-function experiments. Daily feeding–fasting cycles are clearly the most dominant Zeitgebers for liver clocks, although the involved molecular mechanisms remain to be identified. Feeding–fasting cycles could influence the phase of peripheral oscillators through many different molecular pathways. The signals may include hormones secreted upon feeding or fasting (e.g., cholecystokinin, peptide YY, oxyntomodulin, ghrelin, leptin [63], food metabolites, and intracellular redox state (GSH/ GSSG or NAD(P)H/NAD(P)+ ratios). In vitro experiments by Steven McKnight and coworkers have already demonstrated that the NAD(P)H/NAD(P)+ ratio can affect the binding of CLOCK/NPAS2BMAL1 heterodimers to their cognate DNA sequences

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Fig.  34.2  Candidate pathways for the synchronization of liver clocks. Note that the pathways depicted in panels b, c, and e are hypothetical, as they have not yet been examined by loss-of-function genetics. (a) Per1 and Per2 serve as both immediate early genes and core clock genes. Moreover, most examined synchronization pathways activate or repress the transcription of these genes. Hence, in this “cogwheel cartoon” PER1 and PER2 link phase information from systemic cues to local liver oscillators. (b) Towards the end of the daily fasting period (postabsorptive phase), the nuclear receptor PPARa gets activated. It then stimulates hepatic Fgf21 transcription. FGF21 may act through paracrine and humoral mechanisms in liver and other tissues, respectively, to activate the ERK-CREB pathway. This results in the activation of Per gene transcription. In addition, FGF inhibits lipogenesis in the liver. This may increase the NADH/NAD+ ratio and thereby decrease the activity of SIRT1. Feeding augments the concentration of NAD+ through the stimulation of lipogenesis [90]. (c) Oscillations in body temperature drive the expression of heat shock proteins through the cyclic activation of HSF1 and that of the cold-inducible RNA-binding proteins CIRP and FUS/TLS.

HSP25, which strongly oscillates in the nucleus, may elicit oscillations in redox potentials. HSF1 is also activated through feeding. FUS/TLS may repress some CREB target genes (perhaps Per1/2) (see text). (d) The SCN drives the circadian production and secretion of glucocorticoid hormones in the adrenal gland through the hypothalamus–pituitary–adrenal axis (HPA). This leads to the rhythmic activation of the glucocorticoid receptor (GR), a strong activator of Per1 transcription. (e) Cytoskeleton signaling and SRF. The ring finger ubiquitin ligase RNF6 targets LIMK kinases for degradation. LIMK phosphorylates and inactivates cofilin, a regulator of actin disassembly, and thereby promotes actin polymerization. Free actin represses the transcription activation potential of serum response factor (SRF), which is itself an activator of actin gene transcription. Conceivably, SRF also activates Per1/2 transcription. This would be in keeping with the observation that a serum shock strongly stimulates Per1/2 transcription in cultured fibroblasts. It remains to be shown whether circadian systemic cues act directly on SRF through cytokines and metabolites, the expression of RNF6, or both. Adapted from [94] with permission of SAGE publications

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[64]. The NAD+-dependent histone deacetylase Sirtuin 1 (SIRT1) might be another candidate for connecting cellular metabolism to circadian gene expression [33, 65]. SIRT1 not only deacetylates N-terminal histone tails but also various transcription factors and coactivators [66]. Genetic and biochemical experiments suggest that SIRT1 indeed influences the circadian expression of several clock genes in a significant fashion [33, 65]. Taken together, the data indicate that SIRT1 forms stable complexes with the CLOCKBMAL1 heterodimer and deacetylates BMAL1 and PER2. The deacetylation of PER2 renders this protein less stable, and PER2 thus accumulates to higher levels in Sirt1 knockout cells. In turn, the higher expression of PER2 in SIRT1-deficient cells might lead to a reduced transcription of the genes encoding PER1, PER2, and RORg, and the attenuated expression of RORg (an activator of Bmal1 transcription) is accompanied with a decrease in Bmal1 expression [33]. In addition, SIRT1 could also reduce circadian gene expression more directly by deacetylating H3 and H4 histone tails in the chromatin encompassing these genes [65].

Glucocorticoid Signaling The plasma levels of glucocorticoid hormones display robust daily oscillations in laboratory rodents and humans [67]. Moreover, the glucocorticoid receptor (GR) agonist dexamethasone acts as a strong phaseshifting agent for circadian oscillators in cultured fibroblasts and several peripheral organs, including the liver [43, 46]. Using genetic loss-of-function experiments, Le Minh et al. demonstrated that glucocorticoid hormones act indeed as Zeitgebers at physiological concentrations. These authors monitored circadian gene expression in several tissues of mice with a hepatocyte-specific disruption of the GR gene (GRAlfpCre mice) [45]. Although the loss of GR did not affect the steady-state phase in liver (because of the redundancy of synchronization pathways), it dramatically accelerated the kinetics of feeding-induced phase inversion. The following interpretation was offered by the authors of this study: since the SCN keeps its phase upon changing the feeding regimen, the SCN-regulated signals might be in conflict with Zeitgeber signals emanating from inverted feeding. If so, removing a counteracting SCN-driven pathway like GR signaling should speed up food-related phase inversion, which

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was indeed observed. The Per1 gene contains two highly conserved glucocorticoids-responsive elements (GREs) in its promoter and its first intron, and glucocorticoid-mediated phase resetting may thus act through glucocorticoid-driven Per1 transcription [45].

The Genome-Wide Identification of Putative Immediate Early Genes The large variety of signaling pathways affecting the phase of circadian oscillators in cultured cells suggests that molecular mechanisms accounting for the synchronization of peripheral oscillators in intact animals are very complex and their investigation will be challenging. Kornmann et al. [7, 11] have taken a novel approach for the identification of putative signaling components that may be implicated in the phase entrainment of liver clocks. They engineered a mouse strain in which hepatocyte oscillators can be switched on and off at will and then used genome-wide transcriptome profiling to identify about 50 genes that maintained circadian expression in the absence of functional hepatocyte clocks. The cyclic expression of these genes is likely to be driven by systemic cues emanating directly or indirectly from the master clock in the SCN. At least some of these systemically driven genes probably function as immediate early genes in the synchronization of hepatic oscillators. According to this scenario, these genes sense intraand extracellular rhythmic signals and convey the phase information to core clock components of local oscillators. Intriguingly, Per2 was one of the genes whose rhythmic expression persisted in a nearly unperturbed fashion in the absence of functional hepatocyte oscillators, and Per2 transcription can thus be driven by both systemic cues and local oscillators. PER2 is, therefore, expected to play a prime function in conveying SCNdriven systemic signals to hepatocyte oscillators. Other interesting genes that emerged from this screen and possibly involved in the phase entrainment of hepatocyte will be discussed in the following sections.

FGF21 and PPAR Signaling The system-driven cyclic expression of fibroblast growth factor 21 (FGF21) is particularly intriguing. This protein belongs to a small, atypical FGF subfamily composed of the three members FGF15/19 (FGF19

34  Hepatic Clocks

is the human ortholog of murine FGF15), FGF21, and FGF23 (for review, see [68]. Despite their names, these proteins act as hormones rather than growth factors since, in contrast to other FGFs, they have only low affinity for heparin and are thus not trapped by the extracellular matrix surrounding the producing cells. FGF15/19 and FGF21 are synthesized by intestine and liver, respectively, and both require the coreceptor bKlotho in addition to the canonical FGF receptors FGFR1c (for FGF21) and FGFR4 (for FGF15/19). These systemically acting FGFs control metabolism in various target tissues. FGF15/19 production is induced by bile acids in the gut and dampens hepatic bile acid synthesis by decreasing Cyp7a1 expression [69]. Thereby, FGF15/19 establishes a negative feedback loop that tunes bile acid de novo synthesis to bile acid recycling from the gut to the liver. FGF21 expression is induced by starvation, supposedly through the activation of the PPARa [70, 71]. The phases of diurnal PPARa and Fgf21 mRNA accumulation are indeed very similar and in keeping with a fasting-dependent regulation. Both Ppara and Fgf21 mRNA reach zenith levels at the end of the postabsorptive phase. According to recent studies, FGF21 enhances fat oxidation and glucose uptake in adipocytes and hepatocytes and suppresses de novo lipogenesis in liver [72]. It is, thus, tempting to speculate that FGF21 might participate in peripheral phase entrainment through multiple actions. For example, by inducing glucose uptake and by preventing lipogenesis it may influence the ratio of reduced to oxidized nicotinamide adenosine dinucleotides [NAD(P)H/NAD(P)+] and thereby affect the activity of SIRT1 [33, 65] and/or CLOCK/NPAS2 [64]. In addition, FGF21 is expected to stimulate the mitogenactivated protein kinase (MAPK) pathway and thus elicit the activity of cyclic response element-binding protein (CREB), a major regulator in the circadian synchronization of circadian clocks in SCN neurons and peripheral cells ([73] and references therein). PPAR receptors and coactivators are intimately connected to circadian hepatic gene expression. All three PPAR isoforms (PPARa, PPARb/d, and PPARg) accumulate in daily cycles with different phase angles [74]. They not only serve as hands of the clock to modulate circadian lipid metabolism (for review, see [75, 76]), but also affect the transcription of core clock genes. For example, PPARa and BMAL1 reciprocally activate transcription of their genes and thus establish a feed-forward loop within the hepatic clock [77]. Hence, feeding–fasting rhythms could influence the phase of liver clock gene expression through the

507

circadian production of PPARa ligands. Moreover, the PPARg coactivator PGC-1a cooperates with nuclear orphan receptors of the ROR family to activate Bmal1 and Rev-erba transcription. Since PGC-1a is a sensor of energy metabolism, it may also be an important player in synchronizing hepatic clocks to feeding–fasting rhythms [78].

Signaling Through Components of the Cytoskeleton Other systemically driven genes encode constituents involved in cytoskeleton organization [7, 11]. These include actin (a, g), tubulin (a4, b6), and dynein (LC1) isoforms, and RNF6, an E3 RING finger ubiquitin ligase. RNF6 polyubiquitinates the LIM domain kinases, LIMK1 and LIMK2, and thereby targets them for proteasome-mediated degradation. These two protein kinases phosphorylate and inactivate the actindepolymerizing factors ADF/cofilin and, hence, promote actin polymerization [79]. Thus, the rhythmic accumulation of RNF6 is expected to lead to a corresponding cycle of actin depolymerization and therefore to an oscillation of free-actin (G-actin) levels. In turn, G-actin represses the activity of serum response factor (SRF), an immediate early transcription factor induced by serum treatment in cultured cells [80]. The activation of SRF may be one of the many signaling pathways through which high serum concentrations reset the circadian oscillators in cultured cells, and it is conceivably that cytoskeleton/SRF signaling may also be operative in peripheral tissues of intact animals.

Signaling Through Ubiquitin Ligases The screen for system-driven transcripts also revealed four subunits of various ubiquitin ligase complexes: the two ring finger ubiquitin ligases, RNF6 (see above) and MARCH7, and the two F-box substrate receptors, FBXL20 and FBXO21. Ubiquitin ligase complexes target proteins for proteasome-mediated degradation via polyubiquitination or modulate the activity of proteins through monoubiquitination. The F-box protein FBLX3 has recently been shown to play an important role in the mammalian circadian timing system through the destabilization of cryptochrome proteins [81–83]. Moreover, the F-box proteins beta-TrCP1 and beta-TrCP2 target

508

PER2 for proteasome-mediated degradation. Thus, the downregulation of beta-TrCP1 expression by RNA interference and the overexpression of a dominant negative beta-TrCP1 version increases the stability of PER2 and lengthens t in cultured fibroblasts [84]. Moreover, knocking down the synthesis of both beta-TrCP1 and beta-TrCP2 by shRNAs strongly attenuates circadian gene expression in such cells [85].

Body Temperature Rhythms as Zeitgebers The presence of multiple heat shock protein (HSP) genes (Hspa1a, Hspca, Hspa4, Hspa8, Hsp110, and Stip1) among systemically driven genes suggests a role of body temperature rhythms in driving rhythmic gene expression [7, 11]. The phase of HSP expression is in accordance with this postulate, as zenith levels are observed when body temperature is maximal. In keeping with this observation, the activity of heat shock transcription factor 1 (HSF1), the major temperaturesensing transcription factor regulating transcription of HSP genes, is activated in a robustly circadian manner [86]. Moreover, the expression of CAMKIIB mRNA, encoding a kinase that stimulates the transactivation potential of HSF1, also follows a system-driven daily cycle [87]. The elevated expression of HSPs at maximal body temperature may be a defense mechanism against proteotoxic stress. However, chaperoneassisted protein folding may also play a role in circadian oscillator function. A recent study showed that feeding (the major Zeitgeber for hepatocyte oscillators) can also induce HSF1-mediated HSP transcription in the liver [88]. Perhaps, feeding activates HSF1 through food-borne electrophilic xenobiotics causing oxidative stress [89] or through fat synthesis, which generates an oxidative environment by depleting reduced nicotinamide adenine dinucleotides [NAD(P) H] [90]. As outlined above, the ratio of oxidized to reduced NAD cofactors may influence circadian gene expression by modulating the activities of CLOCKBMAL1 and the histone deacetylase SIRT1. Transcripts, whose accumulation may be temperature dependent, also specify RNA-binding proteins such as CIRP (and its paralog RBM3), FUS/TLS, and DDX46, a DEAD box RNA helicase. The accumulation cycles of these proteins display a phase opposite to that of HSP expression and reach zenith values when body temperature is minimal. Moreover, in

U. Schibler et al.

cultured cells the expression of CIRP, RBM3, and FUS/TLS is induced by lowering the incubation temperature by a few degrees [91, 92]. Hence, the expression of these proteins may indeed be governed by body temperature rhythms. Remarkably, FUS/TLS has recently been shown to repress the action of CREB on the promoter of the human cyclin D1 (CCND1) gene when bound to noncoding RNAs specified by CCND1 promoter sequences [93]. Conceivably, this mechanism is also operative on the CREs present in Per1 and Per2 promoter/enhancer sequences and thereby connects body temperature-dependent with CREB-dependent synchronization pathways. cAMP/ CREB signaling has recently be shown to be an integral component of circadian oscillators in both the SCN and peripheral tissues, such as the liver [73].

Summary

›› The mammalian circadian timing system has

a complex hierarchical architecture. It is composed of a master pacemaker in the SCN of the ventral hypothalamus and peripheral oscillators in virtually every cell of the body. The molecular makeup of central and peripheral clocks is similar; however, while the oscillators in SCN neurons are coupled through synaptic and paracrine signals the ones operative in peripheral cell types do not communicate with each other. The central pacemaker is synchronized daily to light– dark cycles through photic cues perceived by the retina and transmitted to SCN neurons via the retinohypothalamic tract. In turn, the SCN  synchronizes peripheral oscillators by employing a puzzling variety of pathways, including feeding–fasting cycles, body temperature rhythms, and cyclically produced hormones such as glucocorticoids (Fig. 34.3). Several candidate pathways discussed in this chapter emerged from a circadian liver transcriptome analysis aimed at the identification of proteins whose expression and/or activities are driven by systemic signals rather than by local oscillators. Among these are FGF21, structural and regulatory components of the cytoskeleton, and proteins whose activities depend upon body temperature rhythms.

34  Hepatic Clocks

509

Fig. 34.3  Synchronization of SCN and liver clocks. The SCN is composed of neurons whose cellular oscillators are coupled through synaptic and paracrine cues. The SCN emits rhythmic signals that are interpreted by immediate early genes in hepatocytes (and other peripheral cells). The coupling of immediate early gene expression with clock gene expression leads to the synchronization of hepatocyte oscillators. The latter drive the cyclic expression of clock-controlled genes specifying enzymes and regulators of metabolism

SCN SCN

Systemic Signals feeding-fasting cycles hormones (e.g. glucocorticoids) metabolites redox potential body temperatures

CCG CCG IEG photoperiod

Multiple Choice Questions 1. The molecular oscillators in SCN neurons differ from oscillators in peripheral cells in that they are: (a) Self-sustained (b) Cell-autonomous (c) Coupled 2. Rest–activity cycles (driven by the SCN) play an important direct or indirect role in the synchronization of hepatic clocks: (a) Wrong (b) Correct (c) Only if the activity phase exceeds the resting phase 3. According to the currently held model, the circadian oscillator relies on (a) Transcriptional mechanisms (b) Translational and posttranslational mechanisms (c) Transcriptional and posttranslational mecha­ nisms Remark: You cannot fail with answer C3, since it includes all conceivable mechanisms 4. All known signalling substances involved in the phase-resetting of circadian clocks act through (a) Transmembrane receptors

CCG

liver

CCG

(b) Nuclear receptors (c) Transmembrane and nuclear receptors 5. Hepatocytes of mice with a liver-specific knockout of the glucocorticoid receptor gene synchronize more rapidly to inverted feeding–fasting cycles than hepatocytes of wild-type mice. Therefore, (a) Glucocorticoid signaling is one of the foodinduced signaling pathways synchronizing hepatic clocks (b) Glucocorticoid signaling is both necessary and sufficient for the synchronization of hepatic clocks (c) Glucocorticoid signaling counteracts feeding– fasting-dependent synchronization of hepatic clocks when feeding cycles are inverted

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Answers

 1  2  3  4  5  6  7  8  9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34



1e 1e 1c 1e 1e 1d 1c 1d 1b 1c 1b 1b 1d 1e 1b 1c 1a 1e 1e 1b 1d 1b 1e 1e 1b 1a 1d 1b 1c 1c 1a 1c 1a 1c

2d 2e 2a 2c 2d 2b 2b 2a 2e 2e 2c 2b 2b 2e 2c 2c 2a 2e 2c 2a 2c 2e 2b 2e 2c 2a 2d 2a 2c 2e 2b 2a 2b 2b

3e 3a 3b 3e 3a 3c 3b 3e 3e 3b 3a 3a 3e 3d 3c 3c 3a 3d 3c 3a 3a 3c 3e 3d 3e 3c 3b 3b 3a 3b 3b 3a 3a 3c

4a 4e 4a 4d 4a 4b 4e 4c 4d 4c 4b 4d 4a 4d 4c 4a 4b 4b 4e 4b 4a 4d 4d 4e 4a 4a 4e 4e 4e 4e 4b 4c 4b 4c

5e 5b 5e 5d 5b 5a 5c 5c 5d 5b 5d 5c 5d 5e 5c 5c 5e 5a 5d 5c 5c 5c 5c 5c 5e 5d 5c 5e 5e 5e 5d 5d 5c 5c

513

Index

A ABCA1 protein, 294. See also MYC protein Acetaldehyde adducts (AA), 132 Acetaminophen (AAP)-induced liver injury, 218–219 Acetaminophen and liver injury, 73. See also Kupffer cells (KC) Acetaminophen (APAP), application, 296 Acetylation, of HIF-1, 406 Acetyl-CoA carboxylase (ACC), 275, 290 Acidic sphingomyelinase activating domain (ASD), 159 Activated stellate cells, senescence, 59 Activator protein 1 (AP-1), 26 Activin A, role, 49 ADAM molecules, 99. See also Extracellular matrix (ECM) Adenine nucleotide translocator (ANT), 441 Adenomatous polyposis gene product (APC), 365 Adenosine receptor, role, 55 Adenylate cyclase enzyme, 27 Adipokine receptors, role, 55–56. See also Stellate cells Adiponectin receptor 1 (adipoR1), 277 Adult liver repair, Hh-producing and responsive cell types, 392–393. See also Hedgehog (Hh) signal pathway Aflatoxin B1 (AFB1), 470 Akt/protein kinase B (Akt/PKB), 305 Alagille syndrome, 5 Alcoholic hepatitis and TLRs, 154. See also Toll-like receptor (TLR) Alcoholic liver disease (ALD), 74, 83, 132–133. See also Hepatic sinusoidal endothelial cells (HSEC); Kupffer cells (KC); Liver disease Alcohol-induced ER stress, 290–292. See also Liver disease Alpha-fetoprotein (AFP), 5, 344 Amino acid and ammonia metabolism, in hepatocyte, 14 (see also Hepatocyte) in hVps34 activation, 261 metabolism, PPARa, 305 5-Amino-4-imidazole-carboxamide (AICA), 274 AMP-activated protein kinase (AMPK), 262, 273–274 in liver activation, 274 metabolic effects, 274–277 in liver diseases hepatocellular carcinoma and inflammation, 278

liver steatosis, 277–278 type 2 diabetes, 277 a-naphthylisothiocyanate (ANIT), 128 Anterior endomesoderm (AE), 372 Antigen presenting cells (APC), 117 Antimitochondrial antibody (AMA), 131 AP-endonuclease (APE), 349 Apical sodium-dependent bile acid transporter (ASBT), 27 Apoptosis, 437–438 cholesterol and sphingolipids, 443–444 death receptor mediated, 438–440 and liver diseases hepatocarcinogenesis, 446 ischemia/reperfusion liver injury, 445–446 steatohepatitis, 444–445 mitochondrial Ros and Gsh, 442–443 proapoptotic proteins, mitochondrial membrane release cytochrome c mobilization, 440–441 Omm permeabilization, 441–442 regulation, p53, 349–350 Apoptosis inducing factor (AIF), 440 Apoptosis protease activating factor-1 (APAF-1), 162 Apoptosis signal regulated kinase 1 (ASK1), 288 Arginine vasopressin (AVP), 50 Arrest-defective-1 (ARD1), 406 Aryl hydrocarbon receptor nuclear translocator (ARNT), 401 Ascites formation, VEGF, 427–428 Ascorbate deficiency, in ER stress, 297 ATF6 modulation, of UPR transcription, 287 ATP-binding cassette (ABC), 18, 315 Atrial natriuretic peptide (ANP), 74 Autistic clocks, 500–501. See also Hepatic circadian clock; Liver clocks, synchronization Autoimmune hepatitis (AIH), 130 Autoimmune liver diseases (AILD), 128, 130–132. See also Liver disease B Base excision repair (BER), 349 Basic fibroblast growth factor (bFGF), role, 47 Basic-helix-loop-helix (bHLH), 401 Basolateral efflux pumps, in hepatocyte, 18 Basolateral hepatocellular bile acid uptake, 318–319 515

516 Basolateral or sinusoidal domain, 6. See also Hepatocyte Bax inhibitor-1 (BI-1), 296 B-cells, 128 Bcl-2-mediated inhibition, of Bax, 350 Benign liver neoplasms, Wnt/b-catenin pathway, 375–376 Bile acid activated nuclear receptors in bile acid metabolism, secretion and cholestasis, 315 hepatobiliary transport regulation, 318–320 phase I and II bile acid metabolism regulation, 317–318 synthesis regulation, 316–317 therapeutic principles, 320–321 for glucose metabolism, 322–323 for lipid metabolism HDL metabolism, 321 triglyceride and fat metabolism, 321–322 non alcoholic fatty liver disease, 323 Bile acids (BAs), 315 Bile acid synthesis by NRs, regulation, 316–317 Bile duct differentiation, b-catenin, 373 Bile duct epithelial cells (BECs), 130 Bile duct ligation (BDL), 393 Bile duct tumors, Wnt/b-catenin pathway, 378 Bile metabolism, in hepatocyte, 10–11. See also Hepatocyte Bile salt excretion, in hepatocyte, 18 Bile salt export pump (BSEP), 18 Biliary epithelial cells cAMP, 27–28 cytosolic Ca2+, 28–31 expression, 153 MAPK signaling, 31–32 membrane receptors, 25–27 pathological conditions, 33–34 PI3-kinase signaling, 32–33 PKC, 31 Biliary pole, of hepatocyte, 6 Ble acid response element (BARE I), 316 Blood clearance, role, 82. See also Hepatic sinusoidal endothelial cells (HSEC) Bmal1 gene, 500 Bone morphogenetics proteins (BMP), 3 Bone morphogenic protein (BMP) signaling, 366 Branched-chain amino acids (BCAAs), 261 Bromosulphophathalin (BSP), 17 Brown adipose tissue (BAT), 323 C Ca2+-calmodulin activated protein kinase (CaMKII), 456 Cadherins, types, 371 Ca2+-induced Ca2+ release (CICR), 29 Calcium signaling calcium in hepatocyte nucleus, 456–457 Ca2+ oscillations in hepatocytes, 456 and cell death in liver, 457–458 hepatocyte Ca2+ oscillations, 453–454 and hepatocyte proliferation, 457 intercellular Ca2+ waves, 455–456 intracellular Ca2+ waves, 454–455 and ischemia-reperfusion injury, 458 and viral hepatitis, 458

Index Calmodulin-dependent protein kinase (CaMK), 26 Calmodulin-dependent protein kinase II (CamKII), 368 cAMP-response element binding protein (CREB), 26, 337 Canalicular/apical domain, 6–7. See also Hepatocyte Canalicular bile acid excretion, 319–320 Cancer and MYC network, transcriptome, 357–360 Cannabinoids receptors, role, 58. See also Stellate cells Cannabis sativa, 58 Canonical WNT pathway signaling, 366–368. See also Wnt/b-catenin pathway Carbohydrate response element-binding protein (ChREBP), 275 Carnitine palmitoyltransferase 1 (CPT1), 337 Carnitine palmitoyltransferases (CPTs), 305 CAR retention protein (CCRP), 334 Casein kinase (CSK), 391 b-Catenin in bile duct differentiation, 373 gene, 376 in hepatocyte maturation, 373 in liver biology, 374 Cathepsin D (ctsD), 439 CCAAT/enhancer-binding protein (C/EBP), 165, 410 C/EBP homologous protein (CHOP), 289 Cell death in liver and calcium signaling, 457–458 Cell–matrix interaction, pathways. See also Extracellular matrix (ECM) ADAM molecules, 99 DDR2, 99 growth factors, 99 integrin family, 98–99 Cellular lipid uptake and PPARa, 304 Cellular proteins and HBV replication, 464–466. See also Hepatitis B virus (HBV) Cellular transcription and HBx, 467–468 Chaperones and RT–pgRNA interaction, 465 Chemokines, role, 81, 126 Chenodeoxycholic acid (CDCA), 317 Cholangiocarcinoma growth, 32 Hh pathway, 395–396 (see also Hedgehog (Hh) signal pathway) Cholangiocytes. See Biliary epithelial cells Cholestasis, 33. See also Liver disease and bile acid, in ER Stress, 296 Cholestatic disorders, treatment, 320 Cholestatic liver disease, 182–183 Cholesterol 7a-hydroxylase (CYP7A1), 316 Cholesterol and sphingolipids (SLs), in cell death, 443–444. See also Apoptosis Chromatin immunoprecipitation (ChIP), 358, 468 Chromogranin A, occurrence, 28 Chronic cholestasis, platelets, 112–113. See also Platelets Chronic hepatitis B, interferon signaling, 194 Chronic hepatitis D, interferon signaling, 194 Chronic liver disease, platelets. See also Liver disease alterations, 111 in chronic cholestasis, 112–113 and liver fibrosis/cirrhosis, 112 in nonalcoholic steatohepatitis, 112 and viral hepatitis, 111–112

Index Circadian rhythm, molecular model, 500 Circadian timing system, 499–500. See also Hepatic circadian clock; Liver clocks, synchronization Cirrhosis, 83–85. See also Hepatic sinusoidal endothelial cells (HSEC) Cold ischemia, 107–108. See also Platelets Concanavalin A (ConA), 125, 165 Connective tissue growth factor (CTGF), 49, 406, 485 Constitutive androstane receptor (CAR), 315, 331 biology, 333–334 endobiotic homeostasis and disease, 336–337 gene regulation, 334–335 as therapeutic targets, 338 xenobiotic metabolism, 335–336 Copper excretion, in hepatocyte, 19 Covalently closed circular (CCC), 464 C-reactive protein (CRP), 15, 307 CREB binding protein (CBP), 367 Cristae, role, 9. See also Hepatocyte C-terminal functional domain (CTD), 465 C-terminal transactivation domain (C-TAD), 402 CXC chemokine ligand 12 (CXCL12), 26 C-X-C motif chemokine receptor 4 (CXCR4), 26 Cyclic adenosine diphosphate (ADP)-ribose (cADPr), 29 Cyclic Adenosine 3’, 5’-Monophosphate (cAMP), 27–28 Cyclic-AMP response element binding protein (CREB), 295, 505 Cyclin-dependent kinase inhibitor (CDKI), 348 Cyclo-oxygenase-2 (COX-2), 52, 425 Cyclophilin D (CypD), 441 CYP3A4 gene, 334 CYP2B6 gene, 334 Cysteine rich domains (CRD), 159 Cystic fibrosis transmembrane conductance regulator (CFTR), 27 Cytokine receptors, role, 54–55. See also Stellate cells Cytomegalovirus (CMV), 125 Cytosolic Ca2+, 28–31. See also Biliary epithelial cells Cytotoxic T-lymphocyte (CTL), 111, 122 D Death effector domain (DED), 161 Death-inducing signaling complex (DISC), 161, 179, 437 Death receptor 5 (DR5), 469 Death receptor mediated apoptosis, 438–440 Dehydroepiandrosterone sulfate (DHEA), 16 Dendritic cells (DCs), 120–123 expression, 153 Discoidin domain receptors, 99 Divalent metal transporter 1 (DMT1), 15 DNA binding domain (DBD), 331 DNA methyltransferase (DNMT), 474 DNA synthesis, 5 Drug-induced liver disease, 296. See also Liver disease E Ectopic expression, of WT p53, 350 eEF2 kinase, role, 262 eIF-4E, role, 32 Electron transport chain (ETC), 442

517 Endobiotic homeostasis and disease, 336–337 Endoplasmic reticulum (ER), 286. See also Liver disease in hepatocyte, 7–8 signaling pathways with ER stress response, 288–289 unfolded protein response, 286–288 stress in liver, 289 alcohol-induced ER stress, 290–292 drug-induced liver disease, 296 in genetic disorders, 295–296 in nonalcoholic fatty liver disease and insulin resistance, 293–294 in viral infection, 294–295 Endothelial cells, expression, 153 Endothelial dysfunction, in cirrhosis, 84 Endothelial massage, definition, 79. See also Hepatic sinusoidal endothelial cells (HSEC) Endothelial PAS domain protein 1 (EPAS1), 402 Endothelial progenitor cells (EPCs), 85 Endothelin converting enzyme-1 (ECE-1), 49 Endothelin-1 (ET-1), 49 End-stage liver failure (ESLF), 129 Epidermal growth factor (EGF), 26 Epidermal growth factor receptor (EGFR), 26 Epithelial-to-mesenchymal transition (EMT), 394 ER associated degradation (ERAD), 286 ER degradation-enhancing-mannosidase-like protein (EDEM), 295 ER oxidase 1 (ERO1), 289 ER stress response element (ERSE), 287 Erythropoietin (EPO), 16, 401 Eukaryotic elongation factor-2 (eEF2), 276 Extracellular matrix (ECM), 4, 41–43 cell–matrix interaction, pathways ADAM molecules, 99 DDR2, 99 growth factors, 99 integrin family, 98–99 changes, 97–98 components, 93–94 collagen scaffold, 94–95 fibronectin, 95–97 laminin, 95 matricellular proteins, 97 proteoglycans, 95 metalloproteinases and inhibitors, 100–101 stem cell niche, 99–100 Extracellular-signal regulated kinase (ERK), 44, 47, 468 F Factor inhibiting HIF-1 (FIH), 405 Familial advanced sleep phase syndrome (FASPS), 500 Farnesoid X Receptor (FXR), 53, 315, 336 FAS-associated death-domain protein (FADD), 161, 439 Fas (CD95/APO-1), 179 Fas/FasL, 177–178 ligand, 179 in liver diseases, 180–183 signaling, 179–180 Fas ligand (FasL/CD95L), 179 Fasting, PPARa, 305–306

518 Fatty acid binding proteins (FABPs), 304 Fatty acid b-oxidation and PPARa, 304–305 Fatty acid synthase (FAS), 290 Fatty acid transporter 1 (FATP1), 304 Fatty liver and steatohepatitis, 221–222. See also c-Jun NH2-terminal kinases (JNKs) Fatty liver disease, 207 Fibroblast growth factor (FGF), 3 Fibroblast growth factor 15 (FGF15), 317 Fibroblast growth factor 21 (FGF21), 306, 504 Fibronectin (FN), 95–97. See also Extracellular matrix (ECM) Focal adhesion kinase (FAK), 47, 468 role, 56 Focal nodular hyperplasia (FNH), 375 Fz related proteins (FRPs), 366 G G1-arrest of cell cycle, role of p53, 349 Genes of Wnt/b-catenin pathway, 368 Genetic disorders, in ER stress, 295–296. See also Liver disease Glial fibrillary acidic protein (GFAP), 41 Glitazones, role, 307 Glucagon-like peptide 1 (GLP-1), 32, 323 Glucocorticoid receptor, 54 Glucocorticoids-responsive elements (GREs), 504 Glucose metabolism. See also Bile acid activated nuclear receptors AMPK activation, 275 (see also AMP-activated protein kinase (AMPK)) hepatic energy homeostasis, 337–338 in hepatocyte, 11–14 (see also Hepatocyte) PPARa, 305 role of BAs, 322–323 Glucose regulated protein 78 (GRP78), 286 Glucuronidation, of bile acids, 318 Glutamine synthetase, overexpression, 374 Glut-2 transporter, role, 12 Glybenclamide, role, 33 Glycerol kinase (GK), 305 Glycerol-3-phosphate dehydrogenase (GPDH), 305 Glycochenodeoxycholic acid (GCDCA), 296 Glycogen synthase kinase 3b (GSK3b), 365 Glycogen synthase kinase 3 (GSK3), 391, 406–407 Glycosaminoglycans (GAGs), 93 Glycosylphosphatidylinositol (GPI), 71 Golgi complex, in hepatocyte, 8 G6Pase gene, 337 G-protein-coupled-receptor (GPCR), for bile acid, 323–324 Granulocyte macrophage colony stimulating factor (GM-CSF), 121 Granzyme B, 127 Growth factor receptor and integrin signaling, 56–57 GSK3b-binding protein (GBP), 366 GTPase activating protein (GAP), 423 GTP-bound Rheb, role, 261 Guanosine diphosphate (GDP), 25 Guanosine triphosphate (GTP), 25

Index H HBV protein X (HBx), 169 HBx protein and p53, interaction, 348 Heat shock protein (HSP), 506 Heat shock protein 90 (HSP90), 334 Heat shock transcription factor 1 (HSF1), 506 Hedgehog (Hh) signal pathway, 58–59, 391–392. See also Stellate cells hedgehog-producing and hedgehog-responsive cell types, 392–393 in hepatocarcinogenesis cholangiocarcinoma, 395–396 hepatoblastoma, 396 hepatocellular carcinoma, 396 in non-malignant liver diseases biliary fibrosis, 393–394 nonalcoholic and alcoholic fatty liver disease, 394–395 vascular remodeling, 395 Hepatic circadian clock. See also Liver clocks, synchronization circadian timing system, 499–500 communicative and autistic clocks, 500–501 molecular oscillator model, 500 PPARa, 306–307 signaling to peripheral oscillators, 501–502 Hepatic energy homeostasis glucose metabolism, 337–338 lipid metabolism, 337 Hepatic immune cell regeneration, 136–137. See also Liver disease Hepatic inflammation and PPARs, 307–308. See also Peroxisome proliferator-activated receptor alpha (PPARa) Hepatic ischemia-reperfusion injury (IRI), in ER stress, 296. See also Liver disease Hepatic macrophages, role, 74 Hepatic pathophysiology, TNF signaling. See also Tumor necrosis factor-a (TNF) receptors liver regeneration, 165–166 nonalcoholic steatohepatitis, 168–169 toxin-induced liver injury, 166–168 viral hepatitis, 169 Hepatic sinusoidal endothelial cells (HSEC), 117 cellular functions adhesion molecules, expression, 80–81 antigen presentation, 82–83 blood clearance, role, 82 fenestration/filtration, 79–80 metabolism, 82 signaling, 81–82 development and structure, 79 pathobiology aging process, 87 alcoholic liver disease, 83 angiogenesis/hepatic malignancies, 85 cellular rejection, 86 drug toxicity, 85–86 ischemia-reperfusion injury, 86–87 NAFLD, 83 portal hypertension/cirrhosis, 83–85

Index Hepatic stellate cells (HSC), 41–44, 48–49, 117, 308 Hepatitis B surface antigen (HBsAg), 133, 470 Hepatitis B virus (HBV), 348, 463 and HCC genomic and transcriptomic analysis, 471 oncogenesis mechanisms, 471–474 viral epidemiology, 470–471 life cycle and cellular proteins role, 464–466 and liver pathophysiology and apoptosis, 468–470 and lipid metabolism, 470 Hepatitis B virus X protein (HBx), 406, 425 transcription and cell signaling and cellular transcription, 467–468 and signal transduction, 468 and virus, 466–467 Hepatitis C virus (HCV), 294, 308. See also Hepatitis B virus (HBV) clinical consequences of IR and T2D, 485 interference with insulin signaling, 483–484 and interferon signaling, 192–194 perspectives for clinical management, 485–486 and T2D, association, 481–483 Hepatobiliary transport regulation, by NRs alternative basolateral bile acid export, 320 basolateral hepatocellular bile acid uptake, 318–319 canalicular bile acid excretion, 319–320 Hepatoblastomas. See also Hedgehog (Hh) signal pathway Hh pathway, 396 Wnt/b-catenin pathway, 376 Hepatocarcinogenesis (HCC), 446. See also Apoptosis Hh pathway (see also Hedgehog (Hh) signal pathway) cholangiocarcinoma, 395–396 hepatoblastoma, 396 hepatocellular carcinoma, 396 and PPARa, 308 (see also Peroxisome proliferatoractivated receptor alpha (PPARa)) Hepatocellular carcinoma (HCC), 16, 85, 181, 264, 343, 372. See also Hepatocarcinogenesis (HCC) AMPK, 278 and HBV genomic and transcriptomic analysis, 471 oncogenesis mechanisms, 471–474 viral epidemiology, 470–471 Hh pathway, 396 VEGF, 426–427 Wnt/b-catenin pathway, 376–378 Hepatocyte apoptosis and cholesterol, 443–444 (see also Apoptosis) Ca2+ oscillations, 456 development, 3–5 expression, 153 intercellular Ca2+ waves, 455–456 maturation, b-catenin, 373 nucleus, calcium, 456–457 nucleus/polyploidy, 9–10 organelles ER, 7–8 golgi complex, 8 lysosomes, 9

519 mitochondria, 8–9 peroxysomes, 9 physiology acute phase response, 14–15 amino acid and ammonia metabolism, 14 glucose metabolism, 11–14 hepatocyte, 15–16 iron metabolism, 15 lipid/lipoprotein, cholesterol, and bile metabolism, 10–11 protein synthesis, 14 transport-systems, 16–19 plasmamembrane basolateral or sinusoidal domain, 6 canalicular or apical domain, 6–7 lateral domain, 7 tight junctions, gap junctions and desmosomes, 7 proliferation and calcium signaling, 457 structure and renewal, 5–6 Hepatocyte Ca2+ oscillations, 453–454 Hepatocyte growth factor (HGF), 16 Hepatocyte nuclear factor 4 alpha (HNF4a), 316 Hepatocyte nuclear factor (HNF), 10, 465 Hh-interacting protein (Hhip), 393 HIF-1 prolyl hydroxylases (HPH), 403 HIF-related factor (HRF), 402 High-density lipoprotein (HDL), 304 High-fat diet (HFD), 264 High methionine low folate (HMLF), 292 Histone acetyl-transferases (HATs), 303, 406 Histone deacetylase inhibitors (HDACi), 466 Histone deacetylases (HDACs), 406 Human HCC. See also Hepatocarcinogenesis (HCC) pharmacological inhibition of mTOR, 265–266 tumors, upregulation of mTOR signaling, 264–265 Human leukocyte antigen (HLA), 126 Human vacuolar protein sorting 34 (hVps34), 261 Hydroxlation, of HIF-1, 403, 405 5-Hydroxyindole acetic acid (5-HIAA), 106 Hydroxymethylglutaryl-CoA reductase (HMG-CoAR), 10, 446 4-Hydroxy-2,3-nonenal (HNE), 47 5-Hydroxytryptophan (5-HTP), 106 Hyperhomocysteinemia (HHcy), 291 Hypertriglyceridemia, for cardio-vascular disease, 321–322 Hypoxia. See also Liver disease in liver pathologies (see also Hypoxia-inducible factor-1 (HIF-1)) ischemia-reperfusion injury and ischemic preconditioning, 408–409 liver cell carcinomas, 409–410 liver cirrhosis, 409 liver regeneration, 410 other cellular processes, 410 in VEGF expression, 421–422 (see also Vascular endothelial growth factor (VEGF)) Hypoxia-inducible factor-1 (HIF-1) expression in liver, 407–408 identification, 401 regulation acetylation, 406 hydroxlation, 403, 405

520 phosphorylation, 406–407 S-nitrosylation, 407 SUMOylation, 407 structure, 401–403 target genes, 403–404 I IFN stimulated genes (ISGs), 188 IL-1 receptor associated kinase (IRAK), 151 IL-17, role, 131 Impaired glucose tolerance (IGT), 482 Inducible nitric oxide synthase (iNOS), 32, 73 Inflammatory bowel disease (IBD), 338 Inhibitor of apoptosis proteins (IAPs), 161, 440 Inhibitor of kappaB (IkB), 308 Inhibitory PAS domain protein (IPAS), 402 Inner mitochondrial membrane (IMM), 437 Inositol 1,4,5-trisphosphate receptor (InsP3R), 27 Insulin and growth hormone, 234 Insulin-induced PKB/Akt phosphorylation, 264 Insulin-like factor binding protein 1 (IGFBP-1), 13 Insulin-like growth factor (IGF), 260 Insulin receptor (IR), 228 Insulin receptor substrates (IRS), 289, 483 Insulin resistance ER stress, 293–294 (see also Liver disease) response, mTORC1 activation, 263 Insulin response sequence (IRS), 337 Insulin, role, 227–228 Insulin sensitizing agents, application, 485 Insulin signaling, inhibition, 233–234 Insulin, transcriptional regulation, 234–235 Integrin receptors, composition, 98 Integrin signaling and growth factor receptor, cooperation, 56–57. See also Stellate cells Intercellular adhesion molecule 1 (ICAM-1), 81 Intercellular Ca2+ waves, in hepatocytes, 455–456 Interferon alpha/beta receptor (IFNAR), 187 Interferon (IFN) induction of type I, 187–188 signaling and Jak-Stat pathway negative regulators, 190 receptor–kinase complex, 188 refractoriness, 191 STATs, 188–190 signaling in viral hepatitis, 192–194 type I effects antiproliferative effects, 192 antiviral effects, 191–192 interferon regulated genes, 191 Interferon stimulated gene factor 3 (ISGF3), 188 Interleukin-1 receptor-associated kinase (IRAK), 71 Intermembrane space (IMS), 437 Intestinal bile salt transporter (IBAT), 16 Intracellular Ca2+ waves, 454–455 Intrahepatic bile ductal units (IBDUs), 27 Intrahepatic cholangiocarcinoma (ICC), 378 IRE1a-XBP1, role, 286–287 Iron metabolism, in hepatocyte, 15. See also Hepatocyte Iron-responsive elements (IRS), 15 proteins and HCC, 229–230

Index and hepatocyte proliferation, 230–231 insulin signaling pathways, 231–233 members, 228–229 Ischemia-reperfusion injury (IRI), 73–74, 86–87, 219, 296. See also Hepatic sinusoidal endothelial cells (HSEC); Liver disease and calcium signaling, 458 in liver, 408–409, 438, 445–446 (see also Apoptosis) platelets, 106–107 cold ischemia, 107–108 warm ischemia, 108 Ischemic preconditioning, in liver, 408–409 J Jak-Stat pathway and IFN signaling. See also Interferon (IFN) negative regulators, 190 receptor–kinase complex, 188 refractoriness, 191 STATs, 188–190 c-Jun NH2-terminal kinases (JNKs), 47, 164–165, 287. See also Tumor necrosis factor-a (TNF) receptors functions and targets, 213–216 in liver disease AAP-induced liver injury, 218–219 fatty liver and steatohepatitis, 221–222 HCC, 220–221 ischemia-reperfusion injury, 219 liver fibrosis, 220 liver regeneration, 220 TNF-mediated liver injury, 216–218 pathway, 317 Jun-N-terminal kinase (JNK), 370 K Kupffer cells (KC) expression, 153 host defense, neutrophil interaction, 71–73 in liver injury, 73–74 molecular mechanisms, 69–71 portal and pressure, 74 L Laminin, 95. See also Extracellular matrix (ECM) Lateral domain, of hepatocyte, 7. See also Hepatocyte Leptin receptors, signaling, 55 Ligand-binding domain (LBD), 331 Lipid metabolism. See also Bile acid activated nuclear receptors AMPK activation, 275–276 (see also AMP-activated protein kinase (AMPK)) bile acid activated nuclear receptors HDL metabolism, 321 triglyceride and fat metabolism, 321–322 hepatic energy homeostasis, 337 PPARa cellular lipid uptake and transport, 304 and fatty acid b-oxidation, 304–305 and lipoprotein metabolism, 304 Lipopolysaccharide-binding protein (LBP), 168

Index Lipopolysaccharide (LPS), 57 Lipoprotein lipase (LPL), 304 Lipoprotein metabolism and PPARa, 304 Lithocholic acid (LCA), 336 Liver, AMP-activated protein kinase activation, 274 metabolic effects, 274 glucose metabolism, 275 lipid metabolism, 275–276 mitochondrial effects, 276 non-metabolic effects, 276–277 protein synthesis, 276 Liver cancer, NF-kB, 205–206. See also Liver disease Liver cell carcinomas, HIF-1, 409–410 Liver cirrhosis, HIF-1 signaling pathway, 409 Liver clocks, synchronization body temperature rhythms, 506 feeding-fasting cycles and redox sensing, signaling, 502–504 FGF21 and PPAR signaling, 504–505 genome-wide identification, 504 glucocorticoid signaling, 504 signaling by cytoskeleton components, 505 ubiquitin ligases signaling, 505–506 Liver disease, 117–119 adaptive immunity B-cells, 128 T-cells, 126–127 AMPK, 277–278 and apoptosis hepatocarcinogenesis, 446 ischemia/reperfusion liver injury, 445–446 steatohepatitis, 444–445 ER stress, 289 alcohol-induced ER stress, 290–292 drug-induced liver disease, 296 in genetic disorders, 295–296 in nonalcoholic fatty liver disease and insulin resistance, 293–294 in viral infection, 294–295 Fas/FasL, 180–183 immune cell function alcoholic liver disease, 132–133 autoimmune liver disease, 130–132 hepatic immune cell regeneration, 136–137 liver transplantation, 129–130 viral hepatitis, 133–135 immune response, regulation, 128 innate immunity, 119–120 DC, 120–123 NK cell, NKT cells, and Tgd cells, 123–126 JNKs AAP-induced liver injury, 218–219 fatty liver and steatohepatitis, 221–222 HCC, 220–221 ischemia-reperfusion injury, 219 liver fibrosis, 220 liver regeneration, 220 TNF-mediated liver injury, 216–218 miRNA, 494–495 mTORC1 regulation (see also Mammalian target of rapamycin (mTOR)) liver regeneration, 263–264

521 metabolic disorders, 264 mTOR signaling upregulation, in human HCC, 264–265 NF-kB, 205–207 PI3K/PTEN/PKB pathway HCC development, 250–251 insulin resistance, 249–250 phosphatases involvement, 250 viruses, PKB and liver diseases, 251–252 TLR signaling, 119 Liver, ECM components, 93–94 collagen scaffold, 94–95 fibronectin, 95–97 laminin, 95 matricellular proteins, 97 proteoglycans, 95 Liver fatty acid binding protein (L-FABP), 309 Liver fibrosis, 220. See also c-Jun NH2-terminal kinases (JNKs); Liver disease and cirrhosis, VEGF, 424–425 NF-kB, 206–207 and PPARs, 308–309 (see also Peroxisome proliferator-activated receptor alpha (PPARa)) and TLRs, 155 (see also Toll-like receptor (TLR)) Liver fibrosis/cirrhosis and platelets, 112. See also Platelets Liver injury, KC, 73–74 Liver organogenesis, VEGF, 424 Liver pathophysiology and HBV and apoptosis, 468–470 and lipid metabolism, 470 Liver Receptor Homolog 1 (LRH-1), 316 Liver regeneration, 72–73, 165–166. See also Tumor necrosis factor-a (TNF) receptors HIF-1, 410 host defense, 71 immune tolerance, 71–72 mTORC1, 263–264 phases, 136 PKB, 248–249 platelets platelet-derived serotonin, role, 110–111 role, 108–110 VEGF, 424 Liver resident cell, types, 4. See also Hepatocyte Liver steatosis AMPK, 277–278 ER stress, 293 Liver transplantation, 129–130. See also Liver disease VEGF, 428 Liver, Wnt/b-catenin pathway development, 372–374 growth, metabolism, and homeostasis, 374 regeneration, 374–375 Liver X receptor-a (LXRa), 316, 470 Liver X receptor (LXRs), 11, 54 LPS binding protein (LBP), 70 Lymphocytes, occurrence, 117 Lysosomes, in hepatocyte, 9 M Major histocompatibility complex (MHC), 82 Malondialdehyde-acetaldehyde (MAA), 132

522 Mammalian lethal with SEC13 protein 8 (mLST8), 259 Mammalian target of rapamycin complex 1 (mTORC1), 276. See also Mammalian target of rapamycin (mTOR) activation, physiological effects, 259, 261 negative feedback effect, 262–263 positive anabolic effects, 262 regulation in liver disease liver regeneration, 263–264 metabolic disorders, 264 Mammalian target of rapamycin (mTOR), 259 mTORC1 activation in human HCC, 264–265 liver regeneration, 263–264 metabolic disorders of liver, 264 negative feedback effect, 262–263 positive anabolic responses, 262 pharmacological inhibition in HCC in clinical trials for HCC, 266 preclinical studies in animal models, 265–266 signaling pathways activation of mTOR complexes, 260–262 mTOR complexes, 259–260 Manganese superoxide dismutase (MnSOD), 167 MAP kinase phosphatases (MKPs), 214 MAPK signaling, 31–32. See also Biliary epithelial cells Matricellular proteins, role, 97. See also Extracellular matrix (ECM) Matrix metalloproteinases (MMP), 100, 422 Mdm2 gene, 347 Mdm2-p53 interaction, disruption, 348 Megakaryocyte, maturation, 105 Meprin/A5/Mu (MAM), 371 Metabolic syndrome and PPARs, 307. See also Peroxisome proliferator-activated receptor alpha (PPARa) Metalloproteinases and inhibitors, 100–101. See also Extracellular matrix (ECM) Metastasis-associated protein 1 (MTA1), 406 Metformin, usage, 274 Methionine adenosyltransferase-1A (MAT1A), 444 MicroRNAs (miRNA) history, 491 identification and assessment, 492–494 in liver disease, 494–495 mechanism of action, 492 types and biogenesis, 491–492 Mitochondria, in hepatocyte, 8–9 Mitochondrial permeability transition (MPT), 440 Mitochondrial Ros and Gsh, 442–443. See also Apoptosis Mitochondrial stress, in ALD, 292 Mitochondria of superoxide dismutase (Mn-SOD), 442 Mitogen-activated protein kinase (MAPK), 26, 406, 425, 505 Monocyte chemoattractant protein-1 (MCP-1), 51, 55 Mouse embryo fibroblasts (MEFs), 262 Multidrug resistance glycoprotein (MDR), 19 Multidrug resistance (MDR1), 409 Multidrug resistance protein family (MRP), 18 Murine embryonic fibroblast (MEF), 203 MycER protein, 357 MycERT2 expression, 360

Index MycERtransduced cells, chimeric MycER protein in, 357 Myc gene, 357 MYC protein transcriptome network, 357–360 in vivo biological output, 360 MYC-responsive genes, 357 MyD88-dependent signaling, 151 MyD88-independent signaling, 151–152 Myristoylated alanine rich protein kinase C substrate (MARCKS), 46 N Na+ dependent bile salt uptake, in hepatocyte, 16 Na+ independent hepatic uptake, of hydrophilic organic cations and anions, 17–18. See also Hepatocyte Na+-taurocholate cotransporting polypeptide (NTCP), 16 Natural killer dendritic cell (NKDC), 121 Neutral sphingomyelinase activating domain (NSD), 159–160 NF-E2-related factor-2 (Nrf-2), 287 Niemann Pick type C (NPC), 443 Non-alcoholic fatty liver disease (NAFLD), 83, 168, 264, 277. See also Liver disease; Peroxisome proliferator-activated receptor alpha (PPARa) ER stress, 293–294 and PPARs, 309 and TLRs, 155 (see also Toll-like receptor (TLR)) Non-alcoholic steatohepatitis (NASH), 111, 168–169, 183, 207, 249, 264, 277, 309, 394. See also Tumor necrosis factor-a (TNF) receptors platelets, 112 (see also Platelets) Nonbile salt organic anions, excretion, 18 Nonesterified fatty acids (NEFA), 249 Non-malignant liver diseases, Hh pathway. See also Hedgehog (Hh) signal pathway biliary fibrosis, 393–394 nonalcoholic and alcoholic fatty liver disease, 394–395 vascular remodeling, 395 N-terminal transactivation domain (N-TAD), 402 Nuclear export signal (NES), 346, 406 Nuclear factor-kB (NF-kB), 162–164, 439. See also Tumor necrosis factor-a (TNF) receptors activation hepatocyte cell death, regulation, 202–205 in liver disease, 205–207 regulation, 200–202 therapeutic target, 207–208 transcription factor family, 199–200 Nuclear hormone receptors (NHRs), 331 CAR biology, 333–334 PXR biology, 332–333 Nuclear localization signal (NLS), 346 Nuclear receptor co-repressor (NCoR), 331 Nuclear receptor family, 53–54. See also Stellate cells Nuclear receptors (NRs), 315 O O2-dependent degradation (ODD), 402 Open reading frames (ORF), 463 Organelles, in hepatocyte, 7–9. See also Hepatocyte

Index Organic anion transporters (OAT), 17, 318 Organic anion transporting polypeptide family (OATP), 16–17 Organic cation transporter novel type (OCTN), 17 Organic cation transporter (OCT), 17 Orthotopic liver transplantation (OLT), 129 Outer mitochondrial membrane (OMM), 437 Oxygen-regulated protein 150 (ORP150), 294 P Pancreatic neuroendocrine tumors (PNET), 266 Para aminhippurate (PAH), 17 Pathogen associated molecular pattern (PAMP), 119, 149, 187 Pattern recognition receptors (PRR), 119, 187 PEPCK1 gene, 337 Per/ARNT/SIM (PAS), 401 PERK-eIF2a, in mRNA translation, 287–288 Peroxiredoxin-III (Prx-III), 442 Peroxisome proliferator-activated receptor alpha (PPARa), 465 in amino acid metabolism, 305 in fasting, 305–306 in glucose metabolism, 305 hepatic circadian clock, 306–307 in lipid metabolism, 304–305 Peroxisome-proliferator activated receptor-(co-activator 1 a) (PGC1a), 277 Peroxisome proliferator-activated receptor-g (PPAR-g), 52, 54, 470 Peroxisome proliferator-activated receptors (PPARs), 11, 53, 303–304. See also Liver disease in disease hepatic inflammation, 307–308 and hepatocarcinogenesis, 308 and liver fibrosis, 308–309 metabolic syndrome, 307 Peroxisome proliferator response element (PPRE), 303 Peroxysomes, in hepatocyte, 9 Phosphatidic acid (PA), 44 Phosphatidylinositide-3OH kinase (PI3K) pathway, 260 Phosphatidylinositol 4,5-bisphosphate (PIP2), 28, 454 Phosphoenolpyruvate carboxykinase (PEPCK), 248 Phospholipase C (PLC), 25 Phospholipid excretion, in hepatocyte, 18–19 Phosphorylation, of HIF-1, 406–407 Phosphotidylinositol 3-kinase (PI3-K), 32, 45–46 Phosphotyrosine binding (PTB), 261 PI3-kinase signaling, 32–33. See also Biliary epithelial cells PI3K/PTEN/PKB pathway, in liver diseases HCC development, 250–251 insulin resistance, 249–250 phosphatases involvement, 250 viruses, PKB and liver diseases, 251–252 PKB/Akt activation and regulation activating stimuli and upstream kinases, 242–244 interactors, 245–246 negative regulation, 244–245 positive regulation, 244 concept, 241–242

523 PKB substrates, functions apoptosis, regulation, 246–247 cell cycle control, 247 cell size and survival, regulation, 246 in liver regeneration, 248–249 metabolism, 247–248 Planar cell polarity (PCP), 368 pathway, 369–370 (see also Wnt/b-catenin pathway) Plasmamembrane, in hepatocyte, 6–7. See also Hepatocyte Plasminogen activator inhibitor (PAI), 48 Platelet-derived growth factor (PDGF), 44–47, 110 signaling, 57 Platelet-derived serotonin, role, 110–111. See also Platelets Platelets, 105–106 in chronic liver disease alterations, 111 in chronic cholestasis, 112–113 and liver fibrosis/cirrhosis, 112 in nonalcoholic steatohepatitis, 112 and viral hepatitis, 111–112 definition, 105 ischemia/reperfusion injury, 106–107 cold ischemia, 107–108 warm ischemia, 108 liver regeneration platelet-derived serotonin, role, 110–111 role, 108–110 Poly(ADP-ribose) polymerase-1 (PARP-1), 163 Polychlorinated biphenyls (PCBs), 333 Polyinosinic–polytidylic acid (Poly IC), 125 Polymorpholinos (PMO), 492 Polyploidy, in hepatocyte, 9–10 Porin, role, 9. See also Hepatocyte Portal Associated Lymphoid Tissue (PALT), 121 Portal hypertension, VEGF, 425 PPAR isotypes, expression patterns, 304 Pregenomic RNA (pgRNA), 464 Pregnane X Receptor (PXR), 315, 331 biology, 332–333 endobiotic homeostasis and disease, 336–337 gene regulation, 334–335 as therapeutic targets, 338 xenobiotic metabolism, 335–336 Preligand binding assembly domain (PLAD), 160 p70 ribosomal protein S6 kinase (p70S6K), 276 Primary biliary cirrhosis (PBC), 128, 378, 394 Primary sclerosis cholangitis (PSC), 130, 378 Progressive familial intrahepatic cholestasis type 2 (PFIC2), 18 proline-rich PKB/Akt substrate 40 kDa (PRAS40), 259 Proline-rich tyrosine kinase 2 (Pyk2), 468 Prolyl hydroxylase (PHD), 403 Prostaglandin E2 (PGE2), role, 70 Protein-caloric malnutrition, BCAA supplementation, 262 Protein disulfide isomerase (PDI), 287 Protein inhibitor of activated STAT1 (PIAS1), 190 Protein kinase A (PKA), 45, 391 Protein kinase C (PKC), 31, 368 Protein phosphatase 2A (PP2A), 484 Protein phosphorylation, 56

524 Protein structure of p53, 344–346 Protein synthesis AMPK activation, 276 (see also AMP-activated protein kinase (AMPK)) in hepatocyte, 14 (see also Hepatocyte) Protein tyrosine phosphatase 1B (PTP1B), 371 Proteoglycans and ECM, interaction, 95. See also Extracellular matrix (ECM) Proteoglycans, interaction, 99 p53 tumor suppressor, 343 apoptosis, 349–350 cellular stress, 348–349 liver-specific challenges of functions, 351 protein structure, 344–346 regulation of protein levels, 346–348 selectivity in downstream response, 350–351 subcellular localization, 348 transcription factor, 343–344 Pyruvate dehydrogenase complex (PDC), 131 Pyruvate dehydrogenase kinase 4 (PDK4), 305 R Rag GTPases, activation, 261 Rapamycin application, 259 treatment, on liver regeneration, 263–264 Raptor, role in mTORC1, 259 Ras homolog enriched in brain (Rheb), 261 Ras, role, 261 Receptor-interacting protein (RIP), 161, 439 Redox factor 1 (Ref1), 26 Rel homology domain (RHD), 162 Resistin, role, 56 Retinoblastoma (Rb) protein, 348–349 Retinoic acid receptors (RAR), 53 Retinoid receptor X (RRX), 11 Retinoid X receptor alpha (RXRa), 465 Retinoid X receptors (RXR), 53, 303 Rho-associated kinase (ROCK), 370 Ribonucleotide reducatse, importance, 349 ROR-binding elements (ROREs), 500 Rough ER (RER), 7, 8 Ryanodine receptor (RyR), 29 S S-adenosyl-l-methionine (SAM), 444 Second mitochondria-derived activator of caspases (SMAC), 180 Selective PPAR modulators (SPPARMs), 309 Serotonin, metabolism, 106 Serotonin reuptake transporter (SERT), 106 Ser/Thr protein kinase mTOR, 259 Serum response factor (SRF), 505 Severe combined immunodeficient (SCID), 136 Signal transducer and activator of transcription 3 (STAT3), 165 Signal transducers and activators of transcription (STATs), 188–190 Silencer of the death domain protein (SODD), 161 Silencing mediator of retinoic acid and thyroid hormone receptor (SMRT), 331

Index Simian virus 40 (SV40), 346 Sinusoidal endothelial cells, feature, 79 Sinusoid capillarization, definition, 97. See also Extracellular matrix (ECM) Sinusoids, role, 79 Sirolimus, application, 428 Smad proteins, 48 Small Heterodimer Partner (SHP), 316 Smooth ER (SER), 7 S-nitrosylation, of HIF-1, 407 Soluble frizzled related proteins (sFRP), 376 Sphingolipids and cell death, 444. See also Apoptosis Sphingomyelinases (SMases), 439 Sphingosine 1-phosphate (S1P), 444 Src homology 2 (SH2), 261 SREBP cleavage activating protein (SCAP), 289 Stearoyl-CoA desaturase (SCD), 290, 293 Steatohepatitis (SH), 444–445. See also Apoptosis Stellate cells activated stellate cells, senescence, 59 adipokine receptors, 55–56 cannabinoids receptors, 58 cytokine receptors, 54–55 expression, 153–154 growth factor receptor and integrin signaling, cooperation, 56–57 hedgehog signal pathway, 58–59 HSC, 41–44, 48–49 nuclear receptor family, 53–54 PDGF, 44–47 TGF-b receptor superfamily, 47–49 TLRs and HSC, 57–58 TNF receptor superfamily, 51–53 transmembrane domain receptors, 49–51 Stem cells in adult liver, Wnt/b-catenin pathway, 375 niche and ECM, 99–100 (see also Extracellular matrix (ECM)) StepMiner method, usage, 359 Sterol regulatory element binding protein-1c (SREBP-1c), 168, 275, 289, 470 Sterol regulatory element-binding protein family (SREBP), 234 Stress-activated protein kinases/NH2-terminal-Jun kinase (SAPK/JNK), 468 Subcellular localization, of p53, 348 SUMOylation, of HIF-1, 407 Suppressor of cytokine signaling (SOCS), 190, 232, 483 Suprachiasmatic nucleus (SCN), 499 Sustained virological response (SVR), 483 T TATA-box binding protein (TBP), 371 Taurocholic acid (TCA), 31 Taurolithocholic acid (TLCA), 31 Tauroursodeoxycholic acid (TUDCA), 31 T cell factor/lymphoid enhancing factor (TCF/LEF), 367 T cell factor-1 (TCF1), 365 T cell protein tyosine phosphatase (TcPTP), 190 T-cells, 126–127 5¢-Terminal oligopyrimidine tract (5¢-TOP), 263

Index TGF-b receptor superfamily, 47–49. See also Stellate cells Thiazolidinediones (TZD), usage, 274 Thioredoxin 1 (TRX1), 26 Thioredoxin-2 (Trx-2), 442 Thrombin (THR), 50 Thrombocytopenia, 111 Thrombopoietin (TPO), 15, 105 Tissue inhibitor of metalloproteinase (TIMP), 48, 55, 100–101, 159 TNF-activated factor 6 (TRAF-6), 71 TNFR-associated death-domain (TRADD) protein, 161, 439 TNF receptorassociated factor-2 (TRAF2), 288 TNF receptor-associated factor 6 (TRAF6), 151 TNF receptors (TNFR), 438 TNF-related apoptosis-inducing ligand (TRAIL) receptors, 438 TNFR1, role, 52 Toll-like receptor (TLR), 119 adapters and signaling, 150–152 clinical implications, 154–155 expression in liver, 153–154 future perspectives, 155–156 and HSC, 57–58 (see also Stellate cells) and ligands, 149–150 negative regulation, 152 signaling, 119 Total parenteral nutrition (TPN), 322 Toxin-induced liver injury, 166–168. See also Tumor necrosis factor-a (TNF) receptors Toxoplamsa gondii, 150 TP53 gene, 345 Transarterial chemoembolization (TACE), 427 b-Transducin repeat-containing protein (bTrCP), 366 Transferrin receptors (TfR), 15 Transforming growth factor-b-activated kinase (TAK1), 274 Transforming growth factor b (TGFb), 372 Transient receptor potential (TRP), 30 Transmembrane domain receptors, 49–51. See also Stellate cells Tryptophan hydroxylase (TPH), 105 Tuberous sclerosis complex protein 2 (TSC2), 261 Tumor necrosis factor-a (TNF) receptors in hepatic pathophysiology liver regeneration, 165–166 nonalcoholic steatohepatitis, 168–169 toxin-induced liver injury, 166–168 viral hepatitis, 169 molecules and structure, 159–160 signaling pathways c-Jun N-terminal kinase, 164–165 intracellular death signaling complex, 160–161 mitochondrial amplification, 161–162 NF-kB, 162–164 Tumor necrosis factor related apoptosis inducing ligand (TRAIL), 202 Tumor necrosis factor (TNF), 278 receptor superfamily, 51–53 (see also Stellate cells) Type 2 diabetes (T2D). See also Hepatitis C virus (HCV) AMPK, 277 and HCV, 481–483

525 Type I interferon effects antiproliferative effects, 192 antiviral effects, 191–192 interferon regulated genes, 191 induction, 187–188 U Ubiquitin specific peptidase 18 (USP18), 190 Unfolded protein response (UPR), 286–288. See also Endoplasmic reticulum (ER) 3¢-Untranslated region (3¢ UTR), 49, 492 Ursodeoxycholic acid (UDCA), 31, 319 V Vascular endothelial growth factor receptor (VEGFR), 27 Vascular endothelial growth factor (VEGF) biological functions, 419–420 gene and splice variants, 420 gene expression, 421–422 protein family, 420–421 receptors neuropilin-1 and 2, 423 receptor signaling, 423–424 VEGFR-1, 422–423 VEGFR-2, 423 VEGFR-3, 423 role, 47 signaling in liver conditions ascites formation, 427–428 hepatocellular carcinoma, 426–427 liver fibrosis and cirrhosis, 424–425 liver organogenesis, 424 liver regeneration, 424 liver transplantation, 428 portal hypertension, 425 viral hepatitis, 425–426 Vascular permeability factor (VPF), 419, 427 Vascular pole, definition, 6 Vasoconstrictors, definition, 50 VEGF gene, 420 Veno-occlusive disease (VOD), 86 Very low-density lipoproteins (VLDL), 8, 11, 304 Viral hepatitis, 133–135, 169. See also Hepatitis B virus (HBV); Hepatitis C virus (HCV); Liver disease; Tumor necrosis factor-a (TNF) receptors and calcium, 458 interferon signaling, 192–194 (see also Interferon (IFN)) NF-kB, 207 and platelets, 111–112 (see also Platelets) and TLRs, 154 (see also Toll-like receptor (TLR)) and VEGF, 425–426 Viral infection and ER stress, 294–295 (see also Liver disease) and HBx, 466–467 (see also Hepatitis B virus X protein (HBx)) Virus–cell interactions, in hepadnaviral life cycle, 464 Vitamin D Receptor (VDR), 54, 315 Vitamin D3 up-regulated protein 1 (VDUP1), 502

526 W Warm ischemia, 108. See also Platelets White adipose tissue (WAT), 264, 323 Wilson’s disease, 183 Wnt/b-catenin pathway, 365–366 in liver benign liver neoplasms, 375–376 bile duct tumors, 378 development, 372–374 growth, metabolism, and homeostasis, 374 hepatoblastomas, 376 hepatocellular cancer, 376–378 pathologies, 378 regeneration, 374–375

Index in stem cell biology development, 375 therapeutic implications, 378–379 transduction pathway alternative WNT signaling pathways, 368–370 b-catenin-E-cadherin interactions, 370–371 canonical WNT pathway, 366–368 interactions, 371–372 WNT/Ca2+ pathway, 368–369. See also Wnt/b-catenin pathway Wnt inhibitory factors (WIFs), 366 X X box-binding protein 1 (XBP-1), 286 Xenobiotic metabolism, 335–336 Xenobiotic-responsive enhancer module (XREM), 334