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Hepatotoxicity
Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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Hepatotoxicity From Genomics to in vitro and in vivo Models
Editor
SAURA C. SAHU US Food and Drug Administration, Laurel, MD, USA
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C 2007 Copyright
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Dedicated to My parents, Gopinath and Inchhamani, for their gift of life, love and living examples My teachers for their gift of education My wife, Jharana, for her life-long companionship, love and support for following my dreams My children, Megha, Sudhir and Subir, for their love and care
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Contents
Contributors Preface Acknowledgements SECTION 1
MODELS FOR HEPATOTOXICITY TESTING
1
Current in vitro Models to Study Drug-Induced Liver Injury Julio C. Davila, Jinghai J. Xu, Keith A. Hoffmaster, Peter J. O’Brien and Stephen C. Storm
2
Utilization of an in vitro Hepatotoxicity Test in the Early Stage of Drug Discovery Ikuo Horii, Hiroshi Yamada, Rie Kikkawa, Toshinori Yamamoto, Tamio Fukushima and Kaori Tomizawa
3
4
5
6
xi xvii xix
3
57
Use of Hepatocytes for Characterizing a Candidate Drug’s Metabolism and Drug Interaction Potential Srikanth C. Nallani, John M. Strong and Shiew Mei Huang
69
Human-Based in vitro Experimental Systems for the Evaluation of Human Drug Safety Albert P. Li
89
Hepatocytes as a Model for Screening Food-Related Hepatotoxins and Studying Mechanisms of their Toxicity Saura C. Sahu Some Experimental Models of Liver Damage Pablo Muriel
105 119
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BIOMARKERS OF HEPATOTOXICITY
Biomarkers of Mycotoxin Exposure in Liver Toxicity Angela J. Harris
SECTION 4 9
HEPATOCYTE CULTURES
Application of Short- and Long-Term Hepatocyte Cultures to Predict Toxicities Gregor Tuschl, Jens Hrach, Philip G. Hewitt and Stefan O. Mueller
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MECHANISMS OF HEPATOTOXICITY
Mechanisms of Toxic Liver Injury Nora Anderson and J¨urgen Borlak
191
10
A Role of Cytochrome P450 in Quinone-Induced Hepatotoxicity Yasuhiro Ishihara and Norio Shimamoto
287
11
A Mechanistic View of Troglitazone Hepatotoxicity Rawiwan Maniratanachote and Tsuyoshi Yokoi
299
12
Role of the Kupffer Cell in Hepatotoxicity and Hepatocarcinogenesis James E. Klaunig, Stacy M. Corthals, Lisa M. Kamendulis and Binu K. Philip
313
13
Sinusoidal Cells in Liver Injury and Repair Carol R. Gardner and Debra L. Laskin
341
14
Cytokines in Liver Diseases Pablo Muriel
371
15
Bile Acids as Modulators of Apoptosis Rui E. Castro, Susana Sol´a, Clifford J. Steer and Cec´ılia M.P. Rodrigues
391
16
Drug-Induced Intrahepatic Cholestasis by Interaction with the Hepatic Bile Salt Export Pump (BSEP) Christoph Funk, Johannes No´e, Ren´ee Portmann, Ruben Alvarez-S´anchez, Florian Klammers, Christiane Lamy, Axel Paehler and Michael Pantze
SECTION 5 17
421
GENOMICS OF HEPATOTOXICITY
Application of Toxicogenomics in Predicting Hepatotoxicity – Potentials and Challenges Wen Lin, Guoxiang Shen, Tin Oo Khor and Ah-Ng Tony Kong
449
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18
Genomic Profiling of Liver Injury Kevin Gerrish and David E. Malarkey
465
19
Use of DNA Arrays in Understanding Hepatic Test Systems Angela J. Harris and Daniel A. Casciano
489
20
Prediction of Hepatotoxicity Based on the Toxicogenomics Database Tetsuro Urushidani
507
21
Relationship between N-acetyltransferase-2 Gene Polymorphism and Isoniazid-Induced Hepatotoxicity Yasuo Shimizu, Kunio Dobashi and Masatomo Mori
SECTION 6 22
GENDER DIFFERENCES IN HEPATOTOXICITY
Human and Animal-Based Differences in Hepatic Xenobiotic Metabolism and Toxicity Peter J. O’Brien, Katie Chan and Raymond J. Poon
SECTION 7
Hepatotoxicity in Oncology Drug Development Wei Chen, Kenneth Hastings and John K. Leighton
24
The Potent Rat Hepatocarcinogen Methapyrilene: An Hypothesis Regarding its Hepatotoxicology Daniel A. Casciano
25
26
Index
565
577
HEPATOTOXICITY AND BOTANICAL SUPPLEMENTS
Botanical Supplements and Hepatotoxicity Shabana Khan, Ikhlas Khan and Larry Walker
SECTION 9
539
HEPATOTOXICITY AND HEPATOCARCINOGENICITY
23
SECTION 8
531
591
RISK ANALYSIS OF HEPATOTOXINS
Physiologically Based Pharmokinetic Modeling and Risk Assessment of Hepatotoxicants Kannan Krishnan
609
635
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Contributors
Ruben Alvarez-S´anchez F. Hoffman-La Roche Ltd, Non-Clinical Development – Drug Safety, PRBN-D 69/154, 4070 Basel, Switzerland Nora Anderson Medical School of Hannover, Center for Pharmacology and Toxicology, Carl-Neuberg-Strasse 1, D-30625 Hannover, Germany Jurgen ¨ Borlak Fraunhofer Institute of Toxicology and Experimental Medicine, NikolaiFuchs-Strasse 1, D-30625 Hannover, Germany and Medical School of Hannover, Center for Pharmacology and Toxicology, Carl-Neuberg-Strasse 1, D-30625 Hannover, Germany Daniel A. Casciano Department of Pharmacology and Toxicology, University of Arkansas for Medical Sciences, Little Rock, AR 72205, USA and Dan Casciano and Associates, 47 Marcella Drive, Little Rock, Little Rock, AR 72223, USA Rui E. Castro Centro de Patog´enese Molecular, Faculty of Pharmacy, University of Lisbon, Av, das For¸cas Armadas, Lisbon 1600-083, Portugal Katie Chan Graduate Department of Pharmaceutical Sciences, Faculty of Pharmacy, University of Toronto, 144 College Street, Toronto, ON M5S 3M2, Canada Wei Chen Division of Drug Oncology Products, Office of Oncology Drug Products, US Food and Drugs Administration, FDA/CDER/OODP/DDOP, 10903 New Hampshire Avenue, Silver Spring, MD 20993, USA Stacy M. Corthals Center for Environmental Health, Department of Pharmacology and Toxicology, Indiana School of Medicine, 541 Clinical Drive, Indianapolis, IN 46202, USA
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Julio C. Davila Pfizer Inc., PGRD, Saint Louis Laboratories, 700 Chesterfield Parkway North TA1, St Louis, MO 63017, USA Kunio Dobashi Gunma University School of Health Sciences, Gunma University, 3-39-15 Showa-machi, Maebashi, Gunma 371-8511, Japan Tamio Fukushima Worldwide Safety Sciences, Pfizer Global Research and Development, Nagoya Central Research Laboratories, SS-Nagoya, Pfizer Japan Inc., 5–2 Taketoyo, Aichi 470-2393, Japan Christoph Funk F. Hoffman-La Roche Ltd, Non-Clinical Development – Drug Safety, PRBN-D 69/154, 4070 Basel, Switzerland Carol R. Gardner Department of Pharmacology and Toxicology, Rutgers University, 160 Frelinghuysen Road, Piscataway, NJ 08854, USA Kevin Gerrish Laboratory of Molecular Toxicology – Maildrop D2-04, Environmental Stress and Cancer Group, National Institute of Environmental Health Sciences, 111 Alexander Drive, PO Box 12233, Research Triangle Park, NC 27709, USA Angela J. Harris Center for Toxicology and Environmental Health, LLC, 615 W. Markham, Little Rock, AR 72201, USA Kenneth Hastings Office of New Drugs, Center for Drug Evaluation and Research, US Food and Drug Administration, 5600 Fishers Lane, Rockville, MD 20857, USA Philip G. Hewitt Molecular Toxicology, Institute of Toxicology, Merck KGaA, 64271 Darmstadt, Germany Keith A. Hoffmaster Pfizer Inc., Research Training Center (RTC), 01/5R206 ADME Biology, DTC 308, 620 Memorial Drive, Cambridge, MA 02139, USA Ikuo Horii Worldwide Safety Sciences, Pfizer Global Research and Development, Nagoya Central Research Laboratories, SS-Nagoya, Pfizer Japan Inc., 5–2 Taketoyo, Aichi 470–2393, Japan Jens Hrach Molecular Toxicology, Institute of Toxicology, Merck KGaA, 64271 Darmstadt, Germany Shiew Mei Huang Office of Clinical Pharmacology and Biopharmaceutics, Center for Drug Evaluation and Research, Food and Drug Administration, WO21, HFD-850, 10903 New Hampshire Avenue, Silver Spring, MD 20993, USA Yasuhiro Ishihara Faculty of Pharmaceutical Sciences at Kagawa Campus, Tokushima Bunri University, 1314-1, Shido, Sanuki, Kagawa 769-2193, Japan
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Lisa Kamendulis Center for Environmental Health, Department of Pharmacology and Toxicology, Indiana School of Medicine, 541 Clinical Drive, Indianapolis, IN 46202, USA Ikhlas Khan National Center for Natural Products Research, Research Institute of Pharmaceutical Sciences and Department of Pharmacognosy, School of Pharmacy, University of Mississippi, PO Box 1848, University, MS 38677, USA Shabana Khan National Center for Natural Products Research, Research Institute of Pharmaceutical Sciences, School of Pharmacy, University of Mississippi, PO Box 1848, University, MS 38677, USA Tin Oo Khor Department of Pharmaceutics, Ernest Mario School of Pharmacy, Rutgers University, The State University of New Jersey, 160 Frelinghuysen Road, Piscataway, NJ 08854, USA Rie Kikkawa Worldwide Safety Sciences, Pfizer Global Research and Development, Nagoya Central Research Laboratories, SS-Nagoya, Pfizer Japan Inc., 5–2 Taketoyo, Aichi 470-2393, Japan Florian Klammers F. Hoffman-La Roche Ltd, Non-Clinical Development – Drug Safety, PRBN-D 69/154, 4070 Basel, Switzerland James E. Klaunig Center for Environmental Health, Department of Pharmacology and Toxicology, Indiana School of Medicine, 541 Clinical Drive, Indianapolis, IN 46202, USA Ah-Ng Tony Kong Department of Pharmaceutics, Ernest Mario School of Pharmacy, Rutgers University, The State University of New Jersey, 160 Frelinghuysen Road, Piscataway, NJ 08854, USA Kannan Krishnan D´epartement de Sant´e Environmental/Sant´e an Travail, Universit´e de Montr´eal, 2375 Cote Ste Catherine, Bureau 4105, Montr´eal, QC H3T 1A8, Canada Christiane Lamy F. Hoffman-La Roche Ltd, Non-Clinical Development – Drug Safety, PRBN-D 69/154, 4070 Basel, Switzerland Debra L. Laskin Department of Pharmacology and Toxicology, Rutgers University, 160 Frelinghuysen Road, Piscataway, NJ 08854, USA John K. Leighton Division of Drug Oncology Products, Office of Oncology Drug Products, US Food and Drugs Administration, FDA/CDER/OODP/DDOP, 10903 New Hampshire Avenue, Silver Spring, MD 20993, USA Albert P. Li The ADMET Group LLC, In vitro ADMET Laboratories LLC and Advanced Pharmaceutical Sciences Inc., 15235 Shady Grove Road, Suite 303, Rockville, MD 20850, USA
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Wen Lin Department of Pharmaceutics, Ernest Mario School of Pharmacy, Rutgers University, The State University of New Jersey, 160 Frelinghuysen Road, Piscataway, NJ 08854, USA David E. Malarkey National Toxicology Program Pathology Group, Laboratory of Experimental Pathology – Maildrop B3-06, National Institute of Environmental Health Sciences, 111 Alexander Drive, PO Box 12233, Research Triangle Park, NC 27709, USA Rawiwan Maniratanachote Drug Metabolism and Toxicology, Division of Pharmaceutical Sciences, Graduate School of Medical Sciences, Kanazawa University, Kakumamachi, Kanazawa 920-1192, Japan Masatomo Mori Department of Medicine and Molecular Science, Gunma University, 3-39-15 Showa-machi, Maebashi, Gunma 371-8511, Japan Stefan O. Mueller Molecular Toxicology, Institute of Toxicology, Merck KGaA, 64271 Darmstadt, Germany Pablo Muriel Secci´on Externa de Farmacologia, Cinvestav-IPN, Apdo, Postal 14-740, M´exico 07000, DF, M´exico Srikanth C. Nallani Office of Clinical Pharmacology, Center for Drug Evaluation and Research, Food and Drug Administration, WO21, HFD-870, 10903 New Hampshire Avenue, Silver Spring, MD 20993, USA Johannes No´e F. Hoffman-La Roche Ltd, Non-Clinical Development – Drug Safety, PRBN-D 69/154, 4070 Basel, Switzerland Peter J. O’Brien Pfizer Inc., Drug Safety Research and Development, Ramsgate Road, SSEU Europe 380/1.025, IPC 339, Sandwich, CT13 9NJ, UK Peter J. O’Brien Graduate Department of Pharmaceutical Sciences, Faculty of Pharmacy, University of Toronto, 144 College Street, Toronto, ON M5S 3M2, Canada Axel Paehler F. Hoffman-La Roche Ltd, Non-Clinical Development – Drug Safety, PRBN-D 69/154, 4070 Basel, Switzerland Michael Pantze F. Hoffman-La Roche Ltd, Non-Clinical Development – Drug Safety, PRBN-D 69/154, 4070 Basel, Switzerland Binu K. Philip Center for Environmental Health, Department of Pharmacology and Toxicology, Indiana School of Medicine, 541 Clinical Drive, Indianapolis, IN 46202, USA Raymond Poon Environmental and Occupational Toxicology Division, Health Canada, Ottawa, ON K1A 0L2, Canada
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Ren´ee Portmann F. Hoffman-La Roche Ltd, Non-Clinical Development – Drug Safety, PRBN-D 69/154, 4070 Basel, Switzerland Cec´ılia M. P. Rodrigues Centro de Patog´enese Molecular, Faculty of Pharmacy, University of Lisbon, Av, das For¸cas Armadas, Lisbon 1600-083, Portugal Saura C. Sahu Division of Toxicology, Office of Toxicology, Office of Applied Research and Safety Assessment, Center for Food Safety and Applied Nutrition, US Food and Drug Administration, 8301 Muirkirk Road, Laurel, MD 20708, USA Guoxiang Shen Department of Pharmaceutics, Ernest Mario School of Pharmacy, Rutgers University, The State University of New Jersey, 160 Frelinghuysen Road, Piscataway, NJ 08854, USA Norio Shimamoto Faculty of Pharmaceutical Sciences at Kagawa Campus, Tokushima Bunri University, 1314-1, Shido, Sanuki, Kagawa 769-2193, Japan Yasuo Shimizu Department of Medicine and Molecular Science, Gunma University, 3-39-15 Showa-machi, Maebashi, Gunma 371-8511, Japan Susana Sol´a Centro de Patog´enese Molecular, Faculty of Pharmacy, University of Lisbon, Av, das For¸cas Armadas, Lisbon 1600-083, Portugal Clifford J. Steer Departments of Medicine and Genetics, Cell Biology and Development, University of Minnesota Medical School, 6–160 Jackson Hall, 321 Church St. SE, Minneapolis, MN 55455, USA Stephen C. Strom Department of Pathology, University of Pittsburgh, 200 Lothrop Street 450 South BST, Pittsburgh, PA 15261, USA John M. Strong Laboratory of Clinical Pharmacology, Office of Testing and Research, Food and Drug Administration, WO64, 10903 New Hampshire Avenue, Silver Spring, MD 20993, USA Kaori Tomizawa Worldwide Safety Sciences, Pfizer Global Research and Development, Nagoya Central Research Laboratories, SS-Nagoya, Pfizer Japan Inc., 5–2 Taketoyo, Aichi 470-2393, Japan Gregor Tuschl Molecular Toxicology, Institute of Toxicology, Merck KGaA, 64271 Darmstadt, Germany Tetsuro Urushidani Department of Pathophysiology, Faculty of Pharmaceutical Sciences, Doshisha Women’s College of Liberal Arts, Kodo, Kyotanabe, Kyoto 610-0395, Japan and Toxicogenomics Project, National Institute of Biomedical Innovation, Saito-Asagi, Ibaraki, Osaka 567-0085, Japan
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Larry Walker National Center for Natural Products Research, Research Institute of Pharmaceutical Sciences and Department of Pharmacology, School of Pharmacy, University of Mississippi, PO Box 1848, University, MS 38677, USA Jinghai J. Xu Pfizer Inc., Research Training Center (RTC), 01/55564 Systems Biology, DTC 367, 620 Memorial Drive, Cambridge, MA 02139, USA Hiroshi Yamada Worldwide Safety Sciences, Pfizer Global Research and Development, Nagoya Central Research Laboratories, SS-Nagoya, Pfizer Japan Inc., 5 – 2 Taketoyo, Aichi 470-2393, Japan Toshinori Yamamoto Worldwide Safety Sciences, Pfizer Global Research and Development, Nagoya Central Research Laboratories, SS-Nagoya, Pfizer Japan Inc., 5 – 2 Taketoyo, Aichi 470-2393, Japan Tsuyoshi Yokoi Drug Metabolism and Toxicology, Division of Pharmaceutical Sciences, Graduate School of Medical Sciences, Kanazawa University, Kakuma-machi, Kanazawa 920-1192, Japan
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Preface
The main purpose of this book is to assemble up-to-date information on hepatotoxicity in a single edition. There have been several books published during the last three decades on hepatotoxicity and drug-induced liver disease. However, this text will probably be the first to collate most of the genomic technologies, including the latest toxicogenomics. The book is designed primarily for those research scientists currently engaged in the field of liver toxicity. However, it will also be of interest to toxicologists in general and to students and teachers in toxicology in particular. In addition, it will be of interest to industry, as well as to government regulators and risk assessors of foods, drugs, environmental and agricultural products. Saura C. Sahu
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Acknowledgements
I am indebted to my mentors from whom I have learned so much throughout my professional career and who have paved the way for this book. I owe special thanks to the following scientists who have contributed directly or indirectly to this text and I am deeply grateful to each one of them. I thank Dr Thomas A. Cebula for his inspiration, enthusiasm, leadership, endless energy and passion for science while I also thank Dr Joseph E. LeClerc and Dr Richard B. Raybourne for their encouragement and support. In addition, I wish to thank Dr Daniel A. Casciano for his professional advice and contribution and Dr Philip W. Harvey, Covance Laboratories, UK, for his encouragement and support. Finally I thank all of the scientists who have contributed to this book in their own areas of expertise.
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Section 1 Models for Hepatotoxicity Testing
Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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1 Current in vitro Models to Study Drug-Induced Liver Injury Julio C. Davila, Jinghai J. Xu, Keith A. Hoffmaster, Peter J. O’Brien and Stephen C. Strom
1.1
Introduction
In a published report on the root causes of failed drugs over a 10-year period, it was concluded that the pharmaceutical industry as a whole is still facing the same challenge of selecting drug candidates with high efficacy and low toxicity (Schuster et al., 2005). Among all human organ toxicities, hepatotoxicity and cardiovascular toxicity were the two most prominent causes, accounting for two out of three market withdrawals in the last decade (Schuster et al., 2005). In clinical drug development phases, about half of attrition were due to insufficient efficacy and one third were due to toxicity (Schuster et al., 2005). While regulatory animal toxicity testing was able to identify more than 70 % of human toxicities in a retrospective analysis, hepatotoxicity in humans had the poorest correlation with regulatory animal toxicity tests (Olson et al., 2000). In only half of new pharmaceuticals that produced hepatotoxicity in the clinical stage was there any concordant signals in animal toxicity studies (Olson et al., 2000). Obviously, identifying better models to predict human hepatotoxicity is a critical need for the pharmaceutical industry. Drug-induced liver injury (DILI) can be broadly classified into two categories, based on incidence, animal model predictability and dose-dependency. DILI type 1 (DILI-1) is characterized by relatively higher incidence, reproducible in at least one animal species and dose-dependent increase in incidence and severity of the observed injury. Acetaminophen is a classic case of DILI-1 and accounts for nearly half of acute liver failure in the United States (Larson et al., 2005). It can be modeled in more than one strain of rodents (Mehendale, 2005)
Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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Hepatotoxicity
and has a clear dose-dependency in both animals and humans. In humans, the reported ratio of median dose leading to acute liver failure to maximally recommended therapeutic dose is 2 for unintentional overdose to 6 for intentional overdose (Larson et al., 2005). DILI-2, on the other hand, has a relatively lower incidence, occurring at therapeutic dosesfrom 1 in every 1000 patients to 1 in every 100 000 patients (Lee, 2003). It is typically not predicted by the classic animal models of rodents, dogs and monkeys. Because of its rare incidence and large patient-to-patient variability, there is no clear indication of dose-dependency, although within the same susceptible patient, an underlying concentration–response may be present. DILI-2 is also referred to as idiosyncratic hepatotoxicity. The term ‘idiosyncratic’ derives from the Greek meaning ‘mixture of characteristics’ and in the context of drug toxicity refers to the combination of genetic and non-genetic factors that make a patient susceptible to drug injury. Numerous epidemiological studies have identified increasing age (Andrade et al., 2005), females (Ostapowicz et al., 2002) and certain disease-associations (Boelsterli, 2003) as the top demographic factors associated with higher incidence of DILI. While the reasons for such drug idiosyncracy are not entirely clear, most likely there are multiple factors that encompass both toxicokinetic, and toxicodynamic or adaptive reasons. In terms of toxicokinetic reasons, there have been numerous cases where factors such as age, gender, disease states, enzyme induction and inhibition, genetic polymorphism, food and gut micro-organisms have been implicated in causing variability in the pharmacokinetics of drugs. These factors can easily cause several-fold variations in drug concentrations among individuals given the drug at the same dose (Tam, 1993). In addition, the practice of polypharmacy (patients taking multiple drugs) tends to increase with age, which may surpass the predictability of well-controlled clinical trials on drug–drug interaction studies that are typically conducted on a limited number of drug pairs (FDA, 1999). In addition, the presence of disease states, including inflammation and infection, can affect both drug metabolism (Renton, 2004) and drug transport (Fernandez et al., 2004; Hartmann et al., 2002). In terms of adaptive reasons, the human liver has a remarkable ability to regenerate. Indeed, livers that underwent partial hepatectomy can regenerate and patients can go on to live normal lives. In addition, most of idiosyncratic hepatotoxicity was evident only after weeks of drug therapy, not upon a single dose of drug administration. The liver’s adaptive response after the initial drug ‘insult’ is probably crucial to the final manifestation of the full-blown liver injury. These adaptive responses almost certainly involve tissue repair (or lack thereof) (Mehendale, 2005), and the innate and adaptive immune systems (Holt and Ju, 2006). Numerous factors, including genetic polymorphism in tissue repair and immune responses, systemic and/or tissue inflammation, disease states, and continued challenge by polypharmacy, can affect a particular patient’s response to drug-induced liver injury. The mechanisms of such adaptive responses, including signal transduction pathways (Jaeschke and Bajt, 2006; Schwabe and Brenner, 2006), interactions between hepatocytes and cells of the host immune systems (Minagawa et al., 2004) and cellular decision-making (Malhi et al., 2006), will likely be an important and fruitful area of research in the post-genomic era. In vitro cellular models of drug toxicity have unique and important roles to play in order to understand both the mechanisms of initial drug injury and the many signal transduction pathways involved in tissue repair. This present chapter will focus primarily on the use of in vitro models of drug-induced liver injury. The better known mechanisms of drug-induced liver injury have been reviewed in previous publications (Begriche et al.,
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Models to Study Drug-Induced Liver Injury
5
Table 1.1 Major mechanisms of drug-induced liver injury Mechanism
Prototypical drug
Reference
Altered lipid metabolism causing fatty liver
Amineptine, amiodarone, doxycycline, tetracycline, tianeptine, pirprofen Cyclosporine A, estradiol-17 beta-d-glucuronide, taurolithocholate, ethinyl estradiol Flucloxacillin, diclofenac, tienilic acid, sulfamethoxazole, halothane
Letteron et al., 2003, and references therein Crocenzi et al., 2003a,b; Micheline et al., 2002; Roman et al., 2003 Aithal et al., 2004; Carey and van Pelt, 2005; Robin et al., 1996; Sanderson et al., 2006 Amin and Hamza, 2005; Reid et al., 2005
Decreased bile salt clearance causing cholestasis Formation of protein adduct causing immune reaction Increased oxidative stress leading to cellular injury Decreased mitochondrial function leading to apoptosis or necrosis Cytotoxic T cell-mediated cell killing Incomplete or dysregulated tissue repair
Sodium diethyldithiocarbamate, diclofenac, ketoconazole, acetaminophen and drugs that cause steatohepatitis Nimesulide, amiodarone, tamoxifen, stavudine, zidovudine and drugs that cause steatohepatitis Sulfamethoxazole, lidocaine, carbamazepine, lamotrigine, phenindione Acetaminophen, carbon tetrachloride, chloroform, thioacetamide, trichloroethylene, allyl alcohol
Begriche et al., 2006; Tay et al., 2005 Sanderson et al., 2006, and references therein Mehendale, 2005, and references therein
2006; Jaeschke et al., 2002a; Kaplowitz, 2002; Lee, 2003). These are summarized in Table 1.1. A single small-molecule drug may invoke a multitude of such mechanisms (e.g. amiodarone, perhexiline, diclofenac and acetaminophen). This has led some researchers to postulate the ‘multi-hit’ hypothesis of drug hepatotoxicity (Begriche et al., 2006; Letteron et al., 2003; Pirmohamed et al., 2002). Cellular models based on human liver tissues have unique capabilities for studying such DILI mechanisms, as illustrated by the examples shown in Table 1.2. However, before describing the applications of in vitro models, we will first explore the various types of in vitro models that are available to researchers today.
1.2 1.2.1
In Vitro Models to Study DILI Primary Hepatocytes
Currently, the primary hepatocytes system is the in vitro model of choice for studying drug metabolism and hepatotoxicity of new drugs (Castell et al., 2006; Davila et al., 1998;
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Table 1.2 Representative applications of cell-based assays to study drug-induced liver injury Applications
Methodology
Multiparameter cytotoxicity
Combine several different readouts including multi-spectral cytometric analysis Neutral lipid stain (e.g. Oil Red O)
Steatosis Cholestasis Phospholipidosis Reactive metabolite Oxidative stress
Mitochondria damage Identify targets of toxicological importance
Uptake and efflux of taurocholate Phospholipid accumulation in cytoplasm or lysosomal stain GSH adduct formation; GSH depletion (e.g. monochlorobimane) Redox-sensitive dyes
Mitochondria membrane potential dyes (e.g. TMRM) RNAi technology and/or specific inhibitors
Representative reference O’Brien et al., 2006 Amacher and Martin, 1997 Kostrubsky et al., 2006 Gum et al., 2001; O’Brien et al., 2006 Lilius et al., 1996; Thompson et al., 1998) Lautraite et al., 2003; LeBel et al., 1992; Wang and Joseph, 1999 Haskins et al., 2001 Lee and Sinko, 2006; Pichler et al., 2005; Tan et al., 2005; Xu et al., 2005
Gebhardt et al., 2003; Gomez-Lechon et al., 2004; Guillouzo et al., 1977; LeCluyse, 2001; Sinz and Kim, 2006). Primary hepatocytes represent a unique system since they are able to retain, under a refined cultured condition, Phase I and II enzyme activities, as well as their inducibility by xenobiotics. However, primary hepatocytes that fail to preserve and/or re-establish cell polarity and the expression of liver-specific genes are less responsive to drugs and therefore do not accurately reflect the metabolic potential of the liver tissue (Berthiaume et al., 1996; Kocarek et al., 1992; LeCluyse et al., 1996; Tuschl and Mueller, 2006; Waring et al., 2003) It is well established that primary hepatocytes cultured on plastic surfaces and allowed to form an epithelial monolayer (2-dimensional configuration) lose not only up to 75 % of total CYP450 during the first 24-h period, but also liver-specific functions and differentiation processes (Davila and Morris, 1999; Farkas and Tannembaum, 2005; Gomez-Lechon et al., 2004; LeCluyse, 2001); therefore, it is critical that cell culture conditions be refined for studying liver metabolism and toxicity of drugs in humans. Several optimized culture conditions have been reported to maintain liver-specific functions and gene responsiveness (co-factors expression) to levels comparable with those in vivo; these include the use of r extracellular matrix or complex substrata (e.g. collagen and/or Matrigel , sandwich configuration) (Farkas and Tannembaum, 2005; LeCluyse, 2001; Schuetz et al., 1988; Sidhu et al., 1994), chemically defined culture conditions (e.g. the use of low concentrations of insulin and dexamethasone in a serum-free amino acid-rich culture medium) (Enat et al.,
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1984; LeCluyse, 2001; Sidhu et al., 1994) and co-culture hepatocytes with other cell types (e.g. sinusoidal cells, kidney epithelial cells, kupffer cells) (Begue et al., 1984; Donato et al., 1994). r Primary rat hepatocytes cultured and overlaid with Matrigel in an optimized media enables the hepatocytes to retain and maintain longer-term viability and expression of more stable differentiated liver function, such as Phase I and II metabolizing enzymes, and responr siveness to drugs and chemicals (Davila and Morris, 1999; Davila et al., 1998). Matrigel is basically a solubilized basement membrane preparation extracted from the Engelbreth– Holm–Swarm (EHS) mouse sarcoma, a tumor rich in extracellular matrix (ECM) proteins. This complex matrix is mainly composed of laminin, collagen IV, heparan sulfate proteoglycans and entactin, which polymerizes at room temperature to produce biologically active matrix material resembling the mammalian cellular basement membrane. We have previously reported on the development of a modified sandwich configuration model initially described by Dunn (Dunn et al., 1991) and others (Kocarek et al., 1992; LeCluyse, 2001; Schuetz et al., 1988; Sidhu et al., 1994), where freshly isolated hepatocytes are resuspended, r r plated and cultured in a Matrigel –Matrigel system with serum-free media containing insulin and hydrocortisone (10−7 M) (Davila and Morris, 1999). One advantage of using r this model is that the concentration of Matrigel can be easily monitored before and after the initial plating. The concentration of ECM used for culturing is known to affect the morphology and functionality of the hepatocytes. In preliminary studies, we found that rat r hepatocytes, cultured and overlaid with Matrigel at 0.35 mg/ml, acquire a 3-dimensional (3D) configuration and are reorganized as acinar structures; cells become more cuboidal with a distinctive canalicular network and the polarized phenotype and function of normal hepatocytes are well preserved (Davila and Morris, 1999) . Under these culture conditions, the basal levels of a variety of liver genes are maintained and can be induced/stabilized by xenobiotics to levels that are comparable to those achieved in vivo. The mRNA levels of most of the rat CYP450 enzymes (e.g. CYP1A, CYP2B, CYP3A and CYP4A) can be induced 2–4 h after the initial plating. This is an improvement from the traditional methods using primary hepatocytes where cells have been reported to be unresponsive to inducers during the first 24–48 h (Kocarek et al., 1992; Richert et al., 2002; Silva et al., 1998). Freshly isolated rat hepatocytes cultured under this refined culture condition are proven to be a valuable and important in vitro toxicological approach to assess the chemical-induced changes in expression of rat liver CYP450 and Phase II conjugating enzymes (Davila and Morris, 1999; Davila et al., 1998). However, the ultimate species of interest in predicting liver metabolism and toxicity of drugs is human. 1.2.1.1
Refinement of a Primary Human Hepatocyte System for Drug Metabolism and Toxicity
Primary human hepatocytes are a useful model system to potentially predict the metabolism and toxicity of compounds and allow a better understanding of the mechanism of actions of drugs in the human population. Caveats in using primary human hepatocytes are the limited supply of human liver tissues and the ability of such cultures to loose their phenotype after culturing. Despite their limitations, primary human hepatocytes are currently the system of choice to predict human risk before new chemical entities (NCEs) are first tested in Phase I clinical trials. In this section, we describe the refinement of a feasible, reproducible and simple cell culture system that could be used for routine toxicological applications early during drug discovery and development. This model has been specifically refined for assessing the
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metabolism and toxicity of drugs. Validation studies supporting the use of this optimized human hepatocyte model for metabolism-mediated toxicity studies include: (1) microscopy analysis to identify hepatocytes in a 3D configuration with well define intracellular structures characteristic of normal and functional hepatocytes; (2) the measurement of several constitutive and inducible expressions of human Phase I (CYP1A1, CYP1A2, CYP2B6, CYP2C9, CYP2D6, CYP2E1, CYP3A4, CYP4A11 and fatty acyl CoA oxidase, FACO) and Phase II (uridine diphosphate-glucuronosyltransferase, UDPGT2B7; sulfotransferase, SULT2A1; glutathione-S-transferase, GSTYa) biotransformation enzymes at the mRNA and protein and activity level; (3) in vivo and in vitro comparison of relative Phase I enzyme mRNA levels; (4) the differential expression of CYP450 enzymes in human and rat hepatocytes following treatment with various inducing agents. 1.2.1.2
Procurement, Isolation and Culturing of Human Hepatocytes
Human donor livers used in these studies were received from the National Disease Research Interchange (NDRI, PA) within 16 h of patient procurement. Livers were screened at the NDRI for evidence of infectious agents and preserved with ViaSpan (Dupont/Pharma, Wilmington, DE) upon removal from donors. Livers were only accepted from donors with minimal fat content < 10 %, normal liver enzyme levels, low visible fibrosis, medications which would not be considered as potent liver enzyme inducers (particularly, CYP450s) and patients ranging from 14 to 60 years old. Freshly isolated human hepatocytes were obtained from The University of Pittsburgh, Department of Pathology (Stephen Strom’s laboratory) within 24 h after cell isolation and from our laboratory as indicated below. The isolation of human hepatocytes from the whole liver was performed by a three-step perfusion technique using liver sections in situ, as described by Strom (Strom et al., 1996). The culturing of freshly isolated human hepatocytes was then processed as indicated for rat hepatocytes (Davila and Morris, 1999) with some modification. Briefly, after isolation, liver cells were filtered through a sterile nylon mesh (100 μm) and collected in several 50-ml conical test tubes. Cells were centrifuged twice at 30 × g for 2 min and then the final pellet was resuspended with 30 ml Williams E solution containing hydrocortisone and insulin (10−7 M) and glutamine (4 mM). Ten ml of diluted Percoll (30 vol %) were placed in the bottom of each tube and centrifuged at 1100 × g for 5 min. The pellets were washed with Williams E at 30 × g for 2 min and then resuspended in 20 ml Williams E solution. Viability of the isolated hepatocytes used in these studies was typically greater than 90 %. r The final cell suspension was resuspended in diluted Matrigel solution (0.35 mg/ml) to 5 a final cell density of 4 × 10 cells/ml and plated in 12 (1 ml) or 6 (2 ml) well plates, as previously indicated for rat hepatocytes (Davila and Morris, 1999). 1.2.1.3
Cell Culture Treatment
Hepatocytes were treated for two consecutive days at 24 h and 48 h after initial plating with optimal concentrations of phenobarbital, PB (100 μM), rifampicin, RIF (10 μM), 3-methylcholanthrene, 3MC (1 μM), clofibrate, CLO (50 μM), tamoxifen (TAM, 10 μM), omeprazole (OMZ, 20 μM), pregnenolone-16α-carbonitrile (PCN, 1 μM) and hydrocortisone (HC, 20 μM). Compounds were dissolved in dimethyl sulfoxide (DMSO) and then diluted with Williams E prior to administration to the cultures (final DMSO concentration was 0.1 vol %). The concentrations of the inducers used in these studies were selected
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based on preliminary experiments to determine the concentration of inducer producing the highest levels of enzyme expression in the absence of toxicity (data not shown). When the enzyme was not inducible, a maximum non-toxic concentration was used (50–100 μM). The concentrations of inducers used in this study were not toxic to the hepatocytes as indicated by morphological examination, MTT reduction assay and recovery of total RNA (data not shown). 1.2.1.4
PCR and Fluorogenic cDNA Probes
Messenger RNA was determined using real-time PCR. Primer sets specific for human cyclophylin (CYC, house-keeping gene), CYP450 1A1, 1A2, 2B6, 2C9, 2D6, 2E1, 3A4, 4A11, FACO, UDPGT2B7, GSTYa and SULT2A1, and TaqMan fluorogenic probes specific for each primer set are described in Tables 1.3 and 1.4, respectively. The TaqMan fluorogenic probes were designed in accordance to the guidelines issued by Perkin-Elmer. The cDNA samples were analyzed using an ABI Prism 7700 (ABI) sequence detection system (Perkin Elmer Biosynthesis, Foster City, CA). The results generated by the ABI system are expressed as cycle threshold (Ct), representing the PCR cycle at which an increase in TaqMan probes fluorescence is detected (specific signal) above the baseline signal for each amplicon (Table 1.5, A–F). The target gene Ct (Table 1.5, C) for a sample is then normalized by subtracting the target gene Ct from its specific cyclophilin Ct (Table 1.5, B), therefore adjusting all individual samples to the expression of a common housekeeping gene (Ct, Table 1.5, D). The resulting value for each sample is then further expressed as the difference in Ct for the sample compared to the difference in Ct for the vehicle control (Ct, Table 1.5, E) with the lowest Ct value across experiments. Ct provides a value that is equal to how many cycles each sample is from the experimental control. The final calculation yields a relative quatitation of mRNA expression for each sample by expressing the results as fold induction above the experimental level control. All studies were repeated a minimum of five times, unless otherwise indicated, and representative data are presented. 1.2.1.5
Statistical Analysis
Data are presented as ± SE of the mean and statistically analyzed, using Dunnett’s T-test (P < 0.05). Each determination was performed in triplicate (n = 3 wells) and the data presented are from one of five separate experiments, unless otherwise indicated, giving qualitatively similar results. 1.2.1.6
Results and Discussion
Microscopy analysis. Phase contrast and electron micrographs of three-day old primary r human hepatocytes cultured with Matrigel in a well defined serum-free culture medium r are orare presented in Figure 1.1. Primary human hepatocytes cultured with Matrigel ganized as acinar structures and acquire a 3-dimensional (3D) configuration, cells retain cell polarity and phenotype (Figure 1.1(a)) and viability for several weeks. This technique, where freshly isolated hepatocytes are suspended and plated in a culture media containing r diluted Matrigel (0.35 mg/ml) and then overlaid with a second layer of Matrigel, allows the development of a 3D configuration. Cells cultured in this fashion reorganize into structures that are characteristic of the tissue of origin, adopting an in vivo-like morphology (Berthiaume et al., 1996; Davila and Morris, 1999; Farkas and Tannembaum, 2005;
5 Sense primer (5 to 3 ) CTTgTCCATggCAAATgCTg gATgAgAACgCCAATgTCCAg ACAACgCTgAATggCTTC CACTCATCAgCTCTgTATTCg TgCTATgAgACTTgAgAgg TATggCCTTCACCACAACC ATgAAgCAACCCgAgACACCA AATggACATgAACAACCCTCA CgTgACATCgAAgTACAgg gTTCgAgCAAgTgAggCAC gCATCTACgAggCAATCTACCA gAgATTgATgggATgAAgC CTgAgTTATgAggAgCTg
Cyclophilin (CYC), #Y00052 CYP1A1, #K03191 CYP1A2, #NM 000761 CYP2B6, #X13494 CYP3A4, #D11131, #M18907 CYP4A11, #NM 000778 CYP2E1, #J02625 CYP2C9, #M61855 CYP2D6, #M33388 Fatty acyl-coA oxidase (FACO), #U07866 UDP-glucuronosyltransferase (UDPGT-PB), #J05428 Glutathione-S-transferase (GST)-Ya, # M14777 Hydroxysteroid sulfotransferase (ST), #X84816
179 175 266 355 236 243 234 158 261 294 343 268 196
Fragment size (bp)
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gTgATCTTCTTgCTggTCTTgC CTgCCAATCACTgTgTCTAgC gCTgAACTCCAgTTgCTgT gTAgACTCTCTCTgCAACATgAg gCAAACCTCATgCCAATgCAg TCAACACAAgTCgTgCAATgg AACAACTCCATgCgAgCCAg CTCAgggTTgTgCTTgTCgT gAgAAgCTgAAgTgCTgCAgC CAAgCACAgAgCCAAgTgTC CACATCCAAAgAgTggTACTg AggTAgTCTTgTCCATggCTC CTCAgAAgTTgTgCTTTgTCC
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Table 1.4 Fluorogenic probe sequences for human CYC, CYP450s, FACO, UDPGT, GST and ST. TaqMan probes were designed in accordance to the guidelines issued by Perkin-Elmer Target gene
Fluorogenic probe (TaqMan)
Cyclophilin (CYC) CYP1A1 CYP1A2 CYP2B6 CYP3A4 CYP4A11 CYP2E1 CYP2C9 CYP2D6 Fatty acyl-coA oxidase (FACO) UDP-glucuronosyltransferase (UDPGT-PB) Glutathione-S-transferase (GST)-Ya Hydroxysteroid sulfotransferase (ST)
CCACAATATTCATgCCTTCTTTCAC CTTggATCTTTCTCTgTACCCTgg CAgCATCATCTTCTCACTCAAgg CCAAggACCTCATCgACACCTACCT CCAAgCTATgCTCTTCACCgTgAC TCTgCTCAACACAgCCACgCTTTC ACAgTCgTAgTgCCAACTCTggACTC ACAAgTCAACTgCAgTgTTTTCCAAgCT CACTCATCACCAACCTgTCATCggT CAACCAAAgCAACAgCATCTgAgC CAATCCAgAAgACTgCTCgATCCA CAgACCAgAgCCATTCTCAACTAC CTCAAgAACAgCTCCTTTCAgAgC
LeCluyse et al., 2005) As indicated by electron microscopy (Figure 1.1(b)), these cells exhibit distinctive structural characteristics of in vivo hepatocytes, including numerous mitochondria, bile canaliculi, nuclei, glycogen, gap junctions and a rich smooth and rough endoplasmic reticulum. This suggests the presence in this cell system of the machinery needed for the cells to express a wide array of liver-specific functions
(A)
(B)
n
er
g bc
m
gj
Figure 1.1 Phase contrast (A, ×320) and electron micrographs (B, × 1200) of human hepaR tocytes plated and overlaid with Matrigel for three days. Freshly isolated human hepatocytes suspended, plated and cultured with Matrigel at (0.35 mg/ml) are reorganized as acinar strucutures and acquired a 3D configuration. The electron micrograph shows a well-defined nucleus (n), endoplasmic reticulum (er), mitochondria (m), bile canaliculi (bc), glycogen droplets (g) and gap junctions (gj)
18.2 17.2
17.5
Control Sample A
Sample B
26.83
32.45 28.45
2−Ct (Control) = 1.00 2−Ct (Sample A) = 8.00 2−Ct (Sample B) = 30.27
Ct Control − Ct Control = 0.00 Ct Sample A − Ct Control = −3.00 Ct Sample B − Ct Control = −4.92
(Control)C − B = 14.25 (Sample A)C − B = 11.25 (Sample B)C − B = 9.33
F Relative quantitation
E Ct
D Ct
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Measurement of several Phase I and II biotransformation enzyme expressions in primary human hepatocytes. In this study, we found that both the constitutive and inducible expression of the major families and subfamilies of human mRNA CYP450 (CYP1A1, CYP1A2, CYP2B6, CYP2C9, CYP2D6, CYP3A4, CYP2E1, CYP4A11) and Phase II (UDPGT2B7, SULT2A1, GSTYa) enzymes, as well as the rate of protein synthesis, are maintained in culture for several days. The rank ordering for the average basal mRNA expression levels of human CYP450 and Phase II isoforms expressed in this primary cell culture system from the lowest to the highest was as follows: CYP4A11 < CYP2B6 < CYP1A1 < SULT2A1 < CYP3A4 \ < CYP2D6 < CYP1A2 < CYP2E1 < GSTYa < CYP2C9 < UDPGT2B7 < FACO. The expression of Phase I and II enzymes at the mRNA, protein and activity levels (e.g. CYP1A and CYP3A) in three-day old cell cultures are reported in Figures 1.2, 1.3 and 1.4, respectively. In agreement with other investigators (Bowen et al., 2000; LeCluyse, 2001; Strom et al., 2001), we found that CYP1A1, CYP1A2, CYP2B6 and CYP3A4 are inducible by 3MC, and PB and RIF, respectively, in human hepatocytes (Figures 1.2 and 1.3). Interestingly, the basal r r levels of CYP3A4 and CYP2B6 in human hepatocytes cultured by a Matrigel –Matrigel system did not disappear within the 48 h in culture as previously reported by others (Kocarek et al., 1992; Richert et al., 2006). One explanation would be the difference of the substrata and the culture media components used in these studies. However, in agreement with others (Donato et al., 1995; Silva et al., 1998), some variability in the induction of CYP3A and CYP2B among the donors was observed. Seven out of ten liver donors responded to RIF at 10 μM, while two out of ten cell cultures responded to PB at 100 μM. Furthermore, we have also found that RIF induces CYP1A1 mRNA in all of the human hepatocytes tested. CYP2C9, CYP2D6 and CYP2E1 were found to be unaffected by the inducing agents used in this study. This lack of responsiveness of these enzymes has been previously documented and may be explained by genetic polymorphism, differences in substrate specificity or the high constitutive levels of the enzyme (Elkins and Wrighton, 1999; Guengerich, 1997; Mimura et al., 1993; Waxman et al., 1991). In addition, the disruption of circulating levels of growth hormone patterns in vitro may have altered the constitutive levels and inducibility of some enzymes, e.g. of CYP2D6 and CYP2C9 (Gonzalez, 1996; Neuman et al., 1993; Prough et al., 1996; Waxman et al., 1991). These secondary contributors to enzyme induction were not studied as part of this investigation. Moreover, we found that CLO was not a good inducer of CYP4A11 and FACO; however, we have found that WY14,643 and cyprofibrate at 20 μM are good inducers of CYP4A11, 20- and 7-fold higher than controls, respectively (data not shown). To further characterize this in vitro human hepatocyte system, we studied the effects of liver enzyme inducers on the expression of several Phase II enzymes, UDPGT2B7, GSTYa, and SULT2A1 at the mRNA level (Figure 1.2(g–l)). In contrast to the expression of CYP450 enzymes, Phase II enzyme expression and activity have been scarcely studied in human liver samples. One reason is the lack of specific substrate probes for the various isoenzymes and the lack of sensitive analytical methodology (Burchell and Coughtrie, 1989; Eaton and Bammler, 1999). However, RT–PCR technology allows us to assess the ability of a drug to modulate hepatic Phase II gene expression, thus offering a specific and rapid alternative to quantification of gene induction by immunodetection or substrate metabolism. UDPGT2B7, GSTYa and SULT2A1 were found to be expressed at a high basal
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Relative mRNA expression (arbitrary units)
Human CYP1A1
Human CYP 1A2 mRNA
(a) 120
120
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*
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RIF
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60 30
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Human CYP4A11 mRNA (g)
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Human FACO mRNA
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RIF
Human ST2A1 mRNA
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(k) 40
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0 CON 3MC
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Human CYP2E1 mRNA
0 CON 3MC
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CLO
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*
* RIF
Human CYP2D6 mRNA
Human CYP2C9 mRNA
50 40
30
* PB
0 CON 3MC
*
(d)
*
30
*
0
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(c)
(b)
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Human CYP3A4 mRNA
Relative mRNA expression (arbitrary units)
Human CYP2B6 mRNA 150
150
150
CON 3MC
Relative mRNA expression (arbitrary units)
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Human GSTYa mRNA (l)
*
0 CLO
CON 3MC
PB
RIF
CLO
Figure 1.2 Analysis of mRNA (TaqMan) levels for the analysis of CY450 expression in human hepatocytes. Data represent the results obtained from the analysis of Phase I and II biotransforR mation enzyme expression in human hepatocytes cultured with diluted Matrigel following exposure to prototypical CYP450 inducing agents (3MC, 1 μM; PB, 100 μM; RIF, 10 μM; CLO, 50 μM). The effects of CYP450 enzyme inducers on the expression of CYP1A1 (a) and CYP1A2 (b), CYP2B6 (c), CYP3A4 (d), CYP2C9 (e), CYP2D6 (f), CYP4A11 (g), FACO (h) CYP2E1 (i), UDPGT2B7 (j), ST2A1 (k) and GST-Ya (l) are presented. Data presented are expressed as the mean (± SE) relative mRNA expression levels for each enzyme (fold induction). Data are from one of five separated experiments (N = 3 dishes) using five different donors giving qualitatively similar results
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3MC
PB
15
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RIF
CYP1A1/A2
CYP2B6 CYP3A4 CYP4A11 CYP2E1
Figure 1.3 Analysis of CYP450 apoprotein expression in human hepatocytes cultured in the R presence of diluted Matrigel using Western immunoblotting. Microsomes were prepared from cells exposed to either vehicle (CON, DMEM/0.1 % DMSO) or prototypical CYP450 enzyme inducers (3MC, 1 μM; PB, 100 μM; RIF, 10 μM; CLO, 50 μM) for 48 h. Cells were lyzed and microsomal proteins analyzed for expression of CYP1A1/CYP1A2, CYP2B6, CYP3A4, CYP4A11 and CYP2E1 protein by Western immunoblotting. Data are from one of three separated experiments using three different donors giving qualitatively similar results
(a) 28
CYP1A activity
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CYP3A activity
0.8
0.6
0.4
0.2
0
0 Control
3MC 10 µM
Control
RIF 10 µM
Figure 1.4 Analysis of CYP1A (a) and CYP3A (b) activity levels by HPLC and fluorometric R analysis in primary human hepatocytes cultured with Matrigel after treatment with RIF and 3MC, respectively
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level in this culture system but not markedly induced by chemical treatment. We found that UDPGT2B7 was inducible by 3MC, PB and RIF, and that SULT2A1 was inducible by 3MC and RIF, and that GSTYa was inducible by only 3MC (Figure 1.2(g–l)). 3MC and PB are known inducing agents of Phase II enzymes in the liver (Eaton and Bammler, 1999; Wormhoudt et al., 1999). Data presented here support the potential utility of this in vitro system and RT-PCR for assessing the induction of Phase II conjugating enzymes in human hepatocytes by xenobiotics. In vivo and in vitro comparison of Phase I and II enzyme mRNA levels. As part of the validation study, in vivo and in vitro comparison of Phase I and II enzyme mRNA levels were performed. In these studies, mRNA levels were measured from one hundred human liver tissues and from fourteen separated human hepatocyte cultures (3 day old cells) using real-time PCR. The mean of the target genes was calculated and reported as Ct values (see Table 1.5, D). As indicated in Table 1.6, the mRNA basal levels of CYP1A1, CYP1A2, CYP2C9, CYP2D6, CYP2E1, GSTYa and UDPGT2B7 enzymes in primary hepatocytes were highly maintained for three days and comparable to those of liver tissues. However, CYP2B6, CYP3A4, CYP4A11, FACO and ST2A1 mRNA basal levels were found to be decreased but returned back to levels comparable to liver tissues when CYP450 inducers such as rifampicin (RIF, 10 μM), phenobarbital (PB 100 μM) and WY-14,643 (20 μM) were administrated to the cell cultures. The establishment and validation of a cell culture system which mimics drug biotransformation in vivo is particularly important when considering the elimination of NCEs that generate toxic or reactive intermediate metabolites early in the development process. Differential expression of CYP3A by RIF, OMZ, TAM, PCN and HC in rat and human hepatocytes. The last set of validation studies were designed to examine the differential expression of CYP450 enzymes (CYP1A, CYP2B, CYP3A, and CYP4A) in human and Table 1.6 In vivo and in vitro comparison of relative CYP450 enzyme (mRNA) levels in human liver (data are presented as Ct values) Enzyme phase I–II 1A1 1A2 2B6 2C9 2D6 2E1 3A4 4A11 FACO ST2A1 GSTYa UDPGT2B7
-Ct in vivo
-Ct in vitro
8.9 4.5 6.4 0.9 2.9 −2.3 0.9 4.2 1 1 1.8 −0.2
8.5 2.5 8.7 1.0 3.4 2.0 5.2 10 4.6 4 1.1 0.11
-Ct in vitro induced cells — — 6.2 (Phenobarbital) — — — −0.4 (rifampicin) 3.8 (WY-14,643) 1.7 (WY-14,643) 3 (Phenobarbital)
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Figure 1.5 Comparison of CYP3A induction by rifampicin (RIF, 10 μM), omeprazole (OMZ, 20 μM), tamoxifen (TAM, 10 μM), pregnenolone-16α-carbonitrile (PCN, 1 μM) and hydrocortisone (HC, 20 μM) in rat (a) and human (b) hepatocytes using RT-PCR and fluorogenic cDNA probes (TaqMan). Data presented are expressed as the mean (± SE) relative mRNA expression levels for the enzyme (fold induction). Data are from one of three separate experiments using three different donors giving qualitatively similar results
rat hepatocytes. Results from our studies demonstrate a clear species difference in the induction of these enzymes in rat and human hepatocytes. In particular, CYP3A expression was found to be differentially induced by a number of known CYP3A inducers (Figure 1.5). RIF and TAM were found to markedly induce CYP3A expression in human hepatocytes, whereas OMZ, PC and HC were found to induce CYP3A expression in rat hepatocytes but not in human hepatocytes. These differences in expression of CYP3A, CYP2B and CYP4A enzymes following exposure to xenobiotics have been attributed in part to the
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ability of xenobiotics to differentially activate several members of the nuclear hormone receptor (NHR) superfamily such as PXR/SXR, CAR and PPAR, respectively (Giguere, 1999; Peraza et al., 2006; Roberts, 1999; Schuetz et al., 2000; Xie et al., 2000). The quantity or the quality of these nuclear hormone receptors and the presence of a truncated, inactive form of the receptor, such as in the case of PPARα (Gonzalez and Lee, 1996; Peraza et al., 2006; Peters, 2005), have been suggested as potential factors responsible for such species differences in responses to chemicals. In addition, it has been reported that the species origin of the receptor, rather than the promoter structure of the gene, e.g. CYP3A, dictates the species-specific pattern of CYP3A inducibility (Luo et al., 2004; Xie et al., 2000). The ability of these primary rat and human hepatocytes to respond to a variety of chemicals and to detect differences in CYP450s induction suggests the presence in this cell system of multiple regulatory pathways and of transcription factors associated with functional receptors (Li and Chiang, 2006). Variations in drug metabolism by different species represent a challenge to early metabolism and safety screening. Differential drug-induced expression of liver cytochrome P450 (CYP450) enzymes has been reported to be responsible in part for variations in metabolism and toxicity among species (Hengstler et al., 1999; Kocarek et al., 1995; Lewis et al., 1998). These striking differences in drug-induced CYP450 expression among animal species make it difficult to rely solely on the use of laboratory animals as surrogate species for predicting drug metabolism and toxicity in humans. Therefore, human cellbased systems are important tools in assessing and predicting human drug metabolism and toxicity. r Primary human hepatocytes cultured in a refined cultured condition using Matrigel provides an attractive screening strategy for testing the qualitative and quantitative induction of human CYP450 enzymes by NCEs and to extend our understanding of the role of these NHRs in regulating CYP450 expression by xenobiotics. Results from these studies also highlight the importance of assessing enzyme induction across species as it relates to the potential for drug–drug interactions and metabolism-mediated toxicity. In conclusion, r r we have established a 3D-human cell culture system using a Matrigel –Matrigel configuration and a quantitative RT-PCR method using fluorogenic cDNA technology (TaqMan) to examine the effects of xenobiotics on the expression of a number of human CYP450 and Phase II conjugating enzymes. We have demonstrated that several Phase I and II liver biotransformation enzymes can be induced by xenobiotics in this system. In addition, we have further demonstrated the need for using human cell culture induction systems to assess the metabolism and toxicity of xenobiotics in the human population. 1.2.2
Stem Cell-Derived Hepatocytes
New platforms for discovery, target validation and predictive toxicity screening in therapeutic areas such as arthritis and inflammation and cardiovascular are needed for a positive identification of new therapeutic agents in preclinical development. Included in the testing scheme is the evaluation of compounds or recombinant gene products on in vitro cell systems, such as primary cell cultures, which best reflect the tissue of origin. While primary hepatocytes represent the desired model system for drug metabolism and toxicity, the limited supply of human tissues and the variable quality and interindividual differences hinder its routine use.
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Stem cell technology provides unprecedented opportunities not only for investigating new ways to prevent and treat a vast arrays of diseases but also for changing the way we identify new molecular targets, discover and develop new drugs, as well as test them for safety. Because stem cells are self-renewing population of cells, they can be continuously cultured in an undifferentiated state and give rise to more specialized cells of the human body, such as heart, liver, bone marrow, blood vessels, pancreatic islet and nerve cells. Therefore, it offers an important new tool to develop unique in vitro model systems for testing drugs and chemicals and potentially predict or anticipate toxicity in humans (Davila et al., 2004; Sinz and Kim, 2006). Recent advances in the isolation and culture expansion of multi-potent stem cells from embryonic or adult tissue offer these cell systems as alternative sources of progenitor cells for wide number of tissue-specific cell types. Although human embryonic stem cells (HESCs) retain the potential to differentiate into all of the major cell types of the body, controversy surrounding their research use has limited their application to date. However, adult stem cells, which can be isolated from non-human embryonic tissue, retain their multipotency in culture, and therefore offer a useful cell culture system for the evaluation of compounds during drug discovery and development. Significant and continuing advances in stem cell research to generate an reliable supply of fully functional human hepatocytes has the potential to provide a consistent source of normal and functional human cells that would more closely predict the impact of a new drug on human livers in the body. The ability to obtain an unlimited supply of hepatocytes that retain the expression and activity of drug metabolism enzymes would revolutionize toxicity testing, address the largest bottleneck in new drug research and accelerate the drug development process. While the biological qualities of an optimal hepatocyte source could be debated, there are at least four properties that would seem essential to satisfy most of the requirements of stem cell-derived hepatocytes. The stem cell-derived hepatocyte must: (1) (2) (3) (4)
be routinely available; be easily cryopreserved and restored to culture; provide cells with a metabolic profile observed in adult hepatocytes; provide for a diverse genetic background of donors.
Although there are reports of stem cell-derived hepatocyte-like cells, it is safe to say that there are currently no published reports of cells that fulfill all of the above mentioned requirements. The properties of the ideal stem cell must be kept in mind, as certain cell types such as bone marrow, peripheral blood or amnion, would be expected to be routinely available and in sufficient numbers to provide for a wide range of genetic backgrounds; cell types such as embryonic stem cells (ES) may only be available from a far more limited number of individuals. These considerations could be critically important to eventual drug metabolism and toxicology studies with hepatocytes derived from these stem cell sources. The number of published reports concerning the differentiation of specific cell types into hepatocyte-like cells is too numerous to cite in the context of this limited review. The sources of stem cells reported to differentiate along a hepatic lineage is considerably smaller. By far the most common reports of non-hepatic cells becoming hepatocytes are from the bone marrow, peripheral or placental cord blood or embryonic stem cells of mouse or human origin.
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1.2.2.1
Bone Marrow-Derived Hepatocyte-Like Cells
Petersen et al. (Petersen et al., 1999) were the first to suggest that bone marrow cells could differentiate to hepatocytes. Because putative hepatic stem or progenitor cells express surface marker antigens such as Thy-1 or c-kit which are also found on hematopoietic cells, it was logical to assume that they might have a common source. This initial report was soon followed by several others confirming and extending the initial observations demonstrating in vitro and/or in vivo differentiation of bone marrow-derived cells to hepatocytes (Alison et al., 2000; Lagasse et al., 2000; Krause et al., 2001). The initial exuberance was soon followed by stunning reports that most of the hepatic characteristics expressed in transplanted cells were the result of cell fusion with recipient hepatocytes (Wang et al., 2003; Vassilopoulos et al., 2003). Conflicting reports still remain in the literature. A consensus position is that there may be some low level differentiation of bone marrow-derived hematopoietic cells to hepatocytes although it is most likely quite limited (Wagers et al., 2002; Cantz et al., 2004; Yamaguchi et al., 2006). In vitro differentiation of bone marrow cells to hepatocytes would seem to be devoid of the ‘fusion artifact’ and might prove a useful method to derive hepatocyte-like cells from bone marrow. There are now several reports of the generation of hepatocyte-like cells from bone marrow cells propagated with growth factors such as hepatocyte growth factor (HGF) and/or basic fibroblast growth factors (bFGF) (Avital et al., 2001, 2002; Fiegel et al., 2003; Miyazaki et al., 2002, 2004; Kawasaki et al., 2005; Saji et al., 2004). While several hepatic characteristics such as the expression of albumin, alpha-1-antitrypsin, tyrosine aminotransferase, and urea production, were cited as evidence of hepatic differentiation, there are no reports of the production of cells with a complete set of adult hepatocyte characteristics. These initial results are encouraging, and if full hepatic differentiation could be completed from hematopoietic cells in vitro, they could be an extremely useful future source of hepatocytes. It is interesting that the cell type thought to be responsible for fusion with recipient hepatocytes following bone marrow cell transplants are the myelomonocytic cells in the bone marrow (Willenbring et al., 2004). Investigators have tried to induce hepatic differentiation in cultured monocytes (Ruhnke et al., 2005a,b). These authors report albumin secretion and urea production at rates lower than, but similar to, human hepatocytes and ethoxycoumarin-O-deethylase activity which was inducible with 3-methylcholanthrene and low but measurable CYP3A4 mediated drug metabolism. Tissue such as bone marrow is complex and contains many cell types. In addition to the hematopoietic cells, there are other cell types in bone marrow. In general, hematopoietic cells from bone show limited proliferation in culture and grow as a non-adherent cell type. A second cell type found in bone marrow grows as adherent cells and shows much greater potential to proliferate. This adherent fraction can be isolated from bone marrow, umbilical cord blood and even peripheral blood. Although not necessarily the same cell type, the adherent fraction is alternately referred to as stromal cells, fibroblasts or mesenchymal stem cells (MSCs). There are several reports of the differentiation of mesenchymal cells to hepatocyte-like cells, both in vitro and in vivo (Beerheide et al., 2002; Kakinuma et al., 2003; Kogler et al., 2004; Newsome et al., 2003; Schwartz et al., 2002; Nonome et al., 2005). Markers identified were mainly markers of early hepatic differentiation. In a careful study of MSC-derived hepatocyte-like cells, Ott and coworkers concluded that although the cells expressed some hepatic markers following transplantation into the liver, the MSC-derived
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cells did not express several crucial hepatic markers such as cytokeratin expression (Sharma et al., 2005). The lack of cytokeratin expression would suggest that the cells remained mesenchymal and did not fully differentiate into an epithelial cell type such as a parenchymal hepatocyte. However, more recent publications keep the idea of mesenchymal stem cellderived hepatocytes alive (Aurich et al., 2006). More information on the expression mature liver genes such as the CYP450 genes and adult hepatocyte metabolic activities are needed to determine the actual state of differentiation of these hepatocyte-like cells (Hengstler et al., 2005; Teramoto et al., 2005) 1.2.2.2
Embryonic Stem Cell-Derived Hepatocytes
It might seem that the derivation of hepatocytes from embryonic stem (ES) cells might be the major focus of researchers trying to generate hepatocytes from stem cells. However, this is not the case. More publications report hepatic differentiation of stem cells from sources other then ES cells. There is good preliminary data to suggest that hepatocyte-like cells can be derived from ES cells (reviewed in Teramoto et al., 2005, and Lavon and Benvenisty, 2005). While most reports concern mouse ES cells, there are reports of human ES cell-derived hepatocyte-like cells (Lavon et al., 2004; Rambhatla et al., 2003; Shirahashi et al., 2004; Soto-Gutierrez et al., 2006). Each of the reports show the expression of early markers of hepatic differentiation as well as some more differentiated functions such as some limited drug ammonia metabolism or urea production. There are no reports of robust drug metabolism by any ES-derived hepatocyte-like cells. While quite preliminary, these initial results suggest that at least some of the ES cells are able to differentiate along a hepatic lineage. It is not entirely clear from the published work if differentiation is being directed by the experimental design or if spontaneous differentiation of ES cells is occurring in the cultures. More work will be needed to fully characterize the ES-derived hepatocytes and to optimize and direct the differentiation process. Results reported with pluripotent cells like ESs is complicated by the possibility of differentiation along other pathways. Hepatic marker genes such as albumin, alpha fetoprotein and transthyretin are not only expressed in developing liver, they are also expressed in yolk sac. Yolk sac performs many liver-like functions during fetal life. Thus, it is difficult to determine if the expression of hepatic marker genes is evidence of hepatic differentiation or simply differentiation to visceral endoderm of the yolk sac. A recent report by Asahina et al. (Asahina et al., 2004) suggests that the expression of CYP enzymes such as CYP7A1 was the only definitive marker of actual hepatic differentiation and could distinguish between differentiation of ES cells to visceral endoderm (yolk sac) and the definitive endoderm from which the liver and pancreas eventually are derived. By this measure, the expression of CYP7A1 in stem cell-derived hepatocyte-like cells indicated authentic hepatic differentiation. 1.2.2.3
Hepatocyte-Like Cells Derived from other Sources
There are a number of reports to suggest that hepatocyte-like cells can be derived from other tissue sources. Hepatocyte-like cells were generated from pancreatic tumor cells following long-term exposure to dexamethasone (Tosh et al., 2002). A number of liver-enriched genes we detected included glucose-6 phosphatase, carbamoylphosphate synthetase (CPS I) and glutamine synthetase (GS), as well as some CYP450 and Phase II enzymes. If high levels of mature liver enzymes could be induced, the transdifferentiation of other cell types into
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hepatocytes might prove to be a useful cell source. Miki et al. (Miki et al., 2005) reported that cells with characteristics of pluripotent stem cells could be identified and isolated from human amnion. The amnion-derived stem cells could be induced to differentiate into cell types from all three germ layers, including cells with hepatocyte characteristics. The expression of many CYP450 genes and metabolic activity consistent with the expression and induction of CYP1A family proteins was reported (Davila et al., 2004). A significant observation was that CYP7A1 was expressed in amnion-derived hepatocyte-like cells, suggesting that the culture conditions used in the experiments induced differentiation of the stem cells to definitive endoderm and along an authentic hepatic lineage. 1.2.2.4
Hepatic Stem or Progenitor Cells
It is far beyond the scope of this discussion to thoroughly review the topic of hepatic stem cells. Recent reviews have updated this area (Forbes et al., 2002; Walkup and Gerber, 2006). Recent reports suggest the identification and isolation of putative hepatic stem or progenitor cells (Dan et al., 2006; Nowak et al., 2005; Schmelzer et al., 2006). The expression of some mature liver functions and the ability to expand the cells in culture suggest that the hepatic stem or progenitor cells identified in these studies could be a useful source of human hepatocytes (Dan et al., 2006; Nowak et al., 2005). 1.2.2.5
Summary and Suggestions for Future Studies
In here are a number of sources of cells which all give rise to cells with characteristics of hepatocytes. However, to date none of the stem cell sources, including those derived from the liver, have been shown to produce cells with full mature liver functions. Thus, additional effort is warranted with each stem cell type with the goal to improve hepatic differentiation, in vitro. A number of hepatic genes have been used in the studies cited above as markers of hepatic differentiation. In most cases, the reports identified markers of early liver development, such as the expression albumin and alpha 1-antitrypsin, cytokeratins or human hepatocyte antigen. While these markers are useful to show differentiation along a hepatic lineage, the hepatocyte-like cells acquire the expression of these gene products early in the differentiation process. The expression of later markers of mature liver function, such as CYP450 gene expression and metabolic activity, are rarely reported. In addition, in most studies when mature liver genes were reported they were merely detected by qualitative methods such as immunohistochemistry or RT-PCR. For the field to move forward it will be necessary to get quantitative estimates of the expression of a number of mature liver functions in the stem cell-derived hepatocytes which can be compared to authentic adult human hepatocytes. Although not hepatocyte-specific, the CYP450 genes are expressed at higher levels in liver than most other tissues. In addition, because the CYP450 gene products are critically involved in drug metabolism and toxicology, the expression of these genes will be central to the utilization of the stem cell-derived hepatocytes for basic research. The important drug metabolizing genes of the cytochrome P450 (CYP) families are expressed relatively late in hepatic development and, as such, we propose that they are useful markers of hepatic maturation. There are marked differences in the expression of the CYP enzymes between immature and mature hepatocytes. These differences in the expression of specific CYP450 genes can be used as a gage of hepatic maturation. It is believed that the CYP enzymes expressed early in hepatic development are needed and are responsible for the metabolism and elimination
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of many endogenous compounds, such as steroids prostaglandins and retinoic acid. Both fetal and adult liver express genes of the CYP450 1A, 2C, 3A and 7A families; however, the expression of these genes even in fetal liver is generally in the range of a few percentage of normal adult levels. For most CYP enzymes there is a dramatic increase in expression in the postnatal period. In addition, the expression of specific family members differs in characteristic ways between fetal and adult liver which makes the expression of these genes diagnostic for the degree of hepatic differentiation. For example, Fetal liver expresses little of the adult form CYP1A2 (Maenpaa et al., 1993) but expresses high levels of CYP1A1 (Hines and McCarver, 2002; Omiecinski et al., 1990; Shimada et al., 1996; Yang et al., 1994). Adult liver expresses little CYP1A1 as its expression is slowly extinguished in the postnatal period. The expression of CYP1A2 increases after birth, however; it is one of the last of the CYP enzymes to develop in the postnatal period. Recent studies suggest that CYP1A2 levels may only reach 10 % of adult levels in the first postnatal month and may remain at or below 50 % of the adult levels out to one year (Sonnier and Cresteil, 1998; Tateishi et al., 1997). We propose that the expression of CYP1A2, is an indication of ‘true’ hepatic maturation and the ratio of expression of CYP1A1 to CYP1A2 provide a measure of the maturation process. A similar expression profile exists for members of the CYP3A family. Fetal liver expresses high levels of CYP3A7 (> 98 %) and little CYP3A4 (< 2% of total 3A) (Schuetz et al., 1994; Wrighton et al., 1988; Yang et al., 1994) while CYP3A4 is the major CYP enzyme expressed in adult liver. The expression of CYP3A7 is extinguished in most individuals over the first 6 postnatal months (Hines and McCarver, 2002; Stevens et al., 2003). Some CYP3A7 expression remains in 10–20 % of individuals but even in these CYP3A7 would only be expected to account for < 20 % of total CYP3A expression in that individual. Even after birth, CYP3A7 is not immediately extinguished such that the ratio of CYP3A4/3A7 may not equal 1 until 2–5 years of age. There is a change in the ratio of expression of CYP3A4 to 3A7 during development with the ratio of 3A4/3A7 extremely low during fetal development and high in adults. We suggest that the ratios of CYPs 3A4 to 3A7 and 1A2 to 1A1 will provide an estimate of the degree of hepatic differentiation of the stem cells. In addition to mRNA measurements, the metabolic activity of the individual CYP enzymes should be evaluated with probe drugs, and Western Blots should be run for individual CYP proteins. In this manner, direct comparisons can be made between stem cell-derived hepatocytes and authentic human hepatocytes for each endpoint studied. If implemented, suggestions like these will add quantitative analysis to this area of stem cell research. The development of a stem cell-derived hepatocyte system to examine the potential of new chemical entities (NCEs) to cause hepatotoxicity by a metabolism-mediated process early during the discovery process will provide a competitive advantage within the Pharmaceutical Industry. This novel cell-based system will provide an efficient means of aiding rational drug design and selection, selecting species differences in their responses to xenobiotics and, most importantly, for reducing the probabilities of causing unexpected adverse reaction in the liver when the compound reaches the market place. In addition to primary hepatocytes cultured in a sandwich configuration and stem cellderived hepatocytes, several recent developments in biological engineering and cellular biology have enabled a variety of new models for studying hepatotoxicity in vitro. Genetically engineered cells, as well as so called ‘three-dimensional (3-D) hepatocyte bioreactors’, have been established recently and are just now being applied to further our understanding of
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toxic events in the liver. In the case of engineered cells, these systems seek to provide a basis for more mechanistic insight into our current understanding of drug-induced hepatotoxicity, whereas the 3-D perfused models attempt to re-establish the in vivo microenvironment as well as build and improve upon some of the limitations of existing primary cell models. 1.2.3
Cell Lines and Genetically Engineered Cells
Advances in molecular biology have revolutionized our ability to investigate and dissect complex mechanisms of hepatotoxicity. Although primary hepatocytes remain the ‘gold standard’ for investigating the potential of a novel therapeutic agent to elicit a toxic event, several cellular models can be applied to address similar predictions of hepatotoxicity. Cell lines are available for studying various mechanisms of toxicity and these cells and cellderived systems are becoming better defined and understood as new data emerge from the laboratories and into the literature. Some systems have gained application in the field of hepatotoxicity as they derived from liver-based, cancer cell systems (e.g. HepG2, WIF-B9 cells), whereas some systems have been engineered/created to study specific mechanisms of hepatotoxicity where aspects of a given mechanism are understood (e.g. CYP-engineered cells, BSEP-expressing cells or vesicles). The utility and predictability of these systems relies heavily upon understanding how each system is related to the intact liver in vivo. Although each system may not express the full complement of proteins responsible for overall hepatic function, we are learning now that each system may have merit for predicting hepatotoxic events very early on in the drug-discovery process. 1.2.3.1
WIF-B9 Cells
A sub-clone of the WIF 12-1 cell line, WIF-B9 cells are derived from Fao rat hepatoma cells fused with WI38 human fibroblasts. As a result of their multi-species origin, these cells co-express many rat and human liver-specific proteins (Decaens et al., 1996; Ihrke et al., 1993; Shanks et al., 1994). These cells are capable of synthesizing albumin and were shown recently to express an array of rat and human P450 isoforms, some of which are inducible (Biagini et al., 2006a; Decaens et al., 1996). In addition to metabolic capacity, WIF-B9 cells are polarized, such that they establish functional canalicular spaces into which the cells are capable of transporting bile acid-like molecules (Bravo et al., 1998). Interestingly, these cells do not express the major human protein responsible for the uptake of bile acids into the cell (NTCP; sodium-taurocholate co-transporting polypeptide). However, they may express functional rat Ntcp and/or human OATPs (organic anion transporting polypeptides) that also have been shown to uptake bile acids (Konieczko et al., 1998). As in the case of NTCP, WIF-B9 cells do not express the human isoform of BSEP (bile salt export pump), the transport protein responsible for canalicular excretion of bile acids across the canalicular membrane. Therefore, due to the functional expression of rat Ntcp and rat Bsep (demonstrated by the vectorial excretion of a fluorescently labeled bile acid in these cells), the ability of this model to predict transporter-mediated hepatotoxicity (e.g. druginduced cholestasis) would be limited to predictions based upon the rodent genes (Bravo et al., 1998; Gradilone et al., 2005). Although, this system may better reflect transportermediated toxicity relative to that in the rat, WIF-B9 cells have shown promise in the prediction of hepatic toxicity, as measured by general cytotoxicity measurements (Biagini et al., 2006b). Specific evaluation of WIF-B9 cells for determination of alcohol-induced
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liver injury indicated that CYP2E1 was functional and that alcohol dehydrogenase (ADH) and aldehyde dehydrogenase (ALDH) activity was intact and similar to that observed in isolated hepatocytes (Schaffert et al., 2004). Moreover, the recent demonstration of aquaporin expression and localization suggests that this cell line may have utility in the prediction of toxic events that result in retarded bile formation in the liver (Gradilone et al., 2005). 1.2.3.2
HEPG2 Cells
Probably the most studied cell line with respect to hepatotoxic endpoints is the human hepatoma HEPG2 cell line. HEPG2 cells were first identified and isolated from a Caucasian adolescent over two decades ago (Aden et al., 1979; Knowles et al., 1980). Since their discovery in 1980, these cells have been utilized to examine various mechanisms of hepatotoxicity from ethanol-induced liver damage to most recently a high-content-screening approach for predicting hepatotoxicity due to drug administration (Neuman et al., 1993; O’Brien et al., 2006). One characteristic of HEPG2 cells that makes them attractive for use in hepatotoxicity studies is the expression and inducibility of relevant drug metabolizing enzymes. Expression and function of the key Phase I enzymes CYP1A, CYP2B, CYP3A, and CYP2E as well as several Phase II enzymes (e.g. UDPGT and GST) has been demonstrated and several of these enzymes can be induced by prototypical enzyme inducers (Knasmuller et al., 1998, 2004). However, it should be noted that not all drug metabolizing enzymes are present in HEPG2 cells; those that are may not be expressed at levels similar to those observed in vivo and may not respond to inducers similar to that in primary cells (Smith et al., 2005). Nevertheless, some investigators have transfected metabolic enzymes into this cellular system to establish a more realistic model for hepatotoxicity. HEPG2 cell transfection with CYP3A4, followed by administration of compounds known to form toxic metabolites, resulted in P450-specific formation of reactive species, leading to measured cytotoxicity (Vignati et al., 2005). Furthermore, following treatment with acetaminophen, HEPG2CYP2E1 cells generated reactive metabolites that formed protein adducts and damaged mitochondria, hence suggesting that introduction of mechanism-specific enzymes known to cause toxicity into HEPG2 cells may yield a predictive model for evaluating novel molecular entities (Bai and Cederbaum, 2004). In addition to drug metabolizing enzymes, expression of nuclear transcription factors in HEPG2 cells has provided insight into mechanisms of hepatotoxicity. ‘Upregulation’ of adipose differentiation-related protein through PPARγ was demonstrated in HEPG2 cells, offering insight into the mechanism of hepatic steatosis (Motomura et al., 2006). Furthermore, TCDD was able to upregulate CYP1A1 (as an indicator of oncogenesis) in HEPG2 cells through the aryl hydrocarbon receptor (Zhang et al., 2006). However, one must interpret these data carefully as CYP1A1 induction in HEPG2 cells is much more sensitive to that observed in primary hepatocytes and may not translate to the in vivo response. Much like the P450 enzymes, nuclear receptor transfection into HEPG2 cells also has been utilized to understand the involvement of these molecular factors in various mechanisms of hepatotoxicity (McCarthy et al., 2004; Vignati et al., 2004). HEPG2 cells also show promise for prediction of phospholipidosis, mitochondrial toxicity and oxidative stress (Bova et al., 2005; Kessova and Cederbaum, 2005; Sawada et al., 2005). Although these experimental systems have been optimized to predict specific hepatotoxic endpoints, the idea that HEPG2 cells can predict overall hepatotoxicity from a
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mechanism-independent perspective is now becoming accepted. Evaluation of multiple endpoints through the use of high content screening (see Section 1.3 below) in HEPG2 cells suggests that this cellular system can detect human hepatotoxicity with 80% sensitivity and 90% specificity (O’Brien et al., 2006). These results show promise for this cell line to predict overall hepatotoxicity and underscore that even after more than 25 years of research with this cell line, we continue to uncover the underlying biology of the system and develop means by which to exploit the cells to predict in vivo adverse events. 1.2.3.3
BSEP-Expressing Cell Lines
Modulation of normal bile salt export pump (BSEP) activity (e.g. through inhibition) has been identified as a major molecular mechanism responsible for drug-induced cholestasis observed in the clinic (Fattinger et al., 2001; Funk et al., 2001). Several methodologies have been created to assess the potential for novel therapeutic agents to interact with the BSEP and cause a clinical manifestation of cholestasis. Of the systems available, BSEPtransfected cells and vesicles derived from these cells show a high degree of concordance with clinical observations of cholestasis. One of the first and simplest experimental models involved the expression of a rodent BSEP using an Sf9 insect cell-based system, followed by formation of vesicles from the BSEP-expressing cells (Stieger et al., 2000). The addition of the bile acid taurocholate, as well as ATP, to the vesicle system resulted in functional BSEPmediated transport into vesicles. In addition, several known inhibitors (e.g. cyclosporine A, glibenclamide and rifampicin) were able to inhibit the transport of taurocholate by the BSEP. Although the cell lines expressing the rodent isoforms of the BSEP were generated very soon after the full-length clone was isolated, due to the presence of a cryptic bacterial promoter in the BSEP cDNA, the human isoform of the BSEP was much more difficult to clone and express. However, two groups were able to clone and express functional human BSEP into Sf9 cells and demonstrate a high degree of functional similarity of this protein to the rodent isoform (Byrne et al., 2002; Noe et al., 2002). Following the functional expression of the human BSEP, several groups have utilized these Sf9-derived vesicles to explain the observed hepatotoxicity of therapeutic agents such as nefazadone, ritonavir, and saquinavir (Kostrubsky et al., 2006; McRae et al., 2006). Most recently, the transfection of the human BSEP (coupled with human NTCP) into LLC-PK1-polarized cells has enabled the study of vectorial/polarized transport of bile acids across a cell monolayer (Mita et al., 2006). Undoubtedly, as our understanding of the role of specific transporter proteins in various mechanisms of hepatotoxicity continues to expand, the more predictive these systems will be in assessing the potential for a toxic event resulting from the administration of a novel therapeutic agent. 1.2.3.4
CYP-Engineered Cells
In some instances, hepatotoxicity may not be related to the original molecular entity administered to the patient. Rather, the formation of a highly reactive metabolite by CYP450 may contribute to an observed clinical adverse event. To predict the potential for CYP450 involvement in toxicity, cells have been engineered to express each of the major P450 isozymes responsible for the metabolism of xenobiotics. Immortalization of liver epithelial cells with an SV40 T antigen virus followed by transfection with human CYP450 genes yielded a THLE-CYP cell line that has been applied to address metabolism-mediated toxicity (Pfeifer et al., 1993). These transfected cells have been shown to express upwards of
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20-fold higher levels of CYP450 and have low basal expression levels of P450 enzymes, making them an ideal system to study isolated mechanisms of metabolism. The most studied compound in this system, Aflatoxin B1, is able to form DNA adducts that are modulated by the presence of the CYP enzymes (Mace et al., 1997). Moreover, these cells were able to identify the oxidative pathways for diclofenac and derive the contribution of each P450mediated process to the overall metabolism in vivo. (Bort et al., 1999a). Specific CYP isozymes have been expressed in other cell lines. When exposed to 1,3-dichloropropanol or cyclophosphamide, NIH-3T3 cells or V79 cells engineered to express individual CYP isoforms displayed a higher degree of toxicity than the control cells (Bull et al., 2001). This increased sensitivity was hypothesized to be due to the ability of the CYP-expressing cells to form reactive metabolites of these two compounds, thereby enhancing the cytotoxicity. Although these cells have promise for predicting the impact of Phase I metabolism on the hepatotoxic endpoint, these reagents often lack Phase II enzymes, overexpress the P450 of interest and do not respond to inducers of metabolism pathways. As a result, ultimate in vitro/in vivo correlation of these systems remains a challenge for their widespread use and adoption for assessing hepatotoxicity. 1.2.3.5
Other Cell Lines and Engineered Cellular Systems
In addition to the cellular systems mentioned above, several other cell lines have been used, although not as widespread, to address mechanisms of hepatotoxicity. These reagents include cell lines such as HUH-7, HepaRG and HEP3B cells (Aninat et al., 2006; Le Vee et al., 2006; Manov et al., 2002; Xu et al., 1997). Continued evaluation and testing of these cell lines will be a critical step for the acceptance of these as viable reagents to predict toxic events in the liver. 1.2.4
Three Dimensional (3D) Hepatocyte Bioreactors
Outside of using the intact organ ex vivo, one of the major limitations of current in vitro models of liver function has been the static nature of most in vitro systems. Due to the static nature of these experimental models, it is often difficult to re-establish the microenvironment that cells experience under normal in vivo conditions, and thus mimic in vivo liver physiology. It has been well established that flow through the liver sinusoids imparts shear stress onto liver cells that dictates cellular gene expression and function (Hara et al., 2003; Sato et al., 1997, 1999). Moreover, extensive cell-to-cell contact, establishment of cell polarity and the impact of non-parenchymal cells all can be important for re-establishing in vivo hepatic function (Hamilton et al., 2001; Hoebe et al., 2001; Hoffmaster et al., 2004; LeCluyse et al., 2005; Saad et al., 1993). Static systems have been used extensively, and albeit to some degree of success have shown promise for predicting some incidences of hepatotoxicity in the clinic. In contrast to the more frequent use of static cultures of primary hepatocytes and hepatic cell lines to predict hepatotoxicity as mentioned above, little measure of hepatoxic endpoints in these complex, perfused, 3D models of liver function have been performed. Given the in vivo phenotype observed in many of these novel systems, one can only imagine that the predictive nature of each of these models may far exceed our current ability to assess hepatotoxic events before entering the clinic with a promising new chemical entity. Although the application to the field of hepatotoxicity has not yet been
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exploited with these emerging technologies, the background and rationale for several of the leading 3D perfused models is described briefly below. 1.2.4.1
MIT Liverchip
One of the most promising 3D models for recapitulating liver function has been developed by fabricating a series of small channels into a scaffold where primary hepatocytes can be seeded and subsequently perfused with a user-defined cell culture medium. Developed at the Massachusetts Institute of Technology (MIT), this unique microfabricated bioreactor design enables morphogenesis of 3D tissue structures, optimal oxygen transfer to the established tissue and physiologic shear stresses to be imparted upon the cells cultured in the device (Powers et al., 2002a,b). As a result, these in vitro cultures of cells yield morphology similar to that observed in the intact liver, expression of liver-specific proteins and mRNA near physiologic levels and metabolic function for several key Phase I enzymes in line with formation rates and metabolic profiles observed in freshly isolated hepatocytes or native, intact tissue (Sivaraman et al., 2005). For example, whereas cytochrome P450 levels decline rapidly in primary hepatocytes shortly after isolation, the expression of Cyp2c and Cyp3a was maintained in the ‘Liverchip’, and further supported by formation of specific testosterone metabolites of these CYP isoforms. Testosterone hydroxylation rates in these studies also were similar to rates observed in vivo (Sivaraman et al., 2005). One key advantage of this approach is the ability to scale the system and adapt the system to a high-throughput configuration. Since the functional unit of the reactor is a single channel, the mass of tissue in the device can be controlled directly through fabrication of larger or smaller scaffolds containing more or less individual channels. In addition, the device has been recently adapted to a multi-well configuration amenable to higherthroughput and automated liquid handling – ideal design criteria for screening new chemical entities in a drug discovery environment (unpublished data). Some challenges remain before this technology gains wide application in the field of toxicology. As the majority of data generated to date reflects experiments conducted with rat hepatocytes, the introduction and success of human hepatocytes in the ‘Liverchip’ will be very important for translating data from this system to understand the clinical manifestations of hepatotoxicity. The further characterization of cellular function in the device (e.g. bile acid transport, mitochondrial function, etc.), coupled with specific experiments designed to assess different mechanisms of hepatotoxicity will help demonstrate the overall utility of this system. Nevertheless, the results and data generated to date suggest strongly that this system offers superiority over currently available static culture systems for evaluating liver-specific function. 1.2.4.2
Hollow Fiber Reactor
Originally designed as an extracorporeal bridging device for liver transplant patients, the multi-compartment woven fiber bioreactor has shown that hepatocytes spontaneously form 3D structures once implanted into the device (Gerlach et al., 2003). These structures display morphology similar to that observed in vivo, and cells can be maintained upwards of seven weeks in culture, without undergoing major histological changes (Gerlach et al., 1994; Gerlach et al., 1995). The fundamental principal behind this technology is the incorporation of artificial capillaries providing perfustate flow that promote the formation of 3D structures. When cultured for two weeks, freshly isolated cells were capable of maintaining liverspecific function such as urea synthesis, albumin production, P450 function and glucose
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metabolism (Zeilinger et al., 2000, 2002). Recent data also suggests that non-parenchymal cells are able to organize into functional structures complementary to the hepatocytes, and cells may have some proliferative capacity when cultured in the device (Zeilinger et al., 2004). 1.2.4.3
Zonation-Based Bioreactor
A flat-plate reactor that incorporates both flow and the concept of oxygen gradients over the length of the reactor chamber has been fabricated (Allen and Bhatia, 2003; Allen et al., 2005). These researchers constructed a model based upon the concept of zonation of the liver in vivo, and have recreated this environment by varying the degree of oxygen tension over a flat plate of hepatocytes with or without co-culture with fibroblasts (Allen and Bhatia, 2003; Jungermann, 1986; Pang et al., 1994). Once this gradient is established in culture, regional heterogeneity of drug metabolizing enzymes (e.g. CYP3A, CYP2B) can be seen, similar to the patterns observed in intact liver. Cell viability can be maintained in this device for at least five days and P450 enzymes can be induced by prototypical inducing agents such as dexamethasone. The zonation-based bioreactor is one of the few novel perfusion based system where specific toxicity studies have been designed and conducted. Treatment of the bioreactor with a range of acetaminophen concentrations resulted in zonal toxicity of the hepatocytes in the device; toxicity was higher in the low oxygen portion of the culture (Allen et al., 2005). These data are supported by the hypothesis that low oxygen concentrations promote glutathione depletion, an important detoxification pathway for the toxic NAPQI acetaminophen mediated metabolite. However, enzymes other than CYP3A responsible for acetaminophen metabolism (e.g. CYP2E, UGT, SULT, etc.) have yet to be characterized in the reactor system. 1.2.4.4
Microvascular-Based MEMS Bioreactor
The highly branched vasculature of the liver represents a challenge for reconstruction in an in vitro system. Nevertheless, using principles based on photolithography and silicon micromachining, a group of scientists has machined successfully a vascular network onto a silicon wafer using micro-electromechanical system (MEMS) techniques (Fidkowski et al., 2005; Kaihara et al., 2000). Transfer of this network onto a biodegradable polymer (the wafer acts as a micro-mold) and subsequent removal results in the formation of a two-dimensional polymer scaffold similar to the vascular and capillary networks of the liver. The resultant vascular channels can be seeded with endothelial cells; hepatocytes are seeded into an adjacent parenchymal compartment. These compartments are separated by the presence of a highly permeable membrane that allows the movement of small molecules but isolates the cell types within the device. Oxygen exchange can also happen across this membrane and although the cells remain physically separated, inter-cellular communication through soluble factors is accommodated in this design. When multiple monolayers of hepatocytes are layered and adhered to each other, these ‘sheets’ of hepatocytes recreate a three-dimensional tissue structure that is highly vascularized (Kulig and Vacanti, 2004). The cells remain viable for several days in culture and are able to maintain liver-specific function (e.g. albumin synthesis, urea production, etc.). As the initial application of this technology focused on the fabrication of an implantable human liver, the incorporation of biodegradable and biocompatible polymers has been the recent focus for the development of the technology. However, researchers have realized quickly that this model system shows
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significant promise for the evaluation and screening of potential hepatotoxicants in a human cell-based system prior to moving into the clinic. Experiments are just now being designed to address and exploit this opportunity to better understand the potential for hepatotoxicity in this more complex, more physiologic model system.
1.3
Applications of in Vitro Models
For in vitro models to be considered relevant to in vivo situations, several criteria have to be satisfied: (1) the models are well-characterized and understood; (2) the concentrations of the test articles used are considered relevant in vivo; (3) the measured outcomes are considered relevant in vivo; (4) the underlying mechanisms are considered relevant in vivo. Representative applications of cell-based assays currently used to study DILI are summarized in Table 1.2. Although enzyme leakages (ALT, AST, LDH leakages) can be easily measured both in vivo and in vitro, they do not provide any insight into the underlying mechanisms of damage. Recently, high-content analysis (HCA) has emerged as a versatile technology platform with several distinctive advantages and applications. HCA is essentially a multi-parametric analysis of data that often include both biochemical and cytometric measurements. The HCA readouts may be kinetic on live cells, or single timepoint readouts on fixed cells. High-resolution analysis of sub-population of cells in each well, as well as the morphometric measurements of subcellular organelles, is often part of the HCA readouts. These image cytometric measurements compliment the traditional biochemical readouts such as enzyme leakage, ATP content, albumin synthesis or urea synthesis. High-content screening (HCS) is an abbreviated version of HCA, by selecting a few parameters from the HCA that were deemed to be most informative, predictive and robust. By selecting only a subset of the most informative parameters to measure, HCS can significantly increase the throughput of HCA to meet today’s drug discovery needs. 1.3.1
High-Content Screening for Human Hepatotoxicity Potential
Recent studies have demonstrated that the clinical occurrence of drug-induced human hepatotoxicity is highly concordant with in vitro cytotoxicity when assessed in a cell-based model with a novel combination of critical features and using high content screening (HCS; (O’Brien et al., 2006)). In contrast, concordance was low for previous cytotoxicity assays applied to drugs that produce human hepatotoxicity, because of poor assay sensitivity (Xu et al., 2004). Whereas these previous assays were only effective with severely hepatotoxic chemicals and drugs, they had high specificity, with positive test results being more than 90 % indicative of hepatotoxicity potential. HCS technology applies epifluorescence microscopy and image analysis to live cells incubated in microtiter plates under physiological conditions. It rapidly and automatically photomicrographs and quantitates fluorescence of multiple dyes at different subcellular locations (Haskins et al., 2001). This enables sensitive detection and following of the progression of key cytomorphologic and intracellular biochemistry effects of hepatotoxic drugs, such as on cell proliferation, nuclear and mitochondrial DNA, mitochondrial mass and activity, intracellular calcium, lysosomal mass, oxidative stress, and cell structure (Figures 1.6 and 1.7).
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Figure 1.6 Detection of human hepatotoxicity potential with high-content screening. Composite images of hepatocytotoxic changes produced by incubation of HepG2 cells with cerivastatin (25 μM) for three days. 1, increased mitochondrial potential; 2. increased ionized calcium; 3, permeabilized plasma membrane with increased Ca and decreased mitochondrial membrane potential; 4, ruptured cell
There are several critical features of the cell model used with HCS that produce high assay sensitivity and concordance with human toxicity, in contrast to past cytotoxicity models (O’Brien et al., 2006; O’Brien and Haskins, 2006). These include duration of treatment, multiparametric monitoring of individual and live cells, drug metabolic competency and testing at concentrations relevant to efficacious concentrations. Most critical was the need for sufficient time to allow expression of the cytotoxicity (O’Brien et al., 2006; Slaughter et al., 2002; Xu et al., 2004). Three days of incubation of cells was effective for more than 95 % human hepatotoxic drugs, whereas a single day of incubation was frequently ineffective (O’Brien et al., 2006) or produced cytotoxicity at a much higher concentration (O’Brien et al., 2006; Slaughter et al., 2002; Xu et al., 2004). Next most important for predictivity was the number and choice of parameters measured. Early, sublethal effects on cell proliferation, cell morphology and mitochondria occurred consistently and ubiquitously with toxicity and when used collectively were most diagnostic. The occurrence and timing of effects on intracellular calcium concentration, lysosomal mass, oxidative stress or plasma membrane permeability frequently provided additional information indicative of mechanism of toxicity. For example, in Figure 1.7, phospholipidosis from cationic amphiphilic drugs, mitochondrial DNA depletion by nucleoside reverse transcriptase inhibitors that also inhibit mitochondrial DNA polymerase gamma and redox cyclers that produce reactive oxygen species are demonstrated. In addition, in Figure 1.6, the complex mechanism of statin-induced toxicity is demonstrated with early sublethal effects on apoptosis, mitochondrial function and calcium homeostasis (Diaz and O’Brien, 2006). Use of human hepatocytes with potential for drug metabolism is also important for assessment of hepatotoxicity as numerous drugs produce this effect by their hepatic metabolites. Drugs producing idiosyncratic hepatotoxicity and/or toxicity by their metabolites
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(b)
(c)
Figure 1.7 Mechanisms of human hepatotoxicity identified in HepG2 cells by high-content screening. (a) Phospholipidosis was produced by incubating cells with 12.5 μM imipramine for three days and then staining them with Hoechst 33342 for DNA, Mitotracker Far Red for mitochondrial mass and Lysotracker Green for lysosomal mass, as indicated. (b) Mitochondrial DNA depletion by nucleoside reverse transcriptase inhibitors is demonstrated by incubation of HepG2 cells for three days with 1.6 μM zalcitabine and staining nuclear and mitochondrial DNA (arrows) with Picogreen. (c) Oxidative stress produced by menadione. HepG2 cells were treated with 100 μM menadione and stained with Hoechst 3342 and dihydroethidium. Selection of objects for assessment is indicated by the circles and is based on Hoechst 3342 staining of nuclear DNA. Ethidium forms as a result of oxidative product of dihydroethidium, binds nuclear DNA and fluoresce with a different wavelength than the Hoechst. This is seen in the photomicrograph as increased whiteness of the encircled areas (right)
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(Kalgutkar et al., 2005; Kaplowitz, 2005) were detected with HepG2 cells as effectively as drugs producing toxicity directly (Table 1.7). This high concordance contrasted remarkably with the 15 % concordance of seven other conventional assays in which a 50 % effect at 30 μM was considered positive for changes in any of the seven readouts: DNA synthesis, protein synthesis, glutathione depletion, superoxide secretion, caspase-3 activity, membrane integrity and mitochondrial reductive activity. The basis for this difference in sensitivity was not determined but may relate in part to induction of metabolic competence over the three days of exposure in the HCS assay. Finally, assay effectiveness also depended on measurement at the single-cell level in order to allow exclusion of extracellular staining or artifact or dead cells. Determination of the concentration producing cytotoxicity in the HCS sublethal cytotoxicity assay was assessed by 12-point dose response curves (Figure 1.8). A toxic effect was defined as the point when values for the parameter departed from the baseline and negative controls by more than two coefficients of variation. In Figure 1.8, typical dose – responses are illustrated in which all eight parameters measured were affected. The pattern and sequence of changes in the different parameters frequently reflected the mechanism of toxicity. For fenofibrate (Figure 1.8), there was nuclear swelling and inhibition of cell proliferation, followed by mild increases in intracellular calcium with some loss of mitochondrial membrane potential and an increase in membrane permeability, followed by overt oxidative stress with mitochondrial biogenesis. This pattern contrasted with that of cerivastatin (Figure 1.6), where there was first nuclear shrinkage and increased mitochondrial membrane potential, followed by increased intracellular ionized calcium. At higher concentrations, the calcium progressively increased, the mitochondrial potential progressively fell and membrane permeability increased. Virtually all drugs and chemicals cause toxicity at high enough concentrations. Thus, it is critical to assess toxicity at concentrations relevant to those that are used for drug efficacy. Efficacious concentration, as defined as the maximal serum concentration of drug used for treatment (Cmax ), is highly variable, ranging 10 000 000-fold in one study of 187 marketed human drugs from 100 pM to 2 mM, with 90 % values less than 100 μM, 60 % less than 10 μM, 37 % less than 1 μM and 12 % less than 100 nM (Figure 1.8). Most human hepatotoxic drugs (94 % of 102 tested) are cytotoxic in the sublethal HSC cytotoxicity assay at concentrations less than 100-fold Cmax , whereas most non-toxic drugs (96 % of 23 tested) are cytotoxic in this assay at concentrations more than 100-fold Cmax (O’Brien et al., 2006) (Figure 1.9). Hormesis, in which compensatory adaptive changes precede and occur at lower doses than degenerative changes, was detected for half of the toxic drugs for cell proliferation, cell morphology and mitochondria (O’Brien et al., 2006). It is demonstrated for cerivastatin in Figure 1.6 for mitochondrial membrane potential and for fenofibrate in Figure 1.8 for mitochondrial mass and nuclear area. Hormesis could not be assessed for parameters that normally have low values, such as intracellular calcium measured by fluo4 or membrane permeability measured by toto-3, because assay methods were not sufficiently sensitive. However, for calcium, more sensitive dyes, with calcium dissociation constants closer to the physiologic concentration of ionized calcium, have detected biphasic effects on resting calcium (O’Brien et al., 1990). In the sublethal HCS assay, sensitivity and specificity for identification of human hepatotoxicant drugs were 94 % and 96 %, respectively, when testing only hepatotoxicant drugs
1 2 3 4 5 6 7 8 9 10 11 12 13
Acetaminophen Chloramphenicol r Danazol i,r Diclofenac i,r Felbamate r Flutamide i,r Hydralazine r Ibuprofen r Indomethacin r Imipramine r Isoniazide i,r Leflunomide i,r Methyldopa
r
i,r
500↑4000↓ 260 13 16↑126↓ 315 50 67 50 190 0.8↑50↓ — 25 330
− − − − − − ND − − − − + ND 4000 — 50↑100↓ 250 160↑ 50 135↑ — 47 100 50↓ — 330
Mitochondrial potentialc — — 25 — — 50 270 — — 50 — — 330
Ca — — 50 126 — 50 — — — 50 — — 330
Membrane Pote ntial
7800 — 13 126↑ 315 — 75 — 2↑ 50 — — 330
Nuclear area
130 57 0.16 4.2 42 6 5 250 6 0.6 40 340 11
Cmax (uM)
4 5 81 4 4 8 13.5 0.2 0.3 1 1 0.07 30
TI
8 53 — 99 24 90 87 99 90 100 0 99.3 15
PPB (%)
OS OS — OP, Ap IM M IM IM, M IM, OS IM, M IM, OS IM —
Mechanismd
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Table 1.7 Concordance of HCS assay results with idiosyncratic or drug metabolite-implicated human hepatotoxicity (based on O’Brien et al., 2006). Drugs are compared with how they tested in seven conventional cytotoxicity assays (column 2) and how they tested in the HCS assay (columns 3–7). Results for the HCS assay are tabulated for lowest observed adverse effect level (LOAEL) concentrations (in μM) causing effects on cell number, mitochondrial membrane potential, intracellular ionized calcium concentration, membrane permeability and nuclear area, respectively. The maximal plasma total concentrations of drugs associated with efficacy (C max ) are indicated in column 8. The ratio of the lowest cytotoxic concentrations from columns 3–7 to Cmax from column 8 is indicated in column 9 (TI). The percentage of drug that is bound to plasma proteins is also indicated (column 10) and where known the cellular mechanism of cytotoxicity (column 11). The sensitivities of the conventional assays and the HCS assay are indicated in the bottom row. The sensitivity of the HCS assay was determined using a TI cutoff of 100 in column 9. Tacrine is the only drug in this panel that falls just outside of the 100 cutoff. It is of interest to note that tacrine only causes asymptomatic elevations of liver enzymes that do not progress to frank liver injury despite continued use of the drug
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25 50 160 630↑ — 140 — 50↑100↓ 25 6 4000↑
b
Superscript ‘i’, idiosyncratic heptatotoxicity; Superscript ‘r’, reactive metabolite. ND, not determined. c ↑signal increased;↓ signal decreased. d OS, oxidative stress; Ap, apoptosis; IM, imune-mediated; M, mitochondrial.
a
13 12 5 630 50 285 1100↑ 25 100↑ 3 1000
— 50 — 630 100 — — 50 — 6 —
— 50 — — 100 — — 25 25 6 4000 96 %
50↑ 0.7↑ 5↑ 320 13↑ 570↑ — 13↑50↓ 0.4↑25↓ 13 8000
8 6 5 12 9 19 217 0.1 0.4 4 540
2 0.1 1 27 1 8 5 130 1 0.8 2
76 62 99 18 81 94 62 55 98 99 93
IM, M IM, OS, M — IM — IM, OS IM, M OS, M M — OS, M
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Hepatotoxicity Nuclear area
Nuclear area
30
30
25
25
20
20
15
15
10
10
lonized calcium (Fluo 4)
Mitochondrial DNA (Picogreen)
200
3500
150
3000 2500
100
2000
50
1500
0
1000
TMRM
Dihydroethidium oxidation rate 0.18
500 400
0.12
300 0.06
200 100
0.00
0
Plasma membrane integrity (Toto-3)
MitoTracker Deep Red 1200
500 400 300
800
200 100 400
0
Cell count (10× objective, 10 fields)
Cell count (20× objective, 6 fields)
5000
300
4000
250 200
3000
150
2000
2000
500
1000
250
62.5
125.0
31.3
7.8
15.6
3.9
0
2000
500
1000
250
125
62.50
31.25
7.81
15.63
3.91
0 0
50
0
2.0
100
1000 1.95
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Figure 1.8 Use of high-content screening for quantitative determination of dose–response relationships for drug-induced, human hepatocellular toxicity (based on O’Brien et al., 2006). Fenofibrate (25 μM) was assayed in HepG2 cells at doubling concentrations from 2 to 2000 using two different combinations (columns 1 and 2) of fluorescent probes. For both assays, Hoechst 33342 was used to determine nuclear area and cell count. Ionized calcium was assayed using Fluo4, mitochondrial membrane potential using TMRM, plasma membrane integrity using TOTO-3, mitochondrial DNA using picogreen, mitochondrial mass using MitoTracker Deep Red and reactive oxygen species using dihydroethidium. The circles in each dose–response curve indicate the lowest observed adverse effect level (LOAEL) values which were used to tabulate the LOAEL concentrations shown in Table 1.7
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Cumulative frequency In vivo efficacious concentration (C max)
Human hepatoxic drugs
Non-toxic drugs
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Figure 1.9 Safety margins for HCS sublethal cytotoxicity assay concordant with human hepatotoxicity. A frequency histogram (a) for the in vivo efficacious concentration in humans (Cmax ) of 187 marketed drugs demonstrates a wide range and indicates the need to assess cytotoxicity in the context of efficacious concentration. A frequency histogram (b) of the safety margin (in vitro HCS cytotoxicity concentration/Cmax ) for 102 human hepatotoxicant drugs and the 23 drugs for which this ratio was determined indicates that a safety margin of 100 is the most effective discriminator of human hepatotoxicity potential (based on O’Brien et al., 2006)
and non-toxic drugs (O’Brien et al., 2006). However, when other drugs that produce other organ toxicities (e.g. kidney, heart, bone marrow, muscle and pancreas) were tested their cytotoxic effects were not distinguishable from those of drugs causing hepatotoxicity. Thus, cytotoxicity in the HCS assay was concordant with human toxicity but not specific for liver toxicity. An additional caveat in the use of the assay is that it did not detect cholestatic effects of drugs. Nor did it detect other organ toxicities produced by drugs’ effects on proteins not found in hepatocytes. 1.3.2
Primary Hepatocytes and Liver-Specific Toxicities
While hepatic cell lines have several advantages over primary cells, it is still prudent to take a selected subset of compounds into primary hepatocytes. The reasons are mainly the following. primary hepatocytes are more differentiated cells compared to cell lines. They maintain higher levels of metabolic activity, normal p53 status and cell cycle regulation and more normal levels of transporter protein expression (Le Vee et al., 2006; Wilkening et al., 2003). For example, primary hepatocytes are more suitable than HepG2 to study metabolismmediated liver toxicants, such as benzo[a]pyrene, dimethylnitrosamine (DMN),
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2-amino-1-methyl-6-phenylimidazo[4,5-b]pyridine (PhIP) (Wilkening et al., 2003), acetaminophen, diclofenac, cyclophosphamide, disulfiram (Bort et al., 1999b; Wang et al., 2002) and nimesulide (Rainsford et al., 2001). High-content analysis of primary hepatocytes has also been used to study more liver-specific toxicities, such as steatosis, steatohepatitis, cholestasis and reactive metabolites. Steatosis, or accumulation of fatty acids (i.e. ‘fatty liver’), can be caused by a variety of drugs including amiodarone, perhexiline maleate, 4,4 -diethylaminoethoxyhexestrol (DEAH), tetracycline, valproic acid and several antiviral nucleoside analogues, such as fialuridine (Fromenty and Pessayre, 1997). One of the major mechanisms involved in steatosis is the inhibition of beta-oxidation of long-chain fatty acids, either by direct inhibition or indirect inhibition, such as CoA sequestration or mitochondria DNA damage (Fromenty and Pessayre, 1995; Jaeschke et al., 2002b). The resulting fatty acid accumulation can be detected and quantified by staining primary hepatocytes with neutral lipid stains such as Oil red O and performing HCA under the microscope (Amacher and Martin, 1997; McMillian et al., 2001). This is different from using phospholipids stains such as NBD-PC or NBD-PE to study phospholipids accumulation in the cytoplasm (i.e. phospholipidosis) (Gum et al., 2001). In addition, the aldehyde products of lipid peroxidation, 4-hydroxynonenal and malondialdehyde (MDA), can be measured by biochemical readouts (Berson et al., 1998a,b). MDA is a known stimulator of inflammatory responses. Therefore, repeated challenge by steatotic drugs and the resulting lipid peroxidation represent an important mechanism of drug-induced steatohepatitis (Ivanov et al., 1992). While simple steatosis was regarded as benign, research in the past decade suggested that mitochondria dysfunction and reactive oxygen species generation are important factors to differentiate simple steatosis from nonalcoholic steatohepatitis (NASH) (Fromenty et al., 2004). Using primary cultured hepatocytes, it was found that amiodarone, perhexiline maleate and DEAH increased the mitochondrial formation of reactive oxygen species and caused lipid peroxidation (Berson et al., 1998b). Based on these findings, it would be important for high content assays to measure mitochondria function and oxidative stress in addition to simple neutral lipid accumulation. Intrahepatic cholestasis, defined as impairment in bile formation and/or bile flow, is another common manifestation of drug-induced liver disease. In humans, intrahepatic cholestasis most often occurs in the elderly (Feuer and Di Fonzo, 1992). As the population ages and the occurrence of multiple drug therapy in geriatric patients increases, it is expected that jaundice and/or drug-induced intrahepatic cholestasis will become even more prevalent. Bile formation is dependant on the specific transporter proteins in hepatocytes. The expression and the appropriate membrane sorting of these transporters are highly dependent upon the differentiated phenotype of the cells (Hoffmaster et al., 2004; LeCluyse et al., 1994). Sandwich-cultured primary hepatocytes are currently the most well-characterized in vitro model to study the disposition of taurocholate (Liu et al., 1999), bilirubin and its glucuronide (Lengyel et al., 2005) and drug-induced cholestasis (Kostrubsky et al., 2003). As expected, inhibition of important hepatobiliary transporters, such as the bile salt export pump (BSEP), can result in drug-induced cholestasis (Kostrubsky et al., 2006). Although in the cases of rifampicin (Li and Chiang, 2006), it was found to induce bile acid and bilirubin detoxification (Marschall et al., 2005) via PXR activation and this drug has been successfully used to treat cholestatic liver disease. This example highlights the importance of using longer-term primary hepatocyte cultures (> 3 days) that express differentiated and
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polarized liver functions as opposed to the short-term BSEP inhibition experiments (< 1 h), to provide a more complete picture of in vivo outcome. It is well known that many hepatotoxic agents can be metabolized to reactive metabolites that can either be detoxified or react with glutathione, enzymes, nucleic acids, lipids or proteins (Knowles et al., 2000; Pessayre, 1995). These reactive intermediates are electrophilic metabolites or free radicals that are generated during the metabolism of a broad range of functional groups. Reactive metabolite formation and/or oxidative stress induced by drugs are also considered important factors in immune-mediated idiosyncratic drug hypersensitivity (Park et al., 2001). There are several rapid in vitro methods to detect and measure the generation of such reactive intermediates. For example, high-throughput assays for identifying pharmaceutical compounds that produce reactive metabolites have been developed. These methods involve incubating drug candidates with a liver microsomal drug metabolizing enzyme system in the presence of glutathione and detecting glutathione conjugates via tandem mass spectrometry (Chen et al., 2001; Haroldsen et al., 1988; Pearson et al., 1988). However, it is now recognized that reactive metabolite formation per se is not sufficient to cause tissue injury (Park et al., 2005; Sanderson et al., 2006). Hence, cell-based models to evaluate the toxicological consequences of the formation of reactive intermediates (or lack thereof) are becoming increasingly important. In cells, the reduced form of glutathione (GSH) is known to interact with electrophilic compounds/metabolites and free radicals to play a key role in the detoxification of such reactive molecules. Depletion of the reduced form of glutathione was reported to be a marker of hepatotoxicity (e.g. Fouin-Fortunet et al., 1984), suggesting its evaluation is important in toxicological studies. For example, monochlorobimane and chloromethylfluorescein diacetate (CMFDA) have been successfully used to monitor cellular GSH levels by epifluorescence in isolated hepatocytes (Lilius et al., 1996; Thompson et al., 1998). The fluorometric dye, 2 ,7 -dichlorodihydrofluorescein (H2 DCF) can be used to detect oxidative stress induced by various oxidative radicals in human cells including hepatocytes (Lautraite et al., 2003; LeBel et al., 1992; Wang and Joseph, 1999). The diacetate form of H2 DCF freely enters the cell and is hydrolyzed by intracellular esterases to liberate H2 DCF, which then reacts with oxidizing species to produce the highly fluorescent DCF. These cell-based assays are amenable to high-throughput evaluation of chemical compounds and their reactive metabolites and/or oxidative species in drug discovery (Lautraite et al., 2003). 1.3.3
RNAi Technology
Cell-based systems are currently being coupled with new technologies such as RNA interference (RNAi) as a potential tool to identify genes with predictive functions significant to drug development. RNAi represents an evolutionary conserved mechanism developed by nature to protect the genome against endogenous and exogenous stress insults, and regulate gene expression. There are numerous excellent reviews on the use of RNAi as a research tool and as therapeutic agents (Cejka et al., 2006; Cullen, 2006; Dillon et al., 2005; Dykxhoorn and Lieberman, 2005; Dykxhoorn et al., 2006; Hannon and Rossi, 2004; Lee and Sinko, 2006; Shankar et al., 2005; Sledz and Williams, 2005). Briefly, endogenous mammalian RNAi is mediated by small-interfering RNAs (siRNAs) produced from long double-stranded hair pin RNA known as micro RNA (miRNA) through the action of an endonuclease of the ribonuclease-III type, called ‘drosha’ and ‘dicer’. These siRNAs
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or small double-stranded RNAs are incorporated into the RNA-inducing silencing complex (RISC), which contains RNAase activity, to become small single-stranded RNAs. The single-stranded RNA guides RISC to mRNA that has a complementary sequence and mediates gene silencing by targeting RNA. This silencing is caused by either translational repression if there is a mismatch in the sequence of the siRNA and target mRNA or by RNA cleavage if there is an exact match between the siRNA sequence and the target mRNA. Thus, the synthesis of the protein encoded by the mRNA targeted by the siRNAs is prevented, and that protein is selectively depleted from the cell. The enzymatic machinery required to process siRNA is ubiquitously expressed in most eukaryotic cells and can be co-opted by exogenous RNAs to direct the sequence-specific gene silencing. Therefore, siRNA can be chemically synthesized and transfected into cells, tissues and animals after being integrated into specialized delivery systems, such as liposomes. Alternatively, plasmids or viral vectors can be used to express shRNA (short hairpin RNA that structurally mimics a miRNA precursor). Successful ‘knockdowns’ of specific genes allow the investigation of potential drug targets, as well as the genetic basis of physiological and disease processes in mammalian systems. RNA interference (RNAi) technology is widely used in vitro for modulating specific targets and assessing biologic pathways; in addition, this novel technology has the potential to be used for addressing drug-induced liver injury (DILI). Toxicological effects observed upon treatment with a given drug in the presence or absence of a specific RNAi molecule can be evaluated in cell-based systems to determine whether the toxicity associated with chemical entities is mediated by interaction with that specific target. For example, Hep2 cells stably expressing specific small interfering RNA directed against the activating signal cointegrator-2 (ASC-2) have been successfully used to demonstrate that ASC-2 protein is likely to participate in acetaminophen-mediated hepatotoxicity as a transcriptional coactivator of the xenobiotic nuclear receptor CAR in vivo (Choi et al., 2005); mouse hepatocytes silenced against Fas expression were found to be protected from cytotoxicity caused by actinomycin D (Song et al., 2003); Moreover, RNAi technology will further demonstrate participation of the specific cytochrome P450 (CYP450) enzyme in the DILI process. This was illustrated when CHL-3A4 cells with 3A4 siRNAs significantly diminished the cytotoxicity of cyclophosphamide and ifosfamide (Chen et al., 2006). RNAi technology has also been utilized to understand the role of transport proteins such as Mrp1-3, Mdr1 and Bcrp (Lee and Sinko, 2006; Sahi, 2005; Tian et al., 2004; Xu et al., 2005), and the functional analysis of nuclear receptors such as the Farnesoid X receptor and androgen receptor (Plass et al., 2002; Wright et al., 2003) and their co-regulators, such as SRCs, p300, NcoR and SMRT (Debes et al., 2002; Yoon et al., 2003; Zhou et al., 2003) in DILI. At present, the prediction of metabolism-dependent hepatotoxicity is difficult or even impossible because there are no suitable experimental (in vivo/in vitro) models and we do not understand the basic mechanism involved in the toxicity when it does occur in man. Cell-based models coupled with RNAi technology could be useful systems for predicting metabolic activity of drugs in vivo. In vivo delivery is still the greatest challenge to the effective use of this technology in target characterization and investigating adverse pharmacology and chemically mediated toxicity. However, recent advances in in vivo delivery and vector technology make it probable that RNAi can be effectively used for in vivo hypothesis-driven research (Aligner, 2006; Behlke, 2006; Lu et al., 2006).
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Conclusions
We have reviewed various in vitro models currently used or under development to better enable the evaluation of drug-induced liver injury (DILI). Continuing advances in stem cells research, new immortalized cell lines and genetically engineered cells, 3D-hepatocyte bioreactors and our enhanced ability to predict outcomes from primary cells have allowed a better understanding of the mechanisms of toxicity, metabolism of drugs and the species differences in expression of toxicity. Each model described in this review has significant advantages and it is possible to find an appropriate system for any particular toxicological question asked. Further research should be directed towards the refinement of existing methodologies and the development of new alternatives and testing paradigms for human relevancy. This approach coupled with the establishment of new technologies (e.g. HCS, RNAi and ‘omics’) will improve our ability to assess DILI earlier in the drug discovery process and would allow us to identify compounds with decreased risk of hepatotoxicity at later stages of development for eventual human health risk assessment. To fully establish the credibility and relevance of in vitro toxicity evaluation of drugs, it is essential that in vitro/in vivo correlations of toxicity of NCEs or drugs be determined by toxicologists and cell culture scientists.
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2 Utilization of an in vitro Hepatotoxicity Test in the Early Stage of Drug Discovery Ikuo Horii, Hiroshi Yamada, Rie Kikkawa, Toshinori Yamamoto, Tamio Fukushima, and Kaori Tomizawa
2.1
General Introduction
Safety evaluation of pharmaceutical products has been conducted in the form of risk assessment and the study of prediction and prevention of adverse effects on the basis of an extensive scientific area over toxicology, pathology, pharmacology, biochemistry, physiology, etc. With the rapid progress of genomic science, diseases and their causes have become understood at the genetic level. With remarkable advancements both in drug discovery, R and D activities and in concomitant technical supports, drug discovery strategies have become focused on drug safety screening in their early stages. Today, drug discovery efforts start from ‘molecular targeting’ based on pharmacogenomics, probably because the introduction of drug discovery research centered on gene targeting, as well as combinatorial chemistry, has realized the synthesis of many compounds over a short period. The phenomenon suggests the necessity of evaluating the toxicities of various compounds with small amounts of their samples as quickly as possible. This series of toxicological strategies is now commonly called ‘High-Throughput Toxicology’ (HTP-Tox), an essential part of toxicological study in the early phase of drug discovery. Among others, in vitro evaluation systems, particularly as screening systems, play an important role in the early phase of pharmaceutical development. They are also important to clarify the mechanisms of toxicity observed during development. On the other hand,
Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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toxicopanomics technologies (a collective designation for the ‘-omics’, such as toxicogenomics, toxicoproteomics and metabonomics) is expected to be applicable to predictive toxicology and mechanism-based risk assessment in the area of toxicology (Stubberfield and page, 1999, Pennie et al., 2000, Suter et al., 2004). Currently, toxicopanomics technologies are being applied to the development of new in vitro safety evaluation systems.
2.2
Introduction of in vitro Screening Tests
One of the advantages of in vitro tests is that direct effects on cells can be evaluated under experimental conditions, controlled artificially and strictly. Closely designed in vitro tests provide useful information to clarify toxicological mechanisms. Since the quick, efficient measurement of multiple compounds is available simultaneously under the same conditions, in vitro tests are useful for the screening or ranking of compounds. Generally, since most compounds are metabolized in the liver, using hepatotoxicity as the indicator for a screening system can trigger safety evaluation in the early phase of drug discovery. On the other hand, cells never function independently in an organism. Instead, they form close and complicated networks with each other or with the matrix of the organism to realize their functions in a three-dimensional structure. Therefore, in vitro test data should be interpreted in view of the existence form of cells in the organism and the surrounding environment, and the results obtained must be reviewed in an in vivo system. Particularly, how much the cells to be used in an in vitro test maintain normal properties significantly affects the accuracy of extrapolation into the subsequent relevant in vivo system. Although primary cells in culture and tissue slices maintain relatively normal properties, it is difficult to maintain their properties over a long period. Established cell lines keep properties stable for a long time, but it must be noted that they are inconsistent in many aspects with the properties of normal cells. There are many considerations in conducting an in vitro test, including how to allow cells to maintain normal functions and how to integrate the in vivo metabolic factors of compounds into the in vitro system environment. Anyway, this basic approach is applicable to hepatotoxicity evaluation systems as well.
2.3
Correlations between in vitro Systems and in vivo Systems
When an in vitro system is used to predict the hepatotoxicity of compounds, it is necessary to set endpoints carefully and evaluate correlations with the relevant in vivo system. While some screening systems have been reported for in vitro hepatotoxicity evaluation (Groneberg et al., 2002, Farkas et al., 2005) we report prediction of in vivo hepatotoxicity using primary rat liver cell cultures (Kikkawa et al., 2005, 2006). Four compounds known as hepatotoxicity agents were investigated, namely acetaminophen (APAP), amiodarone (AMD), tetracycline (TC) and carbon tetrachloride (CTC). To evaluate hepatotoxicity in rats, 300 or 1000 mg/kg of APAP, 300 or 1000 mg/kg of AMD, 600 or 2000 mg/kg of TC and 0.3 or 1 ml/kg of CTC were orally administered once to rats, and changes in blood biochemical parameters as well as histopathological changes were investigated 6 and 24 h after administration. For the livers of APAP-administered rats, changes in protein expression were investigated by proteomics. It was found, 24 h after
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administration, that all of the compounds had caused histopathological changes, such as inflammatory ones. Immunohistological examinations revealed the expression of oxidative stress-related proteins 6 h after administration. The analysis of changes in protein expression by proteomics detected changes in the oxidative stress-related proteins and mitochondrial metabolism-related proteins, suggesting their usefulness as hepatotoxicity evaluation markers in in vivo systems. Then, using the same APAP, AMD, TC and CTC, toxicity in primary liver cell cultures was evaluated. In investigation, morphological changes, LDH release (as the indicator of cytotoxicity) and changes in WST-1 (as the indicator of effects on mitochondrial respiration) were evaluated. Furthermore, changes in protein expression were analyzed by proteomics. It was found that all of the compounds induced morphological changes and caused a dose-dependent increase in LDH release and a decrease in WST-1. Changes in WST-1 occurred in shorter exposure times than those in LDH, suggesting that mitochondrial respiration ability would be a useful parameter for cytotoxicity in in vitro hepatotoxicity screening. The analysis of changes in protein expression by proteomics in APAP-exposed samples showed changes in the expression of oxidative stress-related proteins and mitochondrial regulation-related proteins (Figure 2.1). Thus, hepatotoxicity evaluation using primary liver cell cultures was found to be useful to predict hepatotoxicity in in vivo systems, and that the oxidative stress-related proteins
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Figure 2.1 Morphological changes of rat liver and primary rat liver cell cultures treated with APAP, AD, TC and CTC. Log EC50 values calculated from the concentration–response curves from the LDH release data and the WST-1 data after exposure to APAP, AD, TC and CTC
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and mitochondrial regulation-related proteins are useful as hepatotoxicity markers was also suggested. It is considered that future development of new biomarkers will help develop more predicable in vitro hepatotoxicity evaluation systems.
2.4 2.4.1
Application of Toxicopanomics Technologies to in vitro Safety Evaluation Systems Selection and Optimization of Evaluation Systems
Selection of cells (primary cells in culture, cell strains, tissue slices, derived tissues, etc.) and setting of culture conditions (monolayer culture, spheroid, co-culture, culture period, etc.) are important issues to be discussed in establishing in vitro evaluation systems. Panomics provide useful data to select and optimize the evaluation systems. For example, rat liver cells vary in gene expression, as shown below, depending, on cell types (primary cultured hepatocyte, BRL3A and NRLclone9 cell lines, or liver slices) and culture time (Boess et al., 2003). Among these cell types, liver slices showed the gene expression profiles closest to those of the rat liver, and primary liver cell cultures (monolayer culture or sandwich culture) also showed gene expression profiles relatively close to those of rat liver cells. However, the profiles of the cell lines used differed greatly from those of the rat liver (Figure 2.2).
Rat liver cell line (NRL) Rat liver cell line (BRL3A)
Gene expression profiles of the cell lines differed greatly from those of the rat liver
Rat liver Rat liver slice Rat primary liver cell (Monolayer) Rat primary liver cell (Sandwich)
Liver slices showed the gene expression profiles closest to those of the rat liver
Primary liver cell cultures (monolayer culture or sandwich culture) showed gene expression profiles relatively close to those of rat liver cells
Figure 2.2 Cluster analysis results for the similarity of gene expression. Modified from Boess F., Kamber M., Romer S., Gasser R., Muller D., Alberini S. and Suter L., ‘Gene expression in two hepatic cell lines, cultured primary hepatocytes and liver slices, compared to the in vivo gene expression in rats: possible implications for toxicogenomics use of in vitro systems’, Toxicological Sciences, 2003, 73(2), 386–402, by permission of Oxford University Press on behalf of the Society of Toxicology
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The gene expression profiles of liver slices and primary cell cultures changed with time, more greatly in the early phase of culture. The expression of P450-related genes decreased with time. In addition, the expression of metabolic enzyme-related genes in primary cell cultures is reported that many of the Phase-I enzyme genes decrease with time while the Phase-II enzyme genes show various changes (Baker et al., 2001). Characteristically, liver slices showed an increase of the expression of inflammation-related genes, whereas primary cells in culture showed an increase of the expression of adaptation-related genes. Thus, it has been demonstrated at the gene level that the properties of cells differ between cell types, and change depending on culture period or culture conditions. It is also known that gene expression in cells exposed to compounds varies depending on the exposure period (Harries et al., 2001). Therefore, this information on gene expression, and information on protein expression as well, are considered to be useful to select an appropriate evaluation system and optimize the system. Of course, a single in vitro system is insufficient to reproduce or predict complicated phenomena occurring through networks in the relevant in vivo system. Using panomics data in establishing evaluation system to find the features of individual systems and combine them appropriately, is expected to lead to the establishment of a high-accuracy evaluation system. Furthermore, panomics data-based accurate estimation of adaptive changes or stress-induced changes intrinsic to individual systems is considered to lead to reliable toxicity evaluation. 2.4.2
Clarification of Toxicological Mechanisms and Development of Screening Systems
Even the same hepatotoxic substance affects cells differently, depending on the compounds. For example, when exposed to a known hepatotoxic substance, HepG2 cells or primary liver cell cultures show gene expression specific to the compound exposed, suggesting its association with toxicological mechanisms (Harries et al., 2001, Burczynski et al., 2000, Waring et al., 2001). Toxicopanomics provides comprehensive information on gene expression or protein expression, thus significantly contributing to the clarification of toxicological mechanisms triggered by compounds. Moreover, establishing a database which integrates various data on the existing toxic substances and performing bioinformatics-driven analysis can help target markers dependent on toxicological mechanisms. However, this process requires an enormous amount of toxicopanomics information, collected under comparable and reproducible conditions. Therefore, collecting in vitro evaluation data obtained according to a unified protocol serves as an effective measure to establish a toxicity prediction database. Then an in vitro screening system using newly identified markers as endpoints will be developed. 2.4.3
Improvement of Extrapolation by Developing Bridging Biomarkers
Accurate extrapolation between test systems or species is a challenge in the safety evaluation of compounds. In many cases, there are quantitative differences in dose–response relationship between humans and the experimental models, and in extreme cases, even qualitative differences in biological response are expected. This reminds us of the importance of developing ‘bridging biomarkers,’ which are used to correlate toxic reactions between
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Figure 2.3
Use of bridging biomarkers to extrapolate from laboratory models to humans
test systems or species (Aardema and MacGregor, 2002) (Figure 2.3), and toxicopanomics is considered to be effective in developing new bridging biomarkers. With reliable bridging biomarkers, it is expected that the development of in vitro screening systems capable of accurately evaluating toxicity in humans or animals is feasible.
2.4.4
Improvement of Evaluation Sensitivity
Toxicity-related changes in gene expression are known to be often acuter than morphological changes (Corvi, 2002). To reduce the size and improve the throughput of an in vitro test system, new sensitive parameters must be specified in addition to the existing ones. Introduction of new sensitive parameters is also required to predict toxicity previously considered to be impossible to detect in in vitro systems. Toxicogenomics is expected to be an effective measure to meet these requirements. It is important to demonstrate marker genes and protein functions from the standpoint of toxicoproteomics, and gene functions must be established to clarify toxicological mechanisms based on genes targeted by toxicogenomics. One of the measures is to use siRNA (small interfering RNA), the effector for RNAi (RNA interference), which recently attracts attention for its medical applications. It is known that gene silencing, eventually control of protein biosynthesis, can be achieved by introducing siRNA to mammalian cells (e.g. HeLa cells) (Caplen et al., 2001, Elbashir et al., 2001). Using siRNA to ‘knock down’ specific genes paves the way for gene function analysis. Genes and proteins do not independently function, but form networks to closely cooperate in vital activities. Therefore, it is required to comprehensively detect and analyze the intracellular activities of genes and proteins. As indicated by nonlinear biological responses not always dependent on quantitative changes of proteins, not only the quantitative changes but the intracellular localization of proteins must be examined. Networks with different types of cells are often significant in the organism, and it is necessary to evaluate the development of functions via network formation. For this purpose, it is recommendable to use cell-based assay technologies (Beske and Goldbarg, 2002) or cell transfection array technologies (Ziauddin and Sabatini, 2001).
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Introduction of Evaluation Systems Using in vitro Hepatotoxicity Tests in the Early Phase of Drug Discovery Examples of Phospholipidosis Evaluation
Phospholipidosis, a pathological condition with phospholipids excessively accumulated in cells, often impedes development in drug discovery. Examples of predicting induction of phospholipidosis based on the evaluation on an in vitro system and the physical properties are presented below. The in vitro experiment was performed based on a method reported by Ulrich et al. (1991), with modification. Primary rat cell cultures were added with compounds and a fluorescent-labeled lipid vesicle, and 24 h later, intake of the fluorescent dye into the cells was observed under the fluorescence microscope. Evaluation with ten compounds known for their ability to induce phospholipidosis and six ‘negative’ compounds yielded results correlating with their in vivo induction – thus, the usefulness of the in vitro system was confirmed (Figure 2.4). Development of ‘in-silico’ prediction systems is also given priority in studies to discover new drugs. A system using two physical property values (ClogP and net charge of a given molecule) is introduced for the prediction system of phospholipidosis (Tomizawa et al., 2006). It is proved that with the two parameters, the system can predict in vitro results with accuracy. 2.5.2
Examples of Biomarker Development
In the hepatotoxicity evaluation of compounds on an in vitro screening system, it is essential to set biomarkers well reflecting in vivo toxicity. Examples of developing hepatotoxicity biomarkers by means of proteomics (Yamamoto et al., 2005, 2006) and metabonomics technologies (Yamamoto et al., 2006) are explained below. The development of hepatotoxicity biomarkers by proteomics technologies used four compounds which induce hepatotoxicity via different mechanisms: acetaminophen (APAP), amiodarone (AMD), tetracycline (TC) and carbon tetrachloride (CTC). To investigate changes in protein expression in the rat liver, 1000 mg/kg of APAP, 1000 mg/kg of AMD, 2000 mg/kg of TC and 1 ml/kg of CTC were orally administered once to rats, and the liver was extracted 24 h later. To investigate changes in protein expression in primary rat liver cell cultures, 10 mM of APAP, 50μM of AMD and 500μM of TC were exposed to cells for 24 h. These samples were analyzed for changes in protein expression by 2-dimensional electrophoresis and mass spectrometry. In the rat liver, eight proteins whose expression commonly changed in response to the administration of the four compounds were identified (2-oxoisovalerate dehydrogenase, 60k Da ‘heat-shock’ protein, adenylate dehydrogenase-4, carbonic anhydrase-3, glutamate dehydrogenase, NADP-dependent malic enzyme, NDRG1-related protein NDRG2b1 and serotransferrin) (Figure 2.5(b)). In the primary rat liver cell cultures, three proteins whose expression commonly changed in response to the exposure to the three compounds were identified (glutathione peroxidase, peroxiredoxin 1, and peroxiredoxin) (Figure 2.5(a)). These proteins are expected to be used as hepatotoxicity biomarkers in in vivo or in vitro evaluation systems. Furthermore, the expression of three proteins (triosephosphate, glutamate dehydrogenase and beta-actin) in APAP, three proteins (fuructose-bisphosphate aldolase B, serotransferrin and short-chain
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Figure 2.4 (a) Fluorescence microscopy images, toluidine blue staining and transmission electron microscopy (TEM) images of primary rat liver cell cultures treated with DMSO (1 %, vehicle control) and amiodarone (2 μg/ml). The fluorescence-labeled lipid appears as fluorescent droplets surrounding the nuclei. Accumulation of dense inclusion bodies were induced by amiodarone (toluene blue staining, ‘black arrow’), while the inclusion bodies formed distinct lamellar structures (EM, ‘white arrow’: ×1800). (b) Correlations between in vitro and in vivo evaluation systems for phospholipidosis
3-hydroxyacyl-CoA dehydrogenase) in AMD, three proteins (plasma retinol-binding protein, serotransferrin and HMG-CoA synthase) in TC and thirteen proteins (elongation factor 1-γ , catalase, senescence marker protein-30, liver carboxylesterase 4, peroxiredoxin 2, d-dopachrome tautomerase, acyl-CoA dehydrogenase (long-chain specific), glutamate dehydrogenase, phenylalanine-4-hydroxylase, glycogen phosphorylase (liver form), pyruvate kinase isoenzymes R/L, hydroxymethylglutaryl-CoA synthase and glutathione peroxidase) in CTC changed commonly between in vivo and in vitro (Figure 2.5(c)). As reliable markers for in vivo hepatotoxicity, these proteins may be used in in vitro hepatotoxicity evaluation systems. In the development of hepatotoxicity biomarkers by metabonomics technologies, 1400 mg/kg of APAP was orally administered once to human hepatocyte transplanted chimeric
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Figure 2.5 Venn diagrams showing the number of differentially expressed spots after treatment with APAP, AMD, TC and CTC: (a) protein expression changes in primary rat liver cell cultures; (b) protein expression changes in rat liver; (c) protein expression changes in primary rat liver cell culture and rat liver
mice, and intravesical urine and serum, collected 1, 4 and 24 h later, were used as samples. Intrinsic metabolites in the urine and serum samples were measured using 600 MHz NMR spectroscopy and metabolite changes were analyzed by spectral principal component analysis. The results showed changes in three metabolites in the urine (Figure 2.6) and serum. These metabolites had been produced by APAP administration and therefore may be used as biomarkers for hepatotoxicity induced by APAP. They may also be used as biomarkers for APAP-induced hepatotoxicity in humans because the animals used in the experiment were a ‘human model’.
2.6
Future Prospects
Focused on reduction in size and improvement of throughput, development of in vitro screening systems has progressed with the aim of collecting single endpoint information for many compounds, efficiently over a short period of time. Such a method can no doubt evaluate an aspect of toxicological profiles of a particular compound group highly efficiently. For high-quality evaluation in view of the complexity of toxicological mechanisms, however, multilateral evaluation tests must be performed for multiple intracellular parameters to be detected simultaneously. From this standpoint, cell-based assays, whose recent technological advancement is remarkable, will be more demanded to be applied to
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Urinary metabolite changes in APAP-administered mice
screening systems, as well as the toxicological mechanism clarification systems mentioned above. To clarify toxicological mechanisms in an in vitro system, technologies which can freely control the networks surrounding target cells, such as the cell–cell or cell–matrix communication environment, and technologies to comprehensively detect intracellular changes are required. For this purpose, development of new technologies by integrating cell-based assay systems with microfabrication techniques and even with nanotechnologies is desired. ‘On-chip’ single-cell analysis systems are considered to be an example of technologies representing this approach (Inoue et al., 2001). These systems are designed to control cell–cell communications under certain conditions and analyze the functions and relations of cells, which are contained in small culture spaces with 20–50 μm in diameter and 5–10 μm in depth on the chip surface. As comprehensive analyses of genes or proteins became available, bioinformatics (information-processing technologies) has long been expected to grow to interpret floods of data created by those analyses. Bioinformatics is currently under energetic study and development as technologies to select and compile data characteristic of specific life phenomena. However, trends toward the next generation have already been found, i.e. system biology,
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a novel study area aimed to understand life phenomena as a system (Kitano, 2001), is attracting attention. With the progress of development and implementation of new technologies represented by panomics, understanding of components ‘constructing’ life, such as genes and proteins has rapidly advanced. System biology is intended to comprehend such information in the dynamics of life phenomena. Results from studies on system biology will assume important roles in developing simulation models (e.g. E-cell (Tomita et al., 1999)) and analyzing life behaviors (including pathological conditions). The area of toxicology will also greatly benefit from system biology, which is expected to grow into ‘system toxicology’. In studies on system biology, in vitro experiments are still an essential process. Cellbased assay systems capable of comprehensive analysis seem to be needed to demonstrate life phenomena (hypotheses) simulated by different methods. As described above, while development of in vitro safety evaluation systems is dramatically advancing, information has become ‘batch-processed’ with the advent of highthroughput and comprehensive analysis systems. In the future, even ‘in-silico systems’ simulating life phenomena and automatically analyzing life behaviors will be developed. Toxicologists can benefit from such novel technologies. Now that various new tools are available to researchers, their ability and sensitivity to achieve extensive, higher-quality toxicity evaluation utilizing those tools will be tested.
References Aardema M. J. and MacGregor J. T. (2002). Toxicology and genetic toxicology in the new era of ‘toxicogenomics’: impact of ‘-omics’ technologies. Mutat Res 499: 13–25. Baker T. K., Carfagna M. A., Gao H., Dow E. R., Li Q., Searfoss G. H. and Ryan T. P. (2001). Temporal gene expression analysis of monolayer cultured rat hepatocytes. Chem Res Toxicol 14: 1218–1231. Beske O. E. and Goldbard S. (2002). High-throughput cell analysis using multiplexed array technologies. Drug Discov Today 7: S131–S135. Boess F., Kamber M., Romer S., Gasser R., Muller D., Alberini S. and Suter L. (2003). Gene expression in two hepatic cell lines, cultured primary hepatocytes and liver slices compared to the in vivo liver gene expression in rats: possible implications for toxicogenomics use of in vitro systems. Toxicol Sci 73: 386–402. Burczynski M. E., McMillian M., Ciervo J., Li L., Parker J. B., Dunn II R. T., Hicken S., Farr S. and Johnson M. D. (2000). Toxicogenomics-based discrimination of toxic mechanism in HepG2 human hepatoma cells. Toxicol Sci 58: 399–415. Caplen N. J., Parrish S., Imani F., Fire A. and Morgan R. A. (2001). Specific inhibition of gene expression by small double-stranded RNAs in invertebrate and vertebtate systems. Proc Natl Acad Sci USA 98: 9742–9747. Corvi R. (2002). Genomics: an in vitro toxicology point of view. ATLA 30: 129–131. Elbashir S. M., Harborth J., Lendeckel W., Yalcin A., Weber K. and Tuschl T. (2001). Duplexes of 21-nucleotide RNAs mediate RNA interference in cultured mammalian cells. Nature 411: 494–498. Farkas D. and Tannenbaum S. R. (2005). In vitro methods to study chemically induced hepatotoxicoty: a literature review. Curr Drug Metab 6: 111–125. Groneberg D. A., Grosse-Siestrup C. and Fischer A. (2002). In Vitro Models to Study Hepatotoxicity. Toxicol Pathol 30: 394–399. Harries H. M., Fletcher S. T., Duggan C. M. and Baker VA. (2001). The use of genomics technology to investigate gene expression changes in cultured human liver cells. Toxicol in vitro 15: 399–405.
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Inoue I., Wakamoto Y., Moriguchi H., Okano K. and Yasuda K. (2001). On-chip culture system for observation of isolated individual cells. Lab Chip 1: 50–55. Kikkawa R., Yamamoto T., Fukushima T., Yamada H. and Horii I. (2005). Investigation of a hepatotoxicity screening system in primary cell cultures – ‘What biomarkers would need to be addressed to estimate toxicity in conventional and new approaches?’. J Toxicol Sci 30: 61–72. Kikkawa R., Fujikawa M., Yamamoto T., Hamada Y., Yamada H. and Horii I. (2006). In vivo hepatotoxicity study of rats in comparison with in vitro hepatotoxicity screening system. J Toxicol Sci 31: 23–34. Kitano H. (2002). System biology: a brief overview. Science 295: 1662–1664. Pennie W. D., Tugwood J. D., Oliver G. J. A. and Kimber I. (2000). The principles and practice of toxicogenomics: Application and opportunities. Toxicol Sci 54: 277–283. Stubberfield C. R. and Page M. J. (1999). Applying proteomics to drug discovery. Exp Opin Invest Drugs 8: 65–70. Suter L., Babiss L. E. and Wheeldon E. B. (2004). Toxigenomics in predictive toxicology in drug development. Chem Biol 11: 161–171. Tateno C., Yoshizane Y., Saito N., Kataoka M., Utoh R., Yamasaki C., Tachibana A., Soeno Y., Asahina K., Hino H., Asahara T., Yokoi T. and Furukawa T. and Yoshizato K. (2004). Near Completely Humanized Liver in Mice Shows Human-Type Metabolic Responses to Drugs. Am J Pathol 165: 901–912. Tomita M., Hashimoto K., Takahashi K., Shimizu T. S., Matsuzaki Y., Miyoshi F., Saito K., Tanida S., Yugi K., Venter J. C. and Hutchison III C. A. (1999). E-CELL: software environment for whole-cell simulation. Bioinformatics 15: 72–84. Tomizawa K., Sugano K., Yamada H. and Horii I. (2006). Physicochemical and cell-based approach for early screening of phospholipidosis inducing potential. J Toxicol Sci 31 r r r– r r r r. Ulrich R. G., Kilgore K. S., Sun E. L., Cramer C. T. and Ginsberg L. C. 1991. An in vitro fluorescence assay for the detection of drug-induced cytoplasmic lamellar bodies Toxicol Meth 1: 89–105. Waring J. F., Ciurlionis R., Jolly R. A., Heindel M. and Ulrich R. G. (2001). Microarray analysis of hepatotoxins in vitro reveals a correlation between gene expression profiles and mechanisms of toxicity. Toxicol Lett 120: 359–368. Yamamoto T., Kikkawa R., Yamada H. and Horii I. (2005). Identification of oxidative stress-related proteins for predictive screening of hepatotoxicity using a proteomic approach. J Toxicol Sci 30: 213–227. Yamamoto T., Kikkawa R., Yamada H. and Horii I. (2006). Investigation of proteomic biomarkers in vivo hepatotoxicity study of rat liver: Toxicityy differentiation in hepatotoxicants. J Toxicol Sci 31: 49–60. Ziauddin J. and Sabatini D. M. (2001). Microarrays of cells expressing defined cDNAs. Nature 411: 107–110.
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3 Use of Hepatocytes for Characterizing a Candidate Drug’s Metabolism and Drug Interaction Potential Srikanth C. Nallani, John M. Strong and Shiew Mei Huang
3.1
Introduction
Drug-induced liver injury (DILI) is a major health problem which can lead to liver failure resulting in liver transplants or death. It is the most frequent reason for withdrawal of drugs from the pharmaceutical market (Temple and Himmel, 2002) and for labeling drugs with a ‘black box warning’ (Fung et al., 2001). Based on the seriousness of this problem, the FDA recently formed a working group along with representatives from the Pharmaceutical and Research Manufacturers of America (PhRMA) the American Association for the Study of Liver Disease (AASLD) and the National Institute of Diabetes and Digestive and Kidney Diseases to provide a forum to discuss pre-clinical, clinical and post-marketing approaches related to minimizing the risk of DILI. The working group initially sponsored a two-day Heptatoxicity Workshop in February 2001. Since that time the working group has met annually to discuss progress towards realizing its goal of minimizing DILI. The FDA working group maintains a website at www.fda.gov/cder/livertox/, which includes preclinical, clinical, and post-marketing white papers and presentations by participants at the working group meetings. Although a few drugs are excreted unchanged into the bile or urine, most drugs are cleared from the body partially or extensively by drug metabolizing enzymes found mainly in the liver or gut mucosa. Drug metabolism can proceed through detoxification pathways leading to elimination of the drug from the body, or in some cases through intoxification pathways resulting in toxicity (Figure 3.1). Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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Hepatotoxicity Drug Detoxification pathways
Elimination
Figure 3.1
Intoxification pathways
Toxicity
Detoxification pathways
Metabolic pathways leading to drug elimination from the body or to hepatotoxicity
The balance between these two pathways in humans can be affected by numerous factors, such as genetic polymorphisms of drug-metabolizing enzymes, environmental, disease state, sex, age and drug–drug interactions. Although some of these factors can be identified and are relatively stable with time, others such as concomitant medications can alter metabolism abruptly through drug-interactions such as inhibition or induction of drugmetabolizing enzymes and are of particular concern. Drug–drug interactions can lead to changed systemic exposure, to the co-administered drugs or their metabolites, resulting in variations in drug response, including adverse drug reactions and efficacy. The relationship between drug metabolism and adverse events, including hepatotoxicity, cannot be overemphasized. This is illustrated by the fact that the FDA has recently published a number of Guidances for Industry addressing this issue (e.g. ‘Guidance for Industry: Drug Interaction Studies – Study Design, Data Analysis and Imapct for Dosing and Labeling’ (FDA, 2006) and ‘Guidance for Industry: Safety Testing of Drug Metabolites’ (FDA, 2005) and a website containing information on drug–drug interactions (http://internet-dev/ cder/drug/drugInteractions/). It is obviously most desirable that the hepatotoxic potential of drugs be discovered as early as possible during pre-clinical screening and specific human testing. When developing strategies to deal with metabolites and their potential role in hepatotoxicity, Guenguerich (2006), Smith and Obach (2005) and Liebler and Guengerich (2005), respectively, have proposed similar definitions of mechanistic categories of toxicity that could be caused by the parent drug or metabolites which should be considered. Their general classifications include: r parent drug or slightly modified metabolite that elicits its effect on the intended target; r parent drug or metabolite that binds to a receptor or enzyme that is not the primary target; r parent drug or reactive metabolite covalently binds to protein that lead to autoimmune responses; r drug bioactivated to reactive compounds that can cause oxidative stress through redox cycling or reactive metabolites, usually electrophiles, that can covalently bind to critical cellular proteins leading to cell death. Traditionally, drug metabolites in general have not been routinely evaluated in crossspecies safety assessments because their specific contribution to the overall toxicological potential of the parent drug has been unknown. In the past, measurements of circulating concentrations of a parent drug in animals were used as an index of systemic exposure in
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humans. Animal toxicology studies are an important component of assessing drug safety for subsequent human exposure; however, recently there has been a clear trend towards decreased reliance totally upon animal studies and increased emphasis upon experiments with human-derived tissues.
3.2
Metabolic Pathway Identification in Animals and Humans in vitro
Studies in laboratory animals do not always reflect human drug toxicity because of interspecies differences. It has been reported that hepatotoxicity has one of the poorest correlation with regulatory animal toxicity tests (Olson et al., 2000) and that only approximately half of the new pharmaceuticals that produced hepatotoxicity in clinical drug development were detected during animal toxicity studies (Olson et al., 2000, 1998). Metabolites may not be adequately assessed during standard nonclinical studies because they occur only in humans (unique metabolite) or they are produced at much higher levels than in animals. The Agency recommends that attempts be made to identify as early as possible during the drugdevelopment process differences in drug metabolism in animals used in nonclinical safety assessments compared to humans. (FDA, 2005). Studies using human and animal liver tissue, including freshly isolated liver slices or hepatocytes and cryopreserved hepatocytes, to identify a drug’s metabolic pathways in both species are generally conducted before initiation of clinical trials. These intact cellular models contain the complete complement of liver enzymes and cofactors as will as the natural orientation for linked enzymes. This type of study can provide information to prospectively design experiments so that the animal species most similar to humans can be chosen, on a case-by-case basis, for each drug and help in the interpretation of data obtained from animal toxicology experiments. The FDA recommends that toxicological testing in animals should be performed for unique human metabolites and those that represent >10 % of the dose (FDA, 2005).
3.3
Characterizing a Candidate Drug’s Metabolic Pathways Using Human Hepatocytes
Identification of molecules with acceptable metabolic profiles early in the drug-development process is crucial for the success of drug-candidate development. Prediction of in vivo pharmacokinetics before human drug administration based on various in vitro systems is a common practice in the pharmaceutical industry. These systems include human liver slices, freshly isolated, cryopreserved hepatocytes and hepatocytes in culture, and subcellular liver tissue, such as microsomes or post-mitochondrial fractions (S9). Freshly prepared or cryopreserved hepatocytes in suspension are the most commonly used tissue for characterizing a drug’s metabolic pathways; however, due to the loss of cell viability, these experiments are limited to 4–6 h (Gomez-Lechon et al., 2004). Hepatocyte cultures can be beneficial for investigating the metabolic stability of drugs over longer periods of time to investigate secondary, tertiary, etc. metabolites which may not be detected in short-term incubations (Zhao et al., 2005). Hepatocytes in culture are also the method of choice for CYP enzyme induction studies.
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It can be reasonably considered that human hepatocytes reflect the heterogeneity of CYP (CYPs 1A2, CY2A6, 2B6, 2C8, 2C9, 2C19, 2D6, 2E1 and 3A4) expression in human liver. The relative contribution of specific CYP activities in human hepatocytes and human liver microsomes was discussed previously (Gomez-Lechon et al., 2004). The good reflection of hepatic CYP enzyme pattern of hepatocytes as compared to human microsomes constitutes an invaluable tool for preclinical metabolic and toxicity testing of candidate drugs. Approaches for studying other non-CYP enzymes, such as flavin-containing monooxygenase (FMOs), monamine oxidases (MAOs), molybdenum-containing oxidases (Mo–CO), and transferase reactions such as UDP-glucuronosyl-transferases (UGTS), and sulfotransferases (STs), methyltransferases, acetyltransferases and glutathione transferases are evolving. For example, Bjorhnsson et al. (2003) discusses methods to differentiate between CYP and non-CYP oxidative metabolites. Metabolites formed from transferase reactions can be identified by their molecular structure. 3.3.1
Hepatocyte Characteristics and Considerations for Experimental Conditions
Hepatocyte cultures in vitro are known for cell ‘de-differentiation’ which is accompanied by the loss of some specific enzymes, especially cytochrome P450 monooxygenases, affecting the metabolic capacity of the system (Morel et al., 1990; Skett et al., 1995). The functional relevance of an in vivo-like extracellular matrix geometry for the biotransformation of primary hepatocytes in vitro was studied by using a variety of drugs (Bader et al., 1995). Tissue architecture and parenchymal cell morphology reflect the functional differentiation of the liver. Hepatocytes are arranged as monolayer plates in vivo, an organization which allows the cells to be highly vascularized. Sandwiching hepatocytes between two layers of collagen has shown to provide the cells with an in vivo-like environment that enables the cells to preserve their specific functionality for several days (Berthiaume et al., 1996). This observation was further substantiated by Kern et al. (1997) who showed that hepatocyte sandwich cultures not only retained their long-term viability but also their drug-metabolizing capacity. Hepatocytes in suspension offer a simple and cost-effective method to study a drug’s metabolic stability and to identify its overall metabolic pathways and can be automated for screening purposes during early drug development They can be procured from several commercial sources which include cell viability data and instructions for performing drug incubations with their product. Cryopreserved hepatocytes are particularly attractive because they are readily available as ‘off the shelf items’ and one does not have to wait for a donor liver to perform their study; however, one should be aware that drug-metabolizing CYP, glutathione-S-transferase and sulfotransferase enzyme activity has been reported to be less when compared to freshly isolated hepatocytes. On the other hand, UDP-glucoronosyl transferase activity is unaltered by cryopreservation (Hengstler et al., 2000). Damage to the cell membrane while thawing cryopreserved hepatocytes and the resulting reduction in the levels of cofactors such as NADPH might partially explain the loss of CYP enzyme activity. Some commonly used experimental conditions for these types of hepatocyte short term experiments are listed in Table 3.1. Following incubation, the samples can be analyzed for metabolites in the medium and cells separately or following disruption of the cells as the total metabolic content in the incubation. The most preferred method of analysis is by liquid chromatography combined with mass spectrometry/mass spectrometry (LC–MS/MS), a highly sensitive and specific technique which can provide both qualitative structural information and quantification
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Table 3.1 Typical experimental conditions for metabolic characterization of drug candidates in hepatocytes Experimental condition Incubation time and medium Hepatocyte Concentration Incubation vial
Incubation volume Incubation conditions
Current practice and rationale For ease of operation, reactions for ≤ 3 h are conducted in balanced salt solution; ≤ 6 h in cell culture medium 1–5 × 106 viable cells/ml provide optimal cell viability and metabolic capacity, low cellular aggregation For qualitative and quantitative metabolic stability, reactions can be carried out in varied capacity: –centrifuge (glass) tubes (<2 h) –polystyrene microtiter plates/petridishes/flasks For adequate oxygenation and reaction volume: 50–100 μl in 96-well plate, 250 μl in 24-well plate, ≤1 ml in 5 ml vial, 5–15 ml in petridishes/flasks –37 ◦ C, 95 % relative humidity; 95 % air/5 % CO2 to simulate physiological conditions –Gentle shaking (90 rpm) to inhibit cell aggregation and adherence
of metabolites identified. If a radiolabeled drug is available, an alternative method is to incubate the radiolabeled drug with the hepatocytes and analyze the incubation mixture by high performance liquid chromatography (HPLC) in combination with a radiochemical detector. A major disadvantage of this alternate technique is that no metabolite structural information is obtained. Before initiating these studies, it is advisable to establish the cell viability using standard biomarkers such as ATP content, neutral red uptake, tetrazolium salt MTT or MTS analysis, etc., as well as CYP metabolizing enzyme characterization. By virtue of extensive characterization using human liver microsomes over the years, selective and specific substrates for CYP enzymes have been identified for characterizing the drug-metabolizing enzyme activity in human hepatocytes. A list of preferable and acceptable chemical substrates for this purpose is given in Table 3.2. If human in vivo data or in vitro metabolic pathway identification studies indicate CYP enzymes contribute > 25 % of a drug’s clearance, studies to identify drug-metabolizing CYP enzyme in vitro are recommended. These latter studies investigate the individual CYP enzymes responsible for each of the metabolites formed and identified using human hepatocytes. These studies use a combination of techniques including human liver tissues, such as post-mitochondrial fraction (S9), cytosolic fraction and microsomes. A detailed discussion of the methods and techniques used for the latter studies are beyond the scope of this chapter and the reader is referred to the following reference for the further information (Bjornsson et al., 2003).
3.4
Prediction of in vivo Clearance of Candidate Drugs
The metabolic stability data obtained from the human hepatocyte studies described in the previous section provides important information if performed in the early stages of drug
1–14 52–94
Midazolam 1-hydroxylation Testosterone 6β-hydroxylation
3A4/5
a
P-Nitrophenol 3-hydroxylation Lauric acid 11-hydroxylation Aniline 4-hydroxylation Erythromycin N-demethylation Dextromethorphan N-demethylation Triazolam 4-hydroxylation Terfenadine C-hydroxylation Nifedipine oxidation
Omeprazole 5-hydroxylation Fluoxetine O-dealkylation Debrisoquine 4-hydroxylation
Note that this is not an exhaustive list. For an updated list, see the following link: http://www.fda.gov/cder/drug/drugInteractions/default.htm
2E1
9–15 0.44–8.5 39–157
(±)-Bufuralol 1’-hydroxylation Dextromethorphan O-demethylation Chlorzoxazone 6-hydroxylation
67–838 1.5–4.5 3.4–52 13–35
2D6
2C19
2C9
2C8
0.30–2.3 13–162 17–23 67–168 5.4–19
7-ethoxyresorufin-O-deethylation theophylline-N-demethylation caffeine-3-N-demethylation tacrine 1-hydroxylation — — Propofol hydroxylation S-mephenytoin-N-demethylation Amodiaquine N-deethylation Rosiglitazone para-hydroxylation Flurbiprofen 4’-hydroxylation Phenytoin-4-hydroxylation
3.3 130 6.3–24 33–88 133–710 234 15 5.1–47
17–26 3.7–104 5.6
0.18–0.21 280–1230 220–1565 2.8, 16 — — 3.7–94 1910 2.4 4.3–7.7 6–42 11.5–117
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Coumarin-7-hydroxylation Nicotine C-oxidation Efavirenz hydroxylase Bupropion-hydroxylation Taxol 6-hydroxylation
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Preferred and acceptable chemical substrates for in vitro experimentsa
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development. The ability to classify the metabolic clearance of a drug as low, intermediate or high is useful when prioritizing further development of candidate drugs. Once the individual metabolites formed by human hepatocytes are identified and the contribution of each metabolite to the overall in vitro clearance of the drug is determined, the total intrinsic drug clearance in vitro, CL,i, is obtained by using the following equation: CL, i =
j n Vmax j
j=1
Km
(3.1)
where Vmax is the maximal velocity of the metabolic reaction, K m is the Michaelis–Menten constant and j represents the number of metabolites quantified in the study. Assuming that the clearance of the drug is independent of the blood flow rate through the clearing organ, in this case the liver, one can predict the in vivo intrinsic clearance of the drug using the simple well-stirred model (Pang and Rowland, 1977) by employing the following equation: CLin vivo =
fuCL, iQ H fuCL, i + Q H
(3.2)
where fu = fraction unbound in plasma and Q H = hepatic blood flow (Davies and Morris, 1993). Scaling of the in vitro intrinsic clearances, CL, i, for calculation of CLin vivo can be accomplished by assuming that approximately 120 million cells are found in 1 g of liver and that the total liver weight is about 25 gm/kg of body weight with an average blood flow of about 21 ml/min/kg (Davies and Morris, 1993). In the past, human liver microsomes or the S9 fractions have been used for this purpose of metabolic stability evaluation. Recently the use of fresh or cryopreserved hepatocytes is gaining acceptance. For example, clearance of substrates by fresh or cryopreserved human hepatocytes via CYP enzyme and UGT transferase reactions in vitro have been used to predict the CLin vivo for these enzymes (Soars et al., 2002). The extent of ‘underprediction’ of CLin vivo was shown to be less that those determined using human liver microsomes. In another study designed to establish the effect of product inhibition of CYP enzymes by a candidate drug’s metabolites, the results indicated that microsomes predicted a lower C L in vivo compared to that predicted using heptatocytes (Naritomi et al., 2003).
3.5
Characterizing Drug Metabolizing Enzyme Iinduction-Potential of Candidate Drug
The average percentage of individual CYP enzymes have been characterized in human liver and hepatocytes by quantifying CYP mRNA expression, protein and enzyme activity. The basal expression and activity of CYP and non-CYP drug-metabolizing enzymes is highly variable between batches of hepatocytes isolated from different donors. In addition, the expression of individual enzymes decreases with time at variable rates during incubation of primary cultures of hepatocytes. 3.5.1
Regulation of Drug Metabolizing Enzyme Expression in Hepatocytes
Many of the drug-metabolizing enzymes and drug transporters can be induced after exposure to specific xenobiotics. Over the past decade, our understanding of mechanisms that
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regulate the gene expression of these drug-metabolizing enzymes and transporters in hepatocytes has greatly expanded. Nuclear receptors, including the aryl hydrocarbon receptor (AhR), pregnane-X-receptor (PXR) or steroid and xenobiotic receptor (SXR), the constitutive androstane receptor (CAR) and peroxisome proliferator activated receptor (PPAR), play a key role in this process (Kliewer et al., 1999). Table 3.3 presents a partial list of drugs (and chemicals) that interact with nuclear receptors expressed in different organs
Table 3.3 A partial list of drugs (and chemicals) that interact with nuclear receptors expressed in different organs known to regulate the expression of target drug metabolizing enzyme and drug transporter genes (Tirona and Kim, 2005; Klaassen and Slitt, 2005; Chang and Waxman, 2006) Nuclear receptor (NR)
Tissuerxpressing NR
Target gene of the NR
β-Naphtoflavone, 2,3,7,8tetrachlorodibenzo-pdioxin
Aryl hydrocarbon receptor (AhR)
Acetaminophen, Phenobarbital, phenytoin, TCPOBOP
Constitutive androstane receptor (CAR)
Lung Thymus Kidney Liver Liver
Clofibrate, ciprofibrate
Peroxisome proliferator-activated receptor α (PPARα)
Liver Heart Muscle Kidney
Clotrimazole, dexamethasone, paclitaxel, phenobarbital, ritonavir, rifampicin
Pregnane X receptor (PXR)
Liver Small intestine Colon Lymphocytes
CYP1A1 CYP1A2 UGT1A1 UGT1A6 CYP2A6 CYP2B6 CYP2C9 CY2C19 CYP3A4 Sult1a1 Sult2a1 Sult2a9 UGT1A1 Mrp2 Mrp3 Mrp5 Mdr1 CYP4A UGT1A9 UGT2B4 Mdr2 Oatp1 CYP1A2 CYP2B6 CYP2C9 CYP2C19 CYP3A4 CYP3A7 SULT2A1 UGT1A1 UGT1A3 UGT1A4 Mdr1 Mrp2 Mrp3
Drug/chemical
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known to regulate the expression of target drug metabolizing enzyme and drug transporter genes. It is important to note that in some cases ‘cross-reactivity’ exists between individual nuclear receptors. For example, phenobarbital induces CYP2B and CYP3A mainly through activation of CAR, whereas dexamethasone and rifampin induce CYP2C and CYP3A enzymes by means of their interaction with PXR (Moore et al., 2000). UDP-glucuronosyl transferases and sulfotransferases are involved in the detoxification and ultimate excretion of a wide variety of structurally diverse endo- and xenobiotics, including many therapeutic agents and endogenous steroids. Emerging evidence strongly suggests that the nuclear receptors PXR, CAR, and PPAR can also regulate these transferases (Sugatani et al., 2005; Runge-Morris and Kocarek, 2005). In general, drug transporters such as the ‘organic anion transporting polypeptide family’ (OATPs) along with ‘organic cation transporter 1’ (OCT1) and ‘organic anion transporter 2’ (OAT2) mediate uptake of a large number of xenoblotics from blood into liver. Conversely, ‘multi-drug resistance proteins’ (MDRs), ‘multi-drug resistance associated proteins’ (MRPs) and the ‘breast cancer resistance protein’ (BCRP) mediate efflux of xenobiotics from liver into bile or blood. The nuclear receptors PXR and CAR have been shown to be involved in the regulation of these transporters (Tirona and Kim, 2005). Coordinate regulation of Phase I and Phase II enzyme induction, along with drug transporter induction, has been reported with the treatment of prototypical enzyme inducers. Taken together, induction of the drug metabolism and drug transporting enzymes in a coordinated fashion provides an important means to protect the body from xenobiotics ‘insults’. A drug that induces a specific drug-metabolizing enzyme can increase the rate of metabolic clearance of a co-administered drug that is a substrate of the induced pathway, resulting in sub-therapeutic blood concentrations. Alternatively, the induced metabolic pathway could lead to increased formation of a bioactivated compound with the potential to produce liver toxicity, e.g. acetaminophen conversion to the putative toxin, N-acetyl-pbenzoquinoneimine. 3.5.2
Experimental Considerations for Evaluating CYP Induction in Human Hepatocytes
Presently, the most reliable method to study a drug’s induction potential is accomplished by using freshly isolated human hepatocytes in culture or cryopreserved hepatocytes that are capable of attachment and incubation in culture. An overview of a typical procedure for conducting an induction experiment is described below; however, more detailed information can be found in a review article (LeCluyse, 2001). Initially, hepatocytes at a density of 100 000–200 000 cells/cm2 are placed on multiwell plates coated with a matrix such as r collagen or Matrigel and allowed to attach for 2–4 h. It is recommended that monolayer cultures or cells with medium are replaced and overlayed (sandwiched) with collagen or r Matrigel to provide a 3-dimensional environment and to minimize variability in cellular integrity. Hepatocytes are then incubated for 2–3 days with medium replacement every 24 h in order to stabilize the hepatocytes. The incubation medium is not critical and several products, such as Chee’s Medium, William’s medium E, Ham’s and F-12/Lebovitz L-15 (1:1), are commercially available. Customized media that include the following supplements, 6 μg/ml insulin, 6 μg/ml transferrin, 6 ng/ml selenium, 2 mg/ml BSA/linoleic acid complex and a glucocorticoid such as dexamethasone (25–100 ng/ml excessive dexamethasone
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Table 3.4 Chemical inducers for in vitro experimentsa
CYP Inducer—preferred 1A2
Omeprazole ß-Naphthoflavone 3-Methylcholanthrene
2A6
Dexamethasone
2B6
Phenobarbital
2C8 2C9
Inducer concentrations Fold Inducer – (μM) induction acceptable 25–100 33–50 1, 2
14–24 4–23 6–26
Lansoprazole
10
10
50
9.4
1000
7.7
500–1000
5–10
Phenytoin
50
5–10
Rifampin
10
2–4
Phenobarbital
500
2–3
Rifampin
10
3.7
Phenobarbital
100
2.6
2C19 Rifampin
10
20
—
—
—
2D6 None identified
—
—
—
—
—
2E1
None identified
3A4
Rifampin
Pyrazole
Inducer concentration Fold (μM) induction
—
—
—
—
—
10–50
4–31
phenobarbital Phenytoin Rifapentine Troglitazone Taxol Dexamethasone
100–2000 50 50 10–75 4 33–250
3–31 12.5 9.3 7 5.2 2.9–6.9
a
Note that this is not an exhaustive list. For an updated list, see the following link: [http://www.fda.gov/cder/drug/ drugInteractions/default.htm].
can induce enzymes), should be sufficient to ensure maintenance of monolayer integrity and hepatocyte morphology for at least 1 week and give reliable results with positive controls. Additional additives, specific for sugar metabolism, amino acids, antioxidants, hormones and heavy metals, are also acceptable in an attempt to improve cellular maintenance. After the hepatocytes have stabilized, the wells containing hepatocytes are divided into three sets and incubated for 2–3 days with medium replacement every 24 h in the presence of (set 1) test drug, (set 2) a known inducer drug listed in Table 3.4 as a positive control and (set 3) no drug and used as a negative control. Inducer-drug concentrations should be utilized, based on the expected human plasma drug concentrations. At least three test drug concentrations spanning the therapeutic range should be studied. If this information is not available, concentrations ranging over at least three orders of magnitude should be studied since the induction response may not be linear with concentration. To assess the integrity of the hepatocyte culture following drug treatment, the viability of hepatocytes should be assessed by visual inspection or by cell-viability assays such as MTT or MTS assays, LDH assay, ATP assay, etc. Following treatment of the three sets of hepatocytes, the medium is replaced and a test probe substrate (Table 3.2), is incubated with the hepatocytes or microsomes prepared from the individual hepatocytes incubations to determine the relative enzyme activity of untreated and treated cells. When conducting the experiments to determine enzyme activity, the following experimental conditions are relevant: microsomal protein concentrations, if used, usually range from 0.01 to 2.0 mg/ml. Typically, the incubation time ranges from 5 to
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30 min. Preferably, <10 % substrate depletion should occur during the incubation time. The relationship between time and amount of product formed should be linear. Solvents typically used to prepare the stock probe substrate solutions, including methanol, ethanol, acetone, acetonitrile and dimethyl sulfoxide, should represent <1 vol% of the incubation medium. When analyzing the results of experiments to determine enzyme activity, the following issues are relevant: (a) Drug probe enzyme activities determined using hepatocytes should be reported as metabolite formed/min/number of hepatocyte cells. (b) Drug probe enzyme activities determined using microsomes should be expressed as metabolite formed/min/microsomal protein/sample. (c) The simplest and most frequently used endpoints to identify enzyme inductions are the fold induction activity: fold induction =
(activity of test drug treated cells) (activity of negative control cells)
(d) A drug that produces a fold-change that is equal or greater than 40 % of the positive control can be considered as an enzyme inducer in vitro: % positive control =
(activity of test drug treated cells) × 100 (activity of positive control cells)
(e) An alternative endpoint is the use of an ED50 value (effective concentration at which 50 % maximal induction occurs), which represents a potency index that can be used to compare the potency of different compounds. Other endpoints than enzyme activities have been reported for evaluating a drug’s induction potential following a hepatocyte induction experiment, including enzyme expression, is generally detected by Western immunoblotting or immunoprecipitation probed with specific polyclonal antibodies. Relative quantification of the specific P450 enzyme protein requires that the electrophoretic system clearly resolves the individual enzymes and/or the primary antibodies be specific for the enzyme quantified. Enzyme antibody preparations are highly variable. Induction of mRNA transcript is easily detected using reverse transcriptase-polymerase chain reaction (RT-PCR). This technique can quantify mRNA expression for a specific CYP enzyme but is not necessarily informative of protein expression levels.
3.6
Bioactivation of Drugs to Reactive Metabolites
It is notable that of 27 drugs eliciting liver toxicity and were withdrawn from the market or whose use has been restricted (Table 3.5) 85 % are known be ‘bio-activated’ to reactive metabolites. Many other drugs have undoubtedly been discarded because of animal or early human evidence of liver toxicity, but the information has never been reported. At first glance, the data in Table 3.5 appear to indicate that reactive metabolites play a causative role in hepatotocity, and as was pointed out by Evans et al. (2004), identifying the bioactivation process is relatively straightforward. In contrast, the downstream consequences of this
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Table 3.5 Drugs eliciting liver toxicity that were either removed from the market or have their use restricted Drug generic name Iproniazid Ibufenac Ticrynafen Oxyphenisatin Benoxaprofen Pirprofen Phenacetin Zomepirac Perhexiline Nomifensin Suprofen Isaxonine Felbamate Tolrestat Sulfathiazole Temafloxacin Bromfenac Ebrotidine Mibefradil Tolcapone Trovafloxacin Troglitazone Nefazodone Pemoline Amodiaquine Zileutin Zimelidine
Reactive metabolite
Dose (mg)
Date of regulatory action
Yes Yes Yes — Yes Yes Yes Yes — Yes Yes Yes Yes Yes Yes Yes — Yes Yes Yes Yes Yes Yes — Yes Yes Yes
— 600 400 — 600 800 900 — 300 125 — 1500 400 400 1000 — 50 800 100 300 200 400 300 120 200 600 300
1959 1965 1979 1972 1982 1982 1982 1983 1985 1986 1987 1992 1994 1996 1997 1997 1998 1998 1998 1998 1999 2000 2001 2005 — — —
processes leading to cellular toxicity are complex and usually not well understood. For example, there is immutable agreement that formation of a reactive electropihilic metabolite, N-acetyl-p-benzoquinoneimine (NAPQI), is a necessary requirement for the observed liver toxicity following an overdose of acetaminophen; however, following more than 30 000 articles concerning these phenomena over the past 40 years (Rumack, 2002), our understanding of the molecular toxicology of acetaminophen was summed up by David when he concluded that ‘hepatocellular damage likely results from the aggregated effect of damage to multiple proteins and simultaneous inhibition of many enzymes and cellular functions’ (David, 2005). Even so, there should be and is the concern when drugs are bioactivated to reactive metabolites (Watkins and Seeff, 2006).
3.7
Identification of Reactive Metabolites
Reactive metabolites are transient and can either react with cellular proteins or undergo detoxification reactions, most commonly via conjugation with cellular glutathione. Their
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relative rates of formation and degrees of reactivity can determine which cellular molecules have the potential for adduct formation and the resulting biological effect: r highly reactive metabolites are usually bound to the enzyme which formed them, i.e. mechanism-based enzyme inhibition; r slightly less reactive metabolites lasting seconds to minutes may remain in the organelle that produced them; r intermediately reactive metabolites may migrate to other critical organelles within the cell, such as the nucleus, mitochondria or hepatocellular transporter proteins r long-lived reactive metabolites can enter the systemic circulation and have the potential to target proteins in cells throughout the body. Since reactive electrophilic metabolites, by definition, are unstable, they are not usually identified as such but as covalent adducts to cellular proteins or as products of detoxification pathways, such as glutathione transferase enzymes, to form glutathione conjugates. In some cases, these conjugates or adducts can be used to assess the exposure to the potentially toxic reactive metabolites, For example, Davern et al. (2006) recently reported that serum acetaminophen–protein adducts reliably identified acetaminophen toxicity in patients with acute liver failure and may be a useful clinical diagnostic test. However, the monitoring of drug–protein adducts is often difficult due to their short ‘half-life’ and lack of sensitive analytical techniques. Glutathione conjugates of reactive electrophilic metabolites are usually identified by LC–MS/MS analysis as their hydrolyzed products, N-acetylcysteineor cytseine-conjugates, excreted in urine and/or bile. The major detoxification of reactive electrophilic metabolites via glutathione transferase to form glutathione conjugates can be used as a tool to screen for reactive metabolites during the drug-development process. The use of glutathione as a ‘trapping agent’ is not new. Rollins et al. (1979) 27 years ago used human adult liver microsomes and fetal liver cells to oxidize acetaminophen by trapping the toxic acetaminophen reactive metabolite with glutathione. In recent years, there has been an increased interest in screening for reactive metabolites which is partly due to recent developments in mass spectrometry instrumentation, i.e. LC–MS/MS, and the availability of human liver tissue which have provided the tools to screen for these potential toxins using human liver tissue in vitro early in the drug-development process (Dieckhaus et al., 2005; Mutlib et al., 2005). There are numerous reports in the literature using human liver hepatocytes and cellular fractions in combination with glutathione as a trapping agent and LC–MS/MS to screen for reactive metabolites. An example of this method involves the antiepileptic drug, felbamate, whose use was severely restricted in 1994 by the FDA due to reports of aplastic anemia and liver failure (Table 3.5). There is evidence that an intermediary reactive metabolite, 2-phenylpropenal (ATPAL), is toxic to freshly isolated human hepatocytes in vitro (Kapetanovic et al., 2002) and may be responsible for the observed hepatotoxicity (Figure 3.2). Major inter-species difference in the metabolism of felbamate leading to formation of ATPAL have been reported (Dieckhaus et al., 2002). Presently a felbamate analog, fluorofelbmate, is undergoing drug development as a replacement for felbamate. Therefore, we used human liver post mitochondrial (S9) fractions in the presence of glutathione as a trapping agent and NADPH cofactor to compare the conversion of a felbamate intermediary metabolite, CCMF, to ATPAL to that formed from the analogous intermediary fluorofelbamate metabolite F-CCMF, Figure 3.2 (Parker et al., 2005). Following incubation, the media
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Felbamate
F-Felbamate O
C
O
CH2 OCNH2 H CH2 OCNH2
C
CH2 OCNH2 F CH2 OCNH2
O HO
O HO
H NH H C O CH2 O C
C
H NH F C O CH2 O F-CCMF C
C
CCMF ATPAL O C H C CH2 O C H C H CH2 GS ATPAL-GS
Figure 3.2 Partial felbamate metabolic pathways and those proposed for fluorofelbamate leading to formation of the reactive metabolite, ATPAL
was analyzed ATPAL by LC–MS/MS. The results of this study revealed that <0.1 % of the F-CCMF was converted to ATPAL when compared to CCMF. The data listed in Table 3.4 appear to indicate that hepatotoxicity is only observed with compounds administered at high clinical doses (i.e. >200 mg/day). Examples that support this hypothesis are the closely related antidiabetic drugs, troglitazone, pioglitazone and rosiglitazone, which all contain a common thiazolidinedione ring (Figure 3.3). After a number of reports that associated troglitazone with acute liver toxicity and failure, leading to death or liver transplants, the drug was removed from the market three years after its introduction in 1997 (FDA, 2000). Very rare reports of liver toxicity with the presently marketed troglitazone analogs, i.e. rosiglitzone and pioglitazone, have been usually milder and reversible compared to troglitazone (Isley, 2003). Surprisingly, it as been reported that the thiazolidinedione (TZD) ring in all three compounds is bioactivated by human liver microsomes through a common novel oxidative scission of the TZD ring system to electrophilic reactive metabolites. This was based on the addition of glutathione to the media to trap the reactive metabolites as their glutathione adducts which then were identified following analysis of the media by LC–MS/MS (Kassahun et al., 2001; Alvarez-Sanchez et al., 2006; Baughman et al., 2005). Loyd et al. recently investigated the hepatotoxicity of troglitazone and rosiglitazone to cryopreserved human hepatocytes from 37 donors (Lloyd et al., 2002) using several cell viability analyses. Based on their results, they concluded that
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O
83
O S NH
HO Troglitazone (dose = 600 mg) N
O
O O S NH
N
Pioglitazone (dose = 45 mg) N
O
O O S NH
Rosiglitazone (dose = 8 mg)
O
Figure 3.3 Chemical structures of three thiazolidinedione antidiabetic drugs and maximum daily doses
the differences in toxicity observed in humans may be due in part to the difference in clinical doses which is consistent with the data shown in Table 3.4 and Figure 3.3 which compares the doses for the three glitazone drugs. Although both troglitazone and rosiglitazone were toxic to human hepatocytes in vitro, a 600 mg troglitazone dose is 75-fold > rosiglitazone and 14-fold > pioglitazone doses. 3.7.1
Screening for Drug-Induced Hepatotoxicity in Cells in vitro
There is considerable evidence that reactive metabolites are formed from drugs known to cause liver toxicity (Walgren et al., 2005) and tools are now readily available to screen for reactive metabolites in vitro and in humans in vivo; however, the mechanism by which these toxic species initiate and propagate cellular injury is poorly understood (Park et al., 2005; Zhou et al., 2005). Recently, there has been a major focus on developing cellular models in vitro and sensitive and specific biomarkers in vivo that can predict the risk for drug-induced liver toxicity. Hepatoxicity can potentially result from cellular interactions of the parent drug or one of its metabolic products and drug–drug interactions can modify human exposure to these molecular entities. Therefore, any cellular model in vitro used to screen for drug-induced hepatoxicity should express the full complement of metabolizing enzymes. Several investigators have used immortalized cells including a human hepatoma cell line (HepG2) (Wilkening et al., 2003; Vignati et al., 2005) and a human hepatocyte derived cell line (Fa2N-4) (Ripp et al., 2006) to screen for hepatotoxins; however, the activity and expression, especially CYP enzymes, have been demonstrated to be extremely low (Rodriguez-Antona et al., 2002; Jover et al., 2001).
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Some investigators have used various cell models in vitro to overcome this deficiency, for example, Vignati et al. (2005) incubated drugs in combination with HepG2 cells in combination with microsomes containing cDNA-expressed CYP 3A, while Dai and Cederbaum (1995) demonstrated the cytotoxicity of acetaminophen in human CYP2E1-transfected HepG2 cells.
3.8
Conclusions
Screening drug candidates early during the drug-development process for their potential to interact with other drugs and/or to produce bioactivation toxic metabolites resulting in hepatotoxicity is of high priority for the pharmaceutical industry. Further research efforts are underway to develop reliable models in vitro to minimize the risk for drug-induced hepatotoxicity. Currently, human hepatocytes are the preferred in vitro model to investigate the total biotransformation and interaction potential of a drug in human liver and to identify a drug’s potential to produce toxicity in vitro.
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Davies B. and Morris T. (1993). Physiological parameters in laboratory animals and humans. Pharm Res 10: 1093–1095. Dieckhaus C. M., Thompson C. D., Roller S. G. and Macdonald T. L. (2002). Mechanisms of idiosyncratic drug reactions: the case of felbamate. Chem Biol Interact 142: 99–117. Dieckhaus C. M., Fernandez-Metzler C. L., King R., Krolikowski P. H., and Baillie T. A. (2005). Negative ion tandem mass spectrometry for the detection of glutathione conjugates. Chem Res Toxicol 18: 630–638. Evans D. C., Watt A. P., Nicoll-Griffith D. A. and Baillie T. A. (2004). Drug–protein adducts: an industry perspective on minimizing the potential for drug bioactivation in drug discovery and development. Chem Res Toxicol 17: 3–16. FDA (2000). ‘Medwatch’, Food and Drugs Administration (FDA), Silver Spring, MD, USA [http://www.fda.gov/medwatch/safety/2000/safety00.htm#rezuli]. FDA (2005). ‘Guidance for Industry: Safety Testing of Drug Metabolites’ Food and Drug Administration (FDA), Silver Spring, MD, USA [http://www.fda.gov/cder/guidance/6366dft.pdf]. FDA (2006). FDA Guidance for Industry: Drug Interaction Studies – Study Design, Data Analysis and Impact on Dosing and Labeling’. Food and Drug Administration (FDA), Silver Spring, MD, USA [http://www.fda.gov/cder/guidance/6695dft.pdf]. Fung M., Thornton A., Mybeck K., Wu J. H., Hornbuckle H. and Muniz E. (2001). Evaluation of the characterisitics of safety withdrawal of prescription drugs from worldwide pharmaceutical markets – 1960 to 1999. Drug Inform J 35: 293–317. Gomez-Lechon M. J., Donato M. T., Castell J. V. and Jover R. (2004). Human hepatocytes in primary culture: the choice to investigate drug metabolism in man. Curr Drug Metab 5: 443–462. Guengerich F. P. (2006). Cytochrome P450s and other enzymes in drug metabolism and toxicity. AAPS J 8: E101–E111. Hengstler J. G., Utesch D., Steinberg P., Platt K. L., Diener B., Ringel M., Swales N., Fischer T., Biefang K., Gerl M., Bottger T., Oesch F. (2000). Cryopreserved primary hepatocytes as a constantly available in vitro model for the evaluation of human and animal drug metabolism and enzyme induction. Drug Metab Rev 32: 81–118. Isley W. L. (2003). Hepatotoxicity of thiazolidinediones. Expert Opin Drug Saf 2: 581–586. Jover R., Bort R., Gomez-Lechon M. J. and Castell J. V. (2001). Cytochrome P450 regulation by hepatocyte nuclear factor 4 in human hepatocytes: a study using adenovirus-mediated antisense targeting. Hepatology 33: 668–675. Kapetanovic I. M., Torchin C. D., Strong J. M., Yonekawa W. D., Lu C., Li A. P., Dieckhaus C. M., Santos W. L., Macdonald T. L., Sofia R. D. and Kupferberg H. J. (2002). Reactivity of atropaldehyde, a felbamate metabolite in human liver tissue in vitro. Chem Biol Interact 142: 119–134. Kassahun K., Pearson P. G., Tang W., McIntosh I., Leung K., Elmore C., Dean D., Wang R., Doss G. and Baillie T. A. (2001). Studies on the metabolism of troglitazone to reactive intermediates in vitro and in vivo. Evidence for novel biotransformation pathways involving quinone methide formation and thiazolidinedione ring scission. Chem Res Toxicol 14: 62–70. Kern A., Bader A., Pichlmayr R. and Sewing K. F. (1997). Drug metabolism in hepatocyte sandwich cultures of rats and humans. Biochem Pharmacol 54: 761–772. Klaassen C. D. and Slitt A. L. (2005). Regulation of hepatic transporters by xenobiotic receptors. Curr Drug Metab 6: 309–328. Kliewer S. A., Lehmann, J. M., Milburn, M. V. and Willson, T. M. (1999). The PPARs and PXRs: nuclear xenobiotic receptors that define novel hormone signaling pathways: 2. Recent Prog Horm Res 54: 345–367. LeCluyse E. L. (2001). Human hepatocyte culture systems for the in vitro evaluation of cytochrome P450 expression and regulation. Eur J Pharm Sci 13: 343–368. Liebler D. C. and Guengerich F. P. (2005). Elucidating mechanisms of drug-induced toxicity. Nat Rev Drug Discov 4: 410–420.
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Lloyd S., Hayden M. J., Sakai Y., Fackett A., Silber P. M., Hewitt N. J. and Li A. P. (2002). Differential in vitro hepatotoxicity of troglitazone and rosiglitazone among cryopreserved human hepatocytes from 37 donors. Chem Biol Interact 142: 57–71. Moore L. B., Parks D. J., Jones S. A., Bledsoe R. K., Consler T. G., Stimmel J. B., Goodwin B., Liddle C., Blanchard S. G., Willson T. M., Collins J. L. and Kliewer S. A. (2000). Orphan nuclear receptors constitutive androstane receptor and pregnane X receptor share xenobiotic and steroid ligands: 8. J Biol Chem 275: 15122–15127. Morel F., Beaune P. H., Ratanasavanh D., Flinois J. P., Yang C. S., Guengerich F. P. and Guillouzo A. (1990). Expression of cytochrome P-450 enzymes in cultured human hepatocytes. Eur J Biochem 191: 437–444. Mutlib A., Lam W., Atherton J., Chen H., Galatsis P. and Stolle W. (2005). Application of stable isotope labeled glutathione and rapid scanning mass spectrometers in detecting and characterizing reactive metabolites. Rapid Commun Mass Spectrom 19: 3482–3492. Naritomi Y., Terashita S., Kagayama A. and Sugiyama Y. (2003). Utility of hepatocytes in predicting drug metabolism: comparison of hepatic intrinsic clearance in rats and humans in vivo and in vitro. Drug Metab Dispos 31: 580–588. Olson H., Betton G., Stritar J. and Robinson D. (1998). The predictivity of the toxicity of pharmaceuticals in humans from animal data – an interim assessment. Toxicol Lett 102–103: 535–538. Olson H., Betton G., Robinson D., Thomas K., Monro A., Kolaja G., Lilly P., Sanders J., Sipes G., Bracken W., Dorato M., Van D. K., Smith P., Berger B. and Heller A. (2000). Concordance of the toxicity of pharmaceuticals in humans and in animals. Regul Toxicol Pharmacol 32: 56–67. Pang K. S. and Rowland M. (1977). Hepatic clearance of drugs. I. Theoretical considerations of a ‘well-stirred’ model and a ‘parallel tube’ model. Influence of hepatic blood flow, plasma and blood cell binding, and the hepatocellular enzymatic activity on hepatic drug clearance. J Pharmacokinet Biopharm 5: 625–653. Park B. K., Kitteringham N. R., Maggs J. L., Pirmohamed M. and Williams D. P. (2005). The role of metabolic activation in drug-induced hepatotoxicity. Annu Rev Pharmacol Toxicol 45: 177–202. Parker R. J., Hartman N. R., Roecklein B. A., Mortko H., Kupferberg H. J., Stables J. and Strong J. M. (2005). Stability and comparative metabolism of selected felbamate metabolites and postulated fluorofelbamate metabolites by postmitochondrial suspensions. Chem Res Toxicol 18: 1842–1848. Ripp S. L., Mills J. B., Fahmi O. A., Trevena K. A., Liras J. L., Maurer T. S. and de Morais S. M. (2006). Use of Immortalized Human Hepatocytes to Predict the Magnitude of Clinical Drug–Drug Interactions Caused by CYP3A4 Induction. Drug Metab Dispos 34: 1742–1748. Rodriguez-Antona C., Donato M. T., Boobis A., Edwards R. J., Watts P. S., Castell J. V. and GomezLechon M. J. (2002). Cytochrome P450 expression in human hepatocytes and hepatoma cell lines: molecular mechanisms that determine lower expression in cultured cells. Xenobiotica 32: 505–520. Rollins D. E., von B. C., Glaumann H., Moldeus P. and Rane A. (1979). Acetaminophen: potentially toxic metabolite formed by human fetal and adult liver microsomes and isolated fetal liver cells. Science 205: 1414–1416. Rumack B. H. (2002). Acetaminophen hepatotoxicity: the first 35 years. J Toxicol Clin Toxicol 40: 3–20. Runge-Morris M. and Kocarek T. A. (2005). Regulation of sulfotransferases by xenobiotic receptors: 1. Curr Drug Metab 6: 299–307. Skett P., Tyson C., Guillouzo A. and Maier P. (1995). Report on the international workshop on the use of human in vitro liver preparations to study drug metabolism in drug development (held in Utrecht, The Netherlands, 6–8 September 1994). Biochem Pharmacol 50: 280–285. Smith D. A. and Obach R. S. (2005). Seeing through the mist: abundance versus percentage. Commentary on metabolites in safety testing. Drug Metab Dispos 33: 1409–1417. Soars M. G., Burchell B. and Riley R. J. (2002). In vitro analysis of human drug glucuronidation and prediction of in vivo metabolic clearance. J Pharmacol Exp Ther 301: 382–390.
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Sugatani J., Nishitani S., Yamakawa K., Yoshinari K., Sueyoshi T., Negishi M. and Miwa M. (2005). Transcriptional regulation of human UGT1A1 gene expression: activated glucocorticoid receptor enhances constitutive androstane receptor/pregnane X receptor-mediated UDPglucuronosyltransferase 1A1 regulation with glucocorticoid receptor-interacting protein 1: 3. Mol Pharmacol 67: 845–855. Temple R. J. and Himmel M. H. (2002). Safety of newly approved drugs: implications for prescribing. JAMA 287: 2273–2275. Tirona R. G. and Kim R. B. (2005). Nuclear receptors and drug disposition gene regulation: 1. J Pharm Sci 94: 1169–1186. Vignati L., Turlizzi E., Monaci S., Grossi P., Kanter R. and Monshouwer M. (2005). An in vitro approach to detect metabolite toxicity due to CYP3A4-dependent bioactivation of xenobiotics. Toxicology 216: 154–167. Walgren J. L., Mitchell M. D. and Thompson D. C. (2005). Role of metabolism in drug-induced idiosyncratic hepatotoxicity. Crit Rev Toxicol 35: 325–361. Watkins P. B. and Seeff L. B. (2006). Drug-induced liver injury: summary of a single topic clinical research conference: 1. Hepatology 43: 618–631. Wilkening S., Stahl F. and Bader A. (2003). Comparison of primary human hepatocytes and hepatoma cell line Hepg2 with regard to their biotransformation properties. Drug Metab Dispos 31: 1035– 1042. Zhao P., Kunze K. L. and Lee C. A. (2005). Evaluation of time-dependent inactivation of CYP3A in cryopreserved human hepatocytes. Drug Metab Dispos 33: 853–861. Zhou S., Chan E., Duan W., Huang M. and Chen Y. Z. (2005). Drug bioactivation, covalent binding to target proteins and toxicity relevance. Drug Metab Rev 37: 41–213.
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4 Human-Based in vitro Experimental Systems for the Evaluation of Human Drug Safety Albert P. Li
4.1
Accurate Assessment of Drug Safety as a Major Challenge to the Pharmaceutical Industry
Drug development is a costly and time-consuming process. DiMassi et al. (2003) have estimated that for R&D initiated in 2001 with approval twelve years later (based on the average time required for approval), the out-of-pocket cost for a single approved drug is estimated to be US $970 million, equivalent to a capitalized cost of US $1.9 billion. A major challenge in drug development is the low clinical phase success rate (estimated to be 23 % by DiMassi et al. (2003)). Lack of efficacy and the occurrence of unacceptable adverse effects are the two principle reasons of failure. Furthermore, marketed drugs continue to be withdrawn or have their use limited due to adverse effects. Although the number of drugs required to be withdrawn is relatively small compared to all marketed drugs, the economical impact on the company with the withdrawn drug can be astronomical. Efficacy and safety are therefore two co-dependent requirements for successful drug development – clinical failure will result if the drug candidate possesses only one of these two properties. Drug discovery should not only emphasize on pharmacological effects, but should also take into account drug safety to allow the selection of drug candidates with the highest probability of success.
Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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Human-Specific Drug Toxicity as a Major Reason for the Failure of Prediction of Human Drug Toxicity with Preclinical Nonhuman Animal Models
The fact that drug candidates would fail in clinical trials due to unacceptable safety illustrates that the old paradigm of drug safety testing, namely, the use of laboratory animals to predict human safety, is not adequate. Safety testing in nonhuman laboratory animals can be used to remove chemical entities which are toxic to both laboratory animals and humans, but would not be useful in the detection of toxicities that are human-specific.
4.2.1
Species–Species Differences in Drug Metabolism
Species-differences in drug metabolism have been extremely well-established. An example of rat–human differences in drug metabolism can be illustrated by a study in our laboratory using rat and human liver microsomes. In this study (Easterbrook et al., 2001), we evaluated the formation of human metabolites using rat and human liver microsomes from substrates of the major human P450 isoforms. Aroclor 1254-induced and uninduced rat liver microsomes were compared with human liver microsomes in phenytoin O-deethylation, coumarin 7-hydroxylation, tolbutamide 4-hydroxylation, s-mephenytoin 4 -hydroxylation, chloroxazone 6-hydroxylation and testosterone 6beta-hydroxylation. We found that both induced and uninduced rat liver microsomes were active in all of the pathways studied with the exception of coumarin 7-hydroxylation. Coumarin 7-hydroxylation was observed with human liver microsomes but not the rat liver microsomes. The results of this study illustrate that species–species differences in metabolism exist, and that a major human metabolite may not be formed in a nonhuman animal. The ramification of this finding is that if the toxicity of a drug is the result of the formation of a toxic metabolite, and if the formation of the toxic metabolite is specific to humans, then the safety data from nonhuman animal species will lead to an underestimation of human toxicity. Similarly, nonhuman animal data may overestimate human drug toxicity and may draw a wrong conclusion on the safety of the drug in humans, if the toxic metabolite is unique to the nonhuman animals and is not formed in humans.
4.2.2
Species–Species Differences in Target Cell Sensitivity to Toxicant Effects
Another possible explanation for human–nonhuman animal differences in drug toxicity is that a toxicant may have species-specific toxicity towards target cells. An example of this is the species differences of the toxicity of various anticancer agents on myeloid progenitor cells. Erickson-Miller et al. (1997) evaluated the toxicity of ercamptothecin (CAM), topotecan (TPT) and 9-aminocamptothecin (AC) to human, canine, and murine myeloid progenitors (CFU-GM) in vitro and found that the murine IC90 values were 2.6-, 2.3-, 10-, 21-, 5.9- and 11-fold higher than human values for CAM lactone (NSC-94600) and sodium salt (NSC-100880), TPT (NSC-609699), and racemic (NSC-629971), semisynthetic and synthetic preparations (NSC-603071) of 9AC, respectively. In contrast, canine IC90 values were the same as, or lower than, the human IC90 values for all six compounds.
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Overcoming Species–Species Differences as an Approach to Better Predict Human Drug Toxicity
Experimental systems that can provide information towards human-specific toxicity therefore should be valuable in the prediction of human drug toxicity in vivo. As in vivo experimentation with humans in vivo during preclinical phases is neither practical nor ethical, surrogates for humans in vivo need to be applied. Recently, human-based in vitro experimental systems have been developed and applied routinely for the evaluation of drug metabolism and drug–drug interactions (Huang et al., 1999; Davit et al., 1999; Li, 2004a). This ready acceptance of the in vitro systems can be attributed to the well-defined disciplines of drug metabolism. As the liver is the major organ for xenobiotic metabolism, in vitro hepatic systems such as liver microsomes and hepatocytes are found to readily reproduce in vivo hepatic metabolism. Regulatory agencies such as the US FDA (FDA, 1997, 1999, 2006) has endorsed the use of in vitro human-based experimental systems in the definition of human metabolic fate and drug–drug interaction potential of drugs. In fact, the US FDA has recommended that in vitro results with human-based systems such as liver microsomes and hepatocytes can be used to define human drug–drug interaction potential, without further clinical trials (clinical trials are recommended only if the results suggest significant drug–drug interaction potential). It is interesting to note that the application of in vitro drug metabolism technologies using human-based experimental systems has been attributed to the virtual elimination of pharmacokinetics as a reason for clinical trial failure. The success in the application of in vitro drug metabolism systems, in combination with data from relevant in vivo animal models, in the prediction of human metabolism suggest that the same approach will also be successful for safety evaluation (MacGregor et al., 2001). 4.3.1
In vitro–in vivo Strategy (IVIVS) for the Prediction of Human Drug Toxicity
Based on the premise that the inability to accurately predict human drug toxicity is due to species–species differences, i. e. there are human-specific drug properties that cannot be revealed by nonhuman animal studies, a safety evaluation strategy is proposed here for the preclinical evaluation of human drug toxicity: (1) Apply of human-based in vitro systems to provide human-specific toxicity data. (2) Select a relevant animal species to develop in vivo parameters. (3) Predict human in vivo drug toxicity via a combination of human-specific information obtained in vitro and in vivo parameters obtained from nonhuman animals in vivo. This IVIVS will require the development of in vitro experimental systems with humanspecific properties to cover the key toxic drug effects in humans and a vigorous set of parameters defining the relevant nonhuman animal species. 4.3.2
Selection of Human in vitro Experimental Systems
For the in vitro–in vivo paradigm to serve the intended purpose of accurate prediction of human drug toxicity, one has to careful in the choice of experimental systems. The following
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are guidelines for the choice of in vitro systems: (1) Human-specific properties. As the purpose is to derive human-specific information, the system used should have the desired human properties. Examples of human-specific properties include human drug metabolism enzymes, transporters and molecular targets relevant to drug toxicity. (2) Physiologically relevant experimental conditions. In vitro systems, when used under experimental conditions that would not be achieved in vivo, may yield information that is difficult to extrapolate to the in vivo situation. Examples of irrelevant experimental conditions include the use of too high a drug concentration, interaction of the test substance with media ingredients that are not present in human body fluids and the use of cells with properties irrelevant to the target cells in vivo. (3) Appropriate endpoints. The endpoints used in conjunction with a chosen experimental system must be scientifically proven to be representative of the in vivo toxic events Examples of a relevant and a less relevant experimental system for the evaluation of human hepatotoxicity are presented below: Relevant: primary human hepatocytes: r Relevant species: human. r Relevant organ: liver. r Relevant target cells: hepatocytes or liver parenchymal cells are often the cells initially damaged by toxic drugs and reactive metabolites. r Relevant human-specific metabolism: human hepatocytes as freshly isolated cells or cryopreserved freshly isolated cells are well-established to contain full complements of human drug metabolizing enzymes (Li et al., 1999), including Phase I oxidation (e.g. P450 isoforms, monoamine oxidase, flavin-dependent monooxygenases), Phase II conjugation (UDP-dependent glucuronyl transferase, sulfotransferases, gluthathione transferase, N -acetylaminotransferases) and transporter activities (Shitara et al., 2003; Jigorel et al., 2005; Li, 2007). Less relevant: HepG-2 cells (human hepatoadenocarcinoma cell line): r Relevant species: human. r Relevant organ: liver. r Irrelevant target cells: adenocarcinoma cells are not the target of hepatotoxicity in vivo. r Irrelevant human-specific metabolism: Hep-G2 cells are known to have extremely low activities of human metabolizing enzymes, with the embryonic rather the adult phenotypes being expressed (e.g. CYP1A2 and CYP3A7 rather than CYP1A2 and CYP3A4/5) (Rodriguez-Antona et al., 2002). The preferred human in vitro systems are therefore primary cells derived from human organs, used within a period that the cells would retain differentiated functions, thereby serving as surrogates of the similar cells in vivo. The primary cell culture systems that are currently available and the respective organspecific toxicity that can be evaluated include the following: r hepatocytes for liver toxicity; r renal proximal tubule epithelial cells for nephrotoxicity;
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r vascular endothelial cells for vascular toxicity; r neuronal and glial cells for neurotoxicity; r cardiomyocytes for cardiotoxicity. 4.3.3
Application of Human-Based in vitro Systems in Drug Development
In vitro toxicity assays can be applied in various phases of drug development (Li, 2000; 2004a): (1) Early screening of intrinsic toxicity. Cell-based systems are used for rapid screening of drug candidates, especially structural analogs, to allow the selection of less toxic structures for further development. The screening assay can allow logical evaluation of structures responsible for toxicity (toxicophore) which, hopefully, can be separated from structures for pharmacological activity (pharmacophores). Toxicity screening with in vitro systems require only limited amount of test articles, and is rapid and quantitative. Toxicity is most effective when one has an indication for in vivo toxicity (e.g. hepatotoxicity or nephrotoxicity) for a lead molecule, therefore allowing the selection of the most appropriate in vitro system for screening (e.g. hepatocytes for hepatotoxicity and renal proximal tubule cells for nephrotoxicity). (2) Mechanistic evaluations. Besides screening, in vitro experimental systems can be used to define toxic mechanisms. The defined experimental conditions and the availability of reagents and approaches for multiple endpoints allow one to define the key pathways involved in a toxicology phenomenon. Mechanistic understanding is critical to drug development. It allows a better understanding of human health risks, defines potential risk factors and evaluates the relationship between efficacy and adverse effects.
4.4
Overcoming the Major Deficiencies of an in vitro System
An argument routinely raised against the application of in vitro systems in safety evaluation is that toxicity is a complex phenomenon and therefore cannot be adequately modeled by simple in vitro systems such as cell culture assays. The major deficiencies of in vitro experimental systems can be defined as follows: (1) Lack of systemic effects. in vitro experimental systems in general consist of single cell types. Toxic effects are evaluated in the absence of influences from systemic effects that may be critical to drug toxicity. An example is the participation of the immune system in organ toxicity. One hypothesis for idiosyncratic hepatotoxicity, for instance, is the hapten hypothesis which postulates that liver failure arises from the cytotoxicity of antibodies towards antigens developed between the idiosyncratic drug (or its metabolites) on the plasma membrane of the hepatocytes. (2) Absence of chronic dosing. It is generally believe that drug toxicity due to acute cytotoxic events can be studied effectively with in vitro systems. However, toxic effects due to chronic, low-dose treatments may require multiple events that may or may not be obtained with in vitro studies, with cells treated for a relatively short time period (e.g. 24 h). Long-term treatments (e.g. months to years) of cells in culture is
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theoretically possible but in practice near impossible. Furthermore, it is extremely difficult to maintain primary cells, the preferred cell system, in a differentiated state for a long time period. For in vitro systems to be useful, one needs to develop experimental approaches to overcome these deficiencies. 4.4.1
Target Cell Initiation Theory for Drug-Induced Organ Toxicity (TACIT)
It is well-established that the final manifestation of drug-induced organ toxicity may require systemic, multiple organ interactions. For instance, damage to a cell population in an organ may lead to the ‘recruitment’ of inflammatory cells which in turn may secrete cytotoxic cytokines, leading to further damage to the organ. The involvement of inflammation in liver toxicity has been suggested by one line of evidence that hepatotoxicity is enhanced in animals treated with pro-inflammatory agents (Ganey et al., 2002). However, it can be visualized that organ toxicity may be due to a cascade of events, started with a specific cell populations that initiate the cascade. The Target Cell Initiation Theory of Organ Toxicity (TACIT) is proposed here: Drug-induced organ failure is caused by changes in a target cell population which have the potential to initiate a cascade of events, and may or may not involve different cell types within the organ, or mechanisms involving target organs, leading ultimately to extensive damage to the organ in question.
A corollary to the TACIT is that although organ failure may involve multiple cell types and multiple organs, toxicants can be identified in the following are known: r the initiating target cells; r the critical initiation events which would cause the organ-toxicity cascade in the target cells. 4.4.2
Practical Application of TACIT
TACIT states that one can use the target cells in isolation in combination with critical endpoints reflecting initiating events to detect agents that can cause organ failure, regardless of the subsequent pathways which may involve nontarget cells and organs. An example of an application of the hypothesis is the evaluation of hepatotoxic potential using primary hepatocytes: (1) Target cell selection. For most toxic drugs, liver parenchymal cells are the cells that are initially damaged. The parenchymal cells represent the major cell type of the liver, with all of the drug metabolizing enzymes of the liver. The cells can be damaged by the drugs directly (e.g. high, bolus concentration after oral administration) or the metabolites (e.g. cytotoxic or highly reactive metabolites). Primary hepatocytes, as fresh or cryopreserved isolates, represent the most appropriate cell system for the evaluation of hepatotoxicity. (2) Endpoint selection. Hepatotoxicity is known to be initiated for both cytotoxic and noncytotoxic events (Table 4.1). Endpoints based on these known events can be selected to be used with hepatocytes for the evaluation of hepatotoxicity.
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Table 4.1 Mechanisms of action of various types of hepatotoxicants. These mechanisms can be used as endpoints in combination with hepatocytes in vitro as early screening assays for hepatotoxicity Hepatotoxic mechanism
Toxicants
Direct cytotoxicity Toxic/reactive metabolite formation Toxic antibodies towards neoantigens
Cadmium chloride; cis-platinum Acetaminophen; cocaine; cyclophosphamide; ethanol
Cholestasis Steatosis Phospholipidosis
Amineptine; erythromycin derivatives; halothane imipramine; isaxonine; alpha-methyldopa; tienilic acid Rifampin; glibenclamide; cyclosporin A Carbamazepine; cycloheximide; tetracycline Amiodarone; diethylaminoethoxyhexestrol; perhexiline
Direct cytotoxicity. Liver toxicity is known to be preceded with cytotoxicity to the hepatocytes. Cytotoxicity can be using endpoints such as leakage of cytoplasmic enzymes, cellular ATP content, and mitochondrial metabolic activities (Li, 2004a). Correlation of cytotoxicity with human donor properties may lead to the understanding of key factors for hepatotoxicity of a toxicant (Lloyd et al., 2002; Hewitt et al., 2002). (b) Indirect cytotoxic effects. Toxicity of a substance to the liver may result from effects that may not lead to cytotoxicity directly, but will precipitate subsequent events which will lead to severe hepatotoxicity. These prelethal effects include the following: (i) Formation of reactive metabolites. Reactive metabolites formed in the hepatocytes react with cellular macromolecules which may lead to direct toxicity due to inactivation of key biochemical pathways. The reactive metabolites may also react with cellular proteins to form neoantigens, leading to cytotoxic inflammatory events. Reactive metabolites can be identified as glutathione (GSH) conjugates (e.g. using LC/MS) (Hartman et al., 2002; Prabhu et al., 2002; Zang et al., 2005; Cysyk et al., 2006). An indirect indication of reactive metabolite formation is the depletion of cellular GSH in the absence of cytotoxicity. (ii) Steatosis. Microvesicular steatosis, abnormal accumulation of intracellular lipid, is known to occur with exposure to hepatotoxic drugs. Accumulation of lipid vesicles and subsequent rupture of the lipid-ladden hepatocytes is believed to be a key step towards severe liver damage. Steatosis can be measured in cultured hepatocytes via the quantification of intracellular triglyceride accumulation and light microscopical examination of cells with lipid-specific stains such as Nile red (Amacher and Martin, 1997). (iii) Cholestasis. Bile acid retention in hepatocytes as a result of compromised efflux is known to be related to liver damage. The basolateral and apical transporters for bile acids are thought to be targets of drugs that induce cholestasis. Inhibition of bile acid transporter therefore is a relevant endpoint for the screening of drugs with cholestatic potential (Sahi et al., 2006). (a)
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(iv) Phospholipidosis. Impaired phospholipid metabolism is known to be related to inflammatory events and histopathological changes. Some hepatotoxic drugs are known to cause phospholipidosis in hepatocytes which can be detected morphologically using electron microscopy as cytoplasmic lamella bodies. Screening for lipidosis can be performed by also using a dye for intracellular lipids such as Nile Red (Kasahara et al., 2006). Unlike steatosis, the relationship between phospholipidosis and hepatotoxicity is not yet established. However, this endpoint is a frequently used histopathological endpoint used to suggest hepatotoxic potential (Reasor et al., 2006). 4.4.3
High-Content Endpoint Assays
High-content assays, such as toxicogenomics for gene expression, proteomics for protein expression, metabonomics for natural metabolite synthesis and cellomics for cell morphology changes, are relatively novel technologies with potential application in the definition of drug toxicity. It is anticipated that these high-content assays, which allow one to evaluate the effect of a test substance on myriad events (e.g. expression of 20 000 human genes), one may achieve the following: (1) (2) (3) (4) (5)
Discovery of response patterns (response profiles) that can be used to define toxicity. Discovery of new biomarkers for specific types of toxicity. Discovery of early indicators of chronic toxicity. Discovery of indicators of idiosyncratic toxicity. Elucidation of toxic mechanisms.
The high-content assays may be extremely valuable to the TACIT approach with in vitro systems. For instance, prediction of chronic toxicity using in vitro systems may be possible, provided that the key events for chronic toxicity can be defined in the target cells after treatment with the toxicant in vitro. High-content assays performed on target cells with chronic toxicants versus nontoxic analogs may aid the discovery of these key initiating events. Another potential valuable application of high-content assays is to the definition of idiosyncratic drug toxicity. Efforts to use a single theory (e.g. reactive metabolite formation) to universally defined idiosyncratic drug toxicity have proven not yet to be useful. One possible explanation for the illusiveness of the definition of idiosyncratic toxicants is that there may be multiple factors involved (Li, 2002). These factors are proposed to be the chemical properties of the toxicant and the genetic make up of the human victim, as well as the complex environmental factors surrounding the victim. It is projected that the factors that can be readily controlled are the ‘chemical properties’. Therefore, a more practical approach to eliminate an idiosyncratic drug may be the definition of the common properties of the toxicants (Li, 2002). Towards this goal of defining the properties of drugs that would cause idiosyncratic hepatotoxiciy, we have applied toxicogenomics to evaluate troglitazone, a drug that was withdrawn from the market due to its association with idiosyncratic hepatotoxicity. Trogltiazone was compared to the relatively nontoxic structural analogs, rosiglitazone and pioglitazone (Kier et al., 2004). Human hepatocytes from three donors were treated in vitro with the three chemicals, followed by extraction of mRNA and hybridization with cDNA microarrays. The gene expression heat map (using various ‘grades of shading’ to classify the magnitude of
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response) is presented in Figure 4.1. Dramatic differences were observed in gene expression profiles between the toxic troglitazone and the nontoxic structural analogues. Troglitazone was found to selectively ‘down-regulate’ genes with protective functions and ‘up-regulate’ genes with toxification functions. Similar toxicogenomics results with troglitazone were also obtained from the laboratory of Waring and coworkers (Liguori et al., 2005). The results illustrate the application of toxicogenomics in the development of an hypothesis to allow subsequent mechanistic investigations. Toxicogenomics studies with troglitazone and its less toxic analogs, rosiglitazone and pioglitazone, provided clues for the differential hepatotoxicity of these structurally related chemicals. All three chemicals appear to induce genes for oxidative metabolism (e.g. P450 isoform 3A4) which may be responsible for the generation of reactive metabolites (Kassahun et al., 2000). Troglitazone, however, uniquely induced gene expression of the toxification pathways such as apoptosis and inflammation, and suppressed gene expressions of the gene responsible for detoxification pathways such as acute phase proteins, Phase II conjugation, and stress proteins (Kier et al., 2004; Liguori et al., 2005). It is projected that the genes that are affected differentially by troglitazone and its less toxic structural analogs may be used to predict the human hepatotoxic potential of new troglitazone structural analogs. The combination of toxicgonomics and human cells in vitro may be a powerful combination to aid the accurate prediction of human drug toxicity. The key is to develop an in vitro system to incorporate in vivo factors that are key to toxicity. One such system is the Integrated Discrete Multiple Organ Co-culture (IdMOC) system. 4.4.4
IdMOC for Multiple Organ Interactions
One major drawback of an in vitro system is that each cell type is studied in isolation, while in the human body there may be multiple organ interactions that are critical to drug toxicity. The multiple organ interaction is not covered by the TACIT approach, as the initiating events may include effects of a toxicant on a target cell and a nontarget cell. A simple example is a drug which is first metabolized by the liver to form metabolites which may cause toxicity in a ‘distant’ organ, such as the heart. The TACIT approach using the target cell with the parent drug is therefore unlikely to produce the initiating events for the prediction of the ultimate organ toxicity. To overcome this deficiency, we have developed the Integrated Discrete Multiple Organ Co-culture (IdMoc) system (Li et al., 2004). The IdMOC allows the co-culturing of cells from different organs as physically separated cultures that are interconnected by an overlying medium, akin to the blood circulation connecting the multiple organs in the human body. The IdMOC models the multiple organ interaction in the whole organism in vivo, thus allowing the evaluation of organ-specific effects of a drug and its metabolites (Figure 4.2). The IdMOC represents an improved in vitro experimental system for routine screening of ADMET drug properties. The IdMOC involves the ‘wells-in-a-well’ concept. The typical IdMOC plate consists of an outer well containing within which are several wells (Figure 4.3). Cells of different origins (e.g. from different organs) are initially cultured, each in its own specific medium, in the wells. When the cells are established, the inner wells are flooded with an overlying medium, thereby connecting the multiple cell types. The multiple-cell types now can interact via the overlying medium, akin to the multiple organs in a human body interacting via the systemic circulation.
−1.9 −1.6 −2.1 −2.0 −1.8 −1.5 −1.5 −1.9 −1.4 −1.7 −1.5 −1.6 −1.6 −1.8 −1.7
−1.7 −1.6 −2.2 −1.5 −1.4 −1.2 −1.3 −2.2 −1.7 −1.5 −1.4 −1.5 −1.4 −1.5 −1.9
−3.2 −1.8 −1.2 −1.5 −1.6 −1.3 1.0 −1.3 −1.5 −1.4 −1.3 −1.5 −1.5 −1.4 −1.4 −1.7
−3.7 −2.2 −1.4 −2.1 −2.0 −1.9 −1.3 −1.4 −2.4 −1.0 −1.3 −1.7 −1.5 −1.4 −1.2 −1.8
−1.6 −1.6 −1.7 −1.5 −1.6 −1.4 −1.7 −2.4 −1.6 −1.4 −1.7 −1.6 −1.4 −1.5 −1.3 −2.8 −1.4 −1.2 −1.7 −1.6 −1.4 −1.5 −1.7
−1.6 −1.6 −1.8 −1.5 −1.5 −1.6 −1.7 −2.4 −1.7 −1.2 −1.5 −1.6 −1.4 −1.5 −1.4 1.1 −1.6 −1.3 −1.8 −1.6 −1.6 −1.6 −1.8
10 µM
−1.7 −1.5 −1.7 −1.6 −1.3 −1.4 −1.5 −1.7 −1.6 −1.3 −2.0 −1.6 −1.5 −1.6 −1.9
−2.5
−1.2 −1.5 −1.4 −1.3 −1.6 −1.7 −1.5
−1.9 −1.7 −2.2 −1.9 −1.6 −1.4 −1.8 −1.9 −1.8 −1.3 −2.1 −1.8 −1.5 −1.9 −2.2
−2.7
−2.4 −2.0 −1.6 −1.6 −2.1 −1.9 −1.7
20 µM
24 h
−1.4 −1.1 −1.2 1.0 1.1 −1.6 −1.1 1.0 −1.1 −1.6 −1.1 −1.1 −1.2 −1.2 −1.1
−1.5
−1.2 −1.0 −1.1 1.0 1.1 −1.4 −1.1 −1.0 −1.1 −1.5 −1.0 −1.1 −1.1 −1.1 −1.1
−1.4
Pio −1.3 −1.4 −1.2 −1.6 −1.3 −1.2 −1.2 −1.0 −1.2 −1.1 −1.6 −1.2 −1.3 −1.4
−1.3 −1.1 −1.2 −1.0 1.1 −1.4 −1.0 1.0 1.0 −1.4 −1.1 −1.1 −1.2 −1.2 −1.2
−1.4
−1.2 −1.0 −1.0 1.0 1.0 −1.2 −1.1 1.1 −1.1 −1.3 −1.1 −1.1 −1.1 −1.1 −1.0
−1.4
II: 25 µM, 24 h Rosi −1.2 −1.3 −1.1 −1.4 −1.3 −1.2 1.0 −1.2 −1.2 −1.0 −1.4 −1.1 −1.2 −1.4
−2.0 −2.4 −3.2 −2.3 −1.8 −4.7 −2.4 −2.8 −2.8 −5.0 −1.1 −1.7 −1.7 −1.5 −1.6
−1.8
−1.7 −1.8 −1.5 −2.7 −1.9 −2.6 −2.6 1.2 −2.2 −4.3 −1.0 −1.6 −1.5 −1.4 −1.5
−1.3
Tro −1.9 −4.3 −1.6 −2.1 −2.0 −1.7 −1.6 −2.1 −4.4 −2.5 −3.0 −1.9 −2.2 −2.2
Figure 4.1 Heat map summarizing the toxicogenomics results of the treatment of primary human hepatocytes with the hepatotoxic drug, troglitazone(tro), and its less toxic analogs, i.e. rosiglitazone(rosi), and pioglitazone(pio). The results of genes that are differentially suppressed and induced by troglitazone are shown in (a) and (b), respectively. Genes are grouped by physiological mechanisms of action and each line represents the results of a single gene. Significant response is defined as >50 % induction or suppression over solvent control. The microarray used was the TOX-chip from Phase I Molecular Toxicology, Inc.
Translation
Oxidative stress
−1.8 −1.7 −2.2 −1.7 −1.6 −1.7 −1.6 −1.7 −1.7 −2.0 −1.6 −1.8 −1.8 −2.0 −2.0
−1.9 −1.6 −1.7 −1.8 −1.5 −1.6 −1.6 −2.2 −1.7 −2.4 −1.5 −1.8 −1.8 −2.0 −1.9
−2.6
−2.5
−1.6 −1.4 −1.3 −1.2 −1.1 −1.4 −1.3
−1.9 −1.6 −1.5 −1.4 −1.3 −1.0 −1.7
50 µM
20:18
PHASE II metabolism
−2.1
−2.1
−1.6 −1.4 −1.7 −1.4 −1.2 −2.0 −2.0
−1.4 −1.3 −1.5 −1.2 −1.0 −1.5 −1.6
20 µM
I: Troglitazone
October 31, 2007
Mitochondrial function
Acute phase
−1.2 −1.3 −1.9 −1.3 −1.2 −1.8 −1.9
−2.1 −1.7 −1.9 −1.8 −1.4 −1.8 −1.9
10 µM
6h
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Cell growth
Inflammation
1.8 1.2 −1.0 1.4 3.2 1.8 2.8 1.8 2.8 12.4 4.1 7.3 11.4 7.1 5.1 7.1 4.0 3.7 4.0 4.3 3.0 2.4 2.0 2.8 1.8 1.7 2.5 2.1 1.7 2.9 1.8 1.7 2.0 1.6 2.8 2.2 1.2 1.2 −1.1 2.5 1.2 2.2 1.6 2.7 7.6 3.6 4.4 4.0 4.5 3.2 4.3 4.5 3.7 3.0 3.5 2.8 2.1 2.5 1.5 2.0 2.1 2.9 1.5 2.4 2.5 1.4 −1.1 2.5 1.9 2.1 2.2 1.9 1.8 1.3 1.8 1.3 2.5 1.4 2.8 9.7 4.4 5.4 4.3 4.5 4.1 5.5 3.2 1.9 3.6 2.6 2.6 2.3 1.6 1.7 2.3 2.1 2.4 1.9 2.5 2.7 1.7 1.7 2.9 1.8 1.5 1.1 1.0 1.4 1.0 1.5 2.3 2.6 1.2 2.9 5.9 3.6 6.1 5.2 5.3 5.2 5.1 2.1 3.4 2.9 3.7 2.4 2.3 4.4 2.5 2.2 2.7 2.8 2.0 2.4 2.6 1.9 2.6 2.4 3.2 1.5
Figure 4.1
1.3 1.1 −1.2 1.1 1.4 1.6 2.2 1.1 2.7 7.3 3.2 4.7 2.7 4.6 4.0 5.6 2.4 3.2 2.8 3.0 1.8 2.0 3.7 1.4 1.4 1.7 2.5 2.1 2.1 2.2 1.6 1.2 1.8 1.1 1.3
50 µM
24 h
(Continued)
1.9 1.5 −1.1 1.0 1.3 2.1 1.3 1.3 2.4 2.1 1.3 1.2 2.5 3.1 1.5 1.7 1.7 2.4 10.4 10.9 4.5 6.2 5.3 6.8 2.5 4.7 5.1 5.2 3.3 4.7 4.2 6.8 5.4 6.2 3.2 3.1 5.2 5.7 3.7 3.9 3.0 3.1 2.2 2.4 1.1 −1.1 1.7 1.6 1.8 2.0 1.9 2.3 2.1 1.9 2.7 1.8 1.6 1.6 2.2 2.5 1.6 2.1 −1.0 1.5 2.1 1.8 1.9 3.1 2.6 1.7
10 µM 1.1 1.8 −1.4 1.2 3.0 1.6 1.4 1.0 4.1 2.7 −1.4 −1.1 2.3 3.7 2.1 1.6 1.5 2.6 10.1 11.3 4.2 4.2 6.6 7.7 3.7 3.0 4.6 6.6 5.5 6.1 4.9 5.6 4.2 4.1 1.5 1.7 4.2 2.9 3.9 3.5 3.1 2.6 2.7 2.6 1.3 2.1 1.2 1.6 1.1 2.5 1.7 2.2 1.7 1.4 1.8 1.8 1.0 1.6 2.4 1.8 1.9 1.5 1.3 2.0 1.4 1.8 2.3 2.7 1.7 1.9
20 µM
II: 25 µM Pio Rosi 1.1 1.1 1.2 −1.1 1.2 −1.0 1.3 1.0 1.2 1.0 1.1 1.2 1.7 1.3 1.8 1.3 −1.0 1.0 −1.1 −1.0 1.6 1.2 1.4 1.3 1.0 1.6 1.1 1.4 −1.1 −1.1 1.6 −1.1 1.9 −1.0 1.7 1.5 3.2 1.8 1.6 3.6 1.9 1.2 1.5 2.1 2.0 1.4 1.6 1.6 1.8 1.1 2.0 1.2 2.2 1.4 1.8 2.2 −1.2 −1.5 1.2 −1.1 3.3 1.7 1.5 5.0 1.1 1.0 −1.0 1.0 −1.1 −1.3 −1.0 −1.1 1.1 −1.1 1.1 −1.0 1.2 1.6 1.2 1.8 −1.3 −1.0 −1.4 −1.5 1.2 1.3 1.3 1.3 −1.4 −1.4 −1.5 −1.2 1.1 1.3 1.1 1.1 1.1 1.1 1.2 1.1 1.0 1.2 –1.1 1.1 1.2 1.0 1.2 1.0 1.1 1.0 1.1 1.1 1.1 1.1 1.0 −1.1 1.2 1.2 −1.0 −1.2 −1.0 −1.2 −1.1 −1.1 1.1 1.3 −1.0 −1.2 1.1 1.0 1.0 1.0 1.0 −1.1 1.0 −1.0 1.2 1.6 1.3 1.7 Tro 1.3 1.8 1.3 1.5 1.7 1.1 1.5 1.1 2.4 2.2 1.5 1.8 1.2 1.7 −1.0 -1.1 1.5 3.9 −2.5 1.8 1.2 3.5 −1.2 2.6 −1.5 3.3 2.0 2.4 −1.1 1.1 1.0 2.0 1.2 1.6 −1.4 1.4 1.6 1.2 2.5 1.2 3.5 1.5 2.6 1.6 2.0 2.3 2.1 1.3 2.6 3.0 1.6 1.2 1.7 1.2 1.5 1.5 3.2 2.3 2.5 1.3 2.8 1.8 3.1 1.6 2.3 1.5 1.7 1.2 1.7 1.2
20:18
DNA replication and repair
1.8 1.4 1.5 1.2 2.0 3.2 2.8 2.2 4.2 10.6 5.8 6.5 7.4 10.0 5.1 8.6 2.4 4.0 3.4 3.7 3.3 3.1 1.8 2.0 2.4 2.3 2.4 1.7 1.4 2.5 1.5 1.8 2.6 1.7 1.6
20 µM
I: Troglitazone
October 31, 2007
Phase-1 metabolism
1.9 1.4 1.2 1.6 1.7 1.2 2.0 1.9 2.2 6.7 3.3 4.2 3.9 3.3 3.1 3.4 2.7 3.2 3.8 3.9 2.8 2.0 1.6 2.0 1.7 1.7 1.6 1.4 2.4 2.9 1.4 1.4 1.3 1.4 1.7
10 µM
6h
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Immediate–early
Apoptosis
Induced Genes
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(a)
(b)
Overlying medium connecting the multiple organs
Organ A cells (e.g. hepatocytes)
Organ B cells (e.g. kidney proximal tubule cells)
Organ C cells (e.g. vascular endothelial cells)
Figure 4.2 Schematic presentation of the Integrated Discrete Multiple Organ Co-culture (IdMOC) system. The IdMOC system is based on the concept that in the human body there are multiple organs that are physically separated but interconnected by the systemic circulation (a). A toxicant entering the systemic circulation will have interactions with all of the interconnected organs. Results of the interactions, which can be the formation of metabolites or the induction of reactive biomolecules (e.g. cytokines), will have the potential to interact with the multiple organs. The concept is reduced to practice by using a ‘wells-in-a-chamber’ concept, with multiple wells in a containing chamber, allowing the culturing of cells from different organs as physically discrete wells interconnected by an overlying medium (b)
The IdMOC system can be used for the following: (1) Differential cytotoxicity. Evaluation of the toxicity of a substance on different cell types (e.g. cells from different organs) under virtually identical experimental conditions with multiple cell-type interactions. (2) Differential distribution. Evaluation of the differential accumulation/distribution of a substance among multiple cell types. (3) Multiple organ metabolism. Evaluate the ultimate metabolic fate of a substance upon metabolism by cells representing multiple organs with metabolic functions (e.g. liver, kidney, lung).
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Figure 4.3 Photograph of an IdMOC plate based on the format of a standard 96-well plate (IdMOC-96). This plate consists of six individual wells inside a containing chamber. The wells are filled with colored liquids (left) to show that they are physically separated. The chambers are filled (right) to show the connection of the wells. Each chamber therefore represents a single experimental unit, thus allowing each plate to be used for multiple-treatment conditions. For instance, one IdMOC-96 plate can be used to evaluate the effect on six cell types for fifteen test articles (using one chamber as control) at a single concentration, two test articles at four concentrations in duplicates or a single test article at four concentrations in quadruplicates
We have recently evaluated the differential cytotoxicity of tamoxifen using the IdMOC system (Li et al., 2004). Further validation of the system is now in progress in our laboratory.
4.5
Conclusions
Accurate prediction of human drug toxicity represents a major challenge for drug development. The high rate of clinical failure of drug candidates that have been carefully selected from preclinical studies illustrates clearly that the routine, ‘classical’ approach of preclinical safety evaluation is inadequate. It is argued here that species–species differences in drug toxicity is a major reason – human-specific toxicity, by definition, cannot be predicted with nonhuman laboratory animals . It is proposed here that human in vivo drug toxicity can be predicted by using a combination of human-based in vitro experimental systems and in vivo laboratory animals. It is fully recognized that regulatory toxicology has evolved to be a rigid discipline, with efficiency as a major objective, and from the drug manufacturers’ viewpoint, regulatory
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acceptance as the ultimate goal. Investigational approaches will be difficult to implement for a new drug candidate at the final stages of development. A practical approach will be to apply investigational approaches early in drug development, using experimental systems such as the in vitro human-based experimental systems described here, to remove drug candidates with human safety concerns as early as possible. Efforts towards designing in vitro experimental approaches to maximize the strengths, applying within their limitations, and incorporating relevant in vivo factors whenever possible, may be rewarded with a better prediction of human drug toxicity before the drugs are exposed to the human population. The use of primary cells, high content assays and IdMOC are described here as potentially important approaches. The challenge of the accurate prediction of human drug toxicity can only be met if one realizes that classical toxicological approaches as practiced now, although important, are not adequate. This realization, plus the willingness to accept, evaluate and apply new approaches, are keys to our future success. It also needs to be recognized that drug safety evaluation and the ultimate development of safe drugs requires scientific insight and expertise from multiple scientific disciplines (Li, 2004b,c).
References Amacher D. E. and Martin B. A. (1997) Tetracycline-induced steatosis in primary canine hepatocyte cultures. Fundam Appl Toxicol 40:256–263. Cysyk R. L., Parker R. J., Barchi J. J. Jr., Steeg P. S., Hartman N. R. and Strong J. M. (2006). Reaction of geldanamycin and C17-substituted analogues with glutathione: product identifications and pharmacological implications. Chem Res Toxicol 19:376–381. Davit B., Reynolds K., Yuan R., Ajayi F., Conner D., Fadiran E., Gillespie B., Sahajwalla C., Huang S. M. and Lesko L. J. (1999). FDA evaluations using in vitro metabolism to predict and interpret in vivo metabolic drug–drug interactions: impact on labeling. J Clin Pharmacol 39:899–910. DiMassi J. A. Hansen R. W. and Grabowski H. G. (2003.) The price of innovation: new estimates of drug development costs. J Health Econ 22:151–185. Easterbrook J., Fackett D. and Li A. P. (2001). A comparison of aroclor 1254-induced and uninduced rat liver microsomes to human liver microsomes in phenytoin O-deethylation, coumarin 7-hydroxylation, tolbutamide 4-hydroxylation, S-mephenytoin 4 -hydroxylation, chloroxazone 6-hydroxylation and testosterone 6beta-hydroxylation. Chem Biol Interact 134:243–249. Erickson-Miller C. L., May R. D., Tomaszewski J., Osborn B., Murphy M. J., Page J. G. and Parchment R. E. (1997). Differential toxicity of camptothecin, topotecan and 9-aminocamptothecin to human, canine and murine myeloid progenitors (CFU-GM) in vitro. Cancer Chemother Pharmacol 39:467– 472. FDA (1997). ‘Drug Metabolism/Drug Interaction Studies in the Drug Development Process: Studies in Vitro’ (issued 4/1997, posted 4/8/1997), Food and Drug Administration (FDA), r r r, r r r [http://www.fda.gov/cder/guidance/clin3.pdf]. FDA (1999). ‘In vivo Drug Metabolism/Drug Interaction Studies – Study Design, Data Analysis and Recommendations for Dosing and Labeling’ (issued 11/24/1999, posted 11/24/1999), Food and Drug Administration (FDA), [http://www.fda.gov/cder/guidance/2635fnl.pdf]. FDA (2006). Guidance for Industry (Draft Guidance). Drug Interaction Studies – design, data analysis, and implications for dosing and labelling. www.fda.gov/cber/gdlns/interactstud.htm. Ganey PE, Luyendyk JP, Maddox JF, Roth RA (2002). Adverse drug reactions: inflammatory episodes as consequence and contributor. Chem Biol Interact 150:35–51.
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Hartman N. R., Cysyk R. L., Bruneau-Wack C., Thenot J. P., Parker R. J. and Strong J. M. (2002). Production of intracellular 35S-glutathione by rat and human hepatocytes for the quantification of xenobiotic reactive intermediates. Chem Biol Interact 142:43–55. Hewitt N. J., Lloyd S., Hayden M., Butler R., Sakai Y., Springer R., Fackett A. and Li A. P. (2002). Correlation between troglitazone cytotoxicity and drug metabolic enzyme activities in cryopreserved human hepatocytes. Chem Biol Interact 142:73–82. Huang S. M., Lesko L. J. and Williams R. L. (1999) Assessment of the quality and quantity of drug– drug interaction studies in recent NDA submissions: study design and data analysis issues. J Clin Pharmacol 39:1006–1014. Jigorel E., Le Vee M., Boursier-Neyret C., Bertrand M. and Fardel O. (2005). Functional expression of sinusoidal drug transporters in primary human and rat hepatocytes. Drug Metab Dispos 33:1418– 1422. Kasahara T., Tomita K., Murano H., Harada T., Tsubakimoto K., Ogihara T., Ohnishi S. and Kakinuma C. (2006). Establishment of an in vitro high-throughput screening assay for detecting phospholipidosis-inducing potential. Toxicol Sci 90:133–141. Kassahun K., Pearson P. G., Tang W., McIntosh I., Leung K., Elmore C., Dean D., Wang R., Doss G. and Baillie T. A. (2000). Future of toxicology–metabolic activation and drug design: challenges and opportunities in chemical toxicology. Chem Res Toxicol 19:889–893. Kier L. D., Neft R., Tang L., Suizu R., Cook T., Onsurez K., Tiegler K., Sakai Y., Ortiz M., Nolan T., Sankar U. and Li A. P. (2004). Applications of microarrays with toxicologically relevant genes (tox genes) for the evaluation of chemical toxicants in Sprague Dawley rats in vivo and human hepatocytes in vitro. Mutat Res 549:101–113. Li A. P. (2002). A review of the common properties of drugs with idiosyncratic hepatotoxicity and the ‘multiple determinant hypothesis’ for the manifestation of idiosyncratic drug toxicity. Chem Biol Interact 142:7–23. Li A. P. (2004a). In vitro approaches to evaluate ADMET drug properties. Curr Top Med Chem 4:701–706. Li A. P. (2004b). An integrated, multidisciplinary approach for drug safety assessment. Drug Discov Today 9:687–693. Li A. P. (2004c). A comprehensive approach for drug safety assessment. Chem Biol Interact 150:27– 33. Li A. P. (2007). Human hepatocytes: isolation, cryopreservation and applications in drug development. Chem Biol Interact 168:16–29. Li A. P., Lu C., Brent J. A., Pham C., Fackett A., Ruegg C. E. and Silber P. M. (1999). Cryopreserved human hepatocytes: characterization of drug-metabolizing enzyme activities and applications in higher throughput screening assays for hepatotoxicity, metabolic stability and drug–drug interaction potential. Chem Biol Interact 121:17–35. Li A. P., Bode C. and Sakai Y. (2004). A novel in vitro system, the integrated discrete multiple organ cell culture (IdMOC) system, for the evaluation of human drug toxicity: comparative cytotoxicity of tamoxifen towards normal human cells from five major organs and MCF-7 adenocarcinoma breast cancer cells. Chem Biol Interact 150:129–136. Liguori M. J., Anderson M. G., Bukofzer S., McKim J., Pregenzer J. F., Retief J., Spear B. B. and Waring J. F. (2005). Microarray analysis in human hepatocytes suggests a mechanism for hepatotoxicity induced by trovafloxacin. Hepatology 41:177–86. Lloyd S., Hayden M. J., Sakai Y., Fackett A., Silber P. M., Hewitt N. J. and Li A. P. (2002). Differential in vitro hepatotoxicity of troglitazone and rosiglitazone among cryopreserved human hepatocytes from 37 donors. Chem Biol Interact 142:57–71. MacGregor J. T., Collins J. M., Sugiyama Y., Tyson C. A., Dean J., Smith L., Andersen M., Curren R. D., Houston J. B., Kadlubar F. F., Kedderis G. L., Krishnan K., Li A. P., Parchment R. E., Thummel K., Tomaszewski J. E., Ulrich R., Vickers A. E. and Wrighton S. A. (2001). In vitro
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human tissue models in risk assessment: report of a consensus-building workshop. Toxicol Sci 59:17–36. Prabhu S., Fackett A., Lloyd S., McClellan H. A., Terrell C. M., Silber P. M. and Li A. P. (2002). Identification of glutathione conjugates of troglitazone in human hepatocytes. Chem Biol Interact 142:83–97. Reasor M. J., Hastings K. L. and Ulrich R. G. (2006). Drug-induced phospholipidosis: issues and future directions. Expert Opin Drug Saf 5:567–583. Rodriguez-Antona C., Donato M. T., Boobis A., Edwards R. J., Watts P. S., Castell J. V. and GomezLechon M. J. (2002). Cytochrome P450 expression in human hepatocytes and hepatoma cell lines: molecular mechanisms that determine lower expression in cultured cells. Xenobiotica 32:505–520. Sahi J., Sinz M. W., Campbell S., Mireles R., Zheng X., Rose K. A., Raeissi S., Hashim M. F., Ye Y., de Morais S. M., Black C., Tugnait M. and Keller L. H. (2006). Metabolism and transportermediated drug–drug interactions of the endothelin-A receptor antagonist CI-1034. Chem Biol Interact 159:156–168. Shitara Y., Itoh T., Sato H., Li A. P. and Sugiyama Y. (2003). Inhibition of transporter-mediated hepatic uptake as a mechanism for drug–drug interaction between cerivastatin and cyclosporin A. J Pharmacol Exp Ther 304:610–616. Zhang Z., Chen Q., Li Y., Doss G. A., Dean B. J., Ngui J. S., Silva Elipe M., Kim S., Wu J. Y., Dininno F., Hammond M. L., Stearns R.A., Evans D. C., Baillie T. A. and Tang W. (2005). In vitro bioactivation of dihydrobenzoxathiin selective estrogen receptor modulators by cytochrome P450 3A4 in human liver microsomes: formation of reactive iminium and quinone type metabolites. Chem Res Toxicol 18:675–685.
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5 Hepatocytes as a Model for Screening Food-Related Hepatotoxins and Studying Mechanisms of their Toxicity Saura C. Sahu
5.1
Introduction
Liver is an important organ, which plays a central role in the metabolism of toxins entering the body through the digestive tract and transported via the portal vein. It is the primary organ involved in xenobiotic metabolism and is a major target organ for chemicals, drugs and microbial pathogens (Treinen-Molsen, 2001). Hepatotoxicity is also a common adverse health effect of food-related toxins (Miller and Miller, 1986; Willett et al., 2004; Stickel et al., 2005), and it is thus one of the most serious safety concerns for food additives, food contaminants, dietary supplements, food-borne microbial pathogens and other food-related products regulated by the US Food and Drug Administration (FDA). In vivo animal studies for assessing the hepatotoxic potential of food-related toxins are expensive and time-consuming. For these reasons, alternate in vitro models are important to complement and/or supplement the animal studies for toxicological risk assessment (Green et al., 2001; MacGregor et al., 2001). The in vitro models are relatively inexpensive and they provide well-defined and reproducible experimental conditions for toxicity testing. They are excellent systems to study mechanisms of toxicity and structure–activity relationships at the cellular and molecular level. They are also very useful tools for rapid screening of potential toxins.
Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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The US Food and Drug Administration (FDA) is mandated by the NIH Revitalization Act of 1993 (P.L. No. 103-43) in concert with the ICCVAM Authorization Act of 2000 (P.L. No. 106-545) for federal regulatory agencies to: ‘(a) establish criteria for the validation and regulatory acceptance of alternative testing methods and (b) recommend a process through which scientifically validated alternative methods can be accepted for regulatory use’. Criteria for validation of alternative toxicological test methods have been published (ICCVAM, 1997). The use of in vitro cytotoxicity results to predict starting doses for in vivo acute toxicity studies reducing the number of animals for safety evaluation by the regulatory agencies is allowed (Federal Register, 2001). Primary hepatocytes and some hepatocyte cell lines retain many of the metabolic enzymes characteristic of the intact liver in vivo (Hengstler et al., 2000, 2002; Runge et al., 2001). Therefore, they represent an excellent in vitro model for studying liver function, xenobiotic metabolism, pharmacology and toxicology. They are widely used for the evaluation of liver functions and toxicity of chemicals, drugs and microbes (Barsig et al., 1998; Castell et al., 1997; Li et al., 1997; Michalopoulos, 1999; Sahu et al., 2001, 2006).
5.2 5.2.1
Hepatocyte Cultures Hepatocytes in Primary Culture
Hepatocytes for primary culture are isolated by the two-step in situ collagenase perfusion of liver following the method of Seglen (1976). Isolated hepatocytes survive for a few hours, but they can be cryopreserved for a couple of years with somewhat reduced viability and metabolizing enzyme activities (Chesne et al., 1993). Long-term culture of hepatocytes has been achieved using co-culture with other cells (Donata et al., 1994), in dimethyl sulfoxide (DMSO)-supplemented medium (Isom et al., 1985), collagen sandwich (Kono et al., 1995), collagen-coated dishes and serum-free medium supplemented with hormones and growth factors (Ferrini et al., 1997), cell aggregation (Tateno and Yoshizato, 1996), gel entrapment (Guyomard et al., 1996) and spheroid cultures (Walker et al., 2000; Hamilton et al., 2001; Khalil et al., 2001). Rat hepatocytes in primary culture are the most frequently used and best characterized in vitro model for testing liver toxicity of chemicals and drugs. They have a long history of use by many laboratories throughout the world (Carfagna et al., 1996; DiPetrillo et al., 2002; Hill and Roth, 1998; Hussain and Frazier, 2003; Sahu et al., 2001; Susa et al., 1997; Tirmenstein et al., 2000; Tseng et al., 1997). Hepatotoxicity data obtained from the primary rat hepatocytes are consistent with the data obtained in vivo from rats (Kefalas and Stacey, 1993), and thus, the in vitro and in vivo data obtained from rat studies appear to be complementary. In addition, the in vitro data obtained from the primary rat hepatocytes can be validated with data obtained from in vivo studies. Human hepatocytes in primary culture are the ideal in vitro system for hepatotoxicity assessment of regulated products. However, their availability is limited. Cryopreserved primary human hepatocytes are commercially available, but optimal conditions for maintaining human hepatocytes in culture have not been perfected yet. Unlike the primary rat hepatocytes, which have been extensively used in research for almost three decades, the use of primary human hepatocytes is relatively recent. High inter-individual variability in
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phenotypes and genotypes of human liver, as well as large variations in individual consumption of various diets, are important factors for consideration. For example, donated human liver tissues from nine males and nine females were used to evaluate gender differences in the gene expression of human livers (Delongchamp et al., 2005). This study, which tested 31 100 genes, concluded that the gene expression of 224 genes differed between sexes. The observed gender differences in expression were small. False discovery rates exceeded 80 % for every set of genes selected, making it impossible to identify specific genes with gender differences (Delongchamp et al., 2005). In addition, the in vitro data obtained from cryopreserved human hepatocytes would be difficult to validate in vivo. These are examples of some of the unresolved issues and challenges associated with the use of cryopreserved human hepatocytes for obtaining experimental data that can be used for risk assessment by regulatory agencies. However, as continued research clarifies these issues there will undoubtedly be increasing use of this important experimental system for safety evaluation and risk management of regulated products. 5.2.2
Hepatocyte Cell Lines
Hepatocyte cell lines derived from animal and human liver have been used for hepatotoxicity testing. Rat clone-9 hepatocytes originally isolated from the normal liver of a 4-week-old male Sprague–Dawley rat were used as a model to test chemical hepatotoxicity (Barhoumi et al., 2002; Grunne et al., 2001; Neerman et al., 2004; Reeves et al., 2001; Sahu et al., 2006; Thompson et al., 1998). Mouse BNL CL.2 hepatocytes originally isolated from an embryonic liver of a normal BALB/c mouse were used as a model for testing virulence and intracellular growth of Listeria monocytogenes (Haponsaph and Czuprynski, 1996). Human WRL-68 hepatocytes originally isolated from the normal embryonic liver of a human fetus were used to test hepatotoxicity of metals (Lopez-Ortal et al., 1999; Olivares et al., 1997; Ramirez et al., 2000). These hepatocytes are a useful tool for studying hepatitis virus replication in vitro (Tagawa et al., 1995). Human hepatocyte Hep G2 cells originally isolated from the hepatocellular carcinoma of a 15-year-old Caucasian male are widely used in hepatotoxicity testing. Scheers et al. (2001) found a good correlation between the long-term cytotoxicity results of 27 chemicals in Hep G2 cells and their acute human toxicity (Ekwall et al., 1999) in a multiple evaluation of an in vitro cytotoxicity study in Sweden. They concluded that this in vitro model is a good predictor of long-term human toxicity.
5.3
Limitations of the Hepatocyte Culture Model
The major drawbacks with the hepatocyte culture model are the loss of liver architecture and the degenerative changes following the cell isolation which make this in vitro tool unsuitable for long-term studies. However, with the improved culture conditions, hepatocytes can be maintained in culture with cell integrity and high metabolizing capacities for several weeks (Tateno and Yoshizato, 1996; Michalopoulos, 1999). Another limitation for this popular model is the absence of non-parenchymal cells, which makes it unsuitable for assessing hepatotoxicity mediated by non-parenchymal cells. This limitation is overcome by cocultures of hepatocytes with other cells (Guguen-Guillouzo et al., 1983; Donato et al.,
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1994). Marked phenotypic stabilization of hepatocytes has been observed by co-culturing them with liver epithelial cells (Guguen-Guillouzo et al., 1983). An important aspect of an in vitro screening system is its validation by establishing a correlation with results obtained from in vivo studies. This can be done using appropriate animal models. With proper validation the hepatocyte model is a promising in vitro system for use in quantitative risk assessment (Walum et al., 1992).
5.4
Endpoints of Hepatotoxicity
Hepatotoxicity is not induced by any particular single mechanism; it may involve many different changes in cellular function, the detection of which provides sensitive indices of hepatotoxicity (Lee, 2003) as described below. 5.4.1
Cytotoxicity
Injury to the cell membrane leads to leakage of cytoplasmic enzymes. Therefore, cytotoxicity is a common index of this facet of hepatotoxicity. The in vitro cytotoxicity is a good predictor of acute in vivo hepatotoxicity for safety assessment (Federal Register, 2001). It can be used for screening hepatotoxins (Sahu, 2003; Dambach et al., 2005). 5.4.2
Cell Death
Cell death caused by necrosis or apoptosis is the ultimate endpoint of toxicity. Hepatocyte cultures are used to study apoptosis and cell death. Hepatocyte apoptosis is associated with retention of bile constituents in hepatocytes during cholestasis (Patel et al., 1998). The foodborne pathogen Listeria monocytogenes induces apoptosis in infected mouse hepatocytes (Rogers et al., 1996). Oxidative stress induces apoptosis in rat hepatocyte cultures (Haidara et al., 2002). The dietary carcinogen 3-amino-1,4-dimethyl-5H-pyridole[4,3-b]indole induces apoptosis in rat hepatocytes (Ashida et al., 1998). 5.4.3
Oxidative Stress
Reactive oxygen species induce oxidative stress which plays an important role in many human diseases (Sahu, 1991; McCord, 1995; De Maria et al., 1996), in the toxicity of many xenobiotics (Kehrer, 1993), and in viral (Schwarz, 1996) as well as bacterial infections of the liver (Inoue et al., 1995). Aflatoxin B1 (AFB1 ) induces oxidative stress in cultured rat hepatocytes (Shen et al., 1995,1996). AFB1 cytotoxicity in rat hepatocytes is associated with cellular lipid peroxidation (Shen et al., 1995). The flavonoid myricetin induces oxidative stress leading to lipid peroxidation and DNA strand breaks in cultured rat hepatocytes (Sahu et al., 2001). Oxidative stress leading to lipid and protein oxidation occurs in chronic hepatitis C (De Maria et al., 1996). Nitric oxide plays an important role in hepatotoxicity (Muriel, 2000). Hepatocytes express inducible nitric oxide synthatase (Muriel, 2000). The hepatotoxin acetaminophen induces nitric oxide synthatase in rat hepatocytes (Gardner et al., 1998). The human hepatoblastoma HepG2 cell cultures have been used to evaluate the protective effects of antioxidants on oxidative liver injury (Yang et al., 1999). Lipid peroxides (Stacey and Klaassen, 1981), 8-hydroxydeoxyguanosin (Shigenaga et al., 1990)
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and dichlorofluorescein (Yerushalmi et al., 2001) produced by cellular reactive oxygen species are commonly used endpoints of oxidative stress. 5.4.4
Mitochondrial Dysfunction
Mitochondrial dysfunction is an early marker of cell stress. Mitochondrial permeability leads to release of cytochrome-C from mitochondria and induction of apopotic cell death. Hepatocyte cultures are used to screen toxins that induce mitochondrial dysfunction (Rat et al., 1994). 5.4.5
Cytochrome P450 Induction
Cytochrome P450 enzymes catalyze the oxidation of endogenous substrates and xenobiotics. Induction of these enzymes alters cellular metabolism leading to hepatotoxicity (Guengerich, 1991). This has been used to screen hepatotoxins in hepatocyte cultures (Dambach et al., 2005). 5.4.6
Hepatosteatosis
Accumulation of lipids in hepatocytes leads to ‘fatty liver’ or hepatosteatosis. Lipid accumulation can result from altered mitochondrial lipid oxidation or from altered secretion of lipoproteins. The human hepatoma cell line Hep G2 has been used for studying hepatosteatosis (McMillian et al., 2001).
5.5 5.5.1
Application of Hepatocyte Cultures Screening of Food-Related Products for Potential Hepatotoxicity
Hepatocyte cultures have been widely used for screening chemicals, drugs, carcinogens, mutagens, mycotoxins, microbial pathogens and viruses for their hepatotoxic potential (Sahu, 2003; Dambach et al., 2005). Hepatotoxicity is routinely assessed in the safety evaluation of food additives and is a common adverse health effect of food contaminants and dietary supplements. Many of the botanical herbal products labeled as dietary supplements show hepatotoxicity (Willett et al., 2004; Stickel et al., 2005). Dietary supplements such as Chaparral (Sheikh et al., 1997), Ephedra (Bajaj et al., 2003), Germander (Lekehal et al., 1996), Kava (Teschke et al., 2003), LipoKinetix (Favreau et al., 2002; Novak and Lewis, 2003) and pyrrolizidine alkaloids (Stedman, 2002) are known hepatotoxins. Rat hepatocytes have been shown to be a good in vitro model for testing hepatotoxicity of herbal products (Lekehal et al., 1996). The human hepatocellular carcinoma cell line Hep G2 was found to be an excellent model for assessing cytotoxicity of medicinal plant extracts (Ruffa et al., 2002). 5.5.2
Toxicogenomics Studies
Hepatotoxicity involves changes in gene expression, which can be detected by DNA microarrays (Minami et al., 2006). Such studies are used to identify mechanisms of toxicity. Hepatocyte culture has been used for toxicogenomics studies. Harris et al. (2001) have used the cultured human hepatoma cell line HepG2 to study the gene expression changes
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induced by carbon tetrachloride and ethanol. De Longueville et al. (2003) used primary rat hepatocyte culture to study gene expression patterns in eleven different hepatotoxins. Delongchamp et al. (2005) used a primary human hepatocyte culture to evaluate gender differences in hepatotoxicity. This technology has been used to study acute hepatitis C virus infection in liver (Bigger et al., 2001) and to investigate host–microbe interactions (Cummings and Relman, 2000). 5.5.3
Interactive Hepatotoxicity Studies
Humans are exposed to many different potential toxins. Their interaction is a matter of concern. Synergistic enhancement of toxicity is not uncommon. For example, lipopolysaccharide (LPS) has been shown to influence toxicity of xenobiotics (Cebula et al., 1984; Zhou et al., 2000; Wiesenfeld et al., 2006). Rat hepatocytes have been used as an experimental model to study the interactive toxicities of carbon tetrachloride and trichloroethylene (Kefalas and Stacey, 1993). Primary rat hepatocytes have been used to study the interactive hepatotoxicity of flunitrazepam and ethanol (Assaf and Abdel-Rahman, 1999) as well as LPS and allyl alcohol (Sneed et al., 2000). Therefore, hepatocyte cultures can be used as a model to predict interactive hepatotoxicity of food-related products. 5.5.4
Studies on Species Differences in Hepatotoxicity
Hepatocyte cultures are predictive models for metabolism and pharmakokinetics in different species. Good correlation between in vitro and in vivo drug metabolism has been established using animal and human hepatocyte cultures (Sandker et al., 1994; Pathernik et al., 1995). Human, dog and rat hepatocyte cultures have been shown to be valuable in vitro tools for study of species differences in drug metabolism and pharmacokinetics (Bayliss et al., 1999). Differences in sensitivity towards chemical toxicity have been demonstrated in rat and human hepatocyte cultures (Merrill et al., 1995). Similar sensitivity towards chloroforminduced cytolethality is reported in mouse and rat hepatocytes (Amman et al., 1998). Species-dependent differences in glutathione conjugation of perchloroethylene has been studied in rat and mouse hepatocyte cultures (Lash et al., 1998). A similar metabolism of thiabendazole in rat, rabbit, calf, pig and sheep hepatocytes has been reported (Coulet et al., 1998). 5.5.5
Studies on Gender Differences in Hepatotoxicity
Gender difference plays an important role in the toxic responses of the liver (TreinenMoslen, 2001). Females are predisposed to hepatotoxicity and concomitant agents that induce cytochrome P450 enzymes also increase individual susceptibility (Stedman, 2002). Epidemiological evidence shows that women have greater susceptibility to alcohol-induced liver damage than men (Jensen, 1996; Schenker, 1997). Drug-induced acute liver failure occurs more frequently in women than men (Miller, 2001). Adverse reactions to therapeutic drugs are more common in women than men (Kando et al., 1995). The female mouse is more susceptible to fumonisin B1 -induced hepatocellular neoplasm than the male mouse (NTP, 1999). However, cocaine causes more liver damage in male mice than female mice (Visalli et al., 2004). There exists a distinct sexual dimorphism
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in rat liver cytochrome P450 (CYP) activity (Lewis et al., 1998). Intrinsic sexual differences in hepatocytes of males and females appear to result in different levels of responses to CYP. This intrinsic sexual difference in hepatocytes is more pronounced in rats than humans. Successful use of the primary rat hepatocytes in culture as an in vitro model for evaluating the sex differences in hepatotoxicity has been demonstrated (Carfagna et al., 1996). This model has been used to show the gender difference in hepatotoxicity of safingol (Carfagna et al., 1996).
5.6
Conclusions
The hepatocyte model is an established, well-characterized and widely used popular in vitro tool for hepatotoxicity testing (Groneberg et al., 2002). The standardized methods of hepatocyte isolation from a wide range of species, including humans, and their maintenance in culture with high metabolizing capacity for several days under closely controlled and easily manipulated conditions have made this model the most widely used in vitro tool for studies on the metabolism, toxicity and mechanisms of action of chemicals and drugs (El-Sakka et al., 2002; Yamamoto et al., 2002; Dvorak et al., 2002; Shiota et al., 2002; Haidara et al., 2002; Frenzel et al., 2002; Qu et al., 2001; Muriel, 2000), for screening of mutagens, carcinogens and mycotoxins (Uhl et al., 2000; Knasmuller et al., 1997), for virulence assessment of microbial pathogens and viruses (Wing and Gregory, 2002; Galle et al., 1994), for qualitative and quantitative inter-species comparison (Bayliss et al., 1999; Ammann et al., 1998; Coulet et al., 1998; Lash et al., 1998; Merril et al., 1995; Sandker et al., 1994) and for toxicogenomics studies (Bigger et al., 2001; Harris et al., 2001; Cummings and Relman, 2000; De Longueville et al., 2003; Delongchamp et al., 2005; Minami et al., 2006). The hepatocyte model provides a homogeneous preparation of a single cell type, which can be cryopreserved without loss of significant functional activity (Chesne et al., 1993). Numerous studies have reported good correlation between in vitro hepatocytes and in vivo situations (Li, 1997; Li et al., 1997; Pathernick et al., 1995; Sandker et al., 1994). The use of in vitro models in hazard assessment requires careful consideration. At the present time, there is insufficient evidence for their use in quantitative risk assessment. However, they are suitable for use in qualitative hazard assessment, information from which can be used for quantitative risk analysis (Walum et al., 1992). In summary, the hepatocyte cell culture is a good in vitro tool for evaluating hepatotoxic potential of herbal products, dietary supplements, food additives, food-borne toxicants and microbial pathogens. The results obtained from such in vitro screening can be used to guide the development of in vivo studies to assess the safety of test materials of interest.
Acknowledgments I wish to thank Dr Thomas J. Sobotka for his critical review of this manuscript.
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6 Some Experimental Models of Liver Damage Pablo Muriel
6.1
Introduction
The liver is the largest gland in the human body and accounts for approximately 2.5 % of total body weight. The liver weighs almost 1500 kg in the adult. The liver is divided into four lobes. The right and left lobes are separated by the falciform ligament. These lobes are supplied by the right and left branches of the hepatic artery and the portal vein. Bile is drained from the liver by the left and right hepatic ducts. The right lobe is further divided into two smaller lobes: the quadrate and the caudate. The liver is a unique organ anatomically located to serve its dual role as a metabolic and biochemical transformation ‘factory’. The liver receives blood containing substances absorbed or secreted by the gastrointestinal organs including the spleen, intestine, stomach and pancreas. It uses these substances as raw materials and modifies them or synthesizes new chemicals. These are then returned to the blood stream or to bile for excretion. The vulnerability of the liver to injury is as much a function of its anatomical proximity to the blood supply and digestive tract as to its ability to concentrate and biotransform xenobiotics. Xenobiotics in the blood pass through the hepatic artery and portal vein and then drain through the central and the hepatic veins into the vena cava. The principal liver duct joins the cystic duct from the gall bladder to form the common bile duct, which drains into the duodenum. The literature on experimental hepatotoxicity has become enormous (Zimmerman, 1999). A fascinating area in its own right, it includes many facets relevant to human and veterinary medicine. Experimentally induced liver damage allows studies of human liver diseases,
Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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accidental and industrial toxicity, screening of medicinal agents for potential hepatotoxic effects, studies of hepatic physiology, pathology, and regeneration, and the development of diagnostic tools. Models of liver damage can provide useful tools for the study of teaching of hepatic histopathology. Almost all of the known acute and chronic lesions of the liver can be induced experimentally. These include necrosis (zonal, massive or diffuse), steatosis, hepatic venular lesions and cirrhosis of several morphological types. Specific lesions such as sinusoidal ‘free’ acidophilic bodies (apoptotic bodies), nucleolar and nuclear abnormalities and megalocytes can be produced by several experimental models of hepatic injury. Cholestasis can also be produced experimentally. Models of liver damage have used whole animals or various in vitro preparations. Use of whole animals is essential to demonstrate that an agent has an adverse effect on the liver in a setting of physiologic significance. Whole animals also must be employed if the effects of various factors and manipulations on disease are to have meaning for the mechanism of injury and for the pathophysiologic impact of the hepatic injury. In this review, we will focus on in vivo experimental models of hepatic injury. Studies have used a variety of animal species. Most popular have been rats because of their size and relatively low cost. Most of the accumulated information related to liver damage and on modifiers of susceptibility, such as age, sex and diet, applies to the rat (Zimmerman, 1999). The general use of a relatively uniform experimental model permits comparison of results from different laboratories. Mice, guinea pigs, hamsters, rabbits, gerbils, cats, dogs, horses, sheep, etc., have been employed to various degrees. Primates have also been used because of the greater similarity to disease in humans. In the following section, the most common models of liver injury (Figure 6.1) will be described.
Figure 6.1 The most common models of experimental liver damage. Note that in some cases the causative agent is the same as in human diseases while in others it is not: BDL, bile duct ligation
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Carbon Tetrachloride
Carbon tetrachloride (CCl4 ) once was used widely as a solvent, cleaner and degreaser, both for industrial and home use. Today, the scientific database on the effects of haloalkanes is so vast that it is no longer employed for such purposes although it is used as a model of experimental liver injury (Weber et al., 2003). Today, CCl4 proves highly useful as an experimental model for the study of certain hepatotoxic effects (Muriel, 1997; Muriel et al., 2003; Moreno and Muriel, 2006). It consistently produces liver injury in many species, including non-human primates (Kumar et al., 1972; Yoshida et al., 1999). CCl4 -induced toxicity, depending on dose and duration of exposure, covers a variety of effects. At low doses, transient effects prevail, such as loss of Ca2+ homeostasis (Muriel and Mourelle, 1990), lipid peroxidation (Muriel, 1997), release of noxious or beneficial cytokines (Kyung-Hyun et al., 2006; Muriel, 2007) and apoptotic events followed by regeneration. Other effects, with higher doses or longer exposure, are more serious and develop over a longer period of time, such as fatty degeneration, fibrosis, cirrhosis and even cancer (Weber et al., 2003). In addition, acute intoxication with CCl4 at high doses, when the hepatocelular necrosis exceeds the regenerative capacity of the liver, fatal liver failure will ensue. Extreme doses of CCl4 result in nonspecific solvent toxicity, including central nervous system depression and respiratory failure and death (Berger et al., 1986). r CCl4 -metabolism begins with the formation of the trichloromethyl free radical, CCl3 (McCay et al., 1984) through the action of the mixed function cytochrome P450 oxygenase system of the endoplasmic reticulum (Recknagel et al., 1989). This process involves reductive cleavage of a carbon–chlorine bond. Free radical activation of CCl4 in mitochondria has also been observed (Tomasi et al., 1987) and may contribute significantly to its toxicity. The major cytochrome isozyme to execute biotransformation of CCl4 is CYP2E1, but CYP2B1 and CYP2B2 are also able to attack the CCl4 molecule (Gruebele et al., 1996). In humans, CYP2E1 dominates CCl4 metabolism at environmentally relevant concentrations, but at higher concentrations other cytochromes, particularly CYP3A, also contribute importantly r (Zangar et al., 2000). The resulting CCl3 radical is reactive enough to bind covalently to CYP2E1, either to the active site of the enzyme (Manno et al., 1992) or to the heme group (Fernandez et al., 1982), thus causing its own inactivation (Fuji, 1997). r The CCl3 radical reacts with several important biological substances, like fatty acids r proteins, lipids, nucleic acids and amino acids (Weber et al., 2003). CCl3 also acts by abstracting hydrogen from unsaturated fatty acids to form chloroform. DNA adducts is a mechanism for CCl4 -induced carcinogenesis (DiRenzo et al., 1982). 6.2.1
Lipid Peroxidation r The CCl3 , radical in the presence of oxygen, is transformed into the trichloromethyl peroxy r r radical, CCl3 OO . This radical is more reactive and thus shorter lived than the CCl3 radical r (Mico and Pohl, 1983). The lifetime of the CCl3 OO radical is in the millisecond range and disappears from the tissue by reacting with suitable substrates to complete its electron r r pair. CCl3 OO is more likely than CCl3 to abstract hydrogen from polyunsaturated fatty acids (PUFAs) (Forni et al., 1983) leading to lipid peroxidation (Comporti, 1985; Trible et al., 1987). The abstraction of hydrogen from PUFAs starts sequential reactions that finish in the complete disintegration of the fatty acid molecules with the consequent formation
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of aldehydes, other carbonyls and alkanes in a process called lipid peroxidation. It is important to point out that lipid peroxidation may produce injury by compromising the integrity of membranes and by covalent binding of reactive intermediates to important biological molecules. 6.2.2
Aldehyde Toxicity
The products of lipid peroxidation may produce further liver damage (Esterbauer et al., 1982,1991). They may inhibit enzymes (Hruszkewycz et al., 1978), damage DNA (Ueda et al., 1985), and block lipoprotein secretion. Reactive aldehydes first appear in liver about 6 h after CCl4 intoxication, show a maximum at 24 h and disappear by 36 to 72 h latter r r (Hartley et al., 1999). Aldehydes are less reactive than CCl3 or CCl3 OO , and thus they may reach other organs. However, they do not usually reach toxic concentrations. This is because of their unspecific action because they react typically with amino or sulfhydryl groups, and so a large number of groups need to be inactivated before measurable damage occurs (Sundari and Ramakrishna, 1997). The activities of aldehydes displaying a 4-hydroxy-2,3-trans-unsaturated configuration and especially of 4-OH-2,3-trans-nonenal, is favored, and they are formed in high concentrations after exposure of cells to CCl4 (Benedetti et al., 1980, 1984). These compounds inhibit several enzymes, including adenylate cyclase, at pathophysiological concentrations (Poli et al., 1989). There is also evidence that 4-hydroxynonenal decreases cytochrome P450 (Poli et al., 1989) and inhibits the activities of Ca2+ (Parola et al., 1990) and Na+ , K+ (Morel et al., 1998) –ATPases, and phosphatase and protein kinase activities (Omura et al., 1999). The effect of CCl4 -derived haloalkanes on these enzyme activities may contribute to CCl4 -induced toxicity
6.3
Hypomethylation
It has been reported that in connection with CCl4 -induced inhibition of protein synthesis an important deficit in ribosomal RNA methylation occurs (Clawson et al., 1987). The synthesis of polyamines is also reduced by CCl4 intoxication (Rao et al., 1989). Remethylation of CCl4 intoxicated ribosomes recuperates their normal function (Clawson et al., 1987). These researchers suggest that disturbance of cytosolic Ca2+ levels are to blame for the effect. Carbon tetrachloride inhibits DNA methylation (Ruchirawat et al., 1983) and treatment of rats for three weeks leads to global hypomethylation of liver DNA; this deficiency can be corrected by external supplementation of the methyl donor S-adenosylmethionine (SAM) (Varela-Moreiras et al., 1995). Moreover, several alterations observed in CCl4 -induced cirrhosis, mainly on lipids, can be completely prevented by SAM treatment (Muriel and Mourelle, 1992a). 6.3.1
Calcium Homeostasis
It is very well known that CCl4 increases intracellular Ca2+ levels in hepatocytes (Weber et al., 2003). This effect compromises the viability of the cell because the intracellular concentration of Ca2+ is maintained about four orders of magnitude lower than the extracellular space.
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An early effect is observed only 15 min after the administration of CCl4 to hepatocytes from mitochondria (Albano et al., 1985) and endoplasmic reticulum because the mitochondrial uniport system and the Ca2+ -ATPase of the endoplasmic reticulum become inactivated (Tsokos-Kuhn et al., 1985). This results in increased cytosolic Ca2+ (Long and Moore, 1986). However, mitochondria and endoplasmic reticulum may be responsible for determining the level of intracellular calcium in the short term, but these organelles cannot continue to accumulate Ca2+ within themselves indefinitely; then, in the long term, the plasma membrane Ca2+ -ATPase must be responsible for maintaining the ultimate level of Ca2+ within the cell and CCl4 strongly inhibits this ATPase activity (Muriel and Mourelle, 1992b). Increased levels of Ca2+ in the cell destroy cytoskeletal structures and activate several catabolic enzymes such as endonucleases, proteases and phospholipases, whose continued action results in cell death via apoptosis or necrosis (Nicotera et al., 1992; Zawaski et al., 1993). 6.3.2
Acute CCl4 Intoxication
Acute intoxication in rats or mice are frequently used to produce toxic hepatitis in order to test possible hepatoprotective agents, mainly those with antioxidant properties, and to study liver regeneration after a chemical hepatectomy (Muriel and Mourelle, 1992c; Sotelo-F´elix et al., 2002; Agarwal et al., 2006). 6.3.3
CCl4 -Induced Cirrhosis
Lamson and Wing (1926) were the first to report that CCl4 intoxication produced cirrhosis, while later Cameron and Karunaratne (1932) published a systematic study that established the morphology and show the standard experimental conditions of the model. Nowadays, CCl4 -induced cirrhosis is probably the most widely used model to reproduce cirrhosis in rats and mice (Moreno and Muriel, 2006). To produce experimental cirrhosis, it is necessary to give repeated doses of CCl4 , regardless of the animal and route of administration. The time interval between each ‘insult’ should not be too long, since damaged liver can recover, rendering the effect of the toxin noncumulative (Cameron and Karunaratne, 1932). In our laboratory, cirrhosis is induced in male Wistar rats weighing initially around 100 g – CCl4 is dissolved in liquid petroleum and administered three times per week intraperitoneally at a dose of 0.4 g/kg. Cirrhosis is observed after two months and severity of the disease increased after three months of treatment; after four months, mortality is too high (over 80 %) (Muriel et al., 2005). Well-established CCl4 -cirrhosis is characterized by changes involving several abdominal organs. The liver is frequently enlarged; however, in advanced states of the disease, it may be smaller than normal with gross nodularity. Splenomegaly and ascites are usually present. Increased mesenteric fat lobulation and lymph node hyperplasia frequently occur. Two main problems with CCl4 -cirrhosis are that the response of individual animals to the toxin is variable and that mortality is high (30–60 %); interestingly, these factors also occur in human cirrhosis. Experimental CCl4 -induced cirrhosis seems to reproduce the main characteristics of human cirrhosis: the liver is grossly nodular, there is portal hypertension in most of the animals and the normal architecture is replaced by nodules of regenerating liver surrounded by fibrotic tissue septa with proliferated bile ducts. Portocabal anastomosis develops within
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the connective tissue septa (Daniel et al., 1952), as occurs in man. However, a deeper analysis shows important differences between this experimental model and its human counterpart (P´erez-Tamayo, 1983). Alcoholic cirrhosis is frequently preceded or accompanied by centrilobular sclerosing hyaline necrosis and pericelular and interlaminar fibrosis; these characteristics are not observed in the CCl4 model of cirrhosis. In summary, this model of experimental cirrhosis shares several features with alcoholic human cirrhosis but also shows various differences (P´erez-Tamayo, 1983). The CCl4 -induced cirrhosis is the most widely model used to reproduce human cirrhosis in experimental animals to study pathophysiological mechanisms of the disease (Muriel, 1997, 2006, 2007; Muriel and Escobar, 2003; Moreno and Muriel, 2006; Chavez et al., 2006) and many possible antifibrotic drugs are tested in this model before using them in humans (Cedillo et al., 1996; Muriel et al., 2003; Kyung-Hyun et al., 2006).
6.4
Acetaminophen (Paracetamol)
Acetaminophen (paracetamol; APAP) in normal therapeutic doses is generally considered one of the safest aspirin-like analgesics. However, large overdoses can cause severe hepatic damage and even liver failure. Overdose of APAP is the most frequent cause of druginduced hepatic failure in USA and UK (Lee, 2004). In addition to its clinical relevance, APAP experimental intoxication is widely used as a model of liver damage. Recently, two excellent reviews have appeared on APAP-induced hepatic injury; one of them is concerned with the role of inflammation (Jaeschke, 2005a) while the other deals with intracellular signaling (Jaeschke and Bajt, 2006) in the mechanism of APAP-induced liver damage. Therefore, here I am only going to summarize the most prominent features of APAP overdosing. Since APAP toxicity is associated with depletion of GSH (Jollow et al., 1973; Mitchell et al., 1973a,b), the GSH precursor N -acetylcysteine (NAC), introduced in the 1970s, is an effective antidote against APAP hepatic damage. However, no other effective drug is available in the clinic today. The inflammatory response is dependent on the initial damage to the hepatic cells; therefore, intracellular events may offer interesting targets for therapeutic interventions. When an overdose occurs, a fraction of APAP is activated metabolically to the reactive metabolite N -acethyl- p-benzoquinone imine (NAPQI) that first consumes cellular glutathione and then binds covalently to proteins. This leads to increases in the intracellular Ca2+ concentration, ‘Bax’ and ‘Bid’ translocation to the mitochondria and oxidant stress and peroxynitrite formation in the mitochondria. Reactive oxygen species and peroxynitrite induce the membrane permeability transition that causes the collapse of the mitochondrial membrane potential and ‘abolishes’ ATP synthesis. In fact, some free radical scavengers prevent experimental APAP induced liver injury (Muriel et al., 1992; Muriel, 1997). The decreasing of ATP levels appears to prevent caspase activation by the release of cytochrome c and Smac. The apoptosis-inducing factor (AIF) (Susin et al., 2000) and endonuclease G (van Loo et al., 2001) induce chromatin condensation and nuclear DNA fragmentation, respectively. The extensive DNA damage and the rapid elimination of mitochondria, together with activation of intracellular proteases (calpains), conduce to plasmatic membrane breakdown and oncotic necrosis of the hepatic cells. The massive cell death and liver failure after APAP intoxication can be explained by these intracellular events. Many aspects,
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however, require further investigation. Furthermore, APAP-induced hepatic injury can be modulated by changes in the activity of cytochromes P450 and Phase II detoxification enzymes, modulation in the GSH/GSSG levels and redox state of the cell. Experimental APAP intoxication is a useful model to study the pathophysiology and pharmacology of an overdose of the drug in humans. In addition, it provides a useful tool to study necrosis and apoptosis mechanisms and the effects of hepatoprotective drugs. For example, recently, the receptor antagonist (BN52021) to platelet activating factor (PAF) prevented acute APAP toxicity in rats (Grypioti et al., 2006). This was explained because PAF is an endogenous mediator of various proinflammatory processes in the liver.
6.5
Cholestasis
Bile is the exocrine secretion of the liver. Like most other secretions, it is an aqueous solution of organic and inorganic compounds. Bile acids, bile pigments, cholesterol and phospholipids are the major organic compounds. Bile also contains small amounts of protein. Bile is normally secreted from the hepatocyte to the duodenum, a process that normally depends on the function of several membrane transport systems in hepatocytes and cholangiocytes, and on the functional and structural integrity of the bile-secretory apparatus. Cholestasis comes from the Greek ‘chole’ which means bile and ‘stasis’, standing still; thus meaning bile flow stagnation that can be the result of a failure in the transports of hepatocytes or ductular cells or from a physical blockade of the external bile ducts. The former is considered intrahepatic, and the latter extrahepatic cholestasis. 6.5.1
Human Cholestasis
The main etiologic agents of adult intrahepatic cholestasis are drugs and pregnancy. Antibiotics are the most frequent medications that cause intrahepatic cholestasis. In addition, estrogens, cyclosporine A, rifamycin SV, rifampicin, glibenclamide, chlorpromazine, erythromycin and oxypenicillins may produce intrahepatic cholestasis (Rodr´ıguez-Garay, 2003). Obstructive cholestasis, extrahepatic biliary atresia (EBA), primary sclerosing cholestasis (PSC) and primary biliary cirrhosis (PBC) are other causes of human cholestasis. PBC, an autoimmune cholestatic liver disease, is accompanied by reduced levels of anion exchanger 2 immunoreactivity at the bile canaliculus and bile ducts (Medina et al., 1997). A stone or a tumor at the level of the extrahepatic bile ducts usually leads to extrahepatic cholestasis. The use of animals to reproduce cholestatic disease in the human is a useful tool to study the pathophysiology of the disease and also to test potential liver drugs with anticholestatic properties. 6.5.2
Experimental Prolonged Bile Duct Ligation (Biliary Cirrhosis)
The effects of conventional bile duct ligation (BDL) on hepatic morphology have been examined extensively since publication of the earliest histological studies approximately 70 years ago (Cameron and Oakley, 1932; Cameron and Hasan, 1958; Trams and Symeonidis, 1957). Kountouras et al. (1984) reported a systematic study of prolonged BDL as a model for cirrhosis in the rat. They subjected rats to double ligation of the common bile duct,
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with section between the two ligatures. Animals were studied after five to fifty two days of BDL. After five days of BDL, there was bile duct proliferation, mild edema and an acute inflammatory reaction in portal areas. Fibrosis was slight and limited to the regions of new bile duct formation. A light infiltrate of neutrophil leucocytes around the bile ducts was observed. Hepatocytes contained numerous mitotic figures and necrosis was also observed. BDL for ten days or more was accompanied by fibrosis and bile duct proliferation in portal areas. Bile infarcts were occasionally noted, mainly in portal areas. Biliary obstruction for fifteen days induced the production of fibrous septa bridged portal areas extended into the lobules. After twenty eight days of bile flow obstruction, most of the animals showed cirrhosis; concentric fibrosis was observed around the interlobular and septal bile ducts. All rats ligated for thirty or more days showed cirrhosis with ascites (Kountouras et al., 1984). As mentioned before, the most commonly used method to induce experimental cirrhosis is chronic administration with CCl4 which shares several characteristics with human alcoholic cirrhosis but also shows some important differences (P´erez-Tamayo, 1983). In addition, the individual response of animals is variable, mortality is high and the method takes a relatively long time, 8–12 weeks, to produce cirrhosis (Muriel et al., 2005). In addition, nowadays CCl4 intoxication is extremely rare in man. In the model of prolonged bile duct obstruction, the morphological changes are comparable to those in human biliary cirrhosis. In addition, in the BDL model no bioactivation of any external toxin is required as it occurs in the CCl4 model. This obstructive model seems to have some advantages over the CCl4 and may prove to be a useful tool for studying human cirrhosis (Fern´andez-Mart´ınez et al., 2006) and to prove possible antifibrotic drugs (Fern´andez-Mart´ınez et al., 2001). However, both models possess advantages and disadvantages, for example, administration of CCl4 is easier to perform than to carry out a surgery. Reversion studies are easier in the CCl4 model because it is easier to stop giving the noxious stimuli. Morphologically, the CCl4 model resembles alcoholic cirrhosis, while the BDL model is similar to biliary cirrhosis. In CCl4 cirrhosis, oxidant stress in the main mechanism of liver damage (Comporti, 1985; Muriel, 1997), while in the BDL model oxidant stress occurs as a consequence and not as a cause of hepatic injury (Muriel and Su´arez, 1994; Muriel, 1997; Bar´on and Muriel, 1999). Therefore, both models are useful tools to study cirrhosis, for example, if an antifibrotic drug functions in both models it has more possibilities to work in man. 6.5.3
Estrogen-Induced Cholestasis
Estrogens are known to cause intrahepatic cholestasis during pregnancy in susceptible women, who are using oral contraceptives or who are on postmenopausal hormone replacement therapy (Schreiber and Simon, 1983). Intrahepatic cholestasis experimentally produced by intoxication with 17α-ethynylestradiol (EE) in rodents is a widely used model to investigate the pathophysiological mechanisms of intrahepatic cholestasis. Treatment with EE diminishes the ATP-dependent taurocholate transport in the canalicular membrane of the hepatocyte that is thought to be due to impaired expression of the canalicular bile salt export pump (BSEP) (Lee et al., 2000). In addition, EE treatment decreases uptake of bile acids in the sinusoidal membrane by ‘down-regulating’ the expression of the Na+ /taurocholate cotransporting polypeptide (NTCP) (Simon et al., 1996). The mentioned
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evidence indicates that EE produce cholestasis by decreasing the efflux and influx of bile acids in hepatic cells, leading to a decreased bile flow. The homeostasis of bile acids is regulated by several nuclear receptors, including farnesoid X receptor (FXR), pregnane X receptor (PXR), constitutive/androstane receptor (CAR) and estrogen receptor α (ERα) (Francis et al., 2003; Guo et al., 2003; Yamamoto et al., 2006). There is evidence that bile acids decrease bile acid biosynthesis by ‘downregulating’ transcription of the rate-limiting CYP7A1 through the FXR-SHP (short heterodimer partner)-LRHI (liver receptor homolog 1) cascade (Sinal et al., 2000). Responsible for bile acid uptake into the hepatocytes, NTCP is repressed by FXR activation (Sinal et al., 2000). At the same, time FXR ‘up-regulates’ the expression of BSEP that increases bile acid efflux from the liver to the bile (Sinal et al., 2000). Thus, FXR regulates transport of bile acids and avoids accumulation in hepatocytes. Recently, Yamamoto et al. (2006) provided direct evidence that ERα can become a risk factor for the development of hepatotoxicity following estrogen exposures. The administration of EE to rats decreases bile flow and produces impairment of transport mechanisms in canalicular and basolateral hepatocyte membranes (Rodr´ıguez-Garay, 2003). After EE administration, the excretion of bile salts, bilirubin, phospholipids, cholesterol, HCO3 and sulfobromophtalein is reduced (Bosard et al., 1993). EE treatment to rats results in a decreased sinusoidal membrane surface density (Hornstein et al., 1992). This was associated with impairment of sinusoidal transport molecules involved in the uptake of cholephilic substances (Alvaro et al., 1997). The EE effect has been attributed to the endogenous estrogen metabolite estradiol-17β-Dglucuronide (Meyers et al., 1980). This metabolite reduces bile flow and bile acid secretion in rats in a dose-dependent and reversible manner. Both EE and 17β-d-glucuronide increase tight juntional permeability in rat liver, allowing paracellular regurgitation of bile constituents into the blood (Kan et al., 1989). In conclusion, animal models of estrogen-induced cholestasis are necessary for the progressive understanding of EE acquired human cholestasis, as well as for the development of effective therapeutic interventions.
6.6
Ischemia/Reperfusion Liver Injury
Several excellent reviews have appeared recently dealing with this problem (Glantzounis et al., 2005; Pascher and Klupp, 2005; Casillas-Ram´ırez et al., 2006; Boros and Bromberg, 2006; Jaeschke, 2005b; Nieuwenhuijs et al., 2006; Spiegel and Bahde, 2006). Therefore, the reader is referred to those references and only a summary is presented here. Hepatic ischemia/reperfusion (I/R) damage is well known as an important cause of morbidity and mortality in liver resections and transplantations where anoxic or ischemic hepatic damage occurs. In addition, it also occurs as a consequence of systemic hypoxia or with conditions that produce reduction in the blood supply to the liver that results in low perfusion with the resulting hypoxia. This frequently occurs after systemic hypoxia or with conditions that cause low blood flow to the liver. Models of liver I/R damage represent an experimental simplification of the clinical situation. The choice of an appropriate model will depend on the clinical question to be
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answered. Detailed descriptions of experimental models can be obtained in a recent review (Spiegel and Bahde, 2006). Probably one of the best models is the in vivo global liver ischemia and porto-caval decompression. Ex vivo models are mainly suitable for toxicity evaluation and for answering special questions.
6.7
Hepatic Amebiasis
Amebiasis may have been first recognized as a deadly disease by Hippocrates (460 to 377 BC), who described a patient with fever and dysentery. A review of E. histolytica and its laboratory diagnosis appeared recently (Tanyuksel and Petri, 2003). Amebiasis, associated with high morbidity and mortality, continues to be a major public health problem throughout the world. Poverty, ignorance, overcrowding, poor sanitation and malnutrition favor transmission and increases ‘disease burden’. 6.7.1
Models of Experimental Liver Amebiasis
An excellent recent review on this topic has recently been published (Tsutsumi and Shibayama, 2006); therefore, only a summary of the most important features are presented herein. Reinertson and Thompson (1951) were the first to report an animal model for amebiasis – the hamster. They injected trophozoites of E. histolytica either through the portal vein or directly into the hepatic lobe that produced large amebic hepatic abscesses. Tsutsumi et al. (1984) suggested the role of host inflammatory cells in the process of liver injury after inoculation with E. histolytica trophozoites in hamsters. The results were confirmed with electron microscopic (Tsutsumi and Mart´ınez-Palomo, 1988) and immunocytochemical (Ventura-Ju´arez et al., 2002) studies. These studies indicated that liver abscesses are not produced by direct lysis of the trophozoites but by lysosomal enzymes released from disintegrating inflammatory cells that accumulate around the amebas and are killed by this parasite. Hepatic amebiasis can also be reproduced in gerbils, although trophozoites are less virulent in gerbils than in hamsters. Thus, progression of hepatic injury and death of the infected animals takes a longer time, therefore, allowing sequential studies of the hepatic lesions (Shibayama-Salas et al., 1997). In the case of mice in liver amebiasis, the introduction of genetically modified animals, like the severe combined immunodeficient (SCID) mice, has generated important information. When SCID mice were injected intrahepatically with E. histolytica trophozoites, liver abscesses were developed in all animals; however, only one of seven control animals developed an abscess (Cieslak et al., 1992). This study suggests that SCID mice constitute a powerful model for studying the protective immunity in invasive amebiasis. E. histolytica infects only humans and non-human primates; thus, reproducible animal models of amebiasis are limited. Nevertheless, information obtained from different models of hepatic amebiasis has permitted the better understanding of the mechanisms that take place in the damaged hepatic cell. Despite the fact that we still do not have an animal model that mimics the complete cycle of E. histolytica, as in the human disease, several investigations performed with normal or genetically or surgically altered laboratory animals have led to better understanding of the pathogenesis, immunology and pharmacology of the amebic lesion.
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Concanavalin A
Fulminant hepatic failure due to viral infection, intoxication or other causes is an important medical problem. Mechanisms that lead to apoptosis of hepatocytes frequently involved immune responses. Thus, the development of animal models that mimic the immunedependent parenchymal cell death are essential to understand the action mechanisms that conduce to hepatic failure and to develop pharmacological tools to prevent the human disease. Tiegs et al. (1992) developed a T cell-dependent experimental liver injury in mice induced by concanavalin A (Con A). Con A administration to mice produced dose-dependent apoptosis of hepatocyte (Gantner et al., 1995). Con A has high affinity toward the hepatic sinus producing activation of T cells (Gantner et al., 1995). A subset of T lymphocytes, the CD4-positive cells, is activated since anti-CD4 antibodies block activation, and consequently apoptosis of hepatocytes occurs (Tiegs et al., 1992). Polymorphonuclear is also recruited in the hepatic parenchyma probably by CD4-positive cells (Mizuhara et al., 1994). Con A-induced hepatitis in mice is characterized by increased serum levels of transaminases and infiltration, at the same time, of Kupffer cells, T cells and eosinophils into the liver (Jaruga et al., 2003). Natural killer T (NKT) hepatic cells play important roles in the Con A-induced hepatic damage by releasing several cytokines, including interferon gamma (IFN-γ ), tumor necrosis factor alpha (TNF-α), interleukin 4 (IL-4) and IL-5 (Takeda et al., 2000). Several other cytokines are involved in Con A-induced hepatic damage; some of them, like IL-6 (Mizuhara et al., 1994), IL-10 (Louis et al., 1997), IL-11 (Bozza et al., 1999), IL-22 (Radaeva et al., 2004) and IL-15 (Li et al., 2006), depict beneficial properties, whereas IL-12 (Nicoletti et al., 2000) and IL-18 (Faggioni et al., 2000) aggravate Con A-induced hepatitis. Therefore, hepatitis induced by Con A in mice may serve as a good model of viral hepatitis in the human. Obviously, viruses are not involved in this model. In fact, there is no model of viral hepatitis in animals.
6.9
Alcoholic Liver Damage in Rats
The rat constitutes one the most-used animals in the laboratory and alcohol abuse constitutes one of the most common causes of human liver diseases. Therefore, it seems logical to assume that administration of alcohol to rats may be the best model of alcoholic liver damage. However, it is important to consider that there are several characteristics in the rat that make it difficult to establish such a model: (a) natural aversion to ethanol, (b) higher ethanol metabolic rate and (c) greater tolerance to ethanol (Tsukamoto et al., 1990). To overcome these differences a long-term gastric catheter was designed and implanted in male adult rats (Tsukamoto et al., 1984). Currently, a single gastric catheter with a large bore size is used for infusion of ethanol and a diet. This method has been demonstrated to be the first ‘rat-model’ that induced progressive alcoholic liver injury including fatty liver, liver necrosis, inflammation and fibrosis (Tsukamoto et al., 1986; French et al., 1986). The exact control of ethanol and nutrient intake leads to a high reproducibility and low variability. Unfortunately, this model does not produce cirrhosis. However, the model allows investigations for the pathogenetic mechanisms of alcoholic liver necrosis
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and fibrosis. These mechanisms include the importance of sustained blood alcohol level in alcoholic liver disease (ALD), the role of the amount and the type of dietary fat in ALD, association of hypoxia with necrosis involvement of hepatic stellate and Kupffer cells with alcoholic fibrogenesis, as reviewed by Tsukamoto et al. (1990). More recently, the NK-κB factor (inhibited with allopurinol) was discovered to participate in ALD in this model (Kono et al., 2000). The natural aversion to ethanol, higher ethanol metabolic rate and greater tolerance to ethanol of rodents has led scientists to choose other models of liver damage where a catheter is not needed and cirrhosis is easily produced, as in CCl4 or BDL. However, this model has the advantage that the causative agent is very commonly used in humans.
6.10
Concluding Remarks
Liver diseases constitute a major medical problem of worldwide proportions (Williams, 2006). In Africa and Asia, their main causes are viral and parasitical infections. In many countries, alcohol abuse is the major cause of liver diseases. The vulnerability of the liver to chemical injury is as much a function of its anatomical proximity to the blood supply and digestive tract as to its ability to biotransform and concentrate xenobiotics. Xenobiotics in the blood pass through the portal vein and hepatic artery, and then drain through the central and the hepatic veins into the vena cava. Liver diseases are among the first ten causes of death in the world. In sharp contrast, effective liver drugs are very limited. With the exceptions of antiviral drugs effective in the treatment of viral hepatitis, there are very few drugs, if any, able to protect or ‘cure’ the damaged liver in a clinical situation. In this context, fundamental liver pathophysiology and pharmacology is needed; therefore, animal models of hepatic injury are the best tools to investigate liver disease processes and to test drugs with the ability to interfere with the hepatic injury process. Research with this and other models has provided a lot of knowledge in basic pathology which is leading to the discovery of new therapeutical approaches against liver damage and the recent advances promises prompt new discoveries.
Acknowledgements The author expresses his gratitude to Dr Mario G. Moreno for carefully reviewing the manuscript.
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Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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7 Application of Short- and Long-Term Hepatocyte Cultures to Predict Toxicities Gregor Tuschl, Jens Hrach, Philip G. Hewitt and Stefan O. Mueller
7.1
Introduction
Hepatotoxicity leads to a broad variety of liver pathophysiologies, including, for example, steatosis (fatty liver), cholestasis (obstruction of bile secretion), fibrosis (increased production and deposition of extracellular matrix components), hepatitis (inflammation), necrosis (cell death) or the formation of liver tumours. These pathological findings may arise from diseases affecting the liver, but also from xenobiotics, alcohol abuse or undesired drug– drug interactions. Hepatotoxicity is a major issue in pharmaceutical drug development [1] and drug-induced liver injury (DILI) is the major reason for attrition in clinical studies, as well as post-marketing [2]. In addition, 30 to 50 % of acute liver failures and 15 % of liver transplantations are related to chemical-induced hepatotoxicity [3–6]. Unfavourable hepatic reactions are often idiosyncratic, occurring on a background of transient liver injuries in less than 1 in 10 000 patients [7]. This may add to the fact that certain human hepatotoxicities have not been predictable from standard in vitro cytotoxicity assay results [8] or regulatory animal studies [9, 10]. This holds true despite the fact that there is reasonable understanding of the general pathophysiological mechanisms of most drug-induced hepatoxicities [11–14]: inhibition of mitochondrial function, disruption of intracellular calcium homeostasis, activation of apoptosis, oxidative stress, inhibition of specific enzymes or transporters and formation of reactive metabolites that cause direct toxicity or immunogenic response, potentially leading to idiosyncratic effects. Although there are ways to analyse
Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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most of these parameters, prediction of human hepatotoxicity is poor due to strong interspecies variations in data obtained from in vivo studies and the lack of true physiological conditions, especially the complexity of a whole organism, in in vitro experiments. Along with hypersensitivity and skin reactions, hepatotoxicity in humans has the poorest correlation with regulatory animal toxicity tests [9, 10]. On the other hand, conventional in vitro cytotoxicity assays also have low predictive value for the detection of human hepatotoxicity. However, it is worth mentioning that if these assays identified a compound as a liver-toxicant, there is more than 80 % specific correlation to the corresponding findings in humans [8]. The poor sensitivity of standard cytotoxicity assays is due to several reasons [8]. They usually measure lethal events in late stages of toxicity, but toxicity may not be lethal per se. In vitro cytotoxicity may take several days to appear [8, 15–18], making repeated administration of the investigated drug necessary. This, of course, raises the need for long-term in vitro models that facilitate extended exposure times. In addition, standard tests in general investigate one parameter of cell damage, whereas hepatotoxicity has many different mechanisms and is considered a multi-step process. In order to improve sensitivity, it will be necessary to analyse several morphological, biochemical and functional endpoints in parallel. Finally, tests should be performed not only with high concentrations, causing acute toxicity, but also with in vivo pharmacological concentrations. To address the limitations of many in vitro applications, we will present here long-term hepatocyte cultures as a model for the investigation of chronic and low-dose effects and also high-content screening (HCS), as a powerful tool which can be used to study several parameters in parallel at the single cell level. We will give an overview of the main in vitro test systems used for the assessment of hepatotoxicity and describe examples of the analysis of morphological, functional and gene expression endpoints in hepatocyte cultures.
7.2
In vitro Liver Models
The drug development process comprises a variety of steps to assess whether a test compound has adequate bioreactivity, appropriate physicochemical properties, metabolic stability, safety and efficacy in humans. While in vivo models are used to investigate hepatic drug effects in the context of toxicokinetics and systemic influences, cell culture models provide test systems for the investigation of specific mechanisms in a precisely controlled environment [19]. Hepatotoxicity occurs by the direct or indirect interaction of the toxicants with basic hepatocyte constituents such as proteins, lipids, RNA or DNA which can be analysed at the molecular, cellular or organ level. In vivo studies, limited by animal welfare/ethical concerns and difficulties in distinguishing between primary and secondary toxic effects, can be supplemented and partly replaced by in vitro models to accomplish a more detailed view of the mechanism of toxicity. There are many in vivo and in vitro test systems currently in use to predict hepatotoxicity in humans. However, their application is limited due to the frequent idiosyncratic nature of liver toxicity and the inherent differences between the metabolic activity in human and non-human species. The simplicity of some in vitro systems provides the ability to specifically manipulate and analyse a small number of parameters. The most commonly used test systems of the past few decades include, for example, the isolated perfused liver, liver slices, primary hepatocytes in suspension or culture, cell lines, transgenic cells and sub-cellular fractions like S9-mix, microsomes,
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supersomes or cytosol (Table 7.1). The reduction of the complexity of the system and the increase in throughput offers the ability to study specific parameters more closely but creates inherent constraints for each model (Table 7.1). This limits their widespread use and acceptance by the regulatory authorities as an alternative for in vivo screening [20], although studies have shown that in vitro cytotoxicity data can be used to identify appropriate doses for in vivo studies [25]. One major obstacle of some in vitro models is the limitation in metabolic activation of xenobiotics, mainly due to the downregulation of cytochrome P450 (CYP) enzymes over time [26, 27]. This is especially important since Phase I and Phase II metabolic conversion of chemicals has a great influence on their toxicity and can lead to both detoxification and toxification of xenobiotics [28–30]. To overcome these problems, new and innovative strategies are being developed in order to find reliable markers that are involved not only in early toxic responses but also in chronic toxicities, both occurring at sub-lethal doses of a test compound. Furthermore, there is a strong need for a robust longterm in vitro screening system that allows the characterization of drug/chemical-induced toxicities and helps to reduce the use of animals in toxicity testing. 7.2.1
Isolated Perfused Liver
Ideally, an in vitro test system should adequately represent the in vivo situation of drug metabolism and biotransformation in the liver, but there is still a long way to go to accomplish this goal. Most liver specific features are preserved in whole isolated and perfused livers, first developed in 1972 by Gordon and colleagues [31]. Especially, the threedimensional architecture of the liver, with cell–cell, cell–matrix interactions and functional bile canaliculi are maintained, leading to in vivo-like metabolism. Additionally, all liver cell types are present and the communication between them can play an important role in mediating toxicity. Perfused livers from different kinds of laboratory animals have been used for studies of hepatotoxicity, especially for compounds with the potential to affect bile flow. Furthermore, whole isolated liver is the only in vitro model that allows the measurement of haematodynamics. Despite all of these advantages, the isolated perfused liver model is difficult to handle and retains its functional integrity for only a few hours. For that reason, it can only be used for toxicants that are expected to have toxic effects at very early ‘time-points’. Moreover, reproducibility is low, the use of animals is not significantly reduced and human organs are rarely available. 7.2.2
Liver Slices
First used in 1923 by Otto Warburg [32] and improved in the following decades [33– 35], precision-cut liver slices are another in vitro model, with partly conserved liver cytoarchitecture, cell–cell, cell–matrix contacts and the presence of different cell types, which are frequently used in in vitro toxicology [36]. The preparation of slices from different parts of the liver facilitates lobe- and zone-specific analysis of metabolism and toxicity. In addition, since many slices can be prepared from the same human or animal donor liver, reproducibility and throughput can be increased significantly. Another major advantage is the possibility to conduct histopathologic examinations, as well as biochemical and molecular-biological studies from the same tissue. Although the thickness of liver slices is 200–250 μm, resembling 10–20 cell layers, the adequate supply of nutrients and oxygen from the incubation medium is only maintained for the outer cell layers. Therefore, liver
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Table 7.1 Overview of common liver in vitro systemsa Model
Advantages
Disadvantages
Isolated perfused liver
– functions closest to in vivo – in vivo-like expression of drug metabolising enzymes and transporters – three dimensional cytoarchitecture – functional bile canaliculi – lobular structure preserved – collection of bile possible – short-term kinetic studies – in vivo cytoarchitecture preserved – reasonably high throughput – functional drug metabolizing enzymes, transporters and bile canaliculi – zone-specific metabolism and toxicity may be studied – lobular structure preserved, selective effects detectable – human tissue slices more easily available than whole organs – long-term use possible – re-establishment of 3D cytoarchitecture – continuous perfusion with medium – hepatocyte specific gene expression closer to in vivo than in hepatocyte cultures
– not a high throughput system – hepatic function preserved for only a short time (2–3 h) – complicated to use – study of human liver difficult/impossible – best suited for livers of small animals – no significant reduction in the number of animals used – hepatic function preserved for no more than 6 to 48 h – bile cannot be collected and analysed – necrotic cells/scar tissue at edges of the slice – presence of necrotic cells might affect active transport of drugs through the outer cells
Liver tissue slices
3D-Bioreactors (bioartificial liver systems)
spheroids
– re-establishment of 3D cytoarchitecture – presence of non-parenchymal cells on outer layer and extra-cellular matrix throughout the spheroids
Primary hepatocyte cultures
– reasonably high throughput – differentiated function maintained in many short-term and long-term cultures – viability and differentiation preserved for 2 weeks, depending on culture conditions – potential for use of long-term cultures in chronic toxicity, drug metabolism and drug–drug interaction studies
very low throughput
– necrotic, hypoxic cells in centre of spheroids – accumulation of bile in centre of spheroids possible – not usable for long-term investigations (disaggregation and dedifferentiation) – culture may need special supplements in media – survival, differentiation status and function depends on culture conditions – no single system has yet been able to preserve all the different liver specific functions in vitro – difficult to regain cells for FACS analysis
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Overview of common liver in vitro systemsa (Continued)
Model
Advantages
Disadvantages
– relatively easy to use – analysis of human samples possible – functional drug metabolizing enzymes, transporters and bile canaliculi, depending on culture conditions – co-culture with other liver cells possible Hepatocytes in suspension
Liver cell lines
S9-mix Microsomes
Supersomes/ baculosomes
– reasonably high throughput – most drug metabolizing enzymes well-preserved at in vivo levels – easy to use – zone-specific metabolism and toxicity may be studied depending on the method of isolation – cryopreservation possible – analysis of human samples possible – unlimited availability – some liver-specific functions have been shown to be maintained – easy to use – reasonably high throughput – contains microsomal and cytosolic fractions – Phase I and Phase II activities – high throughput system – maintain expression of Phase I enzymes – can be recovered from frozen tissue – production of metabolites for structural analysis possible – use for drug inhibition, covalent binding and clearance studies – available from several species (including human) – high throughput – one or more human enzymes (CYPs, UGTs) can be specifically expressed
– lack of cell polarity limits use for drug transporter studies – lack of functional bile canaliculi – short-term viability (2–4 h) – lack of cell–cell and cell–matrix contacts – variations in samples from different human donors
– lacks in vivo phenotype – only a small set of hepatic functions expressed, at levels different from liver – genotypic instability – cofactors required for activity – lower enzyme activity compared to microsomes or cytosol – lacks Phase II and other cytosolic enzymes – short-term studies – cofactors required for activity – inadequate representation of the diversity of hepatic functions
– cofactors required for activity – UGT-reaction partly impaired
(Continued )
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Table 7.1 Overview of common liver in vitro systemsa (Continued) Model
Advantages
Disadvantages
Mitochondria
– high throughput – analysis of the effect of drugs on respiration, ATP synthesis and fatty acid oxidation
– only very short-term studies
Cytosol
– soluble Phase II enzymes (GST, ST, NAT) can be studied separately, depending on added cofactors – high throughput – one or more human enzymes can be specifically expressed – unlimited cell number
– cofactors required for activity – no CYPs, UGTs
Expression systems (cDNA clones in transient and stable expression systems) The virtual hepatocyte
– mathematical modelling of cellular events – prediction of unknown interactions may be possible
– studies may lack in vivo relevance – no physiologic levels of expressed enzymes – only single (some) enzymes can be analysed – limited computational power – still in experimental stage
a
Adapted from Brandon et al. (2003) [20], Guillouzo (1998) [21], Groneberg et al. (2002) [22], Farkas and Tannenbaum (2005) [23] and Sivaraman et al. (2005) [24].
slices are only useful for short-term toxicology studies due to the limited viability and rapid decline of liver-specific functions. The metabolic activity of tissue slices can be preserved for 1–2 days in culture [37, 38]. Another weakness is the inability to measure bile flow and other functional parameters, such as the portal flow. 7.2.3
Cultures of Isolated Liver Cells
Isolated hepatocytes or whole liver cell suspensions are used as starting material for a variety of in vitro models of different complexity and throughput. These include suspension cultures, adherent cultures on coated surfaces or embedded in extracellular matrix, cocultures with other cell types and ‘3D-Bioreactor’ setups. Fresh liver cells can be obtained relatively easy by different perfusion techniques [39, 40]. All of these procedures involve perfusion of the liver with Ca2+ -free buffers combined with enzymes/proteases, which disintegrate the extra cellular matrix, leading to the separation of the cells from each other. The isolation of liver cells is routinely performed for many species used in toxicity testing, but also with organs from human donors [41, 42]. Partial liver resections and nontransplantable whole livers are the main source of human tissue [43]. Hepatocytes, which make up the largest liver cell population, are highly polarized and differentiated cells, but these features are lost during liver perfusion. Therefore, the most challenging task is to re-establish cell polarity, cell–cell and cell–matrix interactions and to maintain the differentiated state as long as possible in culture [44, 45]. The quality of cultures not only depends on the conditions applied after seeding, but also on cell preparation
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and handling before culturing. Variations in the quality of hepatocyte preparations are particularly distinctive for human cells [43, 46]. For example, the type and concentration of the enzyme used for perfusion, along with its specific protease activity, affect yield and viability of the cell suspension obtained. Low protease activity may result in incomplete separation of cells, resulting in residual tissue ‘clumps’. High protease activity will decrease viability and increase the quantity of dead cells and cell debris in the culture but also lead to a higher number of cells undergoing necrosis during the first 24 h of culture. Moreover, the cell suspension, containing all parenchymal and nonparenchymal liver cells, should be protected from light and stored on ice for as short a time as possible before seeding. The desired cell populations (hepatocytes, endothelial, Kupffer or stellate cells) can be separated by subsequent washing steps and gradient centrifugation [47, 48]. A combined digitonincollagenase perfusion allows the isolation and separation of periportal and perivenous hepatocytes from a single liver [49]. Cryopreservation of hepatocytes offers the opportunity to store the cells for months or even years, making it possible to repeat experiments with cells from the same donor and hence to increase reproducibility [50]. However, cell viability, attachment, liver-specific metabolism and protein neogenesis can be reduced or impaired after thawing [51]. There is no standard protocol for the different culture techniques and each laboratory uses its own standards. The main factors influencing the quality of the culture include media composition, extra cellular matrix environment, cell density and the properties of other cells in co-cultures. There is partial agreement on some of the parameters and we will describe the effects of several different culture methods on hepatocyte characteristics later in this chapter. 7.2.4
Hepatocyte Suspension Cultures
Hepatocyte preparations usually contain about 40–70 × 106 cells/g liver, with 85–95 % viability and less than 5 % contaminating nonparenchymal cells after purification. In suspension, the survival of cells is short and strongly depends on the medium composition, but is normally not longer than a few hours [52]. Although the system is fairly high throughput, easy to use and preserves most of the metabolising enzymes at in vivo levels for a short time, it is only useable for acute toxicology or metabolism studies. By capturing the cells into beads of alginate, the survival-time can be prolonged to 24 h [53]. However, the lack of functional bile canaliculi, cell polarity, cell–cell and cell–matrix contacts limits the use of alginate-embedded cells for drug transporter studies. 7.2.5
Short- and Long-Term Hepatocyte Cultures
Survival time in culture can be increased if hepatocytes are cultured on adhesive surfaces, for example, tissue culture vessels coated with extracellular matrix components. The most commonly used models are the monolayer culture, where hepatocytes are usually attached r and the sandwich culture system, where cells to dried films of collagen I or Matrigel are embedded between two layers of gelled extracellular matrix proteins. Intracellular signalling is closely connected to the interaction between extracellular matrix, cell-adhesion molecules and the cytoskeleton and therefore has major impact on gene expression and the metabolic capacity of cells [54, 55]. Therefore, the application of extracellular matrix to hepatocyte cultures is one step towards the creation of a more in vivo-like environment.
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Unfortunately, hepatocytes still lose many of their specific functions during isolation [56, 57] or during the first few days if maintained in monolayer culture [58]. Today, cultures of primary rat and human hepatocytes are used in a variety of pharmacological and toxicological experiments. For example, the evaluation of hepatic drug uptake and metabolism, drug–drug interactions and hepatotoxicity. The development of long-term primary hepatocyte cultures would also be a step towards the study of chronic effects in vitro. Culturing hepatocytes in a sandwich configuration between two layers of gelled extracelr lular matrix proteins (e.g. collagen I or Matrigel ), has prolonged the time of cultures r displaying hepatocyte-specific functions dramatically [59–62]. Matrigel is a laminin-rich preparation from the Engelbreth–Holm–Swarm mouse sarcoma [63]. The composition of r Matrigel , with extracellular matrix components and growth factors is variable [64], while collagen I, mostly prepared from rat tail tendons, provides more standardised conditions. Another factor with great influence on morphological development and cell survival of hepatocytes in culture is basal medium formulation and the addition/omission of serum, specified hormone mixtures or other supplements [65–67]. Among the most frequently used basal media, Dulbecco’s modified Eagle medium (DMEM), modified Chee’s medium (MCM) and Williams’ medium E (WME), the DMEM/F12 mix seems most appropriate to maintain liver-specific function and rebuild bile canaliculi in cell culture [68]. Another report identified WME and DM-160 medium to be the most suitable with regard to cell attachment, survival and albumin secretion among ten media tested under serum-free conditions [69]. Once the cells are in culture, the addition of the glucocorticoid dexamethasone (DEX) at nanomolar concentrations, is essential for the long-term preservation of hepatocytespecific functions, polygonal hepatocyte morphology, the structural integrity of cytoplasmic membranes and bile canaliculi-like structures [70, 71]. If DEX is added at micromolar concentrations or cultures are supplemented with Rifampicine, Phenobarbital (PB) or 3methylcholantrene, CYP expression is induced and by this means the metabolic activity can be increased [72]. Apart from maintaining CYP expression, PB is also able to suppress hepatocyte growth [73]. Insulin enhances the attachment of hepatocytes and alters their morphology [74]. Selenium, a structural component of the enzyme glutathione peroxidase, which plays an essential role in the neutralization of metabolically generated peroxides, has also been shown to be beneficial when added to the medium [75]. The addition of mitogenic compounds, for example, epidermal growth factor, leads to hepatocyte proliferation, allowing longer culturing, but on the other hand leads to the ‘down-regulation’ of metabolic enzymes and thereby evokes dedifferentiation. Since primary cell cultures are inherently non-sterile, antibiotics have to be added to the culture. Therefore, it has to be taken into account that the antimycotic agent Fungizone (Amphotericin) stresses the cells to a considerable extent and should not be added routinely. Apart from extracellular matrix application and media formulation, cell density also influences the morphology and function of hepatocytes in culture [55]. The formation of bile canaliculi-like structures declines along with the expression and function of MRP2 and MDR1a/b as cell density is reduced [76]. If seeded at very high densities (almost 100 % confluency) cells do not tend to spread out that much in monolayer cultures and display hepatocyte-like morphology for 1–2 additional days, but then they also start to detach from the surface. It has to be taken into account that the same cell density (cells/cm2 ) will result in unequal confluency of cultures seeded on either collagen film or gel, due to the fact
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Figure 7.1 Effect of extracellular matrix environment and media formulation on morphological development and structural integrity in primary rat hepatocyte cultures. Hepatocytes were isolated from male Wistar rats using a modification of the two-step perfusion method described by Seglen [39]. Cells were cultured for the indicated times on collagen monolayer or in a collagen gel sandwich with serum-free or serum-containing medium; media were changed daily. Arrows indicate bile canaliculi-like structures. The white scale bar in the bottom right of each image corresponds to 200 μm
that hepatocytes in a three-dimensional extracellular matrix environment require less area. Since polar differentiation of hepatocytes in culture is only visible in cell aggregates, cell density should always be close to confluency (≥ 90 %). Generally, after being in culture, cells should not be removed from the incubator longer than necessary and not be submitted to direct sunlight. Figure 7.1 gives an overview of the morphological changes occurring over time under different cell culture conditions. Since it is well known that serum enhances the surface attachment ability of hepatocytes [77], cells are generally seeded in medium containing fetal calf serum, regardless of the subsequent culture conditions.
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A morphological distinction between hepatocytes seeded on collagen film (monolayer) or collagen gel (sandwich) is already visible after 4 h. Monolayer-cultured cells adopt their polygonal shape and establish extensive cell-to-cell contacts; whereas in sandwich culture this takes markedly longer, being still mostly spherical and singular after 4 h, probably resulting from the cells’ immersion in the collagen gel (Figure 7.1). In general, monolayercultured cells appear more flattened than sandwich-cultured cells, because of the lack of a three-dimensional extracellular matrix environment. All cells display clear cytoplasm and well-delineated plasma membranes. After overnight incubation, sandwich-cultured hepatocytes also form aggregates and the typical cuboidal cell shape is seen (Figure 7.1). In both culture systems, the formation of structures resembling bile canaliculi (light areas between cells, Figure 7.1, arrows) starts and increased during the following 24 h, but to a greater extent in the serum-free cultures (not shown). This is consistent with the findings of Terry and Gallin [78] who reported that serum is capable of inhibiting the re-establishment of bile canaliculi in hepatocyte cultures. After 72 h of culture on collagen film, there is a strong perturbation of cell morphology in both serum-free and serum-containing cultures (Figure 7.1). Monolayer-cultured hepatocytes spread out and formed fibroblast-like protrusions, the nuclei volume was increased and the cytoplasm appeared granulated. There are no longer well-delineated plasma membrane borders and bile canaliculi-like structures have disappeared almost entirely. The morphological stability of hepatocytes seeded on collagen film can be increased considerably if these cultures are overlaid with collagen gel. However, the time of overlay is crucial for the phenotypic outcome. Since hepatocyte monolayer cultures do not have a steady appearance over time, even in the first days of culture, they should be overlaid within the first 24 h. The best results are obtained if the monolayer/sandwich hybrid is prepared at the initial medium change 4 h after seeding. In this configuration, the morphology at the time of overlay can be preserved for more than 1 week. If the hepatotypic morphology is lost after several days on collagen film, it will not be restored after overlay – only the state at the time of overlay will be preserved (data not shown). By overlaying monolayer cultures of primary hepatocytes with collagen gel, it was possible to obtain cultures displaying some features of collagen sandwich cultures over extended periods of time (e.g. bile canaliculilike structures), although the overall appearance is still different and these hybrid cultures do not last as long as sandwich cultures. For cells cultured in a collagen sandwich, there is no spreading visible; polygonal cell shape and plasma membrane boundaries are still stable after 72 h of culture (Figure 7.1). However, for hepatocytes incubated in serum-containing medium, there is a noticeable deterioration in cytoplasmic integrity and the stability of canaliculi-like structures. In contrast, primary rat hepatocytes in a collagen sandwich and incubated with serum-free media show steady aggregates of polygonally shaped cells with mostly clear cytoplasm and a stable bile canaliculi-like network (Figure 7.1). This is in agreement with the previously reported beneficial effects of an extracellular matrix overlay on cell viability and the preservation of normal morphology [60, 79, 80]. The fact that collagen-sandwich cultures have successfully been used for metabolism and induction studies [81–83] indicates that the collagen overlay does not interfere with the interaction of the cells with the test compounds. In conclusion, only serum-free collagen sandwich cultures of hepatocytes show no signs of deterioration when incubated for more than one week, maintaining polygonal shape, clear cytoplasm and stable bile canaliculi-like structures. The fact that bile canaliculi-like structures are stable
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Figure 7.2 (a) Fibroblast growth in serum-supplemented primary rat hepatocyte monolayer cultures 10 days after seeding. (b) Co-culture of primary rat hepatocytes and Kupffer cells 2 days after seeding. Hepatocytes were isolated from male Wistar rats using a modification of the two-step perfusion method described by Seglen [39]. Kupffer cells were subsequently separated using a modified protocol originally described by Smedsrod and Pertoft [47]. Cells were cultured on collagen monolayer with serum-free medium. Arrows indicate Kupffer cells. The white scale bar in the bottom right of each image corresponds to 200 μm
over time, make these cultures especially valuable for transport studies [84, 85]. Furtherr do not more, hepatocytes seeded on gelled collagen and overlaid with a layer of Matrigel demonstrate superior survival in comparison to cells overlaid with another layer of gelled collagen, exemplified by their lack of stable bile canaliculi-like structures (unpublished data). Hence, using gelled collagen I as the extracellular matrix component in hepatocyte sandwich cultures seems most appropriate for long-term use. The morphological stability of DEX-supplemented serum-free collagen sandwich cultures of primary rat hepatocytes can endure for as long as 4 weeks. DEX can also improve and prolong the differentiation status of serum-containing cultures considerably. However, culturing hepatocytes with serum-supplemented medium may lead to fibroblast overgrowth after several days regardless of the fact whether the cells are cultured in monolayer or sandwich configurations (Figure 7.2(a)). 7.2.6
Co-Cultures
Hepatocytes compose about 60–70 % of the cells in the intact organ; however, liver toxicity may not always originate from these cells. Therefore, co-cultures of hepatocytes with other non-parenchymal liver cells, such as endothelial, Kupffer or stellate cells and also stable cell lines or fibroblasts can be applied to reflect a more physiological situation. Kupffer cells represent the second largest cell population of the liver and they are very important for cytokine signalling within the organ. The non-perenchymal Kupffer cells, which are resident macrophages, are located in the hepatic sinusoids, in between or on top
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of endothelial cells, but also make contact to the parenchymal hepatocytes through their cytoplasmic extensions [86]. Thus, Kupffer cells can act as biosensors affecting hepatocyte and ultimately the livers’ fate through cytokine signalling. For example, the excretion of TNFα or nitric oxide can lead to inflammatory reactions or apoptosis [87, 88]. This may be especially relevant for the development of biologically derived pharmaceuticals. In order to investigate the ‘crosstalk’ between Kupffer cells and hepatocytes and its influence on cytotoxicity in vitro, co-cultures of the two can be used (Figure 7.2(b)). Different studies have shown the improvement of some hepatocyte-specific functions in co-culture with other cell types [89, 90] but also when using non-parenchymal cell conditioned medium for hepatocyte cultures [91]. Co-cultured hepatocytes can survive for several weeks and liver-specific metabolism and expression of Phase I and Phase II enzymes are maintained at higher levels than in pure hepatocyte cultures [92, 93]. 7.2.7
Spheroid Cultures
If a crude liver cell suspension is seeded into cell culture vessels and surface attachment is prevented by continuous shaking, spherical multicellular aggregates (spheroids) will form. Cell-to-cell contacts are re-established, hepatocytes are located on the inside, nonparenchymal cells on the outside and the deposition of extracellular matrix is seen throughout the spheroids. Sometimes, alginate or other materials are added to make up the internal structure of the spheres. Several studies have showed the positive effect of this culture method on the expression of hepatotypic genes and the maintenance of metabolic capacity [94, 95]. However, the fusion of smaller spheres may lead to hypoxic and necrotic cells dying at the centre of larger spheroids (>100 μm). Another problem could arise from the accumulation of bile in the centre of spheroids. The maintenance of prolonged functional activity of spheroids has been related to the restoration and stability of cell polarity and close cell-to-cell contacts [96]. 7.2.8
3D Bioreactor Cultures
Another skilful attempt to mimic a liver-like environment in vitro are bioartificial liver systems (3D-bioreactors). Their major advantage is the re-establishment of the 3D liver cyto-architecture with cell–cell contacts and a three-dimensional extracellular matrix environment, combined with continuous medium perfusion, providing a constant supply of oxygen and nutrients. The first systems established were hollow fiber bioreactors [97]. Today, hepatocyte monolayer and collagen sandwich cultures are being used for bioreactor setups [98]. In addition, spheroids, for example, seeded into channels that are etched into a silicon scaffold, have been reported [99]. It has been shown that cell viability, metabolic activity, including CYP and Phase II reactions, albumin and urea secretion, as well as general hepatotypic gene expression can be sustained over extended culture duration in 3D bioreactors [24]. However, considerable challenges arise from the construction of the bioreactors. For example, composition of the basement structure, flow rate and liquid pressure has to be taken into account. In fact, the shearing power generated by the correctly adjusted flow rate of the medium can improve the metabolic capacity of the culture [100]. Although 3D bioreactors provide high-quality metabolic data and allow the quantitative analysis of pharmacokinetic parameters from key animal species and humans, their use as a tool for high throughput screening is extremely limited due to the large operational effort.
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Cell Lines
To overcome the problem of limited survival and availability of primary cultures, different cell lines are used in a variety of toxicological applications. Since most hepatic cell lines originate from tumours, they have lost the high degree of differentiation of a hepatocyte and their gene expression pattern is distinctively different from normal liver cells. In addition, many cell lines display genetic instability. For example, the frequently used human hepatoma cell line, HepG2, lacks expression of CYP2E1 along with other CYP isoforms and Phase II enzymes, making them insensitive to secondary toxic compounds [101]. To complicate matters, different sources of HepG2 cells can have very different enzyme profiles [102]. Several transfected variants of HepG2 have been constructed which express increased levels of drug-metabolizing enzymes, including CYP1A1, CYP1A2, CYP2E1 and glutathioneS-transferases [101]. These new clones might be particularly useful for the investigation of specific classes of genotoxic compounds and also for mechanistic studies. In vivo relevance may not always be given because expression of the cloned enzymes is not at physiological levels and only single enzyme functions can be analysed. Other cell lines, such as the immortalized but untransformed transgenic hepatocyte cell lines (e.g. MMH-GH [103]), have been used with promising results. Recently, the human hepatoma cell line HepaRG was described [104]. After application of a differentiation protocol, HepaRG cells display hepatocyte morphology and expression of drug-metabolizing enzymes at near in vivo levels [105, 106]. However, these novel cell lines have to be confirmed as a reasonable alternative cell-based assay for use in toxicological studies [107]. 7.2.10
Subcellular Fractions
The simplest liver in vitro models are sub-cellular fractions, such as, organ homogenates, microsomes, mitochondria or nuclei. Most sub-cellular fractions can be prepared and separated relatively easy by homogenization of the tissue and sequential centrifugation. They are available from a large number of species, including humans. Another major advantage is that they can be frozen until needed. Nevertheless, they are only suitable for short-term studies with specific questions such as drug inhibition, covalent binding or clearance studies. For example, liver supernatants (‘S9’) are used as an activation system for xenobiotics in in vitro genotoxicity assays (e.g. Ames-assay [108]). Mitochondria are added for the analysis of drug effects on respiration, ATP-synthesis and fatty acid oxidation. To acquire increased metabolic activity, CYP expression is often induced by treatment with Aroclor 1254 or a Phenobarbital/beta-naphthoflavone (BNF) mixture prior to preparation [109], leading to elevated and unphysiological expression. Apart from the limited viability over time, most systems have to be supplemented with cofactors to preserve enzymatic activity. Other disadvantages include the absence of complete enzyme systems, for example, the lack of Phase II enzymes in microsomes.
7.3
Endpoints for the Analysis and Characterization of Hepatocyte Cultures
The list of tests used by in vitro toxicologists to gain insight into the effect of a test substance on cells is extensive, some of which will be described in detail in this chapter. These range
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from standard tests, e.g. cell viability measurements, to hepatocyte specific activity tests, such as bile production, CYP activity or drug transport, to mention only a few. In addition, the analysis of single genes or proteins with established molecular methods like real-time PCR or Western-blotting is commonly applied. In recent years ‘-omics’ methods have been introduced to generate a more complete understanding of the complexity of toxic events and to integrate this knowledge into Systems Biology approaches. 7.3.1
Morphology – High-Content Imaging
As seen in Figure 7.1, hepatocytes cultured in serum-free collagen sandwich cultures stay morphologically unchanged for a few weeks, offering the ability to investigate alterations in cellular structures induced by chemical treatment with classic light microscopy but also with the use of high-content imaging (HCI), which can be integrated into high-content screening (HCS). HCI is the automated multiparametric analysis of spatial and temporal changes of sub-cellular structures at the single cell level with visible or fluorescent light microscopy techniques [110–113]. The parameters that can be measured include vacuolization, formation of lipid droplets, nuclear morphology, plasma membrane integrity (blebs), cell number (proliferation), size or shape (e.g. neurone outgrowth), but also the analysis of molecular targets like nuclear translocation of transcription factors. By using multi-channel fluorescence detection, alternatively labelled molecular targets can be followed at the same time [114, 115]. Data acquisition is automated and all parameters can be recorded in parallel. Cells can be cultured in several standard tissue cultures formats, e.g. 96- or 384-well plates and many cells are analysed in each well simultaneously. In contrast to standard cytotoxicity assays, where single parameters are averaged over different populations of cells, HCI can create a wealth of data for the detailed analysis of toxic events [113]. This technology is particularly important for the analysis of sub-lethal toxicities. HCS data can also be implemented into Systems Biology approaches and may benefit drug discovery [110, 111]. O’Brien and colleagues demonstrated the ability to detect human hepatotoxicity with high sensitivity and specificity with the use of HCS [114]. 7.3.2
Functional Endpoints
There are large numbers of assays and kits to assess functional and biochemical parameters of cultured cells. In vitro toxicity in hepatocyte cultures is either investigated by nonspecific cytotoxicity or by the measurement of compound-specific endpoints. Chemicalinduced changes in cellular functions may be irreversible, ultimately leading to cell death, where others may be transient. Irreversible endpoints include the induction of apoptosis, which can be investigated by the measurement of increased caspase activity or the loss of plasma membrane integrity. Plasma membrane damage can be analysed by the detection of cytoplasmic enzyme release (e.g. lactate dehydrogenase, LDH) or the uptake of specific dyes such as Neutral Red and Trypan Blue into the cytoplasm. In addition, alterations in general hepatocyte functions like albumin, urea or bile secretion provide information on the impairment of cellular processes. The energy status of the cell is often used to determine cytotoxicity. This is achieved by studying the ATP content of the cells or the mitochondrial or enzymatic capacity to reduce tetrazolium salts (XTT, MTT, WST) [116]. Several compounds may cause oxidative stress, leading to glutathione (GSH) depletion. The induction of general stress proteins can be measured with ELISA (enzyme-linked
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immunosorbent assay)-based assays. Another important hepatocyte specific feature is the uptake of xenobiotics in order to be metabolized and the export of metabolites. Drug transport can be studied by fluorescent dyes or analysis of bile acid transport with high performance liquid chromatography (HPLC) [84, 117]. 7.3.3
Gene Expression
Toxic events, leading to morphological and functional impairment, can also be investigated by analysing the underlying alterations that occur on the gene expression level in hepatocyte cultures. However, recent work has implied that the cell isolation procedure itself leads to gene-expression changes in hepatocytes, especially for specific CYPs [118]. This is caused by changes in the shape of isolated hepatocytes and the induction of ribonuclease activity during perfusion and the first few hours of culture [119]. In addition, one needs to take into account the gene expression changes that occur over time in untreated hepatocyte cultures. Figure 7.3 displays a visualization of the expression changes of 550 hepatotypic genes that occur over time in monolayer and sandwich cultures of primary rat hepatocytes. It is obvious that basal gene expression in liver tissue and freshly isolated hepatocytes is distinctly different from other cultures, regardless of the culture conditions (Figure 7.3 and
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Figure 7.3 Principal Components Analysis (PCA) of gene-expression alterations occurring over time in different primary rat hepatocyte cultures, as well as in intact liver (‘liver’) and fresh hepatocyte suspension (‘fresh’). Cells were cultured as monolayer and sandwich, with (+ serum) or without (− serum). Messenger RNA levels were determined with custom ‘Sentrix BeadChip Arrays’ from Illumina, containing 550 genes. Data from three biological replicates were analysed with the ‘Genedata Expressionist Analyst’ software. Culture samples were taken 2 h, 1 d, 2 d, 3 d, 5 d, 7 d and 9 d after the designated time t0 = 24 h after seeding. The PCA displays a non-supervised dimensionality reduction of the data to show the highest variance in the dataset. (a) Comparison of all four cell culture conditions or (b) only serum-free cultures with freshly isolated hepatocytes and intact liver. Darker shades of grey represent sandwich cultures or intact liver, while lighter shades of grey represent the respective monolayer cultures or freshly isolated hepatocytes
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Figure 7.4 Heat maps, displaying gene-expression alterations occurring over time in different primary rat hepatocyte cultures, as well as in intact liver and fresh hepatocyte suspension. Cells were cultured as monolayer and sandwich with (+ serum) or without (− serum). Messenger RNA levels were determined with custom ‘Sentrix BeadChip Arrays’ from Illumina, containing 550 genes. Data from three biological replicates were analysed with the ‘Genedata Expressionist Analyst’ software. Culture samples were taken 2 h, 1 d, 2 d, 3 d, 5 d, 7 d and 9 d after the designated time, t0 = 24 h after seeding. Each square in a column or line represents the correlation of a given sample relative to another. Darker shades of grey indicate very high correlation, whereas lighter squares mean low correlation. Overall, there is a high similarity between liver and fresh cells (upper left corner, columns left of the arrows). Columns to the right of the arrows display cell culture samples with increasing incubation time
Figure 7.4, arrows). This has also been previously reported for human hepatocytes [120]. In addition, serum-containing cultures are clearly separated from serum-free cultures in the principal components analysis shown in Figure 7.3. Strikingly, serum supplementation is a major modulator of hepatic gene expression. Another important observation is the fact that the strongest gene expression changes were observed during the first days of culture, independent of the culture conditions [120, 121]. However, after an initial adaptation period
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mRNA levels stabilized over time in serum-free cultures, but not in serum-containing cultures (Figure 7.4). These expression levels remained constant for a minimum of four weeks in sandwich cultures without serum. By comparing serum-free collagen sandwich and monolayer cultures ten days after seeding, gene expression in cells cultured between two layers of gelled collagen I appeared to be closer to the in vivo samples or freshly isolated hepatocytes (Figure 7.4). Thus, for stable long-term cultures of primary rat hepatocytes, the serum-free collagen sandwich culture seems most suitable. As well as the gene expression alterations occurring over time in the differentially cultured primary rat hepatocytes, there are differences in mRNA levels between the specific cell cultures at a given time-point [122]. Figure 7.5 shows the gene expression changes of 22 hepatotoxicity-related genes, occurring in dimethyl sulfoxide (DMSO)-treated primary rat hepatocyte monolayer or sandwich cultures, after 72 h in culture (relative to 24 h cultures). DMSO is a very commonly used solvent for test materials designated for the treatment of cells in culture. Su and Waxman [123] reported that the addition of 2 vol % DMSO did not affect CYP1A1 mRNA levels but had great influence on the expression patterns of other CYPs and several liver transcription factors. Furthermore, DMSO was capable of restoring normal CYP2B1, CYP3A1, CYP4A1, HNF4 and CEBPα levels. It also decreased initially elevated HNF6 expression and maintained constant levels of HNF3α, HNF3β and CEBPβ [123]. Moreover, DMSO is used for the hepatocyte differentiation of the hepatoma derived cell line HepaRG [104–106]. However, organic solvents impact the metabolic activity of hepatocytes and therefore the concentration used for dissolving test agents must be used with caution [124–126]. As seen in Figure 7.5, there is a distinct difference in the observed expression patterns for the four cell cultures tested. There is a marked ‘downregulation’ for most of the assayed genes in the serum-containing monolayer culture whereas in the three other cultures almost all genes reached higher abundance levels at 72 h. The observed rise of MDR1 expression over time in primary hepatocytes was also reported by Fardel et al. [127] and Chieli et al. [128]. The increase in MDR1 expression over time is less pronounced in the respective sandwich cultures, which is in agreement with Lee [129]. Cyclins D1 and G1 were elevated over time in the serum-containing sandwich culture (Figure 7.5) as well as in the respective monolayer relative to every other culture condition at 72 h (not shown). This indicates a disturbed cell cycle regulation in normally non-dividing hepatocytes, especially in the serum-containing cultures. Another noticeable feature is the increase in the heat shock factor HSP70 over time in the serum-containing cultures (Figure 7.5). When comparing the cultures at 24 h, there was an increase under serum-free conditions, which was still present at 72 h in the respective collagen monolayer culture but had disappeared in the collagen sandwich. GADD45α expression, although altered only weakly, displayed lower levels in the serum-free monolayer and both sandwich cultures compared to the serum-containing monolayer culture. Since GADD45α is associated with stress signalling, this indicates that the level of cellular stress was highest in the serum-containing monolayer culture, making this culture less appropriate for the detection of toxic effects. Since CYP enzymes are the major monoxygenases involved in liver Phase I metabolism, the expression of these molecules in hepatocyte cultures is especially important. Metabolic conversion of chemicals may lead to altered drug–drug interaction patterns and subsequently cause possible adverse effects. Therefore, the CYP inducing potential of a drug is of great interest. Cultures of primary rat hepatocytes show significant alterations in CYP expression over time (Figure 7.5) but also in relation to each other. In particular, the CYP2C
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Figure 7.5 Gene-expression analysis of 22 hepatotoxicity related genes in primary rat hepatocyte cultures. Cells were cultured as monolayer with (A) or without serum (B) and as collagen sandwich with (C) or without serum (D). Shown are values of fold regulation for the temporal gene expression changes occurring at 72 h compared to the values at 24 h (set as ‘1’) in the respective DMSO-treated cell culture. Messenger RNA levels were determined using quantitative real-time PCR with the ‘TaqMan Low Density Arrays’ (Applied Biosystems). Values were calculated by the efficiency-corrected comparative CT method with 18S rRNA serving as calibrator. Bars illustrate mean values from quadruplicate measurements with standard deviations. Elevated levels of gene expression are indicated by positive figures > + 1 and ‘down-regulation’ is indicated by negative figures < − 1. Please note that the positive, as well as the negative, y-axes are scaled logarithmically (data taken from Tuschl and Mueller (2006) [122]). MRP3, multidrug resistance protein 3; AFP, alpha-fetoprotein; AP, alkaline phosphatase, tissue-nonspecific; ApoE, apolipoprotein E; FABP2, fatty acid binding protein 1; FAS, fatty acid synthase; GADD45α, growth arrest and DNA-damage-inducible 45 alpha; GSTα1, glutathione S-transferase alpha 1; HNF4α, hepatocyte nuclear factor 4 alpha; HSP70, heat shock 70kD protein 1A; MDR1, multidrug resistance transporter 1; COX2, prostaglandin-endoperoxide synthase 2; ACOX, acyl-coenzyme A oxidase 1, palmitoyl; OAT1, organic anion transporter 1; SOD2, superoxide dismutase 2, mitochondrial; SulT1c2, sulfotransferase family, cytosolic, 1C, member 2; Txn2, thioredoxin 2
and CYP4A1 genes are considerably decreased in the serum-containing monolayer culture after 3 days (Figure 7.5). CYP1A1 levels are strongly increased in the serum-free monolayer cultures and to a lesser extent in the sandwich cultures. After 72 h in culture, CYP mRNAs are more abundant in the serum-free monolayer cultures and both sandwich cultures in comparison to the serum-containing monolayer. Indeed, there is evidence in the literature
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that the presence of DEX [130] and a three-dimensional extracellular matrix environment [85] helps to maintain constant levels of CYP expression. The low abundance of the examined CYP mRNAs at 72 h indicates a limited metabolic capacity in serum-containing collagen monolayer cultures, making only short-term applications acceptable [58]. Analysis of CYP expression in primary rat hepatocytes, cultured for 10 days under various conditions, shows distinct patterns for specific CYPs (Figure 7.6). After being relatively constant in the first days of culture, there was a strong reduction of CYP1A2 and CYP2E1 expression after 3 days in all cultures, although the reduction was lowest in the serum-free sandwich. The major Phase I enzymes in rat, CYP2C and CYP3A1, were strongly repressed in the serum-containing cultures over time, displaying stable values close to in vivo in the serum-free cultures until 10 days after seeding. In general, variations in gene expression over time were least pronounced in the serum-free collagen sandwich cultures. Therefore, this culture seems most adequate for the study of gene expression alterations triggered by drugs or chemicals. 7.3.4
The ‘-omics’ Challenge – Systems Biology
The detection of gene expression changes in differentially cultured cells or induced by chemical treatment is no longer limited to several genes at a time, since whole genome arrays became available. The sequencing of the entire chromosomal DNA sequences of human as well as many ‘laboratory’ animal species enabled the synthesis of whole genome arrays. This novel technique created a new field of research, called genomics, often termed transcriptomics, since the molecules that are detected are the transcribed mRNAs. To be able to analyse the whole cellular population of specific molecules, there is now also proteomics, lipidomics, metabolomics, etc. and the development of new methods in the ‘-omics’ fields is rapidly growing. The combination of different ‘-omics’ techniques, called ‘cytomics’, creates an enormous amount of data that could generate a more detailed and complete insight into cellular physiology [131]. However, one of the most challenging tasks of the new ‘-omics’ era techniques is data storage, handling and analysis. Large computational efforts and bioinformatics methods are applied to create algorithms in order to elucidate statistically and biologically significant results. Therefore, rather than looking at the single gene or protein level, profile signatures are determined, with the goal to identify, for example, specific gene patterns that are predictive for toxicity. There is a lot of activity to create databases with compound signatures in order to classify chemicals [132, 133]. This is important for the pharmaceutical industry, especially in the field of toxicology, for hazard identification and characterization of candidate molecules early in development. In addition, toxicogenomics data are valuable for the elucidation of the underlying mechanism of toxicity [134–136]. Many of these data are put together in the Comparative Toxicogenomics Database (CTB) to achieve a better understanding of the etiology of human diseases [137]. Apart from identifying different types of hepatotoxicitiy, there have been attempts to find biomarkers for the early identification of hepatocarcinogenesis with the use of toxicogenomics and proteomics methods [138–140]. There is hope that these new methods will make it possible to detect changes in the molecular pattern of the cell that are indicative of the pathological endpoint, at subtoxic doses, before the endpoint becomes histopathologically detectable [141]. This could facilitate a considerable reduction in the time needed to obtain results and the number of animals used in toxicity testing [142]. In addition, there are several suppliers of
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Figure 7.6 Long-term CYP expression in primary rat hepatocyte cultures. Cells were cultured as monolayers with (M+) or without serum (M−) and as collagen sandwiches with (S+) or without serum (S−). Messenger RNA levels of CYP1A2 (a), CYP2E1 (b), CYP2C (c) and CYP3A1 (d) were determined with custom ‘Sentrix BeadChip Arrays’ from Illumina, containing 550 genes. Data from three biological replicates were analysed with the ‘Genedata Expressionist Analyst’ software. Culture samples were taken 1 d (A), 2 d (B), 3 d (C), 5 d (D), 7 d (E) and 9 d (F) after the designated time t0 = 24 h after seeding. Bars illustrate gene expression changes relative to samples from respective cultures at t0. Non-significant changes are not shown (missing bars)
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commercial databases, with data from in vivo but also in vitro experiments (e.g. GENELOGIC (USA), ICONIX Biosciences (USA)). Although most data are generated from in vivo liver samples, there are efforts to build databases for the screening of hepatotoxicity based on primary hepatocyte cell culture experiments, making it necessary to characterize the cell culture model used, by genomic and proteomic approaches [121, 122, 143]. One step further is the Systems Biology approach, the mathematical modelling of biological processes based on the knowledge of cellular physiology, ultimately leading to in silico biology or the ‘virtual cell’. Although there are various kinds of software for the description and modelling of signalling pathways, the complexity of biological processes limits the application of systems biology, even though there may be an impact on drug discovery in the future [144–146].
7.4 7.4.1
Application of Short and Long-Term Hepatocyte Cultures Cytotoxicity Measurements in Long-Term Hepatocyte Sandwich Cultures
Successive applications are not feasible for cytotoxicity measurements in monolayer cultures. However, repeated dosing of long-term cultures of primary hepatocytes offers the possibility to detect ‘chronic’ effects. This is especially interesting since most in vitro models have very limited functionality over extended periods of time. Rather than generating acute toxic effects with high concentrations of a test compound, the utilization of substance levels close to pharmacologically active plasma concentrations increases the in vivo relevance of the data generated. Figure 7.7 gives an example of long-term cytotoxicity measurement in collagen sandwich cultures of primary human hepatocytes. Cells were treated daily for eight days and the cellular energy status was analysed by measuring the ATP content. The different substances used in this experiment lead to distinct reactions in culture. In comparison to the results obtained after two days, the non-toxic negative control, Metformin, did not show increased cytotoxicity after eight days, which supports the stability of the cultures over time. In contrast, troglitazone displayed increasing toxicity with increasing incubation times. Another feature was displayed by compound ‘X’. After showing considerable toxicity after two days, the IC50 values increased with incubation time, indicating adaptation of the cells to the toxic properties of this substance. This example clearly shows that with long-term cytotoxicity measurements more detailed information on the toxic potential of substances can be obtained. 7.4.2
Cytochrome P450 Induction in Long-Term Hepatocyte Sandwich Cultures
Short-term cultures of primary hepatocytes are frequently used to assess the CYP inducing capability of candidates in preclinical drug development. CYP induction may result in unfavourable drug–drug interactions, perhaps leading to the accelerated/reduced clearance of a co-administered drug. With regard to the more stable expression of CYPs, investigating CYP induction in serum-free collagen sandwich cultures of primary hepatocytes may give more reproducible results. Figure 7.8 shows an example of the analysis for CYP1A1/2 induction, on the mRNA level, in collagen sandwich cultures treated with the model inducers 3-methylcholanthrene (3-MC) and BNF after 3 days in culture. 3-MC and BNF induced
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Figure 7.7 Long-term cytotoxicity measurement in serum-free collagen sandwich cultures of primary human hepatocytes. Cells were treated with the indicated concentrations of the negative control Metformin (A), the test substance compund X (B) and the positive control troglitazone (C) for 8 days with daily medium changes. Cytotoxicity was determined as the cells’ ATP contents relative to the vehicle-treated control and is displayed after 2 days (a) and after 8 days (b) of treatment. The boxes highlight the increased cytotoxicity of the indicated troglitazone concentrations after 8-day treatment while the circles show the decreased sensitivity to compund X after 8 days
CYP1A, where CYP1A1 was more strongly induced in comparison to CYP1A2. These results support the applicability of long-term hepatocyte cultures for CYP-induction studies after extended periods of time. In addition, if treatment is stopped but culturing is continued, this offers the ability to define whether CYP induction can be resolved after a recovery period. It has even been suggested that serum-free collagen sandwich cultures can be used to examine CYP induction of several test compounds consecutively in one culture with recovery phases between treatment stages (PRIMACYT Cell Culture Technology GmbH, personal communication). This would be a step towards higher throughput and also help to further reduce animal usage in preclinical drug development. 7.4.3
Gene Expression Profiling in Short-Term Hepatocyte Cultures – Comparison with in vivo
Although hepatocyte sandwich cultures are clearly superior to monolayer cultures with respect to hepatocyte-like morphology and expression of hepatotypic genes, valuable results can also be obtained from short-term experiments with monolayer cultures. We elucidated whether the drug development candidate and mixed PPARα/γ agonist EMD 392949 (EMD) showed a species-specific induction of the major drug metabolizing enzymes in rats and
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Figure 7.8 CYP induction in serum-free primary rat hepatocyte sandwich cultures. Cells were treated 3 days after seeding with 10 μM 3-MC (A), 10 μM BNF (B) and 20 μM BNF (C). After 24 h treatment, CYP1A1 and CYP1A2 mRNA levels were determined using quantitative ‘TaqMan’ real-time PCR. Values were calculated by the efficiency-corrected comparative CT method with 18S rRNA serving as calibrator. Bars illustrate mean values of fold induction from triplicate measurements of a single PCR run with standard deviations relative to the vehicle control
humans. Furthermore, EMD was compared to fenofibrate, a PPARα agonist, Muraglitazar, a mixed PPARα/γ agonist and pioglitazone, a PPARγ agonist. Compound-specific gene expression changes were examined in primary rat and human hepatocytes, cultured on collagen I coated plates and treated for 24 h and 72 h. Strikingly, alterations in the expression of selected PPARα and PPARγ target genes that were observed in liver samples from a subchronic rat toxicity study were reproduced in primary rat hepatocyte cultures (Figure 7.9). In addition, species-specific reactions could also be detected. For example, the induction of inter-individual and inter-species variability of PPARα- or PPARγ -dependant reactions in rodents or primates, and the CYP-inducing potential of several compounds that was observed in human but not in rats (Figure 7.9). Interestingly, in rat and human hepatocyte cultures some compound-induced alterations were more prominent at the 24 h time-point, where others were more distinct after 72 h. Another feature worth mentioning was the difference in donor-dependent response to the pharmaceuticals in human hepatocyte cultures (data not shown). This clearly shows the necessity to analyse cells from several human donors in order to obtain reliable results.
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Figure 7.9 Gene expression analysis in rat liver samples (a) and serum-free rat (b) and human (c) hepatocyte monolayer cultures. Messenger RNA levels were determined using quantitative real-time PCR with ‘TaqMan Low Density Arrays’ (Applied Biosystems). Values were calculated by the efficiency-corrected comparative CT method with 18S rRNA serving as calibrator. Bars illustrate mean values from three biological replicates/donors with standard deviations. Elevated levels of gene expression are indicated by positive figures > + 1, while ‘down-regulation’ is indicated by negative figures < − 1. Please note that the positive and the negative y-axes are scaled logarithmically in (a) and (b) and linear in (c). Shown are values of fold-regulation for the treatment-induced gene expression changes relative to the vehicle-treated control (set as 1). (a) Treatment of rats with 3 mg/kg (lo) and 100 mg/kg (hi) EMD 392949 for 17 weeks. (b) Rat and (c) human hepatocyte monolayer cultures treated for 72 h with 3 μM (A), 30 μM (B) and 100 μM (C) EMD 392949, 100 μM fenofibrate (D), 3 μM pioglitazone (E) and 1 μM (F) or 30 μM (G) muraglitazar. CPT1A, carnitine-palmitoyltransferase 1A; G6Pc, glucose-6-phosphatase, catalytic subunit; GCK, glucokinase; HNF4α, hepatocyte nuclear factor 4 alpha; PXR, pregnane-X-receptor; CAR, constitutive androstane receptor; ACOX, acyl-coenzyme A oxidase 1, palmitoyl; MDR1, multidrug resistance transporter 1; ADPGK, ATP-dependent glucokinase; PEPCK, phosphoenolpyruvate carboxykinase 1, soluble; OATP2, organic anion transporter 2; OATP8, organic anion transporter 8
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Overall, rat hepatocytes in vitro perfectly reflected the observed effects in vivo for selected deregulated PPAR target genes. These results showed that primary hepatocyte cultures could serve as an appropriate model to predict effects observed in vivo.
7.5
Summary
Many different in vitro models are currently in use with the goal to predict human hepatotoxicity. Each system has specific limitations with regards to availability, throughput, viability of the cells over time and the possibility to analyse a variety of parameters. We have presented here long-term hepatocyte sandwich cultures as a valuable in vitro model for the study of hepatotoxicity. Hepatocyte preparations are available from many species, including humans, allowing the study of species-specific effects. The survival of hepatocytes in sandwich cultures and the maintenance of hepatotypic functions are stably preserved for a few weeks. This offers the opportunity to increase incubation time in order to study long-term effects and also to apply pharmacologically relevant concentrations of the test compound. Since the number of cells needed for the analysis of a specific parameter is usually low, depending on the sensitivity of the method, and experiments can be conducted in various cell culture formats, cultures of primary hepatocytes can be considered a medium-to-high throughput system. Another important feature is the ability to analyse many different methods in parallel, making it possible to obtain various data from the same sample. Optical clarity and stable morphology allow the use of the full range of optical methods, including histopathological methods and HCI. Classic in vitro cytotoxicity measurements, as well as many biochemical assays, can be performed since most hepatotypic functions are preserved. In addition, cell culture samples are compatible with most ‘-omics’ analyses. Hence, a diversity of results can be created from hepatocyte cultures and integrated into HCS and Systems Biology approaches to bring them into biological context. Therefore, we consider long-term hepatocyte sandwich cultures a valuable tool for the assessment of hepatoxicity in vitro, in order to reduce time and the number of animals used in preclinical drug development, and also to provide mechanistic data for the understanding of toxic mechanisms.
Acknowledgements We thank Dr P.-J. Kramer, Merck KGaA for supporting this work and Professor L. Richert for providing human hepatocytes.
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Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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8 Biomarkers of Mycotoxin Exposure in Liver Toxicity Angela J. Harris
8.1
Introducion
Humans are exposed daily to a multitude of environmental and dietary agents capable of causing adverse health effects if absorbed at sufficient doses. Technical advances in the identification and quantification of chemicals and chemical metabolites in biological tissues, as well as recognition that individual chemicals or chemical classes affect biological responses in specific ways, has led to an upsurge in research dedicated to the discovery of biomarkers of chemical exposure. Obviously, the first step in this process is identification of putative biomarkers. The relatively new fields of genomics, proteomics and molecular epidemiology have provided particularly powerful new tools for this step. Both genomics and proteomics technologies allow investigators to determine the effects of chemical or biological agents on expression of hundreds to thousands of genes and proteins simultaneously at any given point in time. By using these tools, not only can an investigator detect altered expression of a single gene or protein of interest as has traditionally been measured using Northern and Western blot techniques, respectively, but can detect changes in global patterns of expression. Thereby, patterns of expression may also be discerned which could serve as a biomarker pattern indicative of any number of biological effects, including exposure, toxicity, genotoxicity, drug efficacy or status of tumor progression. Identification of putative biomarkers is followed by a validation process that is used to verify its predictive value. Biomarkers have been defined as A physical sign or laboratory measurement that occurs in association with pathological process and that has putative diagnostic and/or prognostic utility (Lesko and Atkinson, 2001). There are three basic
Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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types of biomarkers: biomarkers of dose, biomarkers of a biological effect and biomarkers of individual susceptibility. At the present time, there are very few validated biomarkers of exposure to environmental chemical agents. Validation requires that the presence of the biomarker in humans be linked to exposure to the agent in question. This has been useful in occupational monitoring programs for exposure to certain chemicals. Some chemicals or their metabolites can be monitored in the blood and/or urine of workers that are potentially exposed in the workplace. These biomarkers, biological exposure indices (BEIs), are used primarily to quantify or estimate the absorbed dose of a chemical in an occupational setting. However, in many cases, the presence of a biomarker at any concentration in a biological tissue cannot be correlated with the probability that any adverse health effect will occur. This is due to many factors which are related to the half-life of the biomarker in the biological tissues, lack of sufficient information to correlate qualitative or quantitative measure of the biomarker with adverse health effects and a lack of understanding concerning the specificity or mechanism of toxicity. Validated biomarkers are those which have been specifically associated with exposure to the agent, are linked quantitatively to a biological effect and/or can be used to predict the susceptibility of the exposed individual to that agent.
8.2
Mycotoxin and Liver Toxicity
Mycotoxins are a diverse group of naturally occurring toxins produced by various fungal species. Humans are most commonly exposed to mycotoxins through ingestion of foods that have been contaminated with mycotoxin-producing strains and many of these toxins have been shown to be hepatotoxic, hepatocarcinogenic or both. Aflatoxins are produced by Aspergillus flavus and Aspergillus parasiticus and are a worldwide problem in areas where hot, humid conditions promote fungal growth on crops. Aflatoxins cause liver toxicity and are genotoxic. Aflatoxin B1 (AFB1 ) is classified by the International Agency for Research on Cancer (IARC) as a human liver carcinogen. Biomarkers for aflatoxins, particularly AFB1 have been extensively studied and are the most well validated of any mycotoxin biomarker. Fumonisin B1 (FB1 ), a mycotoxin produced by Fusarium verticillioides has also been reported to have toxic or carcinogenic effects on the liver. AFB1 and FB1 are structurally diverse and have different mechanisms of action. Therefore, identification of biomarkers will necessarily be mycotoxin- or mycotoxin class-specific. Biomarker identification for mycotoxin exposure is a growing area of research, in part due to increasing interest in reported health effects in occupants of buildings contaminated with molds capable of mycotoxin production. A summary of crops and food occasionally contaminated with AFB1 or FB1 is found in Table 8.1. Except for AFB1 , there are no well validated biomarkers of exposure to specific mycotoxins. Biomarkers of AFB1 exposure in animals and humans are an example of well validated biomarkers of environmental exposure and should be considered a model for further development of validated mycotoxin biomarkers. 8.2.1
Aflatoxin B1
AFB1 is a potent hepatotoxin and hepatocarcinogen in many animal species and in man. Ingestion of foods heavily contaminated by AFB1 causes aflatoxicosis in both animals
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Biomarkers of Mycotoxin Exposure in Liver Toxicity Table 8.1
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Summary of foods potentially affected by mycotoxins
Mycotoxin
Primary affected crops
Mycotoxin producer
Aflatoxin
Corn, peanuts, sorghum, milk, cottonseed Mill, wheat, nuts Sorghum, rice, beer, mung beans
Aspergillus flavus Aspergillus parasiticus Fusarium verticillioides
Fumonisin
and humans. Severe aflatoxicosis is characterized by hemorrhagic necrosis of the liver, edema, lethargy and death, particularly in children (Cullen and Newberne, 1993). AFB1 is classified as a known human carcinogen and causes primary liver cancer in several animals, including rainbow trout (Sinnhuber et al., 1968), rats (Busby and Wogan, 1984) and tree shrews (Reddy et al., 1976). There is also substantial epidemiological evidence of increased risk for primary liver cancer in individuals living in areas where high concentrations of AFB1 are present in food, especially in areas where hepatitis B infection is also common (Chen et al., 1996; Groopman et al., 1996). Several different aflatoxins are produced by Aspergillus; however, AFB1 is by far the most potent. AFB1 is metabolized by cytochromes P450, primarily CYP1A2 and CYP3A4, to the reactive metabolite AFB1 -8,9 epoxide which readily binds to nucleophilic sites on DNA (Ueng et al., 1995; Gallagher et al., 1996). AFB1 -8, 9 epoxide preferentially binds at the N-7 position of guanine due to the physical characteristics of the metabolite and DNA (Gopalakrishnan et al., 1990). Thus, the predominant DNA lesion formed is the 8,9dihydro-8- (N7-guanyl)-9-hydroxy-AFB1 (AFB1 -N7-Gua) adduct (Essigmann et al., 1977; Lin et al., 1977). It is this lesion that is responsible for the genotoxicity and carcinogenicity of AFB1 . Bulky AFB1 -N7-Gua adducts that are not repaired prior to DNA replication and subsequent cell division may result in mutations at the site of the lesion. The most commonly reported mutations are G → T transversions (Foster et al., 1983; Cariello et al., 1994) although G → C transversions, as well as G → A transitions have also been noted (Levy et al., 1992). The accumulation of mutations in genes playing a critical role in tumor progression or suppression increases the risk of cancer. Since metabolism of AFB1 occurs predominantly in the liver, the highest level of the reactive metabolite, AFB1 -8, 9 epoxide, will also be present in the liver; hence, the liver as the primary target organ for AFB1 induced toxicity and cancer.
8.3
Biomarkers of AFB1 Exposure
The AFB1 -N7-Gua adduct is rapidly removed from DNA by nucleotide excision repair pathways (Bedard and Massey, 2006) and excreted in the urine from which it can be isolated using immunoaffinity chromatography and then quantified by high performance liquid chromatography (HPLC) (Groopman, 1985, 1992a). AFB1 dose after ingestion is well correlated with the concentration of AFB1 -N7-Gua adduct present in the urine of exposed animals (Croy and Wogan, 1981; Groopman et al., 1992a) and humans (Groopman et al., 1992b). The level of AFB1 -N7-Gua adduct in urine is also well correlated with the formation of DNA adducts in the liver (Groopman et al., 1992a) and so is a biomarker
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both of the severity of biological effects and of dose. Since the bulky AFB1 -N7-Gua DNA adduct is rapidly repaired, the urinary AFB1 -N7-Gua biomarker is considered an indicator of recent exposure. AFB1 also forms protein adducts. The AFB1 -8,9 epoxide can be further modified to AFB1 -8,9 dihydrodiol which may be slowly converted to a dialdehyde phenolate ion capable of forming Schiff bases with lysine to form protein adducts (Wild et al., 1986; Sabbioni et al., 1987). The primary serum protein adduct formed is aflatoxin–albumin (serum AF– alb). Unlike DNA adducts, serum protein adducts are not repaired. Therefore, serum AF-alb is a better indicator than urinary AFB1 -7-Gua of cumulative AFB1 exposure. A positive correlation between serum AF-alb and AFB1 dose in animals and humans exposed to AFB1 has been established (Wild et al., 1990a,b, 1992). Epidemiological studies have also evaluated the relationship between AFB1 adduct biomarkers and the presence of primary liver cancer in geographic areas with high dietary intake of AFB1 -contaminated foods. A nested case control study conducted in Shanghai showed that the relative risk (RR) of primary liver cancer (PLC) in men who were positive for AFB1 -N7-Gua was 9.1 (95 % CI = 2.9–29.2) (Qian et al., 1994). Interestingly, the RR was 7.3 (95 % CI = 2.2–24.4) for individuals that were positive for hepatitis B antigen (HBsAg) but negative for aflatoxin urinary biomarkers. However, the RR for PLC was 59.4 (95 % CI = 16.6–212) for individuals who were positive for both biomarkers, indicating a strong synergy between HBV status and dietary AFB1 exposure. Another nested control study in Taiwan showed that the odds ratio (OR) for development of PLC in individuals that were positive for urine aflatoxin biomarkers was 1.7 (95 % CI = 0.3–10.8), compared to an OR of 111.9 (95 % CI = 13.8–905) in individuals positive for both HBsAg and AFB1 -N7-Gua (Wang et al., 1996). These data indicate the effectiveness of agent-specific biomarkers as monitors of exposure in epidemiological studies. Urine and serum biomarkers of AFB1 exposure have also been used to evaluate putative biomarkers of susceptibility in humans. For instance, serum AF-alb and urine AFB1 -N7Gua levels in humans exposed to AFB1 have been compared in individuals with polymorphisms in various Phase I and Phase II metabolic genes (Wild et al., 2000; Sun et al., 2002) and DNA repair genes (Lunn et al., 1999) in order to determine the effects of certain polymorphisms on metabolism of AFB1 and subsequent formation of DNA and protein adducts.
8.4
Genomic Biomarker Identification
AFB1 induced changes in gene expression after short-term exposure in vitro have been reported in cultured primary rat hepatocytes (Harris et al., 1998), Saccharomyces cerevisiae engineered to express human CYP1A2 (Guo et al., 2006), cultured primary human hepatocytes and HepG2 cells (Harris et al., 2004). Investigators have also evaluated altered gene expression in AFB1 -induced tumors in tree shrews (Duan et al., 2005; Li et al., 2004) and rainbow trout (Tilton et al., 2005). At the present time, there are insufficient data to identify any particular gene or pattern of gene expression that is specifically altered in AFB1 -induced toxicity or tumor development. This may be partially due to the selection of the gene array, different dosing regimes, comparison of in vitro versus in vivo effects, different species, endpoint selection and other variables in experimental design.
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Fumonisin
Fumonisin (FB1 ), produced by Fusarium verticillioides (= monoliforme) (Sheldon, 1904) is hepatotoxic in several species (Gelderblom et al., 1988, 1991, 2001; Marasas et al., 1988a) and a hepatocarcinogen in rats and mice (Wilson et al., 1985; Voss et al., 1995; Howard et al., 2001). Fusarium strains affect corn crops worldwide (Table 8.1) and have been linked to increased risk for esophageal cancer in areas with high dietary consumption of FB1 -contaminated foods (Marasas et al., 1979, 1988b). FB1 has been shown to act as a cancer promoter in rats treated with diethylnitrosamine (DEN), a known initiator (Gelderblom et al., 1988). Gamma-glutamyl-tranpeptidase-positive (GGT+ ) foci appeared in the liver of rats after four weeks on a diet containing 0.1 % FB1 . The International Agency for Research on Cancer (IARC) has classified FB1 as possibly carcinogenic to human, class 2B (IARC, 1993). Unlike AFB1 , FB1 is not genotoxic and the mechanism of carcinogenesis is likely epigenetic. At least one outbreak of acute human and animal illness has been attributed to FB1 contaminated sorghum and/or bread (Bhat et al., 1997). In 1995, individuals in 27 villages in India became ill after unseasonal rains caused unusually high mold growth on the sorghum and maize crops. Many villagers who ate unleaved bread prepared from moldy maize and/or sorghum developed abdominal pain, diarrhea and borborygmi, typically within an hour after ingestion. The symptoms resolved once exposure ceased. There were no human fatalities although some domestic animals fed the moldy foods died. There have also been reports of equine leukoencephalomalacia and mild hepatosis in horses that ate Fusariumcontaminated feed (Marasas et al., 1988a). Fumonisins have been shown to disrupt ceramide synthase (Merrill et al., 2001; Riley et al., 2001). As a result, sphingolipid metabolism is disrupted, increasing the levels of ceramide precursors sphinganine (Sa) and sphingosine (So) in tissues, including the liver (Norred et al., 1992) and serum of treated animals (Wang et al., 1992). Disruption of sphingolipid metabolism has significant effects on normal cellular function which is dependent on the regulated synthesis and turnover of complex sphingolipids to maintain a number of signaling pathways. It is hypothesized that disruption of sphingolipid metabolism plays a role in hepatotoxicity; however, the specific pathways by which these events cause toxic and carcinogenic effects in the liver of rodents have not been determined.
8.5
Biomarkers of FB1 Exposure
Unlike AFB1 , there are no well-validated biomarkers for FB1 exposure; however, serum Sa and So levels have been proposed as candidates of exposure and biological effect. The most sensitive candidate biomarker of FB1 exposure reported thus far is serum Sa relative to So, the Sa/So ratio (Riley et al., 1994; Tran et al., 2006). There are a number of studies that have evaluated the use of serum or urine Sa or Sa/So ratios as biomarkers of exposure. A few studies have also attempted to correlate these potential biomarkers with known serum biomarkers of liver toxicity to evaluate their usefulness as biomarkers of biological effects. Some of these data are summarized below.
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Serum Biomarkers in Animals
Serum Sa and So concentrations increased in ponies fed diets containing 44 mg/kg FB1 , even before increased levels of serum biomarkers of liver damage, such as aspartate aminotransaminase or gamma glutamyl transaminase were apparent (Wang et al., 1992). Serum Sa concentration peaked seven days after exposure at serum levels 22-fold higher than before dosing. The serum Sa/So ratio was also increased 6.4-fold. Increased serum biomarkers of liver toxicity typically occurred at about day 10. Tran et al. (2006) reported that an increase in serum Sa could be detected in ducks fed feed containing FB1 as low as 2 mg/kg. A dose-related increase in serum Sa/So was observed when the cumulative FB1 dose in feed was 100–200 mg; however, there was no apparent increase in serum Sa/So above that dose. Tran et al. (2006) also found that effects on serum Sa/So were more apparent in ducks exposed over a short period of time (7–21 days) versus a long period of exposure (42–77 days). A statistically significant dose-related increase in serum Sa/So ratio compared to controls was also reported in vervet monkeys fed a diet with culture material of Fusarium verticilloidides containing FB1 (control mean Sa/So = 0.43; low mean Sa/So = 1.72; high mean Sa/So = 2.57) (Shephard et al., 1996; Gelderblom et al., 2001). Blood serum Sa/So levels were assessed over a 51-day period in vervet monkeys fed a single dose of 1 mg/kg or 10 mg/kg of purified FB1 . Serum Sa/So ratios significantly elevated in the high-treatment group over controls ( p < 0.05) were observed three days after exposure, continued to increase for 27 days and then declined to control values by day 51 (van der Westhuizen et al., 2001a). As reported above for ponies and ducks, the initial rise in serum Sa/So ratios corresponded in general with an increase in serum levels of biomarkers of liver toxicity (GGT and ALT). Vervet monkeys dosed three times a week with 1 mg/kg of FB1 for 51 days had a maximum mean plasma Sa/So ratio of 4.3 after 30 days of treatment (van der Westhuizen et al., 2001b). This level was approximately 3-fold higher than that of non-treated animals. The plasma Sa/So ratio declined by about one third over the course of the remaining 21 days of treatment. These data indicate that the serum Sa/So ratios may be indicative of recent acute exposure to FB1 ; however, a dose–response relationship has not been clearly defined nor has it been clearly correlated with the severity of biological effects.
8.5.2
Urine Biomarkers in Animals
Interestingly, only modest or no differences have been reported by many authors in urine Sa/So ratios observed in animals treated with FB1 (Wang et al., 1992; Shephard et al., 1996; Castegnaro et al., 1998; Garren et al., 2001; van der Westhuizen et al., 2001a,b). However, some studies have reported that dietary FB1 can elevate Sa/So ratios in urine. For instance, rats fed 1–5 mg/kg FB1 on a daily basis had urine Sa/So ratios that ranged from 1.2 to 10 compared to a range of 0.1 to 0.7 in untreated rats (Castegnaro et al., 1996). Some of these seeming inconsistencies may be temporally related to length of treatment and the time of testing relative to treatment. For instance, although an initial rapid increase in the urine Sa/So ratio was seen in vervet monkeys treated with FB1 , it was not sustained and dropped to pre-treatment values very quickly (van der Westhuizen et al., 2001b). In addition, significant interindividual differences in urine Sa/So ratios may obscure subtle effects of FB1 on Sa/So ratio in urine (Garren et al., 2001; Gelderblom et al., 2001).
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Serum Biomarkers in Humans
Methods have been developed and tested for analysis of serum Sa and So in humans (Shephard and van der Westhuizen, 1998; Castegnaro et al., 1998) although only a limited number of studies have evaluated serum Sa/So ratios in humans. No significant differences were seen in serum Sa/So ratios of African males and females, or in individuals living in different geographic areas in Africa (van der Westhuizen et al., 1999). Recently, serum Sa/So ratios have been evaluated as a biomarker for increased risk of neural tube defects in Mexican–American women exposed to FB1 in corn tortillas (Missmer et al., 2006). A dose-related increase was seen in the OR for neural tube defects (NTDs) with increased serum Sa/So ratio and FB1 intake, except at the highest levels. The OR for NTDs was 4.4 (95 % CI 1.2–15.5) in individuals with a serum Sa/So ratio of 0.31–0.35. Control values were ≤ 0.10. The OR for NTDs was 2.3 (95 % CI 1.1–5.1) for pregnant women with a daily dietary FB1 intake of 150.1–650 ng/kg/day.
8.5.4
Urine Biomarkers in Humans
Methods have been determined to analyze the concentration of Sa and So in the urine of humans potentially exposed to dietary FB1 (Castegnaro et al., 1996; Solfrizzo et al., 1997; Qiu and Liu, 2001). These studies report that Sa and So concentrations in the urine of females are higher than in males. It is not known what causes this difference but it is hypothesized to be due to higher number of sloughed cells in the urine of females. Qiu and Liu (2001) evaluated the Sa/So ratio in the urine of human volunteers who consumed a corn diet containing 0.08 to 41.1 mg/kg for one month. These data indicate that ingestion of FB1 affects sphingolipid metabolism in humans since there was a trend toward higher Sa/So ratios in individuals eating foods with the highest contamination by FB1. Males appeared to be more sensitive to this effect than females. A second study compared urinary Sa/So ratios in humans living in areas known to ingest more FB1 in their diets (North Argentina and Brazil) to humans living in areas where maize consumption was very low (Central Argentina) (Solfrizzo et al., 2004). There was a statistically significant difference ( p < 0.001) in the Sa/So ratios from the FB1 -containing diets (mean Sa/So = 1.27) compared to controls (mean Sa/So = 0.36); however, there was also a statistically significant difference in the Sa/So urinary ratios between the two high FB1 areas (South Brazil mean = 1.57 vs. North Argentina mean = 0.69, p < 0.05) even though the mean FB1 intake was similar between the two groups (0.35 mg/kg). These data suggest that in humans the use of the Sa/So ratios may not be useful as a biomarker of exposure except in geographic areas where foods are highly contaminated with FB1 . Similar conclusions were reached in a study of male and female volunteers in rural areas in Africa (van der Westhuizen et al., 1999). Clearly, more research needs to be done in this area to validate the usefulness of this biomarker in the human population for exposure and/or biological effect after exposure to dietary levels likely to occur in humans eating FB1 -contaminated foods. Although the Sa and Sa/So ratios may ultimately be useful only in the case of relatively high, acute exposures, there are some data to suggest that actual measurements of fumonisins in the hair of experimental animals (Sewram et al., 2001) and humans (Sewram et al., 2003) may be useful for monitoring chronic exposure to dietary levels of fumonisins.
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Table 8.2 Gene expression changes in FB1 -exposed mice Gene description
Accession number
Glutathione S-transferase, mu3 (Gstm3) Ras homolog gene family, member u (Arhu) Cyclin-dependent kinase inhibitor 1A (Cdkn1a) Cyclin D1 (Ccnd1) Ras homolog gene family, member Q Glutathione S-transferase, pi 2 (Gstp2) Flavin-containing monooxygenase 5 (Fmo5) Cytochrome P450, family 3 (Cyp3a16) Sulfotransferase-related protein SULT-X2
J03953 AV246963 AW048937 AI849928 AW060401 X53451 U90535 D26137 Sult-x2
8.6
Fold change 15.42 8.27 5.8 5.22 4.72 4.68 4.24 3.16 −73.3
Genomic Biomarker Identification
Limited studies are available that have evaluated the effects of FB1 on gene expression in the liver. Voss et al. (2006) reported that wild type mice fed culture material from F. verticillioides (CM) or purified FB1 for 7 days had increased expression of 180 genes and decreased expression of 193 genes. A group of 48 genes were affected by FB1 treatment only, and not by CM. The expression of 138 genes was altered by both FB1 and CM. Genes with the highest changes in gene expression after FB1 treatment are summarized in Table 8.2. These genes include those associated with cellular proliferation (cyclins), signal transduction (ras) and Phase I and Phase II metabolism. At the present time, there are insufficient data to evaluate their usefulness as potential biomarkers of exposure.
8.7
Summary
Human exposure to mycotoxins is a worldwide problem. Acute toxicity due to overexposure to mycotoxins in the diet can cause symptoms that often include nausea, vomiting, diarrhea and abdominal pain. Hepatotoxicity may also occur. Chronic exposure to some mycotoxins, such as AFB1 and FB1 , is also associated with primary liver cancer. Identification of biomarkers of mycotoxin exposure allows investigators to determine the extent and effects of exposure. They are also invaluable in epidemiological studies designed to determine the effects of exposure in humans. The use of genomics has also been applied in a limited fashion to identify individual genes or patterns of gene expression that may be used as biomarkers of exposure to specific mycotoxins or biological effects (liver toxicity). Although these studies may be eventually used to identify genes that are markers of exposure and/or toxicity to mycotoxins, they will have limited utility in evaluating human exposure or biological effect in the exposed population. Useful biomarkers are those which can be monitored in easily accessible fluids, such as urine, blood or saliva. Therefore, applying proteomic and metabonomics technologies to identify putative biomarkers of mycotoxin exposure is likely to be the most effective strategy for future work in mycotoxin research.
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Missmer SA, Suare L, Felkner M, Wang E, Merrill AH Jr, Rothman KJ and Hendricks KA. (2006). Exposure to fumonisins and the occurrence of neural tube defects along the Texas–Mexico Border. Environ Health Perspect 114: 237–241. Norred WP, Plattner RD, Vesonder RF, Bacon CW and Voss KA. (1992). Effects of selected secondary metabolites of Fusarium moniliforme on unscheduled synthesis of DNA by rat primary hepatocytes. Food Chem Toxicol 30: 233–237. Qian GS, Ross RK, Yu MC, Yuan JM, Gao YT, Henderson BE, Wogan GN and Groopman JD. (1994). A follow-up study of urinary markers of aflatoxin exposure and liver cancer risk in Shanghai, People’s Republic of China. Cancer Epidemiol Biomark Prev 3: 3–10. Qiu M and Liu X. (2001). Determination of sphinganine, sphingosine and Sa/So ratio in urine of humans exposed to dietary fumonisin B1 . Food Addit Contam 3: 263–269. Reddy JK, Svododa DJ and Rao MS. (1976). Induction of liver tumors by aflatoxin B1 in the tree shrew (Tupaia glis), a nonhuman primate. Cancer Res 36: 151–160. Riley RT, Voss KA, Yoo HS, Gelderblom WC and Merril AH Jr. (1994). Mechanism of fumonisin toxicity and carcinogenesis. J Food Prot 57: 638–645. Riley RT, Enongene E, Voss KA, Norred WP, Meredith FI, Sharma RP, Spitsbergen J, Williams DE, Carlson DB and Merrill AH Jr. (2001). Sphingolipid perturbations as mechanisms for fumonisin carcinogenesis. Environ Health Perspect 109(Suppl. 2): 301–308. Sabbioni G, Skipper P, Buchi G and Tannenbaum SR. (1987). Isolation and characterization of the major serum albumin adduct formed by aflatoxin B1 in vivo in rats. Carcinogenesis 8: 819–824. Sewram V, Nair JJ, Nieuwoudt TW, Gelderblom WC, Marasas WF and Shephard GS. (2001). Assessing chronic exposure to fumonisin mycotoxins: the use of hair as a suitable noninvasive matrix. J Anal Toxicol 25: 450–455. Sewram V, Mshicileli N, Shephard GS and Marasas WF. (2003). Fumonisin mycotoxins in human hair. Biomarkers 8: 110–118. Sheldon JL. (1904). A corn mold (Fusarium moniliforme sp.), in Agricultural Experimental Station of Nebraska 17th Annual Report, University of Nebraska, Lincoln, NE, USA, pp. 23–32. Shephard GS and van der Westhuizen L. (1998). Liquid chromatographic determination of the sphinganine/sphingosine ratio in serum. J Chromatogr 710: 219–222. Shephard GS, van der Westhuizen L, Thiel PG, Gelderblom WC, Marasas WF and van Schalkwyk DJ. (1996). Disruption of sphingolipid metabolism in non-human primates consuming diets of fumonisin-containing Fusarium moniliforme culture material. Toxicon 34: 527–534. Sinnhuber RO, Wales JH, Ayres JL, Engebrecht RH and Amend DL. (1968). Dietary factors and hepatoma in rainbow trout (Salmo gairdneri). 1. Aflatoxins in vegetable protein feedstuffs. J Natl Cancer Inst 41: 711–718. Solfrizzo M, Avantaggiato G and Visconti A. (1997). Rapid method to determine sphinganine/sphingosine in human and animal urine as a biomarker for fumonisin exposure. J Chromatogr 692: 87–93. Solfrizzo M, Chulze SN, Mallmann C, Visconti A, De Girolamo A, Rojo F and Torres A. (2004). Comparison of urinary sphingolipids in human populations with high and low maize consumption as a possible biomarker of fumonisin dietary exposure. Food Addit Contam 21: 1090–1095. Sun CA, Wu DM, Wang LY, Chen CJ, You SL and Santella RM. (2002). Determinants of formation of aflatoxin-albumin adducts: a seven-township study in Taiwan. Br J Cancer 87: 966–970. Tilton SC, Gerwick LG, Hendricks JD, Rosato CS, Corley-Smith G, Givan SA, Bailey GS, Bayne CJ and Williams DE. (2005). Use of a rainbow trout oligonucleotide microarray to determine transcriptional patterns in aflatoxin B1 -induced hepatocellular carcinoma compared to adjacent liver. Toxicol Sci 88: 319–330. Tran ST, Tardieu D, Auvergne A, Bailly JD, Babile R, Durand S, Benard G and Guerre P. 2006. Serum sphinganine and the sphinganine to sphingosine ratio as a biomarker of dietary fumonisins during chronic exposure in ducks. Chem Biol Interact 10: 41–50.
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Ueng YF, Shimada T, Yamazaki H and Guengerich FP. (1995). Oxidation of aflatoxin B1 by bacterial recombinant human cytochrome P450 enzymes. Chem Res Toxicol 8: 218–225. van der Westhuizen L, Brown NL, Marasas WF, Swanevelder S and Shephard GS. (1999). Sphinganine/sphingosine ratio in plasma and urine as a possible biomarker for fumonisin exposure in humans in rural areas of Africa. Food Chem Toxicol 37: 1153–1158. van der Westhuizen L, Shephard GS and van Schalkwyk DJ. (2001a). The effect of a single gavage dose of fumonisin B1 on the sphinganine and sphingosine levels in vervet monkeys. Toxicon 39: 273–281. van der Westhuizen L, Shephard GS and van Schalkwyk DJ. (2001b). The effect of repeated gavage doses of fumonisin B1 on the sphinganine and sphingosine levels in vervet monkeys. Toxicon 39: 969–972. Voss KA, Chamberlain WJ, Bacon CW, Herbert RA, Walters DB and Norred WP. (1995). Subchronic feeding study of the mycotoxin fumonisin B1 in B6C3F1 mice and Fischer 344 rats. Fundam Appl Toxicol 24: 102–110. Voss KA, Liu J, Anderson SP, Dunn C, Miller JD, Owen JR, Riley RT, Bacon CW and Corton JC. (2006). Toxic effects of fumonisin in mouse liver are independent of the peroxisome proliferatoractivated receptor alpha. Toxicol Sci 89: 108–119. Wang E, Ross PF, Wilson TM, Riley RT and Merrill AH Jr. (1992). Increases in serum sphingosine and sphinganine and decreases in complex sphingolipids in ponies given feed containing fumonisins, mycotoxins produced by Fusarium moniliforme. J Nutr 122: 1706–1716. Wang LY, Hatch M, Chen CJ, Levin B, You SL, Lu SN, Wu MH, Wu WP, Wang LW, Wang Q, Huang GT, Yang PM, Lee HS and Santella RM. (1996). Aflatoxin exposure and risk of hepatocellular carcinoma in Taiwan. Int J Cancer 67: 620–625. Wild CP, Garner RC, Montesano R and Tursi F. (1986). Aflatoxin B1 binding to plasma albumin and liver DNA upon chronic administration to rats. Carcinogenesis 7: 853–858. Wild CP, Jiang YZ, Allen SJ, Jansen LA, Hall AJ and Montesano R. (1990a). Aflatoxin–albumin adducts in human sera from different regions of the world. Carcinogenesis 11: 2271–2274. Wild CP, Jiang YZ, Sabbioni G, Chapot B and Montesano R. (1990b). Evaluation of methods for quantitation of aflatoxin–albumin adducts and their application to human exposure assessment. Cancer Res 50: 245–251. Wild CP, Hudson G, Sabbioni G, Chabot B, Hall AJ, Wogan GN, Whittle H, Montesano R and Groopman JD. (1992). Dietary intake of aflatoxins with the level of albumin-bound aflatoxin in peripheral blood in The Gambia, West Africa. Cancer Epidemiol Biomark Prev 1: 229–234. Wild CP, Yin F, Turner PC, Chemin I, Chapot B, Mendy M, Whittle H, Kirk GD and Hall A. (2000). Environmental and genetic determinants of aflatoxin–albumin adducts in The Gambia. Int J Cancer 86: 1–7. Wilson TM, Nelson PE and Knepp CR. (1985). Hepatic neoplastic nodules, adenofibrosis and cholangiocarcinomas in male Fischer 344 rats fed corn naturally contaminated with Fusarium moniliforme. Carcinogenesis 6: 1155–1160.
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Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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9 Mechanisms of Toxic Liver Injury Nora Anderson and J¨urgen Borlak
9.1
Introduction
Toxic responses may arise from acute or chronic exposure to chemicals and drugs. Unlike the intentional exposure to therapeutic drugs, exposure to industrial chemicals is accidental. This requires different strategies to investigate toxicity, which always coincides with metabolic dysfunctions and induction of stress-triggered programs. Whether or not an organ will overcome metabolic dysfunction and resume its physiologic function is highly dependent on the circumstances prevailing. In many ways, the liver, as the central organ for detoxification and clearance of exogenous and endogenous substances, constitutes a primary target for toxic reactions, but nonetheless has the remarkable ability to undergo regenerative growth. Several factors can be named that contribute to a high incidence of toxicity in the liver. These include remarkable exposure to high drug concentrations, enormous metabolic activity and the presence of several enzymes held responsible for generation of reactive metabolites, most notably ROS. Possible targets of toxic substances are macromolecular structures or individual molecules such as the bile acid transporters (i) or members of the nuclear receptor family (ii), as well as intracellular lipids (iii), proteins (iiii) or nucleic acids (iiiii) (see Figure 9.1). These targeted molecules become dysfunctional units and activate secondary pathways to result in programmed events, such as apoptosis, necrosis and autophagy, mitochondrial failure and immunological reactions (Kaplowitz, 2002; Lee and Ferguson, 2003). In addition, the functional integrity of a cell can also be disturbed by direct cytolytic reactions, membrane disruption and distortion of trans-membrane transport mechanisms. Even though several different mechanisms might be involved in the onset and progression of hepatotoxicity of a single substance, there are likely to be only a few central mechanisms that are activated within the general toxic response of the liver. Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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Figure 9.1
Cellular sites of drug-related toxicity
The actual sensitivity to certain toxins, however, is related to various determinants, which are also subject to high interindividual variability. Therefore, it is difficult to predict whether hepatotoxicity might occur upon treatment with a new substance, even though when studied in large patient cohorts. The complexity of different interacting pathways and mechanisms that are involved in the toxic responses complicate the distinct assignment of observed phenomena to either adverse effects or, at least in part, to therapeutic drug effects. This might explain why complications due to drug-related hepatotoxicity have such a high incidence, as for example in the USA, where incidences of 1 in 10 000 and 1 in 100 000 patients have been reported (Larrey, 2002). Despite knowledge gained over the past decade, no real progress has been made to face liver toxicity at least from the clinical point of view. Here we wish to discuss hepatotoxicity in a broad sense and follow several lines of evidence. We will particularly focus on the role of oxidative stress and inflammation for liver injury, but also summarize the vast amount of literature on toxicity induced by carbon tetrachloride, amantadine, iron, acetaminophen and halothane, among others. Whenever possible, the molecular pathology of a specific toxicant will be discussed in order to provide insight into mechanisms of the various stages of toxic liver injury. Finally, the needs to direct future research will be emphasized. 9.1.1
Basic Facts about Toxic Liver Injury
Notably, the development of toxic liver injury follows a two-staged course, the first phase of which is characterized by initiation of the injury and may involve direct interaction with a toxicant. This phase may exhibit dose-dependency (Mehendale, 2005). In contrast, the second phase is characterized by progress of the injury in a toxicant-independent fashion that is dominated by secondary events. Among the mechanisms contributing to phase two of toxic liver injury, the three leading mechanisms proposed are contribution of inflammatory cells (i) (Czaja et al., 1994; Laskin and Pendino, 1995; Piguet et al., 1990), oxidative stress and lipid peroxidation
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(ii) (Poli, 1993; Slater, 1984) and leakage of degrading enzymes (iii) (Mehendale and Limaye, 2005; Poli et al., 1987). Furthermore, tissue repair has been determined to be one of the major factors that influence the fate of the liver after toxic injury, which may either be directed towards regeneration or loss of function and necrosis. Therefore, the toxicological response to different toxicants to a significant part obeys similar principles and may be characterized by the same mechanisms, although the acute and first phase of liver injury may be initiated by mechanisms specific to the toxicant. This may also explain why the morphology of toxic or drug-related liver injury usually differentiates into one of the four following phenotypes: hepatocellular injury, often associated with elevated liver enzymes (e.g. isoniazid) (i), cholestatic injury, which due to damage of the bile duct epithelia displays predominantly elevated alkaline phosphatase (AP) (e.g. amoxixillin – clavulanic acid) (ii), mitochondrial injury involving microvesicular steatosis (e.g. valproic acid) (iii) and the often delayed immunologic injury that is associated with fever, rash or eosinophilia (e.g. phenytoin) (iiii) (Navarro and Senior, 2006). Some drugs, as indicated by the examples given in parentheses, typically cause certain morphologies, but do not have a specific correlation to them. Thus, they can induce a mixed injury that is characterized by more than one feature of the different morphologic phenotypes. It is not known why certain drugs induce certain morphologies of liver injury but it is likely that these drugs share similar mechanisms of initiating toxicity in the liver. In the following, mechanisms recognized in different types of liver injury will be discussed, and general features of toxicity will be portrayed in detail.
9.2
Cholestatic Liver Injury
Cholestasis may result from an impairment of bile secretion or an obstruction of bile flow. The mechanistics underlying cholestatic liver injury has been linked to an impairment of bile salt transport by inhibition or downregulation of ATP-dependent bile salt transporters and also to alterations of actin resulting in disruption of the cytoskeleton and an impaired transport of bile along the canalicular system and into the bile ducts (Cullen, 2005). Interaction with bile acid transporters is a mechanism that has been observed under administration of several drugs and toxicants leading to toxic liver injury and is believed to be a central event in the development of cholestatic liver injury (Lewis, 2000). The connection between bile acid transporters and liver injury has first been drawn from observations made after experimental partial hepatectomy, where the acute liver injury was accompanied by regulation of hepatobiliary organic anion transporters (Geier et al., 2002; Trauner et al., 2005). Bile production is important for the clearance of endogenous and exogenous metabolites through the liver, and such is its detergent effect for adsorption and digestion of lipids. The primary bile salts, cholic acid and chenodeoxycholic acid, are derived from cholesterol and released in conjugation with taurine and glycine conjugates (Russell and Setchell, 1992). The uptake of bile salts that underwent enterohepatic circulation is located at the basolateral membrane of the hepatocyte and is mediated by a Na+ -dependent transporter system that communicates with portal vein blood via the space of Disse. This transporter system is
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termed NTCP/Ntcp and belongs to a family of multi-specific organic anionic transporters (OATP/Oatp). Within the hepatocyte, bile acids are transported via diffusion or intracellular membrane vesicles. The canalicular transporters at the apical site of the hepatocyte belong to the family of ATP-dependent ABC transporters, such as the bile salt export pump (BSEP), multidrug transporter 2 (MDR2) and ABCG5/8 transporter (Stieger et al., 1992). Increased extracellular bile concentrations were shown to induce the apoptotic program of cell death in a death receptor-dependent fashion (Higuchi and Gores, 2003). Therefore, deregulation of bile transporters that leads to increased exposure to bile acids may cause serious hepatocellular damage and was furthermore demonstrated to trigger changes in intracellular signal transducers that affect cholangiocyte secretion, proliferation and survival (Xia et al., 2006). Defects of bile acid transporters have been demonstrated to be responsible for development of cholestasis in two of four types of progressive familial intrahepatic cholestasis (PFIC) (the Byler Syndrome (PFIC-2) and PFIC-3) (Jansen et al., 2001). These diseases arise from an impairment of the bile salt export pump (BSEP) and multidrug resistance protein 2 (MRP2) at the apical site of the hepatocytes. Another hereditary disease affecting bile acid transport is the Dubin–Johnson syndrome, in which mutations in MRP2 give rise to high serum bilirubin as a result of the liver’s inability to excrete this metabolite into bile (Paulusma et al., 1997). Carbon tetrachloride (CCl4 ) causes cholestasis and fibrosis and is a toxin frequently used to probe for mechanisms of hepatotoxicity. Indeed, carbon tetrachloride was found to interfere with the regulation of bile acid transporters, causing downregulation of sodium taurocholate co-transporting polypeptide (Ntcp) and organic anion transporting polypeptides 1 (Oatp1) and 2 (Oatp2) (Geier et al., 2002). The mechanisms responsible for this action are still not fully understood; however, pro-inflammatory mediators such as tumour necrosis factor alpha (TNF-α) were suggested to play a central role in CCl4 -induced liver injury and have been demonstrated to exert effects on bile acid transporters (Geier et al., 2002). Such were TNF-α and interleukin 6 (IL-6) shown to inhibit the sodium-dependent uptake of taurocholate by hepatocytes (Green et al., 1994; Whiting et al., 1995). Further in vitro studies have demonstrated that interleukin-1 beta (IL-1β) decreases basolateral Ntcp and Mrp2 promotor activity via reduction of the nuclear levels of transactivating retinoid X receptor/retinoic acid receptor (RXRα/RARXα) (Denson et al., 2000; Li et al., 2002). Moreover, TNF-α and IL-1β were shown to downregulate Oatp2 and MRP2, which are responsible for transport of bilirubin and other organic ions (Zollner et al., 2001). In the case of carbon tetrachloride, the upregulation of pro-inflammatory cytokines, which is frequently observed in cholestatic liver disease, was related to early activation of the early growth response factor-1 (Egr-1) transcription factor. This was supported by studies in Egr-1 knockout mice, where neutrophil accumulation, upregulation of macrophage inflammatory protein-2 (MIP-2) and intercellular adhesion molecule-1 (ICAM-1) in the liver were significantly reduced as compared to wild-type mice (Kim et al., 2006). Moreover, bile acids themselves revealed a stimulating effect on the expression of Egr-1 protein in hepatocytes in an animal model of cholestasis. Thus, elevated bile acids concentrations during cholestasis might cause a secondary downregulation of bile acid transporters by mediation of inflammatory cytokines (Kim et al., 2006).
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Investigations in animal models of hepatic ischemia-reperfusion indicated a crucial role of Kupffer cells for the development of cholestatic liver injury (Kudo et al., 2004). Hepatic ischemia-reperfusion is often followed by cholestasis and, in fact, was associated with downregulation of several bile acid transporting proteins, accompanied by increased levels of TNF-α, IL-1β and IL-6 and decreased levels of the nuclear transcription factor HNF-1α (Tanaka et al., 2006). Interestingly, depletion of Kupffer cells with gadolinium chloride prevented increased expression of TNF-α and effects on HNF-1α, suggesting that alterations in the expression of hepatic transporters in hepatic ischemia-reperfusion injury may be a Kupffer cell-mediated event (Tanaka et al., 2006). By contrast, Kupffer cell involvement in ischemia-reperfusion has been suggested to exhibit protection against liver injury in particular by upregulation of heme degradation and bilirubin excretion. This was implicated by results of a study in which Kupffer cell-depleted rats had displayed significantly elevated the levels of GOT and GPT, as well as serum endotoxin concentrations after ischemia-reperfusion compared to animals with an intact Kupffer cell response (Kobayashi et al., 2002). Direct interaction of the toxicant with bile salt exporter proteins or with genes coding for these is likely to be a primary event of cholestatic liver injury (Yamamoto et al., 2006). The latter has been implied in oestrogen-induced hepatotoxicity that is observed in women during pregnancy or upon contraceptive and postmenopausal hormone therapy (Schreiber and Simon, 1983). The synthetic oestrogen derivative 17α-ethinylestradiol (EE2) causes a decrease of the taurocholate transport at the hepatic canalicular membrane (Lee et al., 2000) and the sodium taurocholate co-transporting polypeptide protein (NTCP) and consequently reduces bile flow (Simon et al., 1996). Oestrogens are ligands of cytosolic receptors that belong to the family of steroid hormone receptors. Once activated these receptors are translocated into the nucleus, where they operate as transcription factors for specific genes. Hence, direct regulation of genes coding for bile transporter proteins would be one anticipated mechanism causing cholestatic liver injury as a result of EE2 administration. Indeed, recent studies in various transcription factor knockout models have recognized oestrogen receptor alpha to be responsible for repression of multiple transporters for hepatic bile and cholesterol, possibly accounting for rising extracellular bile concentrations (Yamamoto et al., 2006). In livers of wild-type mice, the oestrogen receptor alpha (ERα) mediated downregulation of bile acid and cholesterol transporters expression in response to EE2, subsequently decreasing the secretion of bile acids and cholesterol. Moreover, ERα induced a shift of bile acid synthesis towards the acidic pathway by upregulating the expression of CYP7B1 and downregulating the expression of CYP7A1 and CYP8B1. In contrast, Erα null mice were resistant to oestrogen 17α-ethinylestradiol (EE2)-induced hepatotoxicity (Yamamoto et al., 2006). Another possible mechanism in drug-induced cholestasis was found in liver toxicity induced by the immunosuppressive drug cyclosporin A, the antibiotic rifampicin and the antidiabetic drug glibenclamide. These potential liver toxicants cause cholestasis by direct interaction with bile acid transporters and were shown to competitively inhibit the BSEP taurocholate transport (Byrne et al., 2002). In the case of ANIT-induced liver injury, the pharmacodynamics of a drug-transporting bile acid transporter have been emphasized to essentially add to the mechanistics of liver toxicity. Notably, ANIT is conjugated to glutathione in the hepatocyte and then transported across the canalicular membrane by MRP2, where it damages the cholangiocytes lining
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the bile ducts (Liu et al., 2003). Secretion of ANIT and also arsenite by the Mrp2 transporter was demonstrated to induce a cycling of both toxins that was held accountable for a concentration-dependent depletion of GSH. By contrast, Mrp2-deficient rats displayed no GSH excretion and were protected from ANIT-induced cholestasis (Dietrich et al., 2001). 9.2.1
Cholestasis and Nuclear Receptors
Physiologically, bile acid homeostasis is maintained by the enterohepatic nuclear factors farnesol X receptor (FXR), liver X receptor (LXR), pregnane X receptor (PXR) and CAR (Francis et al., 2003; Guo et al., 2003; Karpen, 2002; Xie et al., 2001). Bile salts are natural ligands for several nuclear hormone receptors that regulate transcription of genes involved in bile acid homeostasis. Specifically, the nuclear transcription receptor FXR is an essential regulatory component of cholesterol homeostasis, which is activated by bile acids (Forman et al., 1995a). FXR is responsible for transcription of genes coding for bile acid biosynthesis and transport. Activation of FXR, for example, was shown to induce repression of the basolateral NTCP bile uptake transporters and to mediate upregulation of the bile salt export pump (BSEP), resulting in increased uptake of bile by the hepatocyte and increased export of bile into the bile canaliculi (Ananthanarayanan et al., 2001; Sinal et al., 2000). Moreover, FXR induces expression of the multidrug resistancerelated protein 2 (MRP2; ABCC2), controls expression of the MDR3 (ABCB4) gene, encoding for the canalicular phospholipid flippase (Lambert et al., 2003; Liu et al., 2003), and represses key genes involved in bile acid biosynthesis, such as sterol 12α-hydroxylase (CYP8B1) and cholesterol 7 alpha-hydroxylase (CYP7A1) (Repa and Mangelsdorf, 2000). Knockout models have supported the vital activity of these transcription factors for liver health and have suggested that they play a central role in the development of cholestatic liver injury. FXR-null mice, for instance, were found to exhibit a phenotype similar to that of the inherited cholestatic liver disorder Bylers disease, and indeed further investigations revealed that progressive familial intrahepatic cholestasis type 1 is associated with a decreased activity of farnesoid X receptor (Chen et al., 2004a). The activities of both the human FXR and BSEP promoters were found to be reduced in patients diagnosed with Bylers disease. While type 2 of this disease arises from mutations in BSEP (Strautnieks et al., 1998), as mentioned before, progressive familial intrahepatic cholestasis type 1 (PFIC1) is caused by mutation of the gene encoding a putative aminophospholipid transferase that is required for post-translational modifications enabling the nuclear translocation of FXR (Suchy and Ananthanarayanan, 2006). Therefore, the phenotype of Bylers disease has been suggested to arise from an enhanced ileal uptake of bile salts via upregulation of the apical sodiumdependent bile acid transporter and diminished canalicular secretion of bile salts secondary to downregulation of the bile salt excretory pump due to an altered function of FXR (Chen et al., 2004a). FXR activity is closely related to processes involved in the development of cholestasis. Experimental treatment with the synthetic FXR agonist GW4064 revealed the protective effect of FXR activity against cholestatic liver damage in rat models of extrahepatic and intrahepatic cholestasis (Fiorucci et al., 2005; Rizzo et al., 2005). In ANIT-induced toxic injury, treatment with the FXR agonist resulted in significant reductions in serum alanine aminotransferase aspartate aminotransferase and lactate dehydrogenase. In addition, the extent of inflammatory reaction and necrosis was reduced compared to untreated mice (Liu et al., 2003).
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Besides deregulation of bile acid transport, dysfunctional FXR might also be responsible for interruption of the physiological downregulation of bile acid synthesis. Since bile acids are known to repress the bile acid synthesis by a negative feedback mechanism via downregulation of CYP7A1 transcription (the rate-limiting enzyme of bile synthesis) through the FXR-SHP-LRH1 cascade (SHP = short heterodimer partner) (Goodwin et al., 2000; Lu et al., 2000; Sinal et al., 2000). Despite decreased expression of the bile salt export pump in FXR-null mice, which was suggested to be responsible for their increased sensitivity to bile acid-mediated toxicity (Miyata et al., 2005), it was indicated that FXR-null mice were relatively protected from secondary cholestasis in a bile duct ligation (BDL) model (Stedman et al., 2005, 2006). It was reported that FXR-knockout mice displayed reduced serum and liver bile acid concentrations and less extensive liver injury in response to BDL in comparison to wild-type mice. This protective effect was attributed to increased hepatic expression of the multidrug resistance associated protein (mrp) 4, resulting in an increased transport of bile acids into the blood and consecutively in enhanced urinary excretion (Marschall et al., 2006; Stedman et al., 2006). Thus, it has been hypothesized that the pharmacologically antagonising of FXR may provide a therapeutic benefit in obstructive cholestasis (Stedman et al., 2006). Hence dysregulation of FXR may substantially contribute to the processes involved in development of cholestasis due to toxic liver injury. This is particularly interesting, since repression of FXR activity is not related to direct binding to a xenobiotic ligand, such as seen in other nuclear receptors, including CAR or PXR, implicating a secondary mechanism that may affect the concentrations of the endogenous FXR ligands, as for instance secondary bile acids. 9.2.2
Bile Acid Synthesis
Another possible mechanism that might cause elevation of the extracellular bile acids is altered expression of enzymes involved in bile acid synthesis. This, for instance, has been implicated in liver toxicity induced by ANIT, where cholestatic liver injury was found to be associated with upregulation of NTCP and CYP7A1 expression, thereby increasing bile uptake from the blood in addition to enhanced intracellular bile acid production. Several mechanisms, however, may contribute to the morphology of cholestatic injury, and a wide spectrum of various aspects is available to explain the development of cholestatic liver injury. Notably, toxicant-induced cholestasis is associated with bile duct obstruction that eventually will lead to a decrease of the bile flow. In ANIT-induced cholestasis this has been related to an induction of biliary epithelial cell (BEC) hyperplasia in rats, which was assigned to enhanced BEC proliferation and consecutive bile duct obstruction. The BEC proliferation upon ANIT treatment was correlated to BEC necrosis and bile duct obstruction in a time- and dose-dependent way (Kossor et al., 1995). Another animal model in which cholestasis was induced by feeding lithocholic acid (LCA) has demonstrated that increased concentration of toxic bile acids cause bile duct obstruction and destructive cholangitis accompanied by periductal fibrosis (Fickert et al., 2006). Taken together, these findings suggest that bile duct obstruction in toxic cholestasis may result from bile duct proliferation mediated by increased extracellular bile acid
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Hepatotoxicity Toxicant, chemokines
Bile acid import
Bile acid export Bile
Blood Apoptosis
NTCP
Hepatocyte OATP
Bile acid synthesis
Proliferation
BSEP
Bile acids
Cholangiocyte
ABCG
CYP7A1
Figure 9.2 Possible mechanisms of toxic disruption of bile acid homeostasis, mediated via deregulation of bile acid transporters. An example is the transcriptional downregulation of NTCP and OATP along with an increase of bile acid export by upregulation of BSEP or ABCG. Subsequently increased extracellular bile acids trigger metabolic dysfunctions in hepatocytes and cholangiocytes, causing cell death and bile duct obstruction. Reproduced by permission of the American Society for Biochemistry and Molecular Biology from Yamamoto et al. (2006), Journal of Biological Chemistry, 281, 16625–16631
concentrations which result from toxic disruption of bile acid homeostasis in hepatocytes, as depicted in Figure 9.2. It can be concluded that mechanisms causing cholestasis can be directed either through drugs that interfere with the expression of bile transporters via activation of nuclear transcription factors or through extracellular activation triggered by enhanced cytokine release. The resulting effects, however, affect different types of liver cells, such as hepatocytes, cholangiocytes or hepatic stellate cells. Chronic cholestasis of the liver induces onset of fibrotic processes, which sooner or later will lead to the development of liver fibrosis. In this context, besides the severe metabolic derangements found in hepatocytes, notably the influence of bile acids upon other liver cells has become an important field of research. Specifically, the activation of hepatic stellate cells (HSCs) and portal fibroblasts to myofibroblastic cells is an early and major event of chronic cholestatic liver injury (Hines et al., 1993; Tang et al., 1994; Tuchweber et al., 1996). Mechanisms of cholestatic liver injury in summary: – Obstruction of bile flow (proliferation of bile epithelia) – Inhibition of bile salt transporters→impaired secretion into bile canaliculi (direct interaction or transcriptional downregulation) – Inhibition of physiological downregulation of bile acid synthesis (inhibition of negative feedback mechanisms) – Induction of genes in bile acid synthesis (CYP8B1 and CYP7A1)
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Steatosis
The term ‘hepatic steatosis’ refers to an intracellular accumulation of lipid droplets in the cytoplasm. ‘Primary steatosis’ is often observed in patients displaying symptoms of the metabolic syndrome including obesity, diabetes, hypertriglycerinaemia and insulin resistance. Secondary hepatic steatosis is extrinsically induced by alcohol, several drugs, copper accumulation in Wilson’s disease and other factors (Pessayre et al., 2001). Hepatic steatosis may also be accompanied by hepatocellular necrosis, inflammation and fibrosis, in which case it is termed as ‘non-alcoholic steatohepatitis’ (NASH). The morphology of steatosis has been linked to primary mitochondrial failure and has been implicated for microvesicular steatosis, non-alcoholic steatohepatitis (NASH) and cytolytic hepatitis (Fromenty and Pessayre, 1995). This was concluded from observations that these pathologies were associated with early perturbations of the mitochondrial metabolism and associated with apoptosis induced by mitochondrial pathways. Several events have been demonstrated to cause failure of the mitochondrial metabolism, which eventually will end in cellular necrosis and apoptosis (Kim et al., 2003a).The two main mechanisms by which toxicants cause primary mitochondrial failure are uncoupling of the oxidative phosphorylation and initiation of cell death. Different toxicants, however, have their different ways of reaching these endpoints. For example, uncoupling of the respiratory chain can result from direct interference with mitochondrial enzymes or from dysfunctions caused by mutations of the mitochondrial DNA (mtDNA) (both, for instance, observed effects of iron) (Ramm and Ruddell, 2005). Uncoupling of the mitochondrial oxidative phosphorylation is associated with energy depletion and extensive production of reactive oxygen species (ROS), leading to a vicious circle of cellular stress. In fact, the presence of ROS has been associated with the pathology of fatty liver diseases (Yang et al., 2004b). Increases in intracellular ROS have been observed in alcohol-related NASH and steatohepatitis in Wilson’s disease, accounting for enhanced oxidative degeneration of mitochondrial DNA (Mansouri et al., 1997, 1999). A combination of modified mitochondrial DNA and oxidation of mitochondrial proteins and lipids has also been implicated in increased ROS formation in alcoholic patients (Pessayre et al., 2001). Ethanol-induced hepatotoxicity progresses through different phases, beginning with accumulation of lipids and hepatic steatosis, passing via fibrosis and ending in cirrhosis in a worst-case scenario. Mitochondrial toxicity is often associated with ultrastructural lesions, including crystalline inclusions and formation of oversized megamitochondria, as have been observed in livers from NASH patients (Sanyal et al., 2001). Besides impairment of oxidative phosphorylation, especially alterations of fatty acid ß-oxidation have been suggested to significantly contribute to the development of the steatotic phenotype (Fromenty and Pessayre, 1995). Physiologically, fatty acids are metabolized via mitochondrial and peroxisomal ßoxidation. Disturbance of these pathways through, for instance, dysfunction of mitochondrial key enzymes causes accumulation of the non-esterified fatty acids (NEFAs). This has been demonstrated in patients with NASH and ethanol-induced liver disease, where levels of circulating free fatty acids were found to be elevated and were positively correlated with the severity of the disease (Nehra et al., 2001). On the one hand, fatty acids have been demonstrated to exert toxic effects on the cell and were associated with induction of apoptosis by a mitochondrial and a lysosomal pathway (Martins et al., 2006).
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On the other hand, accumulation of fatty acids results in further perturbations of the lipid metabolism and subsequently non-metabolized fatty acids are esterified to triglycerides. Importantly, cholesterol accumulation in the inner mitochondrial membrane due to chronic ethanol treatment is believed to cause a decrease of GSH transporter activity and thereby selectively depletes mitochondrial GSH. This was demonstrated to be important not only for activation of apoptotic cascades, but also for the redox status of the entire cell, since the mitochondrion is also the most significant cellular organelle for production of H2 O2 (Fernandez-Checa and Kaplowitz, 2005). Alterations of the ß-oxidation cause a gradual remodelling of the normal liver, progressing towards fatty liver. If the lipid fraction constitutes more than 5–10 % by weight of the organ, the morphology is referred to as ‘steatosis’. Impairment of the ß-oxidation may result from the obligatory connection of ß-oxidation and oxidative phosphorylation. Here the generation of reduced NAD+ during ß-oxidation is essential for the operation of the mitochondrial respiration (as demonstrated in the diagram below). Diminished co-substrate NAD may result from interference with the mitochondrial respiration and will consecutively lead to a decreased rate of ß-oxidation. β-oxidation NAD+
NADH mitochondrial respiration
Accordingly, perturbations of the mitochondrial ß-oxidation can derive from dysfunction of several mitochondrial enzymes of ß-oxidation or oxidative respiration. This can be linked to mutations of mitochondrial DNA (mtDNA) or defect-transport mechanisms of nuclear proteins required for the mitochondrial function. Recent investigations revealed increased rates of apoptosis in NASH hepatocytes that were associated with activation of NF-κB, which correlated with the severity of the disease (Feldstein et al., 2003a; Ribeiro et al., 2004). Concomitantly, increased expression of death receptors Fas and TNFR1 and activation of caspases 3 and 7 were observed (Feldstein et al., 2003a; Ribeiro et al., 2004). Further investigations revealed that fatty acids induced apoptosis by two different pathways. One is based on translocation of the antiapoptotic protein Bax to the lysosome, which causes lysosomal destabilization, accompanied by release of the lysosomal cysteine protease cathepsin B (Feldstein et al., 2004). The other pathway is initiated at the site of the mitochondrion and causes cell death by activation of JNK and release of cytochrome c (Malhi et al., 2006a). Release of cathepsin B was furthermore associated with enhanced expression of TNF-α, while inhibition of this lysosomal protease was found to be protective against liver toxicity of free fatty acid, which otherwise resulted in development of hepatic steatosis, liver injury, and insulin resistance (Feldstein et al., 2004, 2006). Studies in steatotic livers of wild-type and cathepsin B-knockout (Ctsb(−/−)) mice have supported the protective effect of cathepsin inhibition against stress-induced apoptosis in a model of cold ischemia–warm reperfusion injury (Baskin-Bey et al., 2005). Mitochondrial dysfunction is a general theme in liver toxicology, and several mechanisms triggered by liver toxicants have been identified, which will be discussed later in detail.
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Mechanisms of steatotic liver injury in summary: – Inhibition of β-oxidation→inhibition of mitochondrial respiration – Accumulation of possibly toxic fatty acids→metabolic dysfunction and activation of lysosomal proteases – Induction of mitochondrial dysfunction and ROS→lipid peroxidation
9.4
Hepatitis
Hepatitis is characterized by the presence of inflammatory cells. In a toxicological context, there are at least two important types of hepatitis. Primary hepatitis results directly from damages induced by the toxicant, while secondary hepatitis has a delayed onset and is mediated by secondary events. Primary hepatitis can be induced by direct and acute reaction with the toxicant, while secondary hepatitis is mediated by intermediate events leading to a delayed form of inflammation. 9.4.1
Halothane-Induced Immune Hepatitis and Bioactivation
Drug-induced immune hepatitis is mostly characterized by a delayed onset and has been described for several substances, such as the anaesthetic halothane, the antidiuretic tienilic acid, the antibiotic sulphamethoxazole (Farrell et al., 2003; Gruchalla and Sullivan, 1991), the neuroleptic carbamazepine (Madden et al., 1996; Naisbitt et al., 2003) and the proteinase inhibitor nevirapine (Park et al., 2005). Intriguingly, all of these drugs share one property – they all undergo bioactivation leading to formation of their hepatotoxic metabolites, which are conjugated to intracellular proteins and recognized by immunocompetent cells, thereby initiating an immunologic response in the liver. The resulting immunoallergic hepatitis has been best described in patients treated with halothane. Halothane-induced hepatitis occurs after the second exposure to the drug, usually on the 5th day after the anaesthesia, and is accompanied by general inflammatory symptoms, such as fever, rash, and eosinophilia (Pohl, 1990). The incidence is rare (1 in 35 000 patients) but the mortality associated with the development of an acute insufficiency of the liver is high (up to 50 %) (Bourdi et al., 1996). In patients diagnosed with halothane hepatitis, the presence of unique antibodies against microsomal proteins is characteristic. Among these are, for instance, antibodies targeting the E2 subunit of pyruvate carboxylase, the 2-oxoglutarate dehydrogenase complex (Frey et al., 1995) and native and isoforms of protein disulfide isomerase (Martin et al., 1993) in humans as well as others (Bourdi et al., 1996). Responsible for the modification of cellular proteins (lysine residues in particular) and formation of neoantigens is the main metabolite of halothane, namely trifluoroacetylchloride (Kenna et al., 1988). Formation of antibodies results from processing of trifluoroacetylated proteins and presentation of neoantigens together with class MHC II by Kupffer or B-cells. Subsequent activation of T-helper cells triggers the transformation of B-cells to plasma cells to produce specific antibodies, although activation of T-killer cells leading to cytolytic reactions might be another possible pathway.
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Note that the exact immunologic mechanism leading to the hepatocellular injury by halothane is not completely understood, even though evidence for a major role of the T-cell response has been provided (Lohse et al., 1990, 1992). Intriguingly, in immunoallergic hepatitis induced by halothane, tienilic acid and antihypertensive dihydralazine antibodies against specific isoforms of the cytochrome enzymes that metabolize these particular drugs, have been identified. These antibodies interact with the CYPs in a way that leads to inhibition of their activity – a feature that has been emphasized to contribute to the toxicity observed under administration of these drugs to a major extent (Park et al., 2005). The reactive metabolite of tienilic acid (ticynafen), for instance, binds covalently to the CYP2C9, by which it is generated, and thereby leads to formation of unique antibodies, termed anti-liver/kidney microsome antibodies type 2 (LKM2) (Bonierbale et al., 1999; Lecoeur et al., 1996; Neuberger and Williams, 1989). In dihydralazine-induced hepatitis, antibodies recognizing CYP1A2 (Belloc et al., 1997; Bourdi et al., 1990) have been identified, while in case of halothane extensive generation of autoantibodies reacting with CYP 2E1 has been detected (Bourdi et al., 1996). Mechanism of autoimmune hepatitis in summary: – Formation of auto antibodies (autoimmune hepatitis)→inhibition of metabolising enzymes and inflammatory reaction
9.5
Liver Fibrosis
Liver fibrosis is characterized by an excessive accumulation of connective tissue, leading to a disturbed liver architecture that is associated with severe pathophysiological consequences. Formation of so-called pseudo-septa and regeneration nodules leads to a diversion of the blood flow and impairs supply with nutrients and oxygen. Furthermore, toxic liver fibrosis is accompanied by both bile ductular proliferation and inflammation under various conditions (Muller et al., 1996). The dynamic processes that are involved in the advance of fibrosis are comparable to those activated during wound healing, including differentiation of local immunocompetent cells and increased production of extracellular matrix (ECM). Hepatocyte layers, sinusoids, portal and central vein tracts determine the distinct architecture of the normal liver. In regions exposed to high levels of shear stress, the extracellular matrix in capsule, septal, perivascular and periductal areas is built of strong collagens I, II, V and fibronectin (Schuppan, 1990). This composition of extracellular matrix is called intestinal matrix and is mainly generated by local fibroblasts. In the perisinoidal space of Disse, most of the intercellular communication between liver and blood cells takes place. Here the extracellular matrix, which is mainly generated by stellate cells, is more delicate and resembles the composition of a basement membrane, containing collagen IV, VI, laminin and fibronectin (Arenson et al., 1988; Hahn et al., 1980). The remodelling within the fibrotic process leads to dramatic metabolic derangements as a result of excessive ECM production and deposition of collagens I and II in the space of Disse. The production and composition of several ECM compounds results from a
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balanced equilibrium of regulatory mechanisms. These involve production of fibrogenic cytokines by Kupffer cells and hepatocytes, including interleukins 1 and 6 (IL-1, IL-6), and participation of pro-fibrogenic growth factors, namely fibroblastic growth factors (FGFs), platelet-derived growth factors (PDGFs), and macrophages colony stimulating growth factor (M-CSF), as well as tumour growth factor (TGF-β) (Poli, 2000). Prevalence of one of the opposing mechanisms regeneration and inflammation within the liver decides the fate of functional liver parenchyma to either regenerate or be inescapably lost (Mehendale, 2005). In particular, the interaction between different liver cell types and between cells and interaction with the extracellular matrix have been recognized to account for both activation of fibrotic processes and regeneration.
9.5.1
Cellular Interactions in Fibrosis due to Toxic Liver Injury
The orchestration of signalling between different types of cells is essential for the secretory, synthetic and metabolic function of the liver. More than 15 different cell types are found in the liver (Malarkey et al., 2005). Hepatocytes are the centre of metabolic and synthetic actions and account for 60 % of the liver cells. The sinusoidal endothelial cells (SECs) are the primary barrier between blood and the hepatocytes filtering blood and the space of Disse (Smedsrod, 2004). These types of cells are particularly susceptible to oxidative stress and energy depletion and have been suggested to play a central role in toxic events (Dobbs et al., 1994; McCuskey, 2006). Kupffer cells are derived from monocytic cells and play a role in immune response and inflammation. Their ability to produce cytokines, such as TNF-α, and radical oxygen species has been implicated to play a crucial role in promoting the progression of toxic liver injury (Videla et al., 2003; Vrba and Modriansky, 2002). Stellate cells (Ito cells) produce extracellular matrix, store and metabolize vitamin A and lipids and are involved in processes that lead to regeneration, fibrosis and cirrhosis (Mabuchi et al., 2004). The interplay between the different cell types of the liver is well balanced. Non-parenchymal cells can influence both positively and negatively the proliferation of hepatocytes (Malik et al., 2002). A deregulation of their interaction due to ischemic or toxic stress can cause severe injury. The liver’s susceptibility to toxic stimuli is variable and shows regional differences. It was demonstrated that the architecture of the liver itself has influence on the changes that occur within physiologic as well as pathologic conditions. Morphologic arrangement of portal vein vessels, bile ducts and arteries, which form the functional units of the acini and the Rappaport fields, located between the branches of the central liver veins, form a 3-dimensional structure that seems to influence the metabolic competence of the liver. Indeed, gradients in protein concentrations, enzyme production and oxygen supply contribute to the variable susceptibility of different regions of the liver to stress and nutrient depletion. These factors seem to play a role for the development of liver injury, but also for the mechanisms of regeneration. Varying regional susceptibility has been observed during toxic responses (Heinloth et al., 2004; Irwin et al., 2004), chemical carcinogenesis (Elba et al., 2002; Garcia-Torres et al., 2003), as well as during development and progression of cirrhosis (Matsuzaki et al., 1997).
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In the past decade, Kupffer cells and stellate cells were found to play a key role in inflammatory signalling and progression of fibrosis. Activation of Kupffer cells and stellate cells is believed to account for profibrotic and inflammatory events observed in development of liver fibrosis. Besides, there is evidence for an involvement of reactive oxygen species (ROS) and oxidative stress in several steps of fibrosis. Specifically, ROS were reported to exert stimulatory effects on cytokine generation by Kupffer cells and biosynthesis of extracellular matrix precursors by stellate cells (Pinzani and Marra, 2001; Refik et al., 2004; Thakur et al., 2006). It is assumed that any kind of liver injury triggers a signal cascade in which macrophages release cytokines and thereby recruit stellate cells, portal fibroblasts and others to initiate differentiation into myofibroblasts, characterized by expression of desmin or smooth muscle alpha actin (SMA), eventually resulting in increased ECM production (Knittel et al., 1999). Moreover, hepatic stellate cells (HSCs) have a central role in regeneration processes of the liver. Activated upon liver injury, HSCs display increased proliferation, motility and extracellular matrix (ECM) production (Marra et al., 1999b). This transformation into the myofibroblastic phenotype is a central event during liver fibrosis and has been related to the activity of extracellular signal-regulated kinase (ERK), which is a member of the protein–serine/threonine kinases known as mitogen-activated protein kinases (MAPKs) (Marra et al., 1999). In response to exposure to the platelet-derived growth factor (PDGF) and ECM, the ERK pathway is activated, induces the ability to increase cell migration (Klemke et al., 1997) and upregulates the expression of procollagen genes (Davis et al., 1996; Svegliati-Baroni et al., 1999). Furthermore, activated hepatic stellate cells produce cytokines, such as TGF-β, which induces immigration of inflammatory cells by chemoattraction and is also able to induce apoptosis of hepatocytes by a paracrine mechanism, notably activating apoptosis by binding to TNFR1 and consecutive caspase activation (Gressner et al., 1996). 9.5.2
Stellate Cells Abide Oxidative Stress in Liver Fibrosis
Generation of ROS has been closely associated to enhanced production of extracellular matrix in ethanol-induced fibrosis in stellate cells. Specifically, CYP2E1 that is induced upon ethanol treatment is a potent generator of reactive oxygen species. The superoxide anion resulting from this reaction forms a hydrogen peroxide which leads to formation of the highly reactive 1-hydroxyethyl radical that is probably the main source of oxidative stress in ethanol-induced liver injury. The strongest indicator for extensive oxidative stress in ethanol-induced liver damage is lipid peroxidation. Indeed, end-products of lipid peroxidation, such as 4-hydroxy-2,3nonenal (HNE) or malonaldehyde (MDA), in particular, were demonstrated to induce upregulation of procollagen I synthesis in stellate cells (Greenwel et al., 2000; Parola et al., 1993). The causal relation of lipid peroxidation and fibrogenesis was demonstrated in experiments in which application of the antioxidant vitamin E ameliorated liver toxicity and fibrogenesis (Parola et al., 1992). In addition, the main metabolite of ethanol itself, namely acetaldehyde, is accountable for H2 O2 production in stellate cells. It was demonstrated that hydrogen peroxide directly induces transcription of procollagen α 1 (I) (Greenwel et al., 2000), although ethanol was shown to account for the major fraction of enhanced α 1 (I) procollagen mRNA expression (Fontana et al., 1997).
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Lipid peroxidation
Lipid aldehydes
ROS Redox sensors
Upregulation of ECM production
Liver cell
Inflammation
205
Stellate cells and fibroblasts
Stellate cell
IL 1, 6 growth factors
Figure 9.3 Central role of lipid peroxidation: toxic stimuli result in production of ROS and lipid peroxidation, which activate enhanced ECM production in fibroblasts and stellate cells
Acetaldehyde was also reported to induce upregulation of the α 2 (I) collagen and fibronectin gene expression, which is believed to involve activation of the protein kinase C and down-stream activation of signal transduction pathways, including phosphatidylinositol 3 -kinase (PI3K) and extracellular-regulated kinase (1/2) (ERK1/2) (Svegliati-Baroni et al., 2001). The importance of acetaldehyde as a profibrotic factor may further arise from its ability to form adducts with the carboxyl-terminal propeptides of procollagen in hepatic stellate cells. This has been demonstrated to result in a reduced feedback of collagen synthesis and hence may be responsible for collagen overproduction in ethanol-induced fibrosis (Ma et al., 1997). In conclusion, upregulation of ECM production is a major finding in fibrosis and is triggered in stellate cells via ROS-dependent signalling. The transcriptional activation of ECM precursors can be induced by several mechanisms, including direct stimulation via ROS, production of cytokines and generation of aldehydic products from lipid peroxidation (see Figure 9.3). Even though, the full extent of pathological processes causing fibrosis is not fully understood, induction of TNF-ß1 and activation of the redox-sensitive transcription factor activator protein-1 (AP-1) are most likely events contributing to the progression and inflammatory reaction of fibrosis (Leonarduzzi et al., 2000). Furthermore, the role of the extracellular matrix composition needs to be emphasized as a mechanism by which activation of hepatic stellate cells may contribute to severe metabolic derailment in liver injury and development of fibrosis. Growth and differentiation of hepatocytes is vitally dependent on interaction with the extracellular matrix (ECM). Lack of ECM was demonstrated to induce a loss of hepatocyte differentiation, which was associated with activation of several stress-signalling pathways, including the MAPK, SAPK/JNK and c-Jun signalling pathways, that caused recruitment of the activated protein-1 (AP-1) complex (Otsu et al., 2001; Sidhu et al., 2004). The effects of ECM on the metabolic competence of hepatocytes has been impressingly demonstrated in in vitro experiments, where the effects of adult ECM, fibroblast ECM and foetal ECM were compared in regard to activity of liver-specific enzymes, proteins and transcription factors, such as hepatic nuclear factor-4 (HNF-4). It was shown that adult hepatocytic ECM supports the expression of adult genes, while foetal hepatocytic ECM induced expression of foetal
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genes. In contrast, fibroblastic ECM inhibited hepatocyte proliferation and tissue-specific gene expression in both foetal and adult hepatocytes (Brill et al., 2002). Moreover, the composition of ECM was shown to modulate other essential functions of liver cells. Exposure to increased collagen concentrations within the ECM has been demonstrated to severely influence the sensitivity to harmful stimuli. Thus, it was shown that in hepatocytes grown on a collagen matrix activation of the heat shock response failed to protect against TNF-α-induced apoptosis (Gosnell et al., 2000). Therefore, activation of hepatic stellate cells may in turn disrupt the hepatocyte’s metabolism in two different ways – first by production of inflammatory cytokines and secondly by an enhanced production of extracellular matrix proteins. Furthermore, apoptosis itself has been proposed to have a central role in activation of this profibrotic cascade. This was, for instance, supported by studies in Fas-deficient mice, where extrahepatic cholestasis was induced by performing bile duct ligation. Three weeks after the intervention, the Fas-deficient mice displayed a significantly reduced amounts of actin and collagen as compared to wild-type mice (Canbay et al., 2002). Moreover, engulfment of apoptotic bodies was demonstrated to induce stimulation of Kupffer cellular generation of death ligands Fas and TNF-α, while caspase inhibition abrogated the latter (Canbay et al., 2003). Therefore, deregulation of apoptotic pathways may contribute to promoting liver fibrosis by an impairment of regeneration processes. This was supported by the finding that fibrosis following chronic hepatocytic apoptosis was associated with inactivation of the pro-apoptotic protein bcl-xl (Takehara et al., 2004). It can be concluded that liver fibrosis arises from a ‘potpourri’ of several mechanisms and profibrotic effects, in which the hepatocellular function and regeneration is altered by inflammation, proliferation of cells of the myofibroblastic phenotype and enhanced production of extracellular matrix. Mechanisms of toxic fibrosis in summary: – Activation of stellate cells→induction of chemoattraction and apoptosis and ROS – Generation of ROS→up-regulation of profibrotic genes – Disturbance of extracellular matrix composition→affecting differentiation and cell metabolism – Presence of apoptosis→enhanced production of death receptors
9.6 9.6.1
General Mechanisms and Features of Toxic Liver Injury Nuclear Receptors, Recognition of Ligands and Xenobiotic Response
The cellular response to xenobiotics and toxicants involves several general pathways that lead to adaptive responses, the first step of which is recognition of an external component. There are different alarming systems to accomplish the latter. Even though some substances are recognized and effective by mediation of extracellular receptors, such as ßblockers or K+ -channel inhibitors, most drugs and toxicants enter the cell unrecognized
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and passively by diffusion, giving rise to wanted and unwanted drug effects. In addition, responses to xenobiotics comprise adaptive reactions of the cellular metabolism, such as enhanced rates of biotransformation, drug transport, proliferation and/or regeneration. Within the past decade, an enormous gain in information regarding cellular xenobiotic sensing systems (the so-called xenosensors) has been achieved. An impressive and highly relevant example for cellular response to drugs and toxicants is the induction of CYP450 enzymes, which play a major role in Phase I reactions of biotransformation. The nuclear receptor family of ligand-activated transcription factors has been discovered to be substantially involved in recognition of drugs and toxicants and regulation of CYP enzyme expression. Nuclear receptors (NRs) are regulators of gene transcription and bind to specific DNA response elements implemented in the flanking regions of genes which they control. The human nuclear receptor superfamily consists of four groups: steroid hormone receptors, dimeric orphan receptors, RXR heterodimers and monomeric receptors. Activation of the RXR heterodimers requires the retinoid X receptor (RXR) as a binding partner. The ligands of the orphan receptors are, as yet, unidentified. The family of ligand-activated nuclear receptors is substantially involved in the xenobiotic response and their impact goes far beyond a simple enhancement of biotransformation. In the following, the role of the constitutive androstane receptor (CAR), the aryl hydrocarbon receptor (AHR), the PXR, FXR, vitamin D receptor and the oestrogen receptor for induction of bioactivation and cellular response to xenobiotics in the liver will be elucidated. 9.6.1.1
The Aryl Hydrocarbon Receptor (AHR)
The aryl hydrocarbon receptor (AHR) is a transcription factor that induces expression of human CYP1A1, CYP1A2 and CYP1B1 (Quattrochi et al., 1994; Whitlock, 1999). In addition, the AHR was determined to be responsible for regulation of genes encoding for enzymes of part II of the biotransformation, such as NADPH:quinone oxidoreductase, GST, and UDP-glucuronosyltransferase (UGT) (Mimura and Fujii-Kuriyama, 2003). Furthermore, there is evidence that the AHR plays an important role in cell cycle regulation, apoptosis and in early embryonic development (Ma and Whitlock, 1996; Nebert et al., 2000; Peters and Wiley, 1995). The inactive form of the AHR is located in the cytoplasm, where it occurs in a complex with heat-shock proteins (hsp90) and a small co-chaperon protein termed p23 (Kazlauskas et al., 1999, 2001). Upon ligand binding, the complex dissociates and AHR translocates into the nucleus, where it forms a heterodimer with the AHR nuclear translocator (ARNT), also known as the hypoxia inducible factor (HIF-1ß) (Hankinson et al., 1991). This heterodimer binds to a DNA region called the xenobiotic response element (XRE, also the AHR-mediated aromatic hydrocarbon response element (AHRE) or dioxin response element (DRE)), which is located in close proximity to genes encoding for the cytochrome enzymes and other enzymes of the biotransformation. Furthermore, the AHR–AHRE complex is responsible for transcriptional regulation of the aryl hydrocarbon repressor (AhRR), which inactivates the complex and therefore serves as a negative feedback mechanism (Mimura and Fujii-Kuriyama, 2003). Ligands for AHR are environmental toxins, including polycyclic aromatic hydrocarbons (PAHs), polyhalogenated biphenyls and dioxins, such as the carcinogen 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) (Whitlock, 1999).
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Due to observations made in AHR-knockout mice that happen to be resistant to TCDD toxicity, it is generally accepted that the AHR is responsible for mediation of toxic, carcinogenic and teratogenic effects of its ligands (Brunnberg et al., 2006; Fernandez-Salguero et al., 1996; Gonzalez and Fernandez-Salguero, 1998; Swanson and Bradfield, 1993). The mechanisms by which TCDD toxicity is further advanced are not completely understood. There is evidence, however, that expression of CYP1A1 is a major event within the development of TCDD toxicity. This cytochrome is not only responsible for biotransformation of AHR ligands, but also exerts its biological effects via metabolism of endogenous substrates, such as arachidonic acid, oestrogens and bilirubin. It also significantly participates in the production of reactive oxygen and has effects on K+ and Ca2+ channels (Rifkind, 2006). In addition, elevated levels of the TGFß1 and TGF-ß3, accompanied by increased apoptosis rates and fibrosis, were found in the portal areas of livers from AHRnull mice (Zaher et al., 1998b), suggesting a role for AHR in the regulation of cytokine signalling and inflammation. Moreover, interaction with other nuclear transcription factors may be relevant for AHRmediated toxicity. Specifically, the AHR was shown to interact with nuclear factor kappa B (NF-κB), which is involved in several physiological and pathological responses, including immune modulation, inflammatory responses and apoptosis. It was demonstrated, for example, that TCDD triggers expression of several apoptotic genes, such as Fas ligand (FasL). This regulation is apparently induced by AHR and subsequent NF-κB activation. Studies in AHR knockout mice had revealed that AHR activation was required for regulation of the FasL promoter activity through NF-κB in thymic stromal cells (Camacho et al., 2005). In addition, crosstalk of AHR with the retinoid acid receptor (RAR), vitamin D receptor (VDR) and peroxisome proliferator-activated receptors (PPARs) has been reported (Fallone et al., 2004). Even though the complete complex of AHR-related actions needs to be unravelled, AHR is an excellent example as to how drug interactions with the cellular metabolism can be induced. 9.6.1.2
The Pregnane X Receptor (PXR)
The pregnane X receptor (PXR, also called the steroid and xenobiotic receptor (SXR) or pregnane-activated receptor (PAR)) is a member of the NR1I subfamily of nuclear receptors, which also includes the vitamin D receptor (VDR), constitutive androstane receptor (CAR) and the frog benzoate X receptors (BXRs) (Kliewer et al., 2002). The pregnane X receptor binds to elements in CYP enzyme drug-responsive enhancers and thereby induces enhanced transcription of CYP3A4 (Bertilsson et al., 1998; Blumberg et al., 1998; Lehmann et al., 1998), CYP3A7 (Pascussi et al., 1999), and the two major isoforms of human liver microsomal carboxylesterases (HC1 and HC2) (Zhu et al., 2000). Furthermore, there is evidence suggesting PXR regulation of the genes coding for CYP2C8, 2C9 and CYP3A11 (Pascussi et al., 2000c; Staudinger et al., 2001a). Interestingly, many target genes of PXR are substantially involved in xenobiotic metabolism and bile acid metabolism (see also Table 9.1). Furthermore, PXR has been implicated in the pathogenesis of cholestatic liver injury and the development of drug-induced hepatomegaly due to its regulatory function in bile acid biosynthesis (cholesterol 7α-hydroxylase) as well as its effect on bile acid and xenobiotic-metabolizing (CYP3A) and -transporting (Oatp2) proteins in vivo (Staudinger et al., 2001b).
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Genes regulated by PXR
Feature
Inducer
Biotransformation Rifampicin, phenobarbital (Phase I) (PB), clotrimazole, 5betapregnane-3,20-dione, organochlorine, organophosphate, pyrethroid pesticides Nonylphenola
Hepatic transporters
b
Gene
References
CYP2B6
Lemaire et al., 2004; Xie et al., 2000
CYP2B9
Hernandez et al., 2006 Ferguson et al., 2005 Chen et al., 2004b; Gerbal-Chaloin et al., 2002 Chen et al., 2003; Gerbal-Chaloin et al., 2001 Durr et al., 2000; Pascussi et al., 2001 Pascussi et al., 1999 Falkner et al., 2001
Dexamethasone Rifampicin, hyperforin, phenobarbital
CYP2C8 CYP2C9
Rifampicin
CYP2C19
Dexamethason, hyperforin
CYP3A4
Rifampicin Biotransformation RU38486b (Phase II) Pregnenolone-16 alpha-carbonitrile (PCN) Flavonoids, chrysin, rifampicin
a
209
CYP3A7 GluthathioneS-tranferase Sulfotransferase
PCN
UDPglucoronosyltransferase Carboxylesterases
PCN, spironolactone
Oatp2
PCN, spironolactone 2-Acetylaminofluorene, rifamipicin, PCN, spironolacton
Mrp3 MRP2
Runge-Morris et al., 1999 Madhu and Klaassen, 1991; Sugatani et al., 2004 Hosokawa et al., 1993 Cheng et al., 2005; Staudinger et al., 2001a Maher et al., 2005 Anapolsky et al., 2006; Johnson and Klaassen, 2002; Kast et al., 2002
Environmental oestrogen. Glucocorticoid receptor (GR) antagonist.
Several substances are known to activate PXR, such as steroid hormones and their metabolites, including progesteron, oestrogen and corticosterone, 5ß-pregnane and androsterol, as well as a variety of structurally varying drugs, such as calcium channel inhibitors, statins, antidiabetic drugs, human immunodefiency virus protease inhibitors and anticancer drugs (Blumberg et al., 1998; Drocourt et al., 2001; Dussault et al., 2001; Goodwin et al., 2002; Jones et al., 2000; Kliewer et al., 1998; Kliewer and Willson, 2002; Liddle and Goodwin, 2002; Synold et al., 2001). One of the most potent inhibitors of PXR is hyperforin, the active component of St. John’s wort (Moore et al., 2000; Wentworth et al., 2000).
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In contrast to AHR and CAR, PXR is located exclusively in the nucleus, where it forms a heterodimer with the 9-cis retinoic acid receptors (RXR; NR2B) after ligand activation (Sueyoshi and Negishi, 2001). Notably, PXR is activated by pregnenolone 16 α-carbonitrile (PCN), a glucocorticoid receptor antagonist that induces expression of the CYP3A family of steroid hydroxylases and modulates sterol and bile acid biosynthesis in vivo. Treatment with PCN caused hepatomegaly due to induction of hyperplasia and hypertrophy in wildtype mice. This effect was completely missing in PXR-null mice (Staudinger et al., 2001a), suggesting an involvement of PXR in the development of drug-induced hepatomegaly. Expression of PXR itself was shown to be stimulated by the glucocorticoid receptor (GR) and is repressed during the acute phase reaction by interleukin-6 (Beigneux et al., 2002; Jover et al., 2002; Pascussi et al., 2000c, 2001). Moreover, PXR is known to interact with other nuclear receptors, such as the hepatic nuclear factor 4 α (HNF-4α). This liver-specific transcription factor is essential for the embryonic development and differentiation of the liver (Watt et al., 2003). By binding to the PXR promoter, HNF-4α regulates transcription of PXR and seems to be involved in a general response of CYP450 (Kamiya et al., 2003; Tirona et al., 2003). In addition, it has recently been demonstrated that PXR is activated by different bile acids, such as secondary bile acid lithocholic acid (LCA) and its 3-keto metabolite. Based on this finding, it has been proposed that cytochrome induction via PXR may protect the liver from bile acid toxicity by enhancing degradation through CYP3A4 (Staudinger et al., 2001b). In fact, this was supported by elevated levels of hydroxylated bile acid products of CYP3A4 found in patients suffering from cholestasis (Araya and Wikvall, 1999; Bremmelgaard and Sjovall, 1979). This information may help to explain why treatment with rifampicin and ursodesoxycholic acid had relieved patients from pruritus associated with intrahepatic cholestasis. In this case, these drugs had caused an enhanced clearance of bile acids by therapeutic induction of CYP3A4 (Bachs et al., 1989; Cancado et al., 1998; Gillespie and Vickers, 1993). 9.6.1.3
The Constitutive Androstane Receptor (CAR)
The constitutive androstane receptor (CAR) is located in the cytoplasm and forms a heterodimer by binding with the retinoid X receptor (RXR) upon activation. This nuclear receptor controls genes involved in Phase I (CYP2B, CYP2C, CYP3A) and Phase II (UGT1A1) of biotransformation and transporters for xenobiotics and bile acids (SLC21A6, MRP2) (Pascussi et al., 2003b). Accordingly, CAR is activated by xenobiotics such as phenobarbital and by endogenous compounds such as bilirubin metabolite(s). Activation of CAR is mediated directly by ligands, such as the phenobarbital-like CYP inducer 1,4-bis [2-(3,5-dichloropyridyloxy)]benzene (TCPONOP) (Tzameli et al., 2000). Upon stimulation, CAR undergoes translocation into the nucleus, where its activity is additionally controlled by protein phosphorylation events (Handshin and Meyer, 2003). The transcriptional regulation of CAR itself was shown to be mediated by the glucocorticoid receptor (GR), which acts via the glucocorticoid responsive element (Pascussi et al., 2000b, 2003a). Thus, it was demonstrated that pro-inflammatory cytokines, such as IL-6 or IL1β, by inducing AP-1 or NF-κB activation and thereby leading to GR inhibition, would inhibit CAR gene expression and phenobarbital-mediated CYP gene expression in human hepatocytes (Pascussi et al., 2003b). Taken together, these findings reveal a fundamental
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link between processes that activate the glucocorticoid receptor, such as inflammatory reactions, and the detoxifying capacity of hepatocytes. CAR has been implicated in liver toxicity induced by acetaminophen (APAP). Here, pretreatment with the CYP-inducers ethanol (isoforms of CYP2E and 3A) (Kostrubsky et al., 1997; Sinclair et al., 1998) and phenobarbital (PB) (isoforms of CYP2B and 3A) (Burk et al., 1990; Pirotte, 1984) had increased toxicity of this antipyretic and pain-relieving drug. The influence of CAR, which is known to mediate PB effects, was assessed by application of toxic APAP doses to CAR knockout mice. These mice were found to be resistant against APAP toxicity, which was indicated by the absence of hepatocellular damage or GSH depletion that was found in wild-type mice (Zhang et al., 2002). 9.6.1.4
The Farnesoid X Receptor (FXR)
The farnesoid X receptor (FXR) belongs to the superfamily of RXR (retinoid X receptor) heterodimers, such as CAR, LXR, PXR and the PPARs. The FXR nuclear transcription factor is activated by bile acids and is located in the liver, kidney, adrenal glands and intestine (Forman et al., 1995). Together with the Liver X receptor (LXR), FXR is an essential regulatory component of cholesterol homeostasis, in which bile acids serve as ligands. The role of FXR has been emphasized particularly in processes involved in the development of cholestasis. Abrogation of alterations and restoring FXR activity by synthetic antagonists was demonstrated to be a promising approach to encountering liver toxicity by ANIT (Liu et al., 2003) and may have introduced a new paradigm for the treatment of hepatotoxicity. 9.6.1.5
The Peroxisome Proliferator-Activated Receptors (PPARs)
The peroxisome proliferator-activated receptors (PPARs) were named for their ability to induce hepatic peroxisome proliferation in response to xenobiotic stimuli. The three human PPAR isoforms (PPARalpha, PPARdelta and PPARgamma) are believed to play a central role in regulating carbohydrate and lipid metabolism. In addition, the PPARs are assumed to possess anti-inflammatory activity. These are exerted by inhibiting the induction of pro-inflammatory cytokines, as well as by stimulating the production of anti-inflammatory molecules (Kostadinova et al., 2005). Xenobiotics, fatty acids and eicosanoids are ligands for this nuclear receptor family (Mehendale, 2000). After binding to the retinoid X receptor, PPARs enfold their transcriptional regulation by binding to specific DNA regions, termed peroxisome proliferator response elements (PPREs), which are found in flanking regions upstream of the target gene sequences (Kane et al., 2006). Among the PPAR target genes are carnitine palimitoyl transferase I, HMG coA synthase 2, fatty acid ω-hydroxylase and apoA-1, all of which are key enzymes of the lipid metabolism. In addition, PPARs participate in the regulation of important genes in cell growth, proliferation and tumour cell aberration, such as c-myc, c-Ha-ras, fos, jun and egr-1 (Corton et al., 2000; Vanden Heuvel, 1999). In the response to xenobiotics, PPARs are believed to play a central role: specifically, they are involved in detoxification, survival and regeneration processes that are activated by xenobiotics. For instance, PPARα induces increased expression of several detoxifying enzymes, such as 7β-hydroxysteroid dehydrogenase type 11 and glutathione S-transferase l1 (GST’1) (Motojima, 2004). This has been connected to its ability to metabolize possible toxic plant compounds. For example, it was demonstrated that feeding PPARα knockout
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mice with sesame caused lethal hepatotoxicity, while in wild-type mice sesame strongly induced Cyp2c29, 2c38 and 2b9 in the intestine and liver (Motojima and Hirai, 2006). The human CYPs corresponding to these mouse CYPs, however, which are CYP2C9, 2B6 and 3A4, are regulated by the constitutive androstane receptor (CAR) (Pascussi et al., 2003c). This finding suggests a possible crosstalk and overlapping of transcriptional programs between PPARα and CAR, such as suggested for the axis of PPARa-RXR-LXR and demonstrated for nuclear vitamin D(3) receptor (VDR), which represses the transcriptional activity of PPARα (Anderson et al., 2004; Motojima and Hirai, 2006; Sakuma et al., 2003). Another member of the PPAR family is the lipid-activated transcription factor PPARγ , which was shown to directly regulate the functional expression of drug transporters, such as the ABCG2 transporter. This member of the ATP-binding cassette transporters is known to perform clearance of endogenous and exogenous toxic agents. Activation of PPARγ and consecutively increased amounts of the ABCG2 transporter protein were shown to significantly increase efflux of xenobiotics in human dendritic cells (Szatmari et al., 2006). In the liver, PPARα has been found to be highly expressed (Motojima and Hirai, 2006). In addition to its function within the lipid metabolism, PPARα was suggested to strongly influence the response to xenobiotics. Specifically, this nuclear receptor has been recognized as the primary site of action for fibrates (Kane et al., 2006), which represent a class of lipid-lowering drugs that have been linked to cases of clinically severe rhabdomyolysis occurring under co-administration with HMG-CoA reductase inhibitors (the statins). High LDL-cholesterol and triglyceride levels represent a major risk factor for cardiovascular diseases. For this reason, this drug–drug interaction has been especially unfortunate, since a combination of these two classes of lipid-lowering drugs had promised to expand the efficacy of lipid-lowering therapy to a new level. There is evidence that these interactions might arise from increasing plasma levels of statins, observed under co-administration with fibrates, in particular gemfibrozil (Kyrklund et al., 2003). Note that clofibrate itself had been connected to clinical cases of liver toxicity (Keller et al., 1990, 1992). It has been proposed that peroxisomal proliferators might cause local toxicity due to inhibition of mitochondrial oxygen uptake. Specifically, this has been demonstrated for the peroxisomal proliferators clofibrate, aspirin, valproate, ethylhexanol, ciprofibrate and perfluorooctanoate (Keller et al., 1992). Interestingly, clofibrate administration was also found to exert a protective effect on liver toxicity induced by acetaminophen (APAP) (Nicholls-Grzemski et al., 1992). Initially, investigations revealed that clofibrate increased the hepatic glutathione (GSH) content and led to the suggestion that enhanced detoxification of APAP by covalent binding to GSH may diminish toxic effects of APAP (Nicholls-Grzemski et al., 2000). New evidence, however, implies that an activation at PPARα might be responsible for this protective effect of peroxisome proliferators, such as gemfibrozil (Mehendale, 2000; Nguyen et al., 1999). The importance of PPARα activation for the protection of peroxisome proliferators from liver injury through APAP have been supported by the finding that gemfibrozil administration in PPARα knockout mice did not result in any protection from liver injury (Chen et al., 2000). Several possible mechanisms by which PPARα activation might protect from liver toxicity have been discussed. These include increased protection against oxidative cell damage by increasing antioxidant levels or induction of oxyradical-degrading enzymes, preventing cell – death through induction of resistance or induction of cell-death inhibitors, as well as stimulation of cell proliferation through responsive mitogenic genes
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(Roberts et al., 1998). The latter might facilitate an enhanced recovery and regeneration from early and reversible liver injury (Soni and Mehendale, 1998). Moreover, it was demonstrated that activation of NF-κB, which is important for induction of inflammation, was reduced by activation of PPARα (Huang et al., 2006). The anti-inflammatory effect of PPARα seems to be dependent on suppression of the DNA-binding activity of NF-κB and consecutive downregulation of TNF-α expression (Ye et al., 2004). Thus, overexpression and activation of PPARα could be one of the first steps leading to a survival pathway that is activated by certain xenobiotics. PPARγ has been implicated in liver toxicity mediated by the insulin-senstizing drug troglitazone, which was approved in 1997 for treatment of Type II diabetes. After initial success, this treatment course was observed to be associated with frequent cases of severe hepatotoxicity that finally led to withdrawal of troglitazone from the market in 2000 (Gale, 2001). No significant elevation of transaminases in pre-clinical trials had predicted the toxicity associated with an extraordinarily increased risk of acute liver failure later observed in patients (Graham et al., 2003). Troglitazone and other glitazones have been recognized ligands of PPARγ and were also shown to selectively induce PPARγ in the liver of diabetic rodents. Furthermore, upon treatment with troglitazone, diabetic mice developed severe microvesicular periacinar steatosis (Bedoucha et al., 2001; Boelsterli and Bedoucha, 2002). The therapeutic effect of glitazones and other PPAR agonists, such as the lipid-lowering drug fenofibrate, however, have been clearly related to activity at the PPARs (Harano et al., 2006; Memon et al., 2000). Specifically, PPARγ activates a number of genes that lead to an enhanced uptake of glucose and lipids, increase glucose oxidation and decrease both free fatty acid concentrations and insulin resistance (Dumasia et al., 2005; Way et al., 2001). Troglitazone was demonstrated to induce a coordinated stimulation of fatty acid uptake, oxidation and oxidative phosphorylation in adipose tissue and muscle of diabetic patients (Boden et al., 2005). In the liver, PPARγ activation was shown to coordinately decrease the expression of genes involved in gluconeogenesis and increase levels of genes involved in fatty acid turnover (Kersten, 2002). It has been proposed that the therapeutic effects of PPAR agonists were due to a reversion of a dysfunction in the lipid metabolism found in diabetes and obesity (Cha et al., 2005). Indeed, obese and diabetic mice inherently display upregulation of PPARy in the liver, which was hypothesized to be responsible for an increased susceptibility towards PPARγ -activating drugs, such as troglitazone (Bedoucha et al., 2001; Memon et al., 2000). There is evidence, however, for involvement of several different mechanisms of liver toxicity induced by troglitazone. For instance, biological activation and generation of electrophilic metabolites associated with consecutive GSH depletion were observed upon treatment with this drug (Smith, 2003). The key role of peroxisome proliferator-activated receptors (PPARs) in the regulation of the lipid and glucose metabolisms, however, has motivated to target different metabolic alterations by pharmacologically activating these receptors. Dys- and hyperlipidaemia are common risk factors for cardiovascular diseases and also characterize the metabolic syndrome, which carries a high cardiovascular risk and destines the majority of patients to develop Type 2 diabetes, along with all its related effects and malignant secondary diseases.
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Notably, even though it displays a relatively low intrinsic activity, the pan PPAR receptor agonist bezafibrate significantly reduces triglycerides and improves insulin resistance, thereby reducing the cardiovascular risk in patients (Tenenbaum et al., 2005). The dual PPAR agonists for PPAR α and δ isoforms, the glitazars, have been tested in advanced phases of clinical studies. Targeting of dyslipidaemia via activation of PPAR α and simultaneously improving the insulin resistance via PPAR δ is expected to considerably advance the life expectancy of Type 2 diabetes patients and patients with metabolic syndrome by reducing their cardiovascular risk (Fievet et al., 2006). As revealed in recent studies, however, dual PPAR agonists are related to toxicity and currently unacceptable adverse reactions (Fievet et al., 2006; Nissen et al., 2005). Therefore, further research regarding the exact functions of PPARs in different organs is a necessity to explore and realize the potential of PPAR agonists to provide benefit to this particular patient cohort. In general, it can be concluded that PPARs play a central role in the xenobiotic response of the liver, where they accomplish distinctive regulatory tasks in different cell types. In chronic liver injury induced by carbon tetrachloride, for instance, a downregulation of PPARα and PPARγ expression could be observed in hepatocytes, while increased levels of these transcription factors were found in Kupffer cells associated with inverse correlation to levels of activated NF-κB (Orfila et al., 2005). Furthermore, activation of PPARα has been strongly correlated with the induction of chemically induced carcinogenesis, as for example by di(2-ethylhexyl)phthalate (DEHP) (Corton et al., 2000). The hypothesis put forward states that mitogenic stimulation and suppression of apoptosis in preneoplastic cells due to undue activation of PPARα might contribute to carcinogenic processes (Melnick, 2001). 9.6.1.6
Liver-Enriched Transcription Factors
Regulation of the expression of liver-specific genes has been primarily attributed to the transcriptional regulation that is mediated by interaction of trans-acting liver-enriched transcription factors with cis-regulatory elements of DNA that share liver-specific motifs. These liver-enriched transcription factors (LETFs) are responsible for maintenance of the tissue-specific phenotype, differentiation and are important for liver regeneration in toxic liver injury (Schrem et al., 2002, 2004). Six families of liver-enriched transcription factors have been characterized so far, including the hepatocyte nuclear factors -1 (HNF-1), -3 (HNF-3, now termed FOXA − (forkhead box subclass A) proteins), -4 (HNF-4), -6 (HNF-6), the basic region leucine zipper (bZIP) family members CCAAT/enhancer-binding proteins (C/EBPα, β, δ and γ ) and the D-binding protein. The hepatocyte nuclear factors have been identified to be essential for regulation processes of differentiation and metabolism (Duncan, 1998). HNF-1 is formed by the heterodimer HNF-1 α and the homodimer HNF-1 β and constitutes a major transactivator of the albumin transcription. It mediates regulation of the glucose-6-phosphatase gene transcription by insulin and glucocorticoids (Streeper et al., 1998). Moreover, HNF-1 is believed to be involved in the downregulation of protein expression upon increased oncotic pressure (Pietrangelo and Shafritz, 1994). The HNF-3 or FOXA subfamily consists of three proteins, namely HNF-3 α, β, and γ , now FOXA3, which play a role in developmental differentiation and were found to decrease
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dramatically in acute phase livers (Jacob et al., 1994). Like HNF-3, the HNF-4 subfamily is composed of three different proteins (α, β, γ ) that bind to fatty acyl-CoA thioesters as ligands (Hertz, 1998). Their potential to transactivate liver-specific genes is dependent on phosphorylation of tyrosine residues. The hepatic nuclear factor-6 (HNF-6) is expressed in liver, pancreas and neurones during early stages of differentiation, implicating a central role in the developmental programs of these tissues (Landry et al., 1997). Expression of HNF-6 can be regulated by growth hormone (GH) mediated by transcriptional regulation of the HNF-6 gene by STAT5 (signal transducer and activator of transcription 5) and HNF-4 (Lahuna et al., 1997, 2000; Rastegar et al., 2000). The hepatic nuclear transcription factors were shown to essentially contribute to the metabolic competence that is specific and unique to the liver. Transfection with HNF-4 cDNA, for instance, was demonstrated to induce activation of liver-specific genes in a dedifferentiated rat hepatoma cell line that displayed a loss of function of HNF-1 and HNF-4 (Deschatrette and Weiss, 1974; Faust et al., 1994; Spath and Weiss, 1997). The CCAAT/enhancer binding proteins or C/EBP transcription factors belong to the family of bZip transcription factors, such as c-jun and c-fos (AP-1). The target-specific DNA enhancer elements are characterized by a complex network of interactions with different transcription factors. C/EBPs have regulatory activities in cell cycle control and development and are also involved in the regulation of biotransformation, as e.g. together with HNF-1 α in control of the human cholesterol 7α-hydroxylase CYP7A1 gene (Antes et al., 2000). Depression of nuclear transcription factors has been implicated to represent an important mechanism in toxicant-induced liver injury. For example, reduced expression of HNF-3 γ was observed upon exposure to carbon tetrachloride (Nakamura et al., 1999). Moreover, the liver-enriched transcription factor HNF-3 β and the C/EBP family are modulated by cytokines. IL-1, IL-6 and IFN-γ were found to exert different effects on the LETFs (Samadani et al., 1995). This is a mechanism that could explain the metabolic response of the liver to inflammatory reaction, such as the protein synthesis switch with pronounced synthesis of acute phase proteins and decreased albumin expression observed during the acute phase response of the liver (Whalen et al., 2004). Indeed, TNF-α treatment in primary mouse hepatocytes as well as TNF-α overexpression was found to induce translocation of C/EBP-β by phosphorylation, which resulted in abrogated transcription of the albumin gene (Buck et al., 2001a). A lack of TNF-α signalling via TNFR-1 in carbon tetrachloride toxicity, as investigated in transgenic TNFR-1-knockout mice, was associated with a decreased DNA binding activity of NF-κB and STAT3, reduced plasma levels of (IL)-6 and liver IL-6 mRNA and significantly reduced DNA binding activity of C/EBP, resulting in a reduced regeneration ability of the liver in these TNFR-1-knockout mice as compared to the wild-type controls (Yamada and Fausto, 1998). Furthermore, activity of C/EBP-β was found to be responsible for the stellate cell proliferation observed in response to administration of the hepatotoxic substance carbon tetrachloride. This was supported by the finding that transgenic animal models presenting a lack in C/EBP-β activity ((C/EBP-β −/−) knockout and nonphosphorylatable mutants) displayed stellate cell apoptosis instead of proliferation upon carbon tetrachloride administration.
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The antiapoptotic effect of C/EBP-β activity was furthermore assigned to inhibition of procaspases 1 and 8 (Buck et al., 2001b). A more recent report in which C/EBP-β transfection of cultured hepatocytes resulted in increased DNA synthesis rates and prolonged cell viability in culture had emphasized the role of the C/EBP-β activity for hepatocellular survival and liver cell regeneration in toxic liver injury. This was documented, in addition, by introduction of the C/EBP-β gene into liver-damaged mice, which hitherto had displayed significantly suppressed serum AST and ALT activities (Isoda et al., 2005). Taken these findings together, it can be concluded that activation of liver-enriched nuclear transcription factors by inflammatory cytokines and probably also by suitable liver toxins may be an important mechanism in liver cell defence and survival in toxic injury. It should be noted that an inflammatory environment is characterized by increased generation of reactive nitrogen and oxygen species and thereby exerts enhanced cellular stress and involves a higher risk of mutagenesis (Iwai et al., 2002). Hence, activation of C/EBP-β and its antiapoptotic effect seem to be predestined to contribute to CCl4 -induced carcinogenesis as well. The transcription factor C/EBP-ξ (also known as GADD153 or CHOP-10) acts as a regulatory control of the C/EBPs. It encodes a C/EBP protein without a functional DNA binding domain which, by dimerization with other C/EBPs, is able to deactivate their DNA-binding activity (Carlson et al., 1993). C/EBP-ξ is induced by mediation of AP-1 and p38 mitogen activated kinase (MAPkinase) upon exposure to DNA-damaging agents, such as methyl methanesulfonate, hydrogen peroxide and UV irradiation (Guyton et al., 1996; Luethy et al., 1990; Wang and Ron, 1996). 9.6.1.7
Vitamin D Receptor
Even though the vitamin D receptor (VDR) is found only rarely in the liver under physiological conditions, effects on liver physiology and drug metabolism exerted by vitamin D3 have been frequently observed. Specifically, VDR has been discovered to be an important sensor of bile acids and is able to prevent toxicity by regulating bile acid metabolism (Makishima et al., 2002). VDR is activated by the hepatotoxic secondary bile acid lithocholic acid (LCA) which is also known to be a potential carcinogen (Nagengast et al., 1995; Ogawa et al., 1998). In association with RXR, the VDR–RXR heterodimer interacts with a response element of the human major cytochrome enzyme CYP3A4 gene. This leads to transcriptional upregulation, enabling CYP3A4 to catabolize LCA (Araya and Wikvall, 1999; Xie et al., 2001). Under this aspect, VDR resembles the function of the evolutionarily related nuclear receptors PXR, FXR and LXR, which are also representatives of the cellular bile acid sensors. In addition, VDR has been proposed to be an active member of the nuclear receptor crosstalk with constitutive androstane receptor and peroxisome proliferator-activated receptor α (Makinen et al., 2002; Sakuma et al., 2003). Moreover, the metabolisms of phenobarbital and vitamin D3 were long known to be connected in some way, indicating other significant functions of VDR for the xenobiotic metabolism. This has been discovered through observations made on the protective effect of phenobarbital against intoxication with vitamin D3 (Gascon-Barre and Cote, 1978).
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As of today, it has been demonstrated that normal nonparenchymal liver cells, such as sinusoidal endothelial, Kupffer and stellate cells, very well express significant amounts of VDR (Gascon-Barre et al., 2003). Nuclear receptors (NR) in liver toxicity in summary: – – – – –
Induction and repression (directly, secondary, e.g. due to inflammation) Induction of apoptosis Disturbance of cholesterol and bile acid homeostasis Reduced activity – reduced biotransformation and accumulation of possible toxins Interference with NR or NR crosstalk in vital cellular functions→reduced metabolic competence
9.6.2
Biotransformation and CYP Induction
Inhibition or induction of enzymes involved in detoxification have a high impact on drug effects and the toxicity of certain substances. Biologic detoxification is located in the liver and is responsible for inactivation and successful elimination of endogenous and exogenous metabolites and toxicants. The metabolic reactions responsible for detoxification comprise Phase I and Phase II reactions. Phase I reactions increase the polarity of the xenobiotics by inserting new functional groups on the drug molecule. Major enzymes performing Phase I reactions are hydrolases and oxidoreductases, such as the cytochrome P450-dependent monooxygenases (CYPs) or the cyclooxygenases. Conjugation to endogenous hydrophilic molecules in Phase II reactions result in potent increases in polarity and water solubility, thereby achieving proper drug elimination. Such reactions are glucuronidation, sulfatation and conjugation to amino acids mediated by enzymes of Phase II of the biotransformation, such as UDP-glucose dehydrogenase (UGDH) and members of the UDP glucuronosyl transferase (UGT) family. Phase III describes transport processes mediated by transmembranous transporter proteins, such as the organic anion transporting polypeptide family (Oatps) along with the organic cation transporter 1 (Oct1) and organic anion transporter 2, which mediate uptake of a large number of xenobiotics from blood into liver, and the multidrug resistance proteins (MDRs), including p-glycoprotein (P-gp) or the multidrug resistance-associated protein (MRP), which mediate the efflux of xenobiotics from the liver into bile or blood (Haimeur et al., 2004; Smith et al., 2005; Wakabayashi et al., 2006). There are major ways in which idiosyncratic toxicity arises from interaction at metabolizing enzymes: one possibility is enzyme inhibition while the other one is enzyme induction. Drug–drug interactions at the site of biotransformation I enzymes are commonly recognized to cause severe clinical complications, such as the ventricular arrhythmia observed upon co-administration of CYP3A4 inhibitors, including terfenadine, astemizole, cisapride or pimozide. Another severe complication is rhabdomyolysis, which has been associated with the co-administration of 3-hydroxy-3-methylglutaryl-coenzyme A reductase inhibitors (‘statins’) and CYP3A4 inhibitors (Dresser et al., 2000). The toxicity of these interactions can be attributed to increasing concentrations of the drug or its activated metabolites due
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to inhibition of their metabolizing enzymes. On the contrary, upregulation of metabolizing enzymes has been associated with organ toxicity, in particular affecting the liver. Enhanced transcription and expression of detoxifying enzymes in the presence of suitable substrates can be considered as a natural adaptive response. Thus, increased substrate supply provokes an increased metabolism rate and results in an enhanced clearance of the particular substance. In regard to CYP monooxygenase reactions, five different types of xenobiotic inducers can be distinguished, named after drugs that induce cytochrome monooxygenases of a typical pattern. These are the phenobarbital-type-inducing CYP2B1, CYP2Cs and CYP3As, the rifampicin-dexamethasone type (CYP3As, CYP2Cs, CYP2B1), the ethanol, isoniazide type (CYP2E1), the clofibrate type (CYP4As) and the polycyclic aromatic hydrocarbons (TCDD) type (CYP1A1, CYP1A2, CYP1B1) (Handschin and Meyer, 2003). Clinically important are, in particular, the inhibitors of cytochrome CYP3A4, such as the fungicides itraconazole and ketoconazole, the antibiotics clarithromycin and erythromycin, the 5HT2A serotonin receptor antagonist and antidepressive drug nefazodone, the proteinase inhibitor ritonavir and grapefruit juice (Dresser et al., 2000). The transcriptional regulation and hence induction of detoxifying Phase I, Phase II and Phase III enzymes has been attributed to the activation of ligand-activated nuclear receptors (Klaassen and Slitt, 2005; Xie et al., 2004). Regulation of the different isoforms of cytochrome-dependent monoxygenases, for example, has been attributed to activation of the nuclear aryl hydrocarbon receptor (AHR: CYP1A1, CYP1A2, CYP1B1) (Quattrochi et al., 1994; Tang et al., 1999; Whitlock, 1999), the pregnane X receptor (PXR: CYP3A4) (Bertilsson et al., 1998; Blumberg et al., 1998; Lehmann et al., 1998), CYP3A7 (Pascussi et al., 1999), the constitutive androstane receptor (CAR: CYP2B6) (Sueyoshi et al., 1999), CYP3A4 (Savkur et al., 2003), CYP2C8 (Ferguson et al., 2005), 2C9 (Al Dosari et al., 2006) and the glucocorticoid receptor (GR: CYP3A5) (Schuetz et al., 1996). Recently, the nuclear PPARα receptor has been found to be involved in the negative regulation of normal and rifampicin-mediated UGDH gene expression (Vatsyayan et al., 2005). UDP-glucose dehydrogenase (UGDH) (EC1.1.1.22) catalyses the NADPH-dependent oxidation of UDP–glucose to UDP-glucuronic acid and is therefore essential for Phase II reactions of the biotransformation. Overexpression of Phase III enzymes, a common resistance strategy found in cancer cells, has recently been demonstrated to be part of the response to chemicals that also increase expression of Phases I and II biotransformation enzymes in the liver, resulting in an enhanced hepatic uptake and biliary excretion due to increased expression of various transporters, e.g. Oatps, MRPs and MDRs via activation of nuclear receptors (Klasssen and Slitt, 2005). The nuclear receptor CAR, for example, was demonstrated to be involved in the regulation of multidrug resistance transporter type 3 (MDR3) (Cherrington et al., 2002), while expression of MDR1 (ABCB1), MRP1 (ABCC1), MRP2 (ABCC2) and the breast cancer resistance protein BCRP (ABCG2) was positively correlated with pregnane x receptor activity (Albermann et al., 2005). Even though the central role of the nuclear receptor superfamily in transcriptional regulation of genes coding for detoxifying enzymes has become apparent, the way in which this response is induced by xenobiotics is not sufficiently elucidated. While the pregnane X receptor (PXR) and constitutive androstane receptor (CAR) have been demonstrated to directly interact with their xenobiotic ligands, other nuclear receptors strongly involved in the induction of detoxifying enzymes are activated by ligands of endogenous origin, such
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as fatty acids and eicanosoids serving as ligands for the peroxisome proliferator-activated receptors (PPARs). In addition, there is evidence for an overlap of transcriptional programmes of different nuclear receptors, such as between PPARα and CAR, LXR or the nuclear vitamin D(3) receptor (VDR) (Anderson et al., 2004; Motojima and Hirai, 2006; Sakuma et al., 2003). The current understanding of the transcriptional regulation of enzymes involved in biotransformation is based on a concept of transcription factor networks that regulate basal and xenobiotic-modulated expression (Pascussi et al., 2003c). This crosstalk between nuclear receptors has been demonstrated for several members of these ligand-activated transcription factors, e.g. for PPARα, RXR and LXR (Anderson et al., 2004). 9.6.2.1
How Does Upregulation of Metabolizing Enzymes Cause Liver Toxicity?
Two major hypotheses have been put forward to explain the mechanisms by which upregulation of detoxifying enzymes can cause hepatotoxicity. The first hypothesis involves generation of bioactive metabolites by certain detoxifying enzymes, such as observed in the oxidative group transfer mediated by cytochrome P450dependent monooxygenases to the narcotic halothane. In this case, activation to the active metabolite trifluoroacetylchloride can cause severe immune hepatitis. Another important example described later in detail is the activation of acetaminophen to the hepatotoxic metabolite N-acetylbenzoquinoneimine (NAPQI) by dehydrogenation of cytochrome P450dependent monooxygenases. In addition, biotransformation can lead to formation of toxic species, such as, for instance, observed in dechlorination of chloroform to phosgene. This toxic metabolite forms a specific adduct with a mitochondrial phospholipids (Di Consiglio et al., 2001) that has been proposed to cause a progressing mitochondrial damage with swelling of the organelles and formation of megamitochondria (Guastadisegni et al., 1999), presumed to induce cell death by activation of apoptosis (Hartig et al., 2005). The second hypothesis is based on the undue generation of reactive oxygen species during biotransformation. This, for instance, takes place during the one-electron quinone reduction via flavoprotein P450-reductase with production of superoxide ions, as is involved in the metabolic activation of paraquat or doxorubicin (Kukielka and Cederbaum, 1990; Ravi and Das, 2004). In addition, dehalogenation can exert reactive radicals that cause toxic reactions, as is observed during biotransformation of carbon tetrachloride formation (Connor et al., 1986). 9.6.2.2
Production of Toxic Metabolites in Hepatotoxicity
An important example of liver toxicity induced by an active metabolite generated in the biotransformation process is acetaminophen. Acetaminophen (APAP) is an analgetic and antipyretic drug that is ubiquitously administered. Overdosing of acetaminophen causes severe hepatotoxicity that is associated with centrilobular necrosis (James et al., 2003a). The active metabolite of APAP is N-acetyl-p-benzoquinone imine (NAPQI), generated in step I of the biotransformation by CYP2E1, CYP1A2 and CYP3A4 (Raucy et al., 1989; Thummel et al., 1993). This bioactive metabolite can interact at different levels and sites of the cellular metabolism and has been associated with affection of transcription factors and gene expression, protein expression and activity.
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The presence of NAPQI was shown to be substantially involved in ubiquitous inhibition of several cellular enzymes, some of which are essential for physiologic metabolic processes. Among these is, for instance, glyceraldehyde-3-phosphate dehydrogenase (GAPDH), which transforms the NAD+ -dependent phosphorylation of glyceraldehyde-3-phosphate to 1,3 bisphoglycerate within glycolysis (Andersson et al., 1990; Burcham and Harman, 1991), or γ -glutamylcysteinyl synthetase, which is the rate-limiting step of glutathione (GSH) synthesis, essential for detoxification of oxidative radicals and toxins such as APAP (Kitteringham et al., 2000). Moreover, NAPQI has been demonstrated to be an effective inducer of cytochrome enzymes, e.g. CYP2E1, which has been associated with increased production of reactive oxygen species (ROS) such as superoxide radicals and H2 O2 (Johansson et al., 1988; Nordmann et al., 1992; Rashba-Step et al., 1993; Wu and Cederbaum, 1994). ROS can activate caspases by direct stimulation of the mitochondrial membrane transition pore and cytochrome c release and also controls the stability of p53, which acts in surveillance of cell integrity (Jaeschke et al., 2003). In addition, oxidative stress was shown to be associated with degradation of actin filaments and dysregulation of Ca2+ homeostasis, which is observed early in APAP-induced hepatotoxicity (Tsokos-Kuhn et al., 1988). Accordingly, it has been proposed that the hepatotoxicity of NAPQI is at least in part due to production of oxidative stress resulting from a futile cycling of P450 leading to reduction of molecular oxygen (Goeptar et al., 1995). ROS is believed to be harmful in particular to APAP-treated hepatocytes, since APAP and its major metabolite NAPQI are both detoxified by conjugation with glutathione (GSH), resulting in depletion of GSH that is normally able to dispose of the oxygen radical metabolites. The connection between induction of cytochrome enzymes by NAPQI and hepatotoxicity of APAP had been confirmed by studies with knockout mice. Specifically, CYP450 1A2and 2E1-deficient mice showed a relative resistance to APAP-induced toxicity compared to wild-type mice. In these mice, a higher dose of APAP was needed to induce hepatotoxicity and an increased overall survival has been observed (Zaher et al., 1998a). The absence of the APAP-metabolizing enzymes P450 1A2 and 3A11 (corresponding to human 2A4) in mice due to a deficiency of the nuclear receptor CAR (constitutive androstane receptor) also resulted in resistance to APAP toxicity (Zhang et al., 2002). It can be presumed that the presence of the bioactive metabolite NAPQI is strongly associated with the striking features observed in APAP-induced hepatotoxicity. For example, formation of NAPQI is involved in the depletion of glutathione (GSH), formation of protein adducts and free oxygen radicals (Gibson et al., 1996; James et al., 2003a), which altogether, to an unknown weighting, contribute to the liver injury observed under overdosage of acetaminophen. The resulting metabolic derangements involve in any case the initiation of an immediate hepatocellular defence response that includes translocation of redox-sensitive transcription factors, such as Nrf-2 and NF-κB. These mediate the orchestration of the adaptive cellular response by enhanced transcription of genes encoding for antioxidant proteins and Phase II drug-metabolizing enzymes (Park et al., 2005). As of today, the mechanism of APAP-induced hepatotoxicity is certainly not completely understood. In particular, the role of the interaction between hepatocytes and other liver cell populations, such as Kupffer cells, has been addressed only in part by now. Evidence is available that the consecutive release of proinflammtory cytokines, such as TNF-α and IL-1 alpha, is associated with certain pathological manifestations of APAP-induced
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hepatotoxicity, but studies with TNF-α-knockout mice are inconclusive, e.g. mice display a liver phenotype similar to wild-type mice treated with APAP, but also exhibit exaggerated hepatotoxicity, delayed recovery of GSH levels and less increase in Akt expression, a downstream target of PI-3K, compared to wild-type mice in response to acetaminophen (Boess et al., 1998; Chiu et al., 2003a,b). Thus, it has been concluded that TNF-α release by macrophages has no central role in acetaminophen-induced liver injury but may be responsible for activation of protective signalling pathways in hepatocytes. The antituberculostatic drug isoniazid (INH) has long been known to cause a spectrum of liver injuries, starting from an asymptomatic elevation of liver transaminases and reaching up to hepatitis or massive necrosis associated with liver failure (Mitchell et al., 1976). Bioactivation of INH and its acetyl radical is assumed to be a significant determining factor for the toxicity caused by this drug. Generation of this reactive electrophile metabolite occurs by acetylation and subsequent hydrolysation (Nelson et al., 1976). In addition, it was demonstrated that isoniazid significantly inhibited CYP2C19 and CYP3A (Desta et al., 2001). The induction of CYP2E1 observed for hydrazine, which is the major metabolite of INH, was suggested to be strongly involved in the hepatotoxicity caused by INH (Yue et al., 2004). Formation of adducts with macromolecules has been put forward as one explanation for the mechanism leading to liver injury induced by isoniazid. The previously discussed involvement of CYP2E1 and enhanced production of oxidative stress upon isoniazid treatment, however, may provide another explanation that is worth being investigated further (Attri et al., 2000; Sodhi et al., 1997). The anti-oestrogen tamoxifen is used in the second-line therapy of breast cancer. During its biotransformation, tamoxifen undergoes α-hydroxylation and sulfonation. The resulting sulfate ester is instable and breaks down to form a genotoxic metabolite, a reactive carbocation. The subsequent formation of DNA adducts in the liver is associated with an increased risk to develop liver cancer. Interestingly, this risk is excessive in rats when treated with tamoxifen, as in rats bioactivation through sulfonation is a major route as opposed to the human liver, where glucuronylation is preferential. Indeed, the glucuronide of αhydroxytamoxifen was found to have a much higher stability compared to the sulfonide (Park et al., 2005). Other clinically relevant cases in which bioactivation leads to formation of hepatotoxic metabolites have been described before, as in the case of halothane-induced immune hepatitis, where formation of antibodies causes inhibition of drug-metabolizing monooxygenases. 9.6.2.3
Knockout Models and Biotransformation in Liver Toxicity
Drug-metabolizing enzymes, in particular cytochrome monooxygenases, show a high rate of polymorphisms associated with marked differences in expression and catalytic activity throughout the population. This is of special interest, since bioactivation of hepatotoxic substances may play an important role in idiosyncratic liver toxicity. Trying to provide evidence of the role of drug-metabolizing enzymes in liver toxicity, the application of knockout and transgenic animal models has exceedingly contributed to the current knowledge of the functional spectrum of CYP450s and Phase II metabolizing enzymes in the physiology and development of toxic liver injury (Gonzalez, 2002). For the interpretation of data derived from experimental studies with knockout or transgenic animals, it is of considerable importance that knockout animals derived in different laboratories display important heterogeneity and that furthermore unintended
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gene compensation may occur during the generation process. For example, generation of CYP1A2-null mice in one laboratory resulted in generation of a mouse line in which the majority of nonviable animals had died after birth displaying an immature respiratory system, while breeding the surviving animals resulted in healthy descendants, suggesting involvement of gene compensation (Liang et al., 1996; Pineau et al., 1995). Table 9.2 summarizes the recent biotransformation knockout models in liver toxicty (see Appendix). 9.6.3
Hepatotoxicity at the Organelle Level
Several toxicants have been reported to cause primary damage of certain cellular organelles and thereby develop their toxic effects. The mitochondrion is a common target of toxicity due to its proximity to ROS production at the oxidative phosphorylation complex. Mitochondrial dysfunction has been observed in APAP- and ethanol-related hepatotoxicity as well as in toxicity induced by antiretroviral nucleoside reverse transcriptase inhibitors (NRTIs) and fatty acid oxidation inhibitors (Szewczyk and Wojtczak, 2002; Vickers et al., 2006; White, 2001). Direct toxic effects on cell organelles, however, may significantly contribute to the specific phenotype and pathology of toxic liver injury. Next to mitochondria, peroxisomes and lysosomes have been identified as primary targets of liver toxicity. Thus, destabilization of lysosomes by fatty acids may contribute to the pathology of steatosis and NASH (Feldstein et al., 2004). Release of lysosomal hydrolases, in particular of cathepsin B, causes severe cellular damage and induces activation of apoptosis. The underlying mechanisms of lysosomal permeability due to fatty acid exposure result from translocation of the pro-apoptotic Bcl-2 family member Bax to the lysosome. In addition, the Bax antagonist Bcl-XL was reported to be suppressed in response to palmitate treatment of hepatocytes. This was confirmed by investigations in which inhibition of Bax activity diminished lysosomal permeability (Feldstein et al., 2006). 9.6.3.1
Phospholipidosis
Drug-induced phospholipidosis (DIPL) is characterized by intracellular accumulation of phospholipids and appearance of so-called lamellar bodies (also: myeloid bodies, dense bodies). This condition can occur in several organs and is induced primarily by cationic amphiphilic drugs. In the liver, DIPL has been associated with fibrotic and inflammatory changes. The assumed target of drugs inducing phospholipidosis is the lysosome. Basically, two hypotheses have been put forward to explain the underlying mechanisms of drug-induced phospholipidosis (Reasor and Kacew, 2001). The first hypothesis assumes that binding of CADs to phospholipids results in indigestible drug–lipid complexes which accumulate and are stored in the form of lysosomal lamellar bodies (Halliwell, 1997), while the second hypothesis is based on the observation that production of lamellar bodies was associated with inhibition of phospholipase activity (Reasor and Kacew, 2001). Furthermore, other lysosomal enzyme activities have been modulated by amphiphilic drugs, notably the activity of cathepsin B protease (Gerbaux et al., 1996). Although some of the drugs that cause DIPL, such as amiodarone and propanolol, also cause hepatotoxicity, it is still unclear whether this is a result of independent adverse effects or due to the phenotype, which has been termed ‘phospholipidosis’.
Acetaminophen
CYP1A2 (−/−)
Flutamide
TCDD, 4-PeCDF, PCB 153
4-Aminobiphenyl
– increased drug plasma levels and alternative metabolite in urine of KO – nutritional GSH depletion and repetitive drug administration caused increased aminotransferase activity and moderate hepatitis in KO not WT→potential toxicity of alternative metabolite
(Continued)
Matsuzaki et al., 2006
Diliberto et al., 1997
Smialowicz et al., 2004 Kimura et al., 1999
Smith et al., 2001
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TCDD
Sinclair et al., 2000 Sinclair et al., 1998; Sinclair et al., 2000 Peters et al., 1999
Tonge et al., 1998
Genter et al., 1998
References
October 8, 2007
– higher mortality in KO mice (less metabolite – more parent substance) – pronounced hepato- and splenomegaly in KO mice – less hepatocellular damage or uroporphyria in KO→KO mice protected from uroporphyria and hepatic injury – KO mice were not resistant to TCDD-induced immune suppression – small differences in malignant foci and involvement of eosinophils – similar extent of ROS stress in KO and WT – no sequestration of TCDD and 4-PeCDF in the liver in KO mice
– less extensive liver injury and kidney damage in KO – similar olfactory mucosal damages in KO and WT – doses of 250 mg/kg lethal only in KO and HT – no difference in liver enzyme and metabolite excretion between WT and KO – no significant differences in liver non-protein thiol concentrations – no differences in DNA adduct formation between KO and WT→no significant role in APAP hepatotoxicity for CYP1A2 – increased resistance in CYP1A2 and CYP2E1 KO – no hepatic uroporphyrin accumulation in KO
Resultsa
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Hexachlorobenzene and iron Phenacetin
Toxicanta
Transgenic model
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Table 9.2 Transgenic animal models of hepatotoxicity
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Ethanol
Acetone
Chloroform Acrylonitrile, acrylamide
Carbon tetrachloride
– higher doses needed to induce toxicity in KO – reduced metabolite excretion in KO mice – KO were more resistant to benzene-induced cyto- and genotoxicity,→benzene metabolism of the liver 95 % dependent on CYP2E1 – no abnormality in liver histology found in KO→KO highly resistant to CCl4 toxicity – protection from toxicity in the liver, kidney and nose in KO – WT excrete epoxides and GSH conjugation metabolites from acrylonitrile (AN) and acrylamide (AM) – urine after cytochrome inhibition and in KO displayed only GSH/AM or AN conjugates→CYP2E1 possibly the only cytochrome P450 involved in AM and AN metabolism – blood acetone levels similar in KO (null) and WT – fasting revealed differences: significantly increased acetone levels in KO compared to WT – both WT and KO showed similar signs of toxicity due to ethanol – no difference in fatty acid metabolism between KO and WT mice (fed choline- and methionine-deficient diet)
Kono et al., 1999 Leclercq et al., 2000
Bondoc et al., 1999
Constan et al., 1999 Sumner et al., 1999
Lee et al., 1996 Sinclair et al., 2000 Valentine et al., 1996 Powley and Carlson, 2001 Wong et al., 1998
Uno et al., 2001, 2004a Uno et al., 2004b
References
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Acetaminophen Benzene
– higher sensitivity of KO compared with WT and HT, with dramatically higher BaP–DNA adduct levels in KO – KO mice were protected from TCDD toxicity (displayed less hepatocyte hypertrophy and less intrahepatocytic lipid and total liver fat accumulation, no uroporphyria, showed reduced lethality and no wasting syndrome such as seen in WT) – thymic atrophy, decreased spleen size and leukocytopenia occurred in both genotypes
Resultsa
October 8, 2007
CYP2E1 (−/−)
Benzo[a]pyrene
CYP1A1 (−/−) TCDD
Toxicanta
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Table 9.2 (Continued)
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NQO1 (−/−)
Ahr (−/−)
Cyp1A2/1B1(−/−)
Menadione
Benzo[a]pyrene
TCDD
– KO more sensitive than HT and WT
(Continued)
Radjendirane et al., 1998
Shimizu et al., 2000
Mimura et al., 1997; Peters et al., 1999
Uno, 2004a
Zaher et al., 1998
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– highly resistant DKO, less GSH depletion and APAP adducts compared to WT – Cyp1a1(−/−) knockout mice are protected, show prolonged survival – higher BaP-DNA adduct levels in Cyp1a1(−/−) than in Cyp1a1(+/−) mice – no induction of CYP1A1 in Ahr–KO mice – resistance against TCDD-induced liver injury in KO (10-fold higher concentrations than WT liver injury dose)→role of CYP1A1 induction – reduced embryotoxicity – reduced teratogenicity in Ahr–KO – no subcutaneous tumours in KO but in WT and HZ
Heidel et al., 1999
Ghanayem et al., 2005
References
October 8, 2007
Benzo[a]pyrene
Acetaminophen
CYP1A2 and 2E1 (−/−) Cyp1A1/1B1(−/−)
– in 6 months, 70 % lymphomas in WT vs. only 7.5 % in KO – generally, increased number of tumours in WT not KO→role of CYP1B1 in DMBA carcinogenesis (bioactivation to DMBA-3,4-diol) – CYP1B1 in bone marrow stromal cells→required for DMBA-induced pre-B-cell apoptosis (role in lymphoblastoma formation)
– highly significant adduct formation between acrylamide–GSH product glycidamide and DNA as well as haemoglobin in KO mice—important role for CYP2E1 in adduct formation/toxicity in acrylamide-induced injury – KO resistant to DMBA toxicity
Acrylamide
Dimethylbenz[a] anthracene
Resultsa
Toxicanta
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CYP1B1 (−/−)
Transgenic model
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Galactosamine/ endotoxin
SOD1 (−/−) and SOD1/GPX1 double (−/−)
Glutathione peroxidase-1 (Gpx1) (−/−)
– higher susceptibility to oxidant stress induced by neutrophils and liver injury in KO than in WT (induced significantly more genes coding for inflammatory response, oxidative stress, growth arrest and responses to DNA damage and/or its repair in KO) – both KO and WT developed apoptosis – increased plasma aminotransferase activities in KO→enhanced sensitivity to secondary cell injury in KO – low survival rates in KO vs. WT mice – increased liver injury, lower activities of liver thioredoxin reductase and catalase and higher liver carbonyl contents in KO
Fu et al., 1999a Fu et al., 1999b
Jaeschke et al., 1999 Bajt et al., 2002 Li et al., 2003
Lei et al., 2006
Zhu and Lei, 2006
Henderson et al., 2000
Dwivedi et al., 2006
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– In contrary to WT, KO and DKO were protected from lethality due to APAP overdosage (prolonged survival time, less cell injury, less hepatic GSH depletion than in WT) – liver injury protection associated with block of the APAP-mediated hepatic protein nitration and reduction of CYP2E1 in KO liver – KO higher resistance to APAP, but more sensitivity to NAPQI toxicity compared to WT – DKO showed lower CYP2E1 activity, but higher glutathione reductase and thioredoxin reductase activities than WT – less peroxynitrite-mediated protein nitration in KO and DKO
– hepatotoxicity in both KO and WT mice – KO show a higher grade of degenerative change (cellular swelling, disarray, vacuolization) surrounding the centrilobular necrotic lesions→role of GSTA-4 in secondary damage – KO highly resistant to APAP-induced hepatotoxicity (difference in recovery of GSH levels in KO, not WT mice)
References
October 8, 2007
Diquat
Acetaminophen
CCl4
GSTP1 GSTA4-4 (−/−)
Resultsa
JWBK195-Sahu
Glutathione S-transferases (GST) P2
Toxicanta
Transgenic model
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Table 9.2 (Continued)
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Methacrylonitrile (MAN) and acrylonitrile (AN) Acetaminophen Acetaminophen CCl4
Acetaminophen
Thioacetamide
Epoxide hydrolase (mEH) (−/−) Deoxyribonuclease 1(−/−)
Hsp70i (−/−)
CAR (−/−)
Retinoid X receptor alpha (−/−)
Metallothionein (MT)-I and MT-II (−/−)
cis-Diamminedichloroplatinum (cisplatin)
Acetaminophen
Oliver et al., 2006
– downregulation of CYP1A2 and CYP3A11 in KO, while upregulated in WT mice – significantly less lipid peroxidation in WT mice than in KO
(Continued)
Liu et al., 1998
Liu et al., 1999
Liu et al., 2000
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– more GSH depletion and reduced liver regeneration in KO – kidney and liver lesions in KO more severe than in WT mice – higher levels of cytokines in KO, more sensitivity to GSH depletion – higher susceptibility to acetaminophen-induced lethality and hepatotoxicity in KO mice compared to WT – increased sensitivity to oxidative stress – higher sensitivity to and pronounced apoptosis in cisplatin-induced toxicity in KO mice
Wu et al., 2004
October 8, 2007
Yamazaki et al., 2005
Tolson et al., 2006
Napirei et al., 2006
El Hadri et al., 2005
References
– KO more sensitive to APAP toxicity – thermal upregulation of HSP decreased hepatotoxicity in both WT and KO, but toxicity was higher in KO – slightly less liver damage in KO – differences pronounced after stimulation with phenobarbital→CAR activation caused CCl4 hepatotoxicity, while CAR inhibition resulted in partial protection against CCl4 -induced hepatotoxicity – resistance against APAP-induced liver injury in KO
– significant reduction in blood cyanide levels in MAN-treated KO→abolished AN to cyanide metabolism in KO mice – necrosis but less affection by overdose of APAP in KO
Resultsa
JWBK195-Sahu
Arsenic
Toxicanta
Transgenic model
JWBK195-09
Table 9.2 (Continued)
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a
– clofibrate treatment showed marked protection against APAP toxicity in WT but not in KO – no reduced hepatic glutathione depletion and arylation of cytosolic proteins in KO vs. WT mice −→ protective role of PPARalpha
– significantly increased expression of tumour necrosis factor-alpha (TNF-α), connective tissue growth factor (CTGF) and reduced IL10 in KO – significantly more severe liver injury and lethality in KO mice – untreated KO: lower UDP-GT expression and NPSH content – no significant differences between KO and WT mice
Chen et al., 2000
Boess et al., 1998
Enomoto et al., 2001
Gardner et al., 2002
TCDD (2,3,7,8-tetrachlorodibenzo- p-dioxin); 4-PeCDF (2,3,4,7,8-pentachlorodibenzofuran – dioxin-like compound); PCB 153 (2,2 ,4,4 ,5,5 -hexachlorobiphenyl).
Acetaminophen + clofibrate
Acetaminophen
Acetaminophen
TNF/lymphotoxinalpha (−/−) PPARalpha (−/−)
– less sensitivity to APAP toxicity in KO
Acetaminophen
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Acetaminophen
– no significant differences in toxicity between KO and WT mice
Acetaminophen
Poly(ADP-ribose) polymerases (PARP) (−/−) gp91 phox (−/−) subunit of NADPH oxidase Inducible nitric oxide synthase (NOS II)
Cover et al., 2005
Xie et al., 2004
October 8, 2007
nrf (−/−)
– greater susceptibility of KO to arsenic-induced pathological changes (elevated LPO levels increased glutathione S-transferase (GST) activity) – enhanced expression of potential oncogenic cyclin D1 only in KO – no difference in liver injuries between KO and WT mice
Arsenic
Mdr1a/1b(−/−)
Zhou et al., 2002
References
JWBK195-Sahu
– resistance to acute alcohol-induced hepatotoxicity in KO – reduced signs of oxidative stress in KO
Ethanol
Metallothionein overexpression
Resultsa
Toxicanta
Transgenic model
JWBK195-09
Table 9.2 (Continued)
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Drug-induced phospholipidosis was recently reviewed by us (Anderson and Borlak, 2006). 9.6.3.2
The Role of Mitochondrial Dysfunction in Hepatotoxicity
Alterations of the mitochondrial metabolism have been demonstrated to play a central role in different kinds of toxic liver injury. Functional impairment of these organelles is associated with energy depletion and extensive production of ROS that lead to a vicious circle of cellular stress. Several events have been demonstrated to cause failure of the mitochondrial metabolism, which eventually will end in cellular necrosis and apoptosis (Kim et al., 2003). The three main mechanisms by which toxicants cause primary functional mitochondrial failure are uncoupling of the oxidative phosphorylation, acivation of the caspases cascade leading to apoptosis, and disturbance of the Ca2+ homeostasis that causes perturbations and dysfunctions of the cellular metabolism, such as a disruption of the wellarranged actin assembly, possibly resulting in disruption of the cell membrane integrity (observed under Amanita phalloides toxin) (Watanabe and Phillips, 1986). In addition, damage of the mitochondrial structure, disruption of the β-oxidation of lipids and alterations of mitochondrial DNA synthesis have been recognized mechanisms in liver toxicity (Cullen, 2005). Different toxicants, however, have their different ways of reaching these endpoints. For example, uncoupling of the respiratory chain can result from direct interference with mitochondrial enzymes or from dysfunctions caused by mutations of the mitochondrial DNA (mtDNA) (both, for instance, observed effects of iron) (Ramm and Ruddell, 2005). Mitochondrial dysfunction results in cell death by apoptosis and is mediated by cytochrome c(1)-dependent activation of the mitochondrial permeability transition pores (MPT-pores) via Ca2+ - or ROS-related signalling (Rauen et al., 2004). This mechanism, for instance, has been discussed to contribute to the hepatotoxic effects of the non-steroidal anti-inflammatory drug salicylic acid (aspirin), the antipyretic drug acetaminophen (Reid et al., 2005; Vendemiale et al., 1996), the narcoleptic valproic acid, toxicity of bile acids (Rolo et al., 2003) and p-hydroxybenzoic acid (parabens) and others (Nakagawa and Moore, 1999; Trost and Lemasters, 1996). In the following, some examples of mitochondrial dysfunction in relation to their morphologic appearance in toxic liver injury will be discussed. The herbicide paraquat is a quatenary nitrogen bipyridinium that causes toxic reactions in the lung, kidneys and liver. In isolated hepatocytes, paraquat caused ultrastructural mitochondrial alterations and cell death in a dose- and time-dependent fashion (Palmeira et al., 1994a). This has been linked to an impressive depletion of ATP and NADPH, accompanied by depletion of glutathione (GSH) and an increase in oxidised glutathione (GSSG) (Palmeira et al., 1994b). More evidence for a central mitochondrial dysfunction was provided by an uncoupling of the respiration chain and partial inhibition of ATPsynthase and the mitochondrial complexes II and IV (Palmeira et al., 1995a). The underlying mechanism in paraquat toxicity has been proposed to result from the formation of oxidative paraquat radicals. By oxidation of critical SH groups at the respiratory complex, these are believed to induce an increased permeability of the proton pore (Castilho et al., 1995). The subsequent decrease of the mitochondrial membrane potential could provide an explanation for an uncoupling of the respiratory complex (Wallace et al.,
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1997). Accordingly, cell death of hepatocytes due to paraquat exposure has been linked to primary mitochondrial failure, resulting in a critical breakdown of the cellular energy supply. Besides, a decreased GSH/GSSG ratio has been suggested to contribute to the progression of irreversible cell injury. Certainly, the decreased GSH/GSSG ratio indicates an imbalance of the cellular redox system, as a result of enhanced intrinsic production of ROS. Investigations regarding the formation of oxygen radicals in the presence of paraquat revealed that an NADH-quinone oxidoreductase embedded in the mitochondrial outer membrane is responsible for an enhanced production of superoxide anions (Hirai et al., 1992). It is, therefore, not surprising that isolated hepatocytes treated with paraquat displayed significant amounts of lipid peroxides, since these are directly formed by reaction of oxidative radicals with intracellular and membranous lipids (Palmeira et al., 1995b). Lipid peroxidation has been demonstrated to cause an impairment of the cellular and metabolic integrity by modification of proteins (Britton, 1996; Khan et al., 2002). Taken together, these findings suggest that metabolic perturbations induced by lipid peroxidation might provide an alternate or additional mechanism contributing to paraquatinduced toxicity. Importantly, formation of undue ROS can be observed in several cases of hepatotoxicity induced by different agents. Even under physiological conditions, mitochondria release as much as 2–4 % of reducing equivalents from the respiratory chain, which produce superoxide anions and hydrogen peroxide from oxygen and water. In addition, radicals can be generated by formation of unstable intermediates at the respiratory chain that are produced when alternate agents serve as electron acceptors. These can carry electrons over to oxygen to produce superoxide radicals (Wallace et al., 1997). Cellular stress, as induced by various physiological and non-physiological conditions, including ageing, toxic agents or inflammation, can exacerbate mitochondrial ROS production (Gao et al., 2004). In addition, undue amounts of ROS can result from diminished clearance through the antioxidative defence system (Pessayre et al., 2001; Vendemiale et al., 1996), which will be discussed in detail later. Increased ROS production is known to induce damage in lipids, proteins and nucleic acids. In mitochondria, it can cause direct uncoupling of the oxidative phosphorylation by increasing the activity of the uncoupling protein UCP2 (Douette and Sluse, 2006). Furthermore, it was demonstrated that alterations of the mitochondrial respiration could be induced by an overload with metals, such as iron and copper, to result in increased generation of ROS and subsequently lipid peroxidation. Adduct formation of cytochrome c oxidase with the lipid peroxidation product 4-hydroxynonenal (HNE) resulted in consecutively decreased activity of cytochrome c oxidase (Chen et al., 1998). This enzyme is the terminal electron acceptor in the respiratory chain and holds an essential role in the production and regulation of energy (Ludwig et al., 2001). Oxidative attack of DNA results in the formation of oxidised bases, such as 8hydroxy-2 -deoxyguanosine (8-oxoG) (Bohr et al., 2002). This 8-OxoG can mispair with 2 -deoxycytidine 5 -triphosphate or with 2 -deoxyadenosine triphosphate during DNA replication, forming C∗ 8-oxoG and A∗ 8-oxoG mispairs. In humans, the MutY repair enzyme is responsible for recognition and repair of this defect. If the repair mechanism fails,
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deletion of C:G and transversion to A:T can lead to further alterations (Parker and Eshleman, 2003). A combination of modified mtDNA and oxidation of mitochondrial proteins and lipids has also been implicated in increased ROS formation in alcoholic patients (Pascussi et al., 2001). Additional mechanisms have been identified to cause mitochondrial dysfunction. For example, an impairment of mitochondrial membrane integrity has been suggested to be part of the toxic response observed under intoxication with aflatoxin B. The major metabolite of aflatoxin was shown to damage the mitochondrial membrane and thereby cause depletion of the mitochondrial respiration (Sajan et al., 1995, 1996, 1997). Nucleoside reverse transcriptase inhibitors (NRTIs) are the cornerstones of the highly active antiretroviral therapy (HAART) against the human immunodefiency virus. The hepatotoxicity associated with these substances clinically appears to be highly variable and ranges from low-grade hepatotoxicity and asymptomatic lactacidaemia to severe liver insufficiency with massive steatosis and life-threatening lactic acidosis (Van Huyen et al., 2006). Investigations revealed that NRTIs can affect the function of mitochondrial polymerase gamma, which replicates mitochondrial DNA. This interaction has been suggested to cause depletion of the mitochondrial DNA or qualitative changes that might relate to the mitochondrial cytopathy found in NRTI-treated patients (White, 2001). The ultrastructural alterations found in mitochondria during NRTI-induced toxicity have recently been associated with a decreased expression of the cytochrome oxidase (COX) subunit I, which is encoded by mitochondrial as well as nuclear DNA (Van Huyen et al., 2006). The exact actions by which toxic effects on mitochondria may affect the cellular metabolism still need to be established. Induction of apoptosis and an impairment of the energy metabolism due to altered oxidative phosphorylation, however, are believed to be the major consequences of mitochondrial dysfunction (Szewczyk and Woitczak, 2002) and current research has focused on elucidating the possible sequences leading to these endpoints.
Toxic effects on mitochondria in summary: – – – – –
Interferences in the oxidative phosphorylation Inhibition of β-oxidation Mutagenic effects on mitochondrial DNA Inhibition of transport mechanisms (BSP) (substrate or cofactor depletion) Induction of apoptosis by MPT and cytochrome c release
9.6.4
Cellular Stress
Oxidative stress. Stress can be defined as any disturbance of the cellular metabolism, which can be induced by various conditions, such as temperature or pH fluctuations, starving or over-alimentation (supernutrition), radiation, ageing and several endogenous and exogenous agents and toxins.
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Oxidative stress is defined as an imbalance between production and clearance of reactive oxygen/nitrogen species (ROS/RNS) (Sies, 1991). Accumulation of oxidative damage has been proposed to represent the ageing process per se. It is implicated in both acute and chronic cell injury and possibly plays a role in chemical carcinogenesis (Klaunig et al., 1998). Reactive oxygen species may be represented by the superoxide anion O− 2 , hydrogen r peroxide (H2 O2 ), the hydroxyl radical OH , organic peroxides or radicals (Fridovich, 1978). The natural abundance of ROS arises from reactions at the inner mitochondrial membrane. Superoxide radicals are produced in reaction with oxygen during electron transfer from the cofactors NADH and FADH2 to the first complexes of the respiratory chain. Thereafter, the mitochondrial manganese superoxide dismutase (MnSOD) transforms superoxides to H2 O2 (Nohl et al., 2005). Nitric oxide is a representative of the reactive nitrogen species (RNS) and provides the main source for formation of peroxynitrate by reaction with superoxide (Beckman, 1996). Peroxynitrate leads to nitration of protein tyrosine residues and is detoxified by GSH. It has recently been implicated to be involved in APAP-induced toxicity, where it was found to be located specifically in areas of necrotic centrilobular liver cells (Hinson et al., 1998). To keep the intracellular equilibrium between oxidative and reductive valences, the presence of antioxidants is of vital importance. The antioxidative defence is provided by nonenzymatic antioxidants, such as glutathione and vitamins E and C, and also by enzymatic antioxidants, including superoxide dismutase (SOD), glutathione peroxidase, glutathione reductase, catalase and hemoprotein peroxidases. Superoxide dismutase and catalase together, for instance, build a functional unit in which SOD captures superoxides and thereby forms H2 O2 , which is still highly reactive and can be transformed to water and oxgen by catalase (Scandalios et al., 2005). Additionally, drug-metabolizing Phase II enzymes such as glutathione-S-tranferase, glucuronosyl transferases and NAD(P)H:quinone reductase, provide direct and indirect protection against oxidative stress. Glutathione-S-transferase, for instance, is responsible for inactivation of electrophilic substances (by conjugation with nucleophilic glutathione) to enable clearance and excretion. An example of indirect antioxidative defence is the removal of possible inducers of ROS, such as is performed by NAD(P)H:chinon oxidoreductase (Sies, 1991). Glutathione (GSH), for instance, is an important intracellular antioxidant that scavenges free radicals and other radical species (e.g. the hydroxyl radical, lipid peroxyl radical, peroxynitrite and H2 O2 ) by direct and indirect enzymatic reactions (Lei, 2002). Two mechanisms allow glutathione to protect the cell from undue oxidative stress: first, it can protect protein thiols from ROS/RNS and secondly, it can reverse oxidative modifications, e.g. through removal of disulfide bonds and nitrosthiols (Han et al., 2006). In addition, glutathione is involved in Phase II reactions of biotransformation, thereby connecting antioxidant defence and the detoxifying system. Possible interferences between these two systems have been implicated in acetaminophen (APAP)-induced hepatotoxicity. Detoxification of the active APAP metabolite NAPQI is achieved by reaction with glutathione (GSH). Thus, it can be observed that toxic overdosage of APAP results in depletion of the GSH reservoir, which can also affect the cellular antioxidant system and causes
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dramatic shifts in the cellular redox state (Davis et al., 1974; Mitchell et al., 1973). Indeed, there is specific evidence for the presence of a significant quantity of ROS in cells exposed to toxic concentrations of APAP. Specifically, lipid oxidation, mitochondrial damage and presence of peroxynitrate, which is formed by reaction of NO with superoxide, have been observed (Gao et al., 2004; Park et al. 2002; Sies, 1991). Glutathione depletion is a common feature in liver toxicity induced by various drugs or xenobiotics. This can result from either detoxification of a drug through conjugation with GSH (APAP) or from exceedingly high production of ROS that can no longer be compensated by GSH. In this context, GSH depletion has different effects depending on the compartment affected. Mitochondrial GSH depletion was found to induce necrosis in primary hepatocyte cultures, while cytoplasmic GSH depletion resulted in sensitisation of proteins to TNF-induced apoptosis by inhibition of NF-κB-dependent survival gene expression (Matsumaru et al., 2003; Nagai et al., 2002). The consequence, however, is an increased potential of oxidative stress and the damage resulting from this. The transcriptional regulation of genes coding for enzymes of the antioxidant defence is a central event within the cellular response to oxidative stress and is mediated by interaction of transcription factors at specific DNA segments to either enhance or silence the expression of a certain gene. The antioxidant response element (ARE) is such an important DNA segment that was discovered in rodents and is relevant for basal expression and monofunctional induction of drug-metabolizing enzymes, such as γ -glutamylcysteine synthetase and heme oxygenase-1 (Friling et al., 1990; Kong et al., 2001; Rushmore et al., 1991). The oxidative stress-sensitive transcription factors Nrf-1 and Nrf-2 were found to specifically interact with AREs and thereby provide a positive regulation of antioxidative genes (Moinova and Mulcahy, 1999). The ARE complex consists of Nrf-1/2 and Maf proteins and was found to be activated by depletion of GSH that involved downstream activation of the PI3 and p38 MAP kinases and activation of the transcription factors AP-1 and NF-κB (Kang et al., 2000; Yang et al., 2005). Besides transcriptional regulation, antioxidant gene regulation is influenced and regulated by phosphorylation and nuclear translocation of transcription factors. The existence of a redox-sensing system has been suggested to be responsible for the detection of oxidative stress and subsequent induction of the anti-oxidative defence. Among these, glutathione and the protein thioredoxin have been proposed to play a key role (Nordberg and Arner, 2001). The mechanism by which GSH acts as a cellular sensor of the intracellular redox situation is regulated by the ratio of unoxidised GSH to the oxidised glutathione disulfide (GSSG). This ratio is maintained by GSSG reductase, which performs an NADPH-dependent reconversion from GSSG to GSH. Hence the GSH ratio is not only dependent on glutathione synthesis (gluamate cysteine ligase and GSH synthetase) and activity of GSSG reductase, but via NADPH dependence is also essentially connected to the energy status of the cell (Han et al., 2006). A shift of GSH/GSSG redox towards an oxidized state was shown to activate important signal transduction pathways, including activation of protein kinase B, protein phosphatases 1 and 2 A, calcineurin, c-Jun N-terminal kinase, apoptosis signalregulated kinase 1 and mitogen-activated protein kinase, AP-1 and NF-κB (Pinkus et al., 1996; Sen, 2000).
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The DNA binding activity of AP-1, NF-κB and Nrf-2 is dependent on the reduction of cysteine residue 506. In this way, the cellular redox can influence these transcription factors (Bloom et al., 2002). The glutathione and thioredoxin (TRX) systems share the feature of controlling the intracellular thiol/disulfide redox environment by thiol-dependent reductions. Even though they have been shown to regulate the cellular redox by different pathways, an overlapping of function has been reported for these two systems (Hansen et al., 2004). Both were demonstrated to function through Nrf-2-dependent signalling, although at different cellular sites. While GSH has recently been demonstrated to control the cytoplasmic dissociation of Nrf-2, TRX was shown to be responsible for the nuclear regulation of Nrf-2/DNA interactions (Hansen et al., 2004). Nrf-2 is a redox-sensitive transcription factor that regulates genes in response to oxidative stress. This is mediated via a cis-acting enhancer sequence: the antioxidative response element (ARE) (Nguyen et al., 2003). The antioxidative response element was shown to be responsible for transcriptional activation of genes involved in glutathione synthesis and regulation (Kwak et al., 2003; Moinova and Mulcahy, 1999; Rushmore and Pickett, 1993; Wild et al., 1998), genes coding for other thiols and other proteins with antioxidative properties (thioredoxin-1 (Kim et al., 2003b) & HO-1 = (Inamdar et al., 1996; Ishii et al., 2000)), as well as genes encoding drug-metabolizing enzymes (Favreau and Pickett, 1991; Jaiswal, 1991; McMahon et al., 2001; Rushmore and Pickett, 1990). Upon stimulation by oxidative stress, Nrf-2 dissociates from its inhibitory complex with the molecule Keap-1 and translocates from the cytoplasm to the nucleus. How activation of Nrf-2 occurs has not been fully elucidated – mechanisms that have been particularly implicated so far are involvement of phospho-dependent signalling cascades dominated by kinases, such as phosphoinositol-3-K (PI3K), MAP-kinase and phosphokinase C (PKC), and also direct phosphorylation (Nguyen et al., 2003). In addition to oxidative stress, ARE is also activated by several xenobiotics, including antioxidants (Rushmore et al., 1991), thiol-containing isocyanates (Bonnesen et al., 2001; Talalay et al., 1995), heavy metals (Prestera and Talalay, 1995) and heme complexes (Inamdar et al., 1996; Ren and Smith, 1995). AP-1. The activator protein-1 (Ap-1) complex is a transcription factor composed of several components from the family of dimeric basic region–leucine zipper proteins, including several proteins belonging to the Jun, Fos, Maf and ATF subfamilies (Shaulian and Karin, 2002). Ap-1 regulates vital cellular functions, such as cell proliferation and differentiation, survival and cell death, and has been implicated to be involved in the response to liver injury induced by toxicants and bile acids (Bernt et al., 2006). Several stimuli are known to induce Ap-1 signalling, however – in the context of toxic liver injury, induction by cytokines and cell matrix–interaction, as well as chemical stress, are of particular interest (Shaulian and Karin, 2002). Nuclear factor-κB. The oxidative stress-sensitive NF-κB has been attributed a central role in mediating inflammation, proliferation and cell death, and it is crucially involved in the regulation of cell survival (Hayden and Ghosh, 2004; Karin, 1999).
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Specifically, involvement of NF-κB in regulation of apoptosis and ROS is of particular interest, since severity and progression of liver injury was shown to depend on the incidence of hepatocytic apoptosis and extent of oxidative stress. Activation of NF-κB represents the major event in the signalling pathway of TNFR1dependent apoptosis. The TNFR1-associated adaptor proteins TRADD, TRAF-2 and RIP are involved in activation of the IκB-kinases that initiate the release of NF-κB from its inhibitory proteins (IκB) (Hayden and Ghosh, 2004). In addition, activation of NF-κB can also prevent apoptosis by downregulation of apoptotic genes or upregulation of antiapoptotic genes, including the cellular inhibitors of apoptosis 1 (IAP-1 and IAP-2), FLICE-inhibitory protein (FLIP), TRAF-1 and TRAF-2, which, for instance counteract apoptotic signalling by caspase inhibition (Poppelmann et al., 2005; Shishodia and Aggarwal, 2002). There is evidence that depletion of GSH may be the possible catalyst for decreased NF–κB activity and contribute to enhanced sensitivity to TNF-induced apoptosis by sustained activation of c-Jun-N-terminal kinase (JNK) (Matsumaru et al., 2003). Changes of NF-κB activity are early events observed after acetaminophen (APAP) administration. It was demonstrated that APAP significantly reduces the DNA binding activity of NF-κB up to complete inhibition 4 h after dosing. The same finding was made for nuclear factor-IL6 (nf-IL-6), which regulates genes involved in the acute phase reaction of the liver (Blazka et al., 1995). It has been proposed that APAP toxicity is characterized by negative regulation of genes that are altered during the toxic response (such as those implicated in hepatotoxicity induced by metals (Cisternas et al., 2005), ethanol-induced liver injury (Yuan et al., 2006), carbon tetrachloride (Liu et al., 1995) and others). For example, APAP was found to prevent the nuclear transition of the NF-κB and NF-IL6, compromising activation of the so-called survival genes (Blazka et al., 1995; Manna et al., 1998). In addition to its antiapoptotic effect, activation of NF-κB has been implicated to promote the second phase of liver injury (in particular inflammation and fibrosis) by induction of cytotoxic cytokine synthesis and initiation of chemotaxis of mononuclear cells, indicating ambivalent functions of NF-κB for the toxic response (Liu et al., 1995). A finding that supported the crucial role of oxidative stress in APAP-induced hepatotoxicity was that administration of the antioxidant thiol agent N -acetylcysteine (NAC), a cysteine precursor, amended APAP-induced toxicity, which was presumably due to replenishment of GSH levels (Bessems and Vermeulen, 2001; Corcoran and Wong, 1986; James et al., 2003b). There is also evidence that NAC influences activation of NF-κB and expression of inflammatory mediators (Bellezzo et al., 1998; Fox and Leingang, 1998; Pahan et al., 1998). The connection between glutathione and NF-κB was suggested to be mediated by thioredoxin, which is reduced by GSH, promotes the reduced state of NF-κB and also provides cysteine which is required for DNA binding of NF-κB (Matsumaru et al., 2003; Matthews et al., 1992). The oxidative response is associated with activation of several pathways of mitogenactivated protein kinases (MAPKs) signalling that are involved in coordinating the response to formation of ROS. MAPKs include a large number of serine/threonine kinases that are involved in the regulation of a wide array of cellular processes, including proliferation, differentiation, stress adaptation and apoptosis (Kolch, 2000). Members of the MAPK subfamilies, such as
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extracellular signal-regulated kinase (ERK), c-jun N-terminal kinases (JNK) and p38, have been shown to be substantially involved in the response to oxidant injury and, therefore, could potentially contribute to a cell’s fate between survival or apoptosis following exposure to oxidative stress (Cuda et al., 2002; Johnson and Lapadat, 2002; Martindale and Holbrook, 2002). Signalling of ROS has particularly been recognized to contribute to TNF-α-dependent activation of NF-κB and JNK, as well as downstream apoptotic pathways, notably, by enhancing the TNFR1 (tumour necrosis factor alpha-tumour necrosis factor receptor 1)dependent pathway and interacting with the redox regulatory protein thioredoxin that acts as regulator of apoptosis signal-regulating kinase 1 (ASK1) (Shen and Pervaiz, 2006). By contrast, opposing observations were made, in which superoxide had an inhibitory effect on death receptor-dependent and independent induction of cellular death (Clement et al., 2005; Clement and Stamenkovic, 1996). Based on these findings, it was proposed that a slight pro-oxidant condition might even provide a survival advantage (Shen and Pervaiz, 2006). 9.6.4.2
How do ROS Harm Cellular Integrity?
Nitric oxide is known to act both as an intra- and extracellular messenger molecule; likewise, the physiological role of ROS for cell development, differentiation and survival has been explored in the past decades (Dalton et al., 1999; Delaunay et al., 2000; Muriel, 2000). Increased amounts of intracellular reactive oxygen species, however, were shown to cause severe damage to lipids, proteins, and DNA (Scandalios, 2005). Protein damage that occurs under conditions of oxidative stress may be found as direct oxidation of proteins by ROS and/or RNS or adduction of secondary products of oxidation, such as sugars (glycooxidation) or polyunsaturated lipids (lipoxidation) (Dalle-Donne et al., 2005; Sayre et al., 2001; Stadtman, 1992). Direct protein damage due to ROS can occur via different mechanisms causing polypeptide chain cleavage, crosslinking, and modification of the side-chain of amino acids (Berlett and Stadtman, 1997; Dean et al., 1997). The modifications induced by oxidative/nitrosative stress may be permanent or reversible (Stadtman and Berlett, 1998). While reversible modifications (usually at Cys residues) modulate the protein function, they may also have a role in protecting the cell from irreversible damage. Irreversible modifications, such as protein–protein crosslinking, are generally associated with permanent loss of function and may lead to either degradation of damaged proteins (Berlett and Stadtman, 1997; Dean et al., 1997; Grune et al., 2003) or their progressive intracellular accumulation into cytoplasmic inclusions, as observed in age-related neurodegenerative disorders (Giasson et al., 2000, 2002; Butterfield and Kanski, 2001). The functional consequences of such reactions are diverse and include partly or complete inhibition of function, enhanced protein aggregation and proteolysis, changes in protein flux, altered immunogenicity or perturbations of cell signal transduction (Han et al., 2006). ROS are known to be involved in activation and inhibition of several signalling pathways, including c-Jun NH2 -terminal protein kinase (JNK), protein kinase C (PKC) and tyrosine kinase signalling. In addition, H2 O2 itself has been suggested to serve as a second messenger in any signalling pathway although no suitable receptor has, as yet, been identified (Han et al., 2006).
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There is indeed cumulating evidence that lipid peroxidation and protein modification due to oxidative stress is involved in the pathogenesis of chronic liver injury and fibrogenesis. This was supported by a study which investigated in vitro effects of oxidized low-density lipoproteins (oxLDL) on collagen and fibronectin synthesis of cultured human and rat hepatic stellate cells (HSC). Specifically, it was demonstrated that oxLDL stimulated the synthesis of collagen Types I and III and fibronectin of cultured HSC in a dose-dependent fashion (Schneiderhan et al., 2001). Moreover, production of reactive oxygen species and of reactive aldehydes from lipid peroxidation is part of the toxic response associated with other liver toxicants known to cause fibrosis, as observed with CCl4 and iron. Iron, in particular, is a potent inducer of reactive oxygen from O2 through the Fenton reaction. This reaction is a commonly accepted mechanism by which free radicals are formed. It consists of two steps, in which ferric iron first is reduced to ferrous iron (Equation (9.1)) to later generate a hydroxyl radical by attacking hydrogen peroxide (Equation (9.2)) (Buda et al., 2003): r− Fe+ −−→ Fe+ (9.1) 3 + O2 − 2 + O2 r − Fe+ −−→ Fe+ 2 + H 2 O2 − 3 + OH + OH
(9.2)
In hereditary hemochromatosis, different types of genetic defects lead to excessive iron absorption, eventually promoting an early onset of liver fibrosis and cirrhosis (Franchini, 2006). Investigations in patients diagnosed with hemochromatosis revealed that blood iron levels were positively correlated with the extent of lipid peroxidation and hence indicated an increased amount of oxidative stress in these patients (Young et al., 1994). A correlation between iron levels and lipid peroxidation supports the central role of ROS and lipid peroxidation in the pathology of hereditary hemochromatosis. Enhanced production of extracellular matrix due to iron provides at least one possible mechanism for initiation of early liver fibrosis associated with this disease. ROS effects onto DNA may lead to formation of specific adducts. These so-called etheno (epsilon)-modified DNA bases are, for instance, generated by reactions of DNA with the major product of lipid peroxidation, namely trans-4-hydroxy-2-nonenal. Even though elimination by base excision repair can be performed via ethenobase-DNA glycosylases in vivo (Gros et al., 2003), a high level of these etheno(epsilon) adducts has been associated with an increased risk to develop cancer. Patients diagnosed with cancer-prone diseases, such as Wilson’s disease and primary hemochromatosis, displayed significantly increased urinary levels of ethenobases, while in patients diagnosed with chronic hepatitis, cirrhosis and hepatocellular carcinoma levels of urinary etheno-deoxyadenosine were measured up to 20–90-fold higher than compared to controls (Bartsch and Nair, 2004). An analogous product, 8-nitroguanine (8-NO(2)-G), is formed by the reactions of guanine, guanosine or 2-deoxyguanosine with nitrogen species (RNS) generated from peroxynitrite. This has also been linked to inflammatory reactions and promotion of carcinogenesis (Ohshima et al., 2006). 9.6.5
Oxidation of Lipids and Lipid Toxicity to the Liver
Lipid oxidation in the liver occurs in association with several liver toxicants, such as carbon tetrachloride, ethanol and the metals iron and copper.
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Increased production of reactive oxygen species (ROS) is responsible for peroxidation of biomembranes and polyunsaturated fatty acids. There are several possibilities by which lipid peroxidation might interfere with important cellular functions. For example, membrane lipid peroxidation products may react with DNA and proteins to cause oxidative modifications (Park et al., 2002). In addition, it was demonstrated that lipid peroxidation destabilizes organelle compartments. Iron-induced lipid peroxidation, for instance, is associated with enhanced lysosomal fragility and inactivation of microsomal enzymes (Barreto et al., 2005; Britton et al., 1990, 1994; Lieber, 1993). Furthermore, mitochondrial dysfunction was associated with lipid peroxidation in iron- and copper-induced liver toxicity (Sokol et al., 1990). The extent of lipid peroxidation in vitro can be assessed by measurement of thiobarbituric acid reactants in the liver. In vivo, ethane in exhaled breath serves as an index for increased intracorporeal lipid peroxidation (Gutteridge and Quinlan, 1983; Riely et al., 1974). Treatment with several different steatogenic hepatotoxicants had linked enhanced lipid peroxidation with the presence of oxidizable fat in the liver. Due to the fact that steatosis is commonly associated with enhanced lipid peroxidation, it has been hypothesized that lipid peroxidation might be the major pathogenic factor in hepatocellular necrosis and mild inflammation observed in steatohepatitis (Letteron et al., 1996). The aldehydes malondialdehyde (MDA) and 4-hydroxynonenal (HNE) are endproducts of the peroxidation of membrane n-6 polyunsaturated fatty acids (Romero et al., 1998). Oxidized fatty acids have an important role in cellular signal transduction, such as in activation of Jun N -terminal kinase (Castello et al., 2005). Indeed, HNE is involved in the regulation of different genes and modulates various enzyme activities. It was demonstrated that increased HNE concentrations activated the protein kinase C-delta and thereby triggered apoptosis in isolated rat hepatocytes (Castello et al., 2005; Chiarpotto et al., 2005; Leonarduzzi et al., 2005). Moreover, HNE was shown to enhance DNA binding of AP-1 transcription factor and to activate macrophage and smooth muscle cells. This chemotactic effect of mediated by activation of the phosphoinositide-specific phospholipase C and leads to induction of the expression and synthesis of monocyte chemotactic protein 1 (MCP-1) and TGFβ1 (Chiarpotto et al., 2005; Leonarduzzi et al., 2005; Rossi et al., 1993). It was also proposed that lipid aldehydes may cause impairment of the cellular function and integrity by covalent binding with proteins (Britton, 1996; Khan et al., 2002). Redox alterations induced by ROS/RNA can induce apoptosis and necrosis in hepatocytes and other cells. Several regulators of cell survival and death are strongly influenced by the intracellular redox, including caspases, NF-κB, JNK and PKC, which have already been mentioned. ROS can activate caspases by direct stimulation of the mitochondrial membrane transition pore and cytochrome c release and also controls the stability of p53, which act in surveillance of cell integrity (Jaeschke et al., 2003). It has been observed that low levels of ROS, in particular, cause apoptosis, while high levels of ROS tend to induce necrosis. This has been explained by the different redox potentials (and therefore different sensitivities) of the diverse proteins that might be involved and also by caspases inhibition that was observed with high levels of ROS (Hampton and Orrenius, 1997). Moreover, oxidative stress-inducing and GSH-depleting agents were shown to influence the response to TNF-α, which is an important inflammatory cytokine and triggers
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apoptosis as well as necrosis (Matsumaru et al., 2003). In addition, the mitochondrial GSH level was shown to be especially important for activation of apoptosis via initiation of MPT. In particular, this has been suggested to be an important mechanism of apoptotic liver cell injury in acetaminophen-, rifampicin- and INH-induced hepatotoxicity (Chowdhury et al., 2006; Masubuchi et al., 2005; Reid et al., 2005). Other investigators had proposed that oxidative stress may be a major determinate of progression of hepatocellular damage, but does not represent a relevant mechanism in APAP toxicity. This was concluded from the finding that co-administration of the antioxidant vitamin E with APAP did not abrogate APAP-induced injury in animals. In addition, massive lipid peroxidation was observed in APAP-induced hepatotoxicity only after the onset of injury, suggesting that LPO may be a consequence rather than a cause of cellular damage (Knight et al., 2003). Another interesting connection has been made between the occurrence of oxidative stress and the degradation of actin filaments observed in early APAP-induced hepatotoxicity (Tsokos-Kuhn et al., 1988). Oxidation-dependent cytoskeletal protein alterations have been observed and investigated in quinone-induced toxicity (Mirabelli et al., 1989). Quinones may induce toxicity by several mechanisms, including alkylation and oxidative stress following redox cycling. The toxicity of redox cycling quinones in isolated rat hepatocytes was accompanied by a dose-dependent increase in the relative abundance of F-actin and an oxidation-dependent actin crosslinking, as well as by dissociation of the α-actinin from the actin network (Bellomo et al., 1990; Thor et al., 1988). The mechanisms responsible for these alterations have been suggested to involve the depletion of cytoskeletal protein sulfhydryl groups and the increase in cytosolic Ca2+ -concentration following the alkylation or oxidation of free sulfhydryl groups in several Ca2+ -transport systems (Bellomo et al., 1990). Loss of intracellular F-actin-containing stress fibres has been linked to cytotoxicity of hepato- and nephrotoxins (Dartsch et al., 1998). Stress fibres are arrangements of myosin and actin filaments that are linked to the plasma membrane via association with focal adhesions (McCue et al., 2004). They mediate adhesion to the extracellular matrix, migration and are produced during shear stress. Stress fibres in fibroblasts have an essential role in wound healing and morphogenesis and also fibrosis. The transdifferentiation of the stellate cell to myofibroblasts is the key for the regeneration ability of the liver and is closely associated with cytoskeletal reorganization and an increased extracellular matrix production in these cells (Kato et al., 1999). Inhibition of stellate cell activation and disturbance of stress fibre formation have been suggested as possible targets for treatment of liver fibrosis and cirrhosis (Imanishi et al., 2004; Matsui et al., 2004; Matsui and Kawada, 2005). As specified above, the impairment that can arise from oxidative damage concerns a wide range of cellular functions. Liver toxicity, for instance, may, in part, be promoted by enhanced production of ROS. In addition, toxicants may cause damage by an impairment of the antioxidative defence. This, for instance, may result from a decreased expression of antioxidants or an impaired function due to oxidative modification. Likewise, ethanol was demonstrated to impair the antioxidant defence system by depleting GSH and promoting its leakage, thereby causing mitochondrial damage and apoptosis (Lieber, 1997; Seitz and Stickel, 2006).
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Both acetaminophen and amiodarone (AD) were reported to induce significant decreases of glutathione peroxidase (GPX), which is normally responsible for scavenging oxidative radicals, although the same study had revealed that the potential hepatotoxic drugs APAP, AD and tetracycline induced upregulation of other antioxidative proteins, including peroxiredoxins 1 and 2 (PRX1/PRX2) in primary cultured rat hepatocytes (Yamamoto et al., 2005). The complexity of the response to oxidative stress has, as yet, not been elucidated, but the interaction with the response to toxic stimuli is most likely to cause exponentiation of the subsequent hepatocellular damage.
Oxidative stress in liver toxicity in in summary: – – – – – – –
Reducing antioxidative defence (e.g. GSH depletion, SOD repression) Alterations of the antioxidatie response Induction of e.g. CYP1E2→ROS ROS→lipid peroxidation, DNA and protein adducts→impaired metabolic function Direct activation of apoptotic programs and cascades by ROS ROS signalling in death and survival ROS→inflammation→fibrosis
9.6.6 9.6.6.1
Programs in the Hepatotoxic Response Apoptosis and Necrosis
Apoptosis and necrosis are assumed to represent two different sequences that arrive at a common endpoint – cellular death. While apoptosis is a highly organized and structured procedure that is crucial in embryonic development and regeneration activity, the common understanding of the term ‘necrosis’ comprehends an acute incidence that is caused by heavy alterations initiated by an overwhelming stimulus. Necrosis. Necrosis is characterized by ATP depletion that is caused by severe metabolic perturbations. Diminished levels of ATP cause a breakdown of the osmotic regulation and integrity of the cytoskeleton, which results in the characteristic morphology of cell swelling accompanied by membrane blebbing (Gores et al., 1990; Nishimura et al., 1998). A subsequent collapse of the residual cellular functions can be observed by loss of the mitochondrial potential and lysosomal stability, finally leading to an irreversible disturbance of plasma membrane permeability (Herman et al., 1988). Even though apoptosis and necrosis are distinct and independent pathways according to the traditional view, recent discoveries have implemented a link between these two events. Mitochondrial permeability transition (MPT), for instance, was demonstrated to play a causative role in induction of both necrosis and apoptosis in ischemia–reperfusion injury experiments in cultured hepatocytes (Kim et al., 2003a). The current view considers the possibility that necrosis and apoptosis share common pathways, for instance beginning at a common death signal, which depending on modifying factors, such as intracellular ATP concentrations, either entail cell lysis or programmed cell death (Lemasters, 1999). Taking
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this into account, John J. Lemasters suggested to introduce a new term, e.g. ‘necrapoptosis’, to preserve the traditional distinction between apoptosis and necrosis (Lemasters, 1999). Apoptosis. Apoptosis is distinctively characterized by morphological alternations, such as chromatin condensation, nuclear fragmentation and DNA degradation, cell shrinking and membrane blebbing, as well as formation of apoptotic bodies (councilman bodies), which are membrane-surrounded vesicles of the cellular content. Due to this localized degradation of single cells, the apoptotic process exerts less effects onto the environment than necrosis, during which release of potassium attracts inflammatory cells, while release of intracellular enzymes, such as proteases and hydrolases, damage the extracellular matrix and surrounding cells. The apoptotic programme is activated by several stimuli, including both exogenous and endogenous sources, such as DNA damage and ER stress, radiation, ROS and deprivation of growth factors or nutrition. Correspondingly, the terms ‘extrinsic’ and ‘intrinsic pathways’ of apoptosis have been formed. The extrinsic pathway is induced by an external ligand that leads to activation of the so-called death receptors. The four best studied of these transmembrane proteins are the Fas receptor (CD95), which is activated by binding to the socalled Fas ligand, tumour necrosis factor receptor 1 (TNF-α receptor, TNFR1) and tumour necrosis factor-related apoptosis-inducing ligand receptors 1 (TRAIL-R1) and 2 (TRAILR2). In the liver, additionally, death receptor 3 (DR3/TRAMP) and death receptor 6 are of importance (Malhi et al., 2006b). In response to binding to one of their dedicated ligands (Fas, TNF-α and TRAIL), these receptors form oligomers, thereby inducing recruitment of adaptor proteins, such as Fas-associated protein with death domain (FADD) or TNFR1associated death domain protein (TRADD) (Faubion and Gores, 1999). A central event of apoptosis is the activation of specific apoptotic enzymes, the cysteine– aspartate-specific proteases (caspases). Binding to specific adaptors leads to activation of certain caspases (-2,-8,-9,-10), inducing a whole cascade that involves autoactivation and downstream activation of caspases that are capable of degrading cellular contents. Upstream caspases are activated by the so-called death-inducing signalling complex (DISC), which consists of oligomerized death receptors, their adaptor proteins, procaspase 8, and procaspase 10. In contrast, within the intrinsic pathway activation of caspases, for instance, results from damage of DNA and mitochondria and subsequent leakage of pro-apoptotic factors into the cytosol. These include, among others, cytochrome c, SMAC/DIABLO (second mitochondria derived activator of caspases/direct IAP (inhibitor of apoptosis) binding protein with low PI), HtrA2/Omi, apoptosis-inducing factor and endonuclease G (Du et al., 2000; Jiang and Wang, 2004; Verhagen et al., 2000; Widlak and Garrard, 2005). These pathways, however, operate independently. There has been evidence that in several cells and tumours the intrinsic and extrinsic apoptotic pathways are connected in a coordinated crosstalk (Khosravi-Far and Esposti, 2004). Furthermore, the complex regulation of apoptosis is mediated by an interplay of pro-apoptotic and apoptotic factors, including members of the bcl-2 family, such as the pro-apoptotic proteins Bak, Bid, Bad, Bim, Noxa, Puma and the antiapoptotic proteins Bcl-2, Bcl-XL and Mcl-1 (Cory et al., 2003). The proteins Bax and Bak cause changes of the mitochondrial membrane integrity and thereby trigger mitochondrial dysfunction (Wei et al., 2001). An illustration of the apoptotic cascade is presented in Figure 9.4.
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FAS
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Figure 9.4 The apoptotic cascade. In the liver, apoptosis can be induced by two distinct pathways. The type 1 pathway (Peter and Kramer, 2003; Scaffidi et al., 1998) is mediated via death receptors and adaptor proteins, such as TRADD and TRAF2. These induce activation of procaspase 8 to caspase 8, which is responsible for activation of the executioner caspase 3. The type 2 pathway involves cleavage of the pro-apoptotic protein Bid by active caspase 8, followed by further activation of Bcl-2 members, triggering mitochondrial dysfunction and release of cytochrome c (Hatano et al., 2000; Li et al., 2002b)
The inevitable death cascade continues with the release of antiapoptotic factors by the mitochondrion, including cytochrome c and SMAC/DIABLO, thus accelerating apoptosis by reducing caspase via inhibition of IAP (Chai et al., 2000). Notably, the mitochondrial transition pore (MPTpores) that provides one important mechanism of cytochrome c release, is regulated by Ca2+ and induces permeabilization of the inner mitochondrial membrane. In consequence, mitochondrial depolarization results in uncoupling of the oxidative phosphorylation and mitochondrial swelling (Zoratti and Szabo, 1995). Cytochrome c subsequently activates caspase 3 and associates with Apaf-1 to a formation called the apoptosome, which subsequently activates caspase 9 (Li et al., 1997). The executioner caspase 3 leads to activation of endonucleases and induces DNA fragmentation (Liu et al., 1997). Apoptosis in liver injury. Apoptosis has been proposed to be a major event in toxic liver injury. An increased incidence of hepatocyte apoptosis has been observed in patients and animal models of different types of toxic liver injury, including those induced by ethanol, copper and autoimmune-mediated mechanisms.
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Elevated levels of the death receptor Fas and its ligand were demonstrated in ethanolinduced liver injury and found to be even more pronounced in non-alcoholic steatosis, where Fas levels correlated with the severity of the disease (Feldstein et al., 2003a; Pianko et al., 2001). In the liver, Fas receptors are expressed in hepatocytes, cholangiocytes, sinusoidal endothelial cells, stellate cells and Kupffer cells (Malhi et al., 2006b). They are mainly located at the Golgi complex and to a smaller extent are also found implemented in the plasma membrane (Bennett et al., 1998). Upon certain stimuli, a cascade is activated involving activation of sphingomyelinase, protein kinase C-zeta and NADPH oxidase. Consecutive superoxide signalling is responsible for activation of src-kinase, subsequently leading to phosphorylation cascades and activation of Fas. This causes the Fas receptor to translocate to the plasma membrane, where it is activated by binding to the Fas ligand, released by cytotoxic T lymphocytes and natural killer cells (Lee and Ferguson, 2003). The physiological role of the Fas receptor has recently been explored in a Fas-knockout mouse strain. Fas-deficient mice displayed substantial liver hyperplasia, accompanied by enlargement of hepatocytic nuclei and massive proliferation of lymphocytes (Adachi et al., 1995). The Fas system was also found to be involved in the pathogenesis of Wilson’s disease, where a hepatic overload of copper induces fulminant hepatic failure. Studies in patients diagnosed with this hereditary disease as well as in vitro investigations revealed that copper overload activates Fas-induced apoptosis and that this factor also contributes to the massive cell death during hepatic failure (Strand et al., 1998). In ethanol-induced liver injury, circulating levels of the Fas receptor and Fas ligand were found to be increased and notably correlated with the severity of the disease (Natori et al., 2001; Taieb et al., 1998; Ziol et al., 2001). Ethanol has been demonstrated to induce apoptosis by two different pathways: first, it induces the mitochondrial permeability transition (MPT) and secondly, it was shown to cause upregulation of the CD95-Fas ligand (Minana et al., 2002). Other studies, however, have revealed an increased expression of the death receptor TNFR1 in acute and chronic liver injury induced by alcohol (Deaciuc et al., 1995). An increased expression of death receptors may sensitize the liver to lymphocytemediated cytotoxicity (Natori et al., 2001; Tagami et al., 2003). An involvement of inflammatory cells has been concluded from findings where enhanced Fas receptor expression and infiltrating neutrophils colocalized in apoptotic liver areas (Jaeschke, 2002a,b). A possible mechanism by which induction of the Fas ligand by ethanol occurs, is a initiation of the corresponding signalling cascades as a result of undue generation of ROS from the mitochondrial acetaldehyde metabolism (Guicciardi and Gores, 2005; Hug et al., 1997). Acetaldehyde is toxic to the mitochondria and aggravates oxidative stress by binding to reduced glutathione and promoting its leakage (Lieber, 2004). The relevance of ROS for ethanol-related apoptosis was supported by investigations that demonstrated the antiapoptotic effect of antioxidants in ethanol-induced liver inury (Kurose et al., 1997). The toxic effects of ROS are believed to be triggered by induction of TNF-α expression and subsequent activation of NF-κB, which is also involved in the principal signalling pathway initiated by activation of TNFR1 (Chan et al., 1999). Moreover, Fas involvement has been proposed to play a role in the pathogenesis of non-alcoholic steatohepatitis (NASH). Similarly, increased apoptosis and expression of
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Fas receptor and TNFR1 were observed in patients diagnosed with NASH (Ribeiro et al., 2004); this was associated with caspase 8-dependent cleavage of Bid and activation of the executioner caspases 3 and 7 (Feldstein et al., 2003a,b). It is still unclear which role the apoptosis-induced mitochondrial dysfunction has in the development of the steatosis. NASH can be induced by drugs or by nutritional factors. Notably, fatty acids occurring along with this condition, presumably as result of a mitochondrial dysfunction, were found to induce the lysosomal pathway of apoptosis and cause mitochondrial dysfunction alone (Feldstein et al., 2004). Hence accumulation of fatty acids could cause predisposition of the liver to cell death induced by death receptors (Feldstein et al., 2003b). In models of cholestatic liver disease, the Fas receptor was shown to play a central role in bile acid-mediated apoptosis. Hydrophobic bile acids and bile salts induce apoptosis in hepatocytes, as demonstrated in experiments with primary hepatocytes in vitro (Faubion et al., 1999; Guicciardi and Gores, 2002; Higuchi et al., 2001). In animal models of extrahepatic cholestasis, the toxicity of hydrophbic bile salts has been investigated intensively. Increased rates of apoptosis have been observed, which was related to a pathway involving the Fas receptor (Miyoshi et al., 1999). From studies with mutant Fas ligand-expressing and Fas-null hepatocytes, it was demonstrated that apoptosis due to deoxycholic acid occurred in a ligand-independent fashion (Qiao et al., 2001), although it was also demonstrated that presence of the Fas receptor was required in bile acid-induced apoptosis (Gupta et al., 2004). While this was not associated with increased expression levels of the Fas receptor, an enhanced trafficking of the latter to the plasma membrane has been demonstrated (Sodeman et al., 2000). Activation of the Fas receptor was suggested to result from autoactivation induced by the increased density of the Fas receptor at the cell surface (Guicciardi and Gores, 2005). Another study had demonstrated the involvement of other death receptor pathways in bile acid-mediated apoptosis, such as the finding that apoptosis in Fas receptor-deficient cells occurred in a TRAIL associated manner (Higuchi et al., 2001). Furthermore, the Fas receptor-initiated apoptosis was implicated in the pathogenesis of viral hepatitis, hepatic fibrosis, hereditary biliary cirrhosis and autoimmue hepatitis, as well as in the carcinogenesis of hepatocellular and cholangiocellular carcinomas (Ghavami et al., 2005). The central role of apoptosis in toxic and cholestatic liver injury, hepatic inflammatory and fibrosis was supported by the protective effect of caspase inhibition on bile acidmediated cytotoxicity, progression of fibrosis, and stellate cell activation (Canbay et al., 2004). This, in turn, had led to development of caspase inhibitors for the therapeutic intervention in liver damage, which was demonstrated to be effective in animal models of extrahepatic cholestasis (Guicciardi and Gores, 2002). In the meantime, the therapeutic concept of caspase inhibition, as for instance, implemented in the case of the pan caspase inhibitor IDN-6556, has been introduced in clinical trials, where it is currently being evaluated (Linton, 2005; Valentino et al., 2003). 9.6.6.2
Chemokines and Cytokines as Mediators of Hepatocellular Toxicity
Inflammatory reactions are mediated by chemokines and cytokines. These play an important role in the pathology of acute toxic hepatitis, immune hepatitis and chronic liver diseases, including fibrosis and cirrhosis.
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In particular, TNF-α and the members of the C–X–C chemokine family, including interleukin-8 (IL-8) in humans and macrophage inflammatory protein-2 (MIP-2) in rodents, have been suggested to play an important role in damage and repair processes associated with various hepatotoxins (Dong et al., 1998). The most important inflammatory mediator, TNF-α, mediator of both local and systemic inflammation and is mainly produced by activated macrophages but also by other cell types, such as lymphoid cells, mast cells, endothelial cells and fibroblasts (Schwabe and Brenner, 2006; Tracey et al., 1987; Wajant et al., 2003). TNF-α is known to influence both apoptotic and survival mechanisms and is thus part of the difficult regulation between anti-apoptotic and pro-apoptotic pathways. Via Fas ligand activation of the Fas receptor, TNF-α induces apoptosis and, on the contrary, deploys a self-limiting action by inducing nuclear transcription factor NF-κB. NF-κB is responsible for the transcriptional regulation of genes that are termed survival genes and include antioxidant and anti-inflammatory proteins (Baichwal and Baeuerle, 1997; Bubici et al., 2006; Xiao and Ghosh, 2005). By stimulating the expression of the inducible NO synthetase (iNOS), NF-κB enhances the production of nitric oxide, which can inhibit the caspase cascade (Bradham et al., 1998). On the other hand, NF-κB upregulates expression of the Fas receptor and FasL genes and may thereby sensitise cells for the extrinsic pathway of apoptosis (Chan et al., 1999). A possible protective role of TNF-α has been implicated in APAP-induced hepatotoxicity. Here, studies in TNF-α receptor TNFR1-deficient mice had revealed that a lack of TNF response resulted in a delayed recovery of intracellular GSH levels, induction of hemeoxygenase-1 and decreased expression of CuZn superoxide dismutase and its catalase activity, both responsible for the reduction of superoxide anions and hydrogen peroxide levels (Chiu et al., 2003a). Next to TNF-α, hydrogen peroxide was demonstrated to induce production of IL-8 and MIP-2 in freshly isolated hepatocytes (Dong et al., 1998a). Thus, increased oxidative stress may directly induce cytokines that contribute to an inflammatory response. Specifically, this is the proposed mechanism by which metals are believed to induce expression of inflammatory cytokines. This was supported by the finding that administration of an antioxidative substance, such as N -acetylcysteine, abrogated metal-induced cytokine expression. A TNF-α receptor knockout mouse model revealed moreover that cadmium chloride (CdCl2 ) and vanadium pentoxide (V2 O5 ) induced expression of inflammatory cytokines independent of TNF-α production (Dong et al., 1998b). 9.6.6.3
Acute Phase Reaction
After acute liver injury, the liver metabolism switches to an acute phase response, which includes transcriptional activation of acute phase genes and enhanced synthesis of acute phase proteins, such as C-reactive protein (CRP) or alpha 1-acid glycoprotein (AGP) (Ezendam et al., 2004). In transgenic mice, high levels of the acute phase protein CRP were demonstrated to protect from lethal sepsis, assumingly by blocking cytokine stimulation induced by bacterial lipopolysaccharides (LPS) (Xia and Samols, 1997). The acute phase response is also activated in toxic liver injury. Within 24 h after exposure to toxicants, plasma AGP concentrations were found to be increased up to 2–3.5 times of the normal levels in toxic liver injury due to CCl4 , allyl alcohol, bromobenzene, acetaminophen or N -nitrosodimethylamine.
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Although not all liver toxicants seem to induce this reaction, ethionine-induced liver injury, for instance, showed no effects on plasma AGP concentrations, whereas galactosamine even markedly decreased AGP levels (Sugihara et al., 1992). In CCl4 -induced liver injury, the rise in plasma AGP concentrations was positively correlated with increased rates of plasma protein binding, which can be explained by the weak basic properties of CCl4 and the acidic properties of AGP (Sugihara et al., 1991). Taken together, these findings suggest that the acute phase response can be considered to be part of a physiological defensive reaction. The acute phase protein CRP, for instance, is known to induce cytokine expression in monocytic cells and was also suggested to act as a mediator of the classical complement system activation (Heijnen et al., 2006). The same cascades that may protect from bacterial invasion, however, may substantially contribute to the development of the ischemia– reperfusion injury of the liver. In an animal model, the complement complex C5b9 and CRP were found to be co-localized in hepatocytes, and in addition it was demonstrated that inhibition of complement was able to protect rats from ischemia–reperfusion liver injury (Heijnen et al., 2006). Studies in animal models, where the acute phase response was induced by injection of Escherichia coli LPS, indicated that several CYP-dependent activities might be suppressed during the acute phase reaction. The study revealed that the maximum velocity (Vmax ) values and the amount of CYP1A1/1A2 and CYP2E1 apoproteins were decreased during the acute phase response (El Kadi, 1998; Saitoh et al., 1999). Further studies revealed that the relation between the nature of the inflammatory reaction has an influence on P450 activity, and that some cytochrome monoxygenases are more (CYP3A6) susceptible than others (CYP1A1 and 1A2) (Bleau et al., 2001). The impact of the acute phase response upon the metabolizing function of the liver was furthermore emphasized by the finding that the acute phase response resulted in decreased gene expression of albumin and glutathione S-transferase, suggesting an impairment of drug transport and neutralization (short-time inactivation), as well as alterations of the Phase II biotransformation (Whalen et al., 2004). 9.6.6.4
TNF-α Antibodies in Acute Liver Failure
Driven by the consideration that an exacerbating inflammatory reaction can cause severe alterations of the liver metabolism, antagonizing the inflammatory mediators was experimentally investigated as a possible therapeutic strategy. Decreasing pro-inflammatory cytokine levels by pentoxifylline, as a TNF-α synthesis inhibitor, immunotherapy with IL-4 or IL-1 receptor antagonists, as well as reactivation of PPAR-α by administration of long-chain n-3 fatty acids have been evaluated in different disease models in which the inflammatory reaction is a central and progressing factor (Prandota, 2005). In fact, antagonizing the effect of TNF-α, which is particularly involved in cell death during fulminant liver failure (Wang et al., 2003), achieved successful results in animal models of LPS-induced liver injury and ischemia-reperfusion injury (Sneed et al., 2000; Yang et al., 2004b; Yee et al., 2003). In contrast, abrogating TNF-α signalling was not able to prevent liver toxicity induced by the mycotoxin Fumonisin B1 (FB1), in which TNF-α was shown to have a central role (He and Sharma, 2005). Nevertheless, further investigation
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may help to assess the potential that lies in antagonizing the inflammatory reaction in toxic liver injury. The inflammatory response is not only transmitted by monocytic cells: a variety of cytokine genes were found to be expressed by hepatocytes in response to inflammatory stimuli, such as LPS (Rowell et al., 1997) or interferon gamma (IF-γ ) (Ren et al., 2002) and includes among others MCP-1 (Wang et al., 1999), cytokine-induced neutrophil chemoattractant (CINC/ Interleukin-8) (Ohkubo et al., 1998) and the liver-specific cytokine CCL16 (Bauer et al., 1989; Nomiyama et al., 2001). The production of cytokines by hepatocytes was further implicated to be modulated by non-parenchymal cells, such as Kupffer cells (Mawet et al., 1996). Intriguingly, the major aldehydic product of lipid peroxidation, HNE, is chemotactic to macrophages and smooth muscle cells by inducing expression of MCP-1 TGF-β1 (Chiarpotto et al., 2005, Leonarduzzi et al, 2005). Thus, hepatocellular damage by oxidative stress is one possible mechanism by which the inflammatory response may be initiated. Furthermore, the inflammatory reaction in response to toxicants involves enhanced Kupffer cell infiltration and neutrophil infiltration that can be observed along with activation of NF-κB and enhanced expression of intercellular adhesion molecule-1 and cytokineinduced neutrophil chemoattractant-1 mRNAs in hepatocytes (Bauer et al., 2000). There is strong evidence for neutrophils to contribute to liver injury, causing cellular damage by burst of ROS and receptor mediated-neutrophil cytotoxicity. In particular, the release of TNF-α and IL-1 α has been suggested to be responsible for pathological manifestations of APAP-induced hepatotoxicity (Blazka et al., 1995). Indeed, inactivation of macrophages was shown to have a protective effect against APAPinduced hepatotoxicity (Goldin et al., 1996; Laskin and Pilaro, 1986). Even though the role of inflammation for the onset and progression of the acute toxic injury and chronic hepatic disease is not completely understood, inflammation greatly contributes to secondary (toxicant concentration-independent) liver injury. This may explain why inhibition of neutrophils protected hepatocytes from secondary toxicity and limited acute injury, but had no effect on prevention of APAP-induced liver injury (Lawson, 2000). 9.6.6.5
Activators of Survival and Repair
The activation of redox-sensitive transcription factors enables adaptive defence responses against toxic stimuli. In the case of APAP, the activation of these transcription factors has been observed to occur even at low and non-toxic concentrations (Goldring et al., 2004). The oxidative stress-sensitive transcription factor NF-κB plays an essential role in the regulation of genes that are involved in tissue damage and inflammation, such as the gene coding for cyclooxygenase-2 or antioxidative heme oxygenase-1 (HO-1). Heme oxygenase-1 is responsible for degradation of the prooxidant heme into carbon monoxide, iron and the antioxidant biliverdin. Induction of MO-1, for instance, has been observed to be part of the adaptive response upon treatment with carbon tetrachloride and has been suggested to be of critical importance for the recovery from hepatocellular injury. This was concluded from the observation that inhibition of heme oxygenase-1 resulted in a sustained increase in microsomal free heme concentrations associated with exacerbated liver injury (Nakahira et al., 2003).
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In studies investigating the role of HO-1 in chemical carcinogenesis it was shown that HO-1 expression increased along with progressive exposure to the mutagenic pdimethylaminoazobenzene. Notably, an inverse relationship between the expression of HO-1 protein and the degree of tumour differentiation in hepatocellular carcinomas was observed, therefore supporting a significant role of this enzyme in tissue repair processes (Caballero et al., 2004). 9.6.6.6
Regeneration
The regenerative activity of the liver has been a mythology itself for thousands of years. Some important molecular insights have highlighted the role of apoptosis, autophagy and proliferation for the enormous ability of the liver to completely regenerate after partial hepatectomy and loss of parenchyma. Experimental partial hepatectomy was demonstrated to result in extensive changes of the liver’s gene expression, which was mainly attributed to modulation of liver-enriched transcription factors (Schrem et al., 2004). Liver regeneration is associated with increased expression of the hepatocyte growth factor (HGF), which was demonstrated to confer a significant mitogenic stimulus. This cytokine is produced by Kupffer cells and sinusoidal endothelial cells, but not by parenchymal hepatocytes, and exerts not only mitogenic, but also cytotoxic as well as growth inhibitory activities, which are presumably mediated by activation of the transcription factor NF-κB and consecutive induction of the liver-enriched transcription factor C/EBP-β (Shen et al., 1997). Physiologically, hepatocyte proliferation and expression of the C/EBP-β transcription factor are exclusive to the postnatal liver development (Buck et al., 1994). Activation of C/EBP-β is required for a normal proliferative response, where it causes cell cycle arrest through inhibition of cyclin-dependent kinases (cdks). During liver regeneration, however, hepatocytes regain these proliferative features by a mechanism that has not been elucidated. After partial hepatectomy, hepatocytes were demonstrated to acquire the ability to produce the antiapoptotic bcl-2 protein and also display a decreased transcription of the three pro-apoptotic members of the family, Bak, Bad, and Bax (Tzung et al., 1997). Similar experiments revealed enhanced expression of the cytokines TGF-β 1, 2 and 3 during liver regeneration (Bissell et al., 1995). Furthermore, evidence was provided that TNF-α expression and interaction at the TNFR1 receptor were required for liver regeneration (Yamada et al., 1997). A possible link between the mandatory involvement of TNF-α has been provided by the finding that TGF-α-dependent activation of C/EBP-β is essential for hepatocellular proliferation via p90 ribosomal S kinase (RSK) (Buck et al., 1999). The TNF cytokines in the liver were attributed the exert effects on cell growth control, inflammation, and cytotoxicity (Diehl et al., 1994). TNF-α induces cell death in cultured hepatocytes and mouse liver in vivo but, surprisingly, inhibition of TNF-α transcription resulted in extensive cell death, which suggested the activation of further antiapoptotic pathways by TNF-α (Bradham et al., 1998). This pro-apoptotic effect has been ascribed to TNF-α-dependent activation of NF-κB (Patel et al., 1998). Even though NF-κB might contribute a positive effect to liver generation,
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recent studies have demonstrated progress of liver regeneration in presence of a NF-κB inhibitor (Laurent et al., 2005). The early growth response-1 transcription factor (Egr-1) is another gene that has been recognized to be part of the immediate-early gene expression response during early liver regeneration, notably by regulating hepatocellular mitotic progression through the spindle assembly cell cycle checkpoint (Liao et al., 2004). 9.6.6.7
MAPK P38 Inhibitors and Liver Toxicity
The p38 mitogen-activated protein kinase (MAP kinase) is a member of the well-studied MAP kinase family, which is known to be involved in the regulation of vital cellular functions, including cell cycle control, differentiation and apoptosis (Ashwell, 2006). Specifically, the JUN N -terminal kinases (JNKs) and p38 were shown to be activated during the cellular stress response and in reaction to pro-inflammatory stimuli, including lipopolysaccaride (LPS), interleukin-1 (IL-1), transforming growth factor beta (TGF-β) and tumour necrosis factor (TNF) (Freshney et al., 1994; Hannigan et al., 1998; Mendelson et al., 1996; Raingeaud et al., 1995; Rouse et al., 1994). Furthermore, p38 was found to be responsible for signal integration of inflammatory pathways. Activation and response of primary murine T-cells was closely related to activation of p38, and in turn inhibition of p38 was also shown to suppress T cell reactivity to certain stimuli (Zhang et al., 1999). Constitutive expression of the mitogen-activated protein kinase kinase 6 (MAPKK6), an important activator of p38, had suggested a role for p38 in thymocyte development as well as in CD8+ -mediated apoptosis (Pedraza-Alva et al., 2006; Rincon, 2001). In addition, it has been demonstrated that cytokine production of IL-1 beta, IL-2, IL-4, IL-5, IL-8, IL-13 and IFN-γ is mediated by activation of p38 (Clark et al., 2003; Dodeller et al., 2005; Pesu et al., 2002; Zhang and Kaplan, 2000). Targeting the p38 MAP kinase with specific inhibitors has contributed new insights into the physiological role of the MAP kinase pathways in apoptosis, proliferation, survival, stress response and inflammatory processes (Dambach, 2005). Such inhibition of the alpha and beta forms of p38 by BRIRB 796 exerted a suppressive effect on inflammatory processes due to administration of LPS, such as activation of peripheral white blood cells, cytokine production and release of acute phase protein C-reactive protein (CRP) (Branger et al., 2002). In contrast, information gained from studies in knockout mice had implicated a vital role of p38 in development and differentiation, for deletion of the P38alpha gene in knockout mice resulted in embryonic death due to defects in the cardiovascular system (Allen et al., 2000). Moreover, several studies had linked p38 activity to proliferation, differentiation and pathophysiological response of heart and skeleton muscle, neurones and blood cells (Behr et al., 2003; Liang and Molkentin, 2003; Piao et al., 2002; Platanias, 2003; Ravingerova et al., 2003; Verma et al., 2002; Zetser et al., 1999). Development of selective p38 MAP kinase inhibitors, however, has involved the hope for a molecular-targeted drug with the ability to prevent activation of cell death and disease progression in inflammatory diseases. Unfortunately, several substances that have been brought forward in pre-clinical and clinical studies have revealed severe adverse effects on different organ systems, as, for instance, VX-745 and BIRB 796, which exhibited severe toxic effects on the central nervous system (CNS) and the liver, respectively (Cirillo et al., 2002).
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In the liver, p38 activity has been related to liver injury induced by ischemia–reperfusion, viral hepatitis and toxicants, such as ethanol and polycyclic aromatic hydrocarbons (Andrysik et al., 2006; Hsu et al., 2006; Thakur et al., 2006; Wang et al., 2004). Although p38 seems to be closely related to the inflammatory component and activation of Kupffer cells in liver injury, it has also been assigned an important role in regeneration processes after partial hepatectomy (Awad et al., 2000; Spector et al., 1997). The prospects to develop a molecular-targeted drug to interfere downstream in the TNF-α pathway are very promising, since progression of liver injury is often related to activation of the pro-inflammatory cascade; however, currently the lack of specificity of p38 inhibitors and an unacceptable safety profile call for further research to augment the knowledge regarding the necessity and function of p38 (Dambach, 2005). 9.6.6.8
Fluoroquinolones and Liver Toxicity
An example of drug withdrawal from the market is Trovafloxacin, a drug that displayed high antistaphylococcal activity in vitro and was a promising candidate for the therapy of oxacillin-resistant staphylococcal endocarditis (Bayer et al., 1998). After several cases of hepatic incidences, including severe hepatitis and fulminant liver failure (Lazarczyk et al., 2001; Lucena et al., 2000), Trovafloxacin in 1999 was withdrawn from the German market and restricted to special indications in the US (Blum et al., 1994). Another fluoroquinolone antibiotic, Temafloxacin, with similar chemical properties as Trovafloxacin, was withdrawn worldwide in 1992, due to several toxic effects including renal failure and hepatic dysfunction (Blum et al., 1994). It has been reasoned that the toxicity of these substances resulted from similar mechanisms based on their chemical structure. Both contain a unique difluorinated side-chain that is different from other quinolones and is responsible for the lipophility of these substances (Lucena et al., 2000). However, the mechanism by which Trovafloxacin and Temafloxacin cause liver toxicity remains unelucidated. A recent study, in which gene expression under Trovafloxacin treatment was investigated in a model that intended to stimulate idiosyncratic liver toxicity by co-application of LPS, revealed that GM-CSF was upregulated in treated rats. Additional upregulation of the pro-inflammatory cytokines IL-6, matrix metalloproteinase 9 and mitogen-activated protein kinase kinase-1 implicated that the toxicity of Trovafloxacin might result from an inflammatory reaction due to involvement of polymarphonuclear (PMN) cells (Waring et al., 2006). This hypothesis was supported by the finding that depletion of PMN was able to prevent hepatocellular injury in LPS/Trovafloxacin-treated rats (Waring et al., 2006). In vitro experiments demonstrated a toxic effect of pefloxacin, ciprofloxacin and ofloxacin in hepatocyte cultures and unscheduled DNA synthesis, as well as a significant dose-dependent increase in chromatid-type breakage were observed in in vivo investigations in rats that had been treated with doses of 20–200 mg/kg/day (Basaran et al., 1993; McQueen et al., 1991; Nordmann et al., 1989). Although clinically adverse effects (e.g. torsades de pointes, hepatotoxicity and dysglycaemias) are rarely observed under therapy of currently available quinolones, such as ciprofloxacin and ofloxacin, it has been suggested that adverse reactions may be confined to “susceptible” patient populations (Bertino and Fish, 2000; Owens and Ambrose, 2005). Therefore, although the safety profile of application of quinolones seems to be acceptable, it is necessary to be aware of the possible occurrence of toxic effects.
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Biotransformation in liver toxicity in summary: – – – – –
Induction, repression and inhibition of metabolizing enzymes Induction of e.g. CYP1E2→ROS Repression of metabolizing enzymes→accumulation of toxins Bioactivation (generation of toxic metabolites during biotransformation) Interference with NR or NR crosstalk in vital cellular functions→reduced metabolic competence – Inhibition of drug transporters→drug accumulation 9.6.7
Genetic Polymorphisms and Drug-Related Toxicity
The different isoforms of drug-metabolizing cytochrome mono-oxygenases have long been recognized to underlie variable expression due to genetic polymorphisms (Pirmohamed and Park, 2003). These polymorphisms may be associated with reduced (poor metabolizer) as well as enhanced (extensive metabolizer) activity of the CYP enzymes (Pirmohamed and Park, 2003). In particular, frequent polymorphisms of the genes encoding for CYP1A2 and CYP2D6 were found responsible for causing a vast majority of adverse drug reactions (Wolf and Smith, 1999a,b). In the case of CYP2D6 polymorphisms, increased liver toxicity was observed after application of the calcium channel antagonist perhexilene to treat ischemic heart disease (Pirmohamed and Park, 2001). Beside CYP polymorphisms, variations of other drug-metabolizing enzymes have been associated with frequent adverse drug effects and individual susceptibility to specific toxic effects. Investigations in patients with Gilbert’s syndrome demonstrated that a reduced activity of uridine diphosphate glucuronosyl transferase (UGT1A1) was often associated with a reduced activity of N -acetylation, rendering these patients slow acetylators (Fengler et al., 1993). Gilbert’s syndrome is characterized by reduced activity of uridine diphosphate glucuronosyl transferase (UGT1A1) due to frequent gene polymorphisms, which leads to impaired glucuronidation with infrequent occurrence of hyperbilirubinaemia and jaundice that is usually harmless (Sampietro and Iolascon, 1999). Multiple alterations of drugmetabolizing enzymes and transporters, however, may be likely to account for a significant part of unexpected idiosyncratic drug reactions (Pirmohamed and Park, 2001). Drug withdrawal from the pharmaceutical market occurs frequently for retrospective safety issues after introduction to the market (Friedman et al., 1999). The common causes for withdrawal are adverse effects on the hepatic, haematological and cardiovascular systems. The major cause, however, is the hepatic reaction that may become noticeable by elevation of liver enzymes, jaundice, hepatitis or severe hepatocellular damage, necrosis and hepatic failure (Fung et al., 2001). 9.6.7.1
Thiopurine Methyltransferase and 6-Mercaptopurine and Azathioprine
The efficient immunosuppressive drug azathioprine is frequently used in therapy regimes after organ transplantation to prevent rejection and is also applied in autoimmune inflammatory diseases, such as bowels disease, autoimmune hepatitis and rheumatoid arthritis. Azathioprine is degraded to its metabolite, the imidazol derivative 6-mercaptopurine (6-MP), and is further metabolized by methylation via thiopurine methyltransferase
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(TPMT), which is under control of common genetic polymorphisms resulting in a deficiency of 0.3 % and a reduced activity in 10 % of the Caucasian population (Sanderson et al., 2004). Azathioprine treatment in patients displaying a TPMT polymorphism may result in toxicity of the liver and severe myelosuppression (Kontorinis et al., 2004; Lennard et al., 1989). In vitro experiments testing azathioprine in rat hepatocytes had confirmed that even therapeutic concentrations of azathioprine and 6-MP caused liver toxicity by a mechanism which was completely reversible by application of xanthine oxidase inhibitor allopurinol and vitamin E analogue trolox, suggesting the involvement of oxidative stress, mitochondrial injury and ATP depletion in azathioprine and 6-MP toxicity (Lee and Farrell, 2001; Tapner et al., 2004). In particular, the incidence of hepatotoxicity in inflammatory bowel disease patients receiving thiopurines was found to be relevant, although it was mainly associated with corticosteroids co-treatment (Bastida et al., 2005). As a consequence, pre-emptive testing of the TPMT activity has been recommended prior to treatment with azathioprine, aiming to guide the drug’s dosage and to possibly prevent toxicity (Krynetski et al., 1996; Sies et al., 2005; Yates et al., 1997). On the other hand, only 15 % to 28 % of patients displaying adverse drug reactions to azathioprine show a TPMT-deficiency and it has been suggested that the remaining cases of adverse effects may be explained by a deficiency of the enzyme inosine triphosphate pyrophosphatase (ITPase). ITPase deficiency results in accumulation of inosine nucleotide, which was found to form the potentially toxic metabolite 6-thioITP in combination with 6-MP (Marinaki et al., 2004a,b; Sumi et al., 2002). A previous study, however, in which ITPase and TPMT genotypes and activity were correlated with the clinical response to azathioprine in 65 liver transplant recipients, had indicated that adverse drug reactions may not be predicted by preventive assessment of the TPMT activity in liver transplant patients (Breen et al., 2005). Even though there is evidence that the mechanism of action of azathioprine and 6MP is primarily associated with the induction of mitochondrial dysfunction, the exact circumstances by which these drugs cause liver toxicity and myelosuppression are unclear. The recent understanding, however, has led to the recognition that elevated 6-MP plasma metabolite concentrations are strongly associated with toxic incidences, whereas reduction and monitoring of these was clearly demonstrated to be beneficial for the patients (Seidman, 2003). Currently, the effect of antioxidant treatment in azathioprine and 6-MP toxicity is being assessed (Raza et al., 2003), signifying that a clinical benefit may result only from a deeper understanding of the underlying molecular processes. 9.6.7.2
Dihydropyrimidine Dehydrogenase (DPD) and Toxicity of 5-Fluorouracil
An ongoing example for the impact of genetic polymorphisms on toxic effects of therapy regimes is 5-fluorouracil (5-FU). 5-FU is a chemotherapeutic used to treat enterohepatic and gynaecological cancers. Adverse effects, such as mucositis, neutropenia, cardiac and neuronal toxicity, have been linked to deficiency of the liver enzyme dihydropyrimidine dehydrogenase (DPD), which is the rate-limiting step of 5-fluorouracil (5-FU) metabolism (Baek et al., 2006; van Kuilenburg et al., 2004). The clinical use of preventive screening for DPD mutations has recently been discussed for 5-FU (van Kuilenburg, 2006) and oral fluoropyrimidine capecitabine, which also is suspicious for severe toxicity in DPD-deficient cancer patients (Ciccolini et al., 2006; Saif and Diasio, 2006). A tragic example of the individual impact of genetic polymorphisms in drug metabolism is a case report about a pharmacogenetic syndrome, in which two genetic polymorphisms
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collided in a patient treated for colorectal adenocarcinoma, where the combined therapy of 5-fluorouracil and irinotecan had a fatal outcome (Steiner et al., 2005). Lethal toxicity arose from heterozygosity and consecutive reduced activity of two drug-metabolizing enzymes, DPD and uridine diphosphate glucuronosyltransferase 1A1 (UGT1A1), which induced an impaired glucuronidation activity (Gilbert’s syndrome) and was linked to toxicity in the treatment with irinotecan (Steiner et al., 2005). Introduction of routine screening for currently known drug-metabolizing enzymes that are important for drug toxicity, e.g. TPMT, UGT1A1 and DPD, prior to treatment may help to prevent cases of severe toxicity under chemotherapeutic drug treatment. The knowledge that a specific defect may cause drug-related organ toxicity allows preventive strategies to be introduced that help to avoid adverse effects, for instance, by pre-selecting suitable patients for the treatment, but also enables the development of new approaches to come up against adverse effects, e.g. by enhancing gene expression of the enzyme in question (Shimizu et al., 2001; Yamaguchi et al., 2006).
9.7
Conclusions
In summary, several mechanisms are involved in the onset and progression of toxic liver injury. In particular, the initiating parenchymal damage is most likely to result from drugor toxin-specific alterations of the cellular homeostasis. These mechanisms depend on the concentration of the toxin and arise from direct interaction with cellular functions, e.g. by interference with functions of synthesis and integrity, xenobiotic metabolism, as well as transcription and transporter activities. It is one of the greatest challenges of the future to elucidate the true origin of the functional alteration induced by certain toxins. On the other hand, a deep understanding of the secondary events involved in toxic liver injury and the stress-related signalling pathways is also necessary for the development of new therapeutic approaches to limit the progression of liver injury and prevent acute liver failure. The key targets for future therapeutic approaches are namely stress-related JNK-p38, the TNF-α and NK-κB pathways, as well as the complement and apoptotic signalling cascades and their organelles of origin, such as the mitochondrion, the lysosome and the peroxisome. Furthermore, addressing the sources of extracellular and intracellular stress, such as the ROS-generating machinery and expression of pro-inflammatory cytokines, may bear a promising approach to encounter liver injury. Nevertheless, it is of never-ending importance that toxicity is as a function of toxin concentration and exposure time. Therefore, the best therapeutic strategy is prevention of toxic liver injury, for which it is necessary to gain further knowledge about mechanisms of drug interactions, the role of individual gene expression and liver regeneration in the susceptibility to liver toxicity.
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10 A Role of Cytochrome P450 in Quinone-Induced Hepatotoxicity Yasuhiro Ishihara and Norio Shimamoto
10.1
Introduction
Quinones are widely distributed in nature, and are found in almost all respiring animal and plant cells. They form an important group of substrates for flavoenzymes and have vital physiological functions in a number of processes, such as photosynthesis and the respiratory chain in mitochondria. Some quinones are used as anticancer, antimalarial or antibacterial drugs (Asche, 2005; Lown, 1983; Vennerstrom and Eaton, 1988). However, their therapeutic use is limited in some cases because the use of most quinones is accompanied by adverse effects derived from their cytotoxicity. Therefore, understanding the toxic mechanisms of quinones would help us to use quinone drugs more appropriately and to develop novel quinone-based drugs. The toxic mechanisms of quinones are still under debate. The majority of xenobiotics, including food additives, cosmetic products, agrochemicals and drugs, are metabolized by xenobiotic-metabolizing enzymes, which are classified into two groups – Phase 1 and Phase 2. Of the Phase 1 group, cytochrome P450 plays a crucial role for the monooxygenation of xenobiotics as well as some endogenous substrates, which is localized largely in the endoplasmic reticulum in hepatocytes. Phase 2 enzymes conjugate substrates that are usually Phase 1 metabolites. The structure of quinone is not modified by Phase 1 enzymes and thus quinone compounds are thought to be metabolized mainly via a conjugation process. In the liver, quinone compounds are conjugated with glucuronic acid, sulfuric acid and reduced glutathione to form soluble conjugates, followed by release into the central vein or bile ducts. In that sense, quinone hepatotoxicity does not seem to be related
Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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directly to xenobiotic enzymes. Very recently, however, we have shown that quinone hepatotoxicity is enhanced under conditions of cytochrome P450 inhibition, indicating clearly the involvement of cytochrome P450 in quinone hepatotoxicity (Ishihara et al., 2006). In this chapter, we revisit the generally accepted mechanisms of quinone hepatotoxicity and propose the importance of cytochrome P450 systems in quinone-induced hepatotoxicity on the basis of our recent work.
10.2
Mechanism of Quinone-Induced Hepatotoxicity
The toxicity of quinones is induced by two principal mechanisms. They are: (i) arylationl/alkylation, reaction with nucleophiles among important biological constituents, and (ii) redox cycle, superoxide anion generation via quinone–semiquinone interconversion, resulting in oxidative stress, as shown in Figure 10.1. 10.2.1
Arylationl/Alkylation
There are many thiols that play important roles in regulating cellular functions. These include low-molecular-mass thiol compounds and protein thiols. Reduced glutathione (GSH) as a representative low-molecular-mass thiol compound is present at millimolar concentrations in many types of cells, especially hepatocytes, and plays a fundamental role in managing cellular redox homeostasis by protecting cells from exogenous or endogenous oxidative stress (Wu et al., 2004). Protein thiols, which usually indicate the presence of cysteine in
(ii) Redox Cycle
(i) Arylation/Alkylation R–SH
– 2
O
Q
NAD(P)+
NADPH–cytochrome P450 reductase NADH–cytochrome b5 reductase
Q
Q–SR
O2
NAD(P)H
Cytochrome P450
GSH depletion Enzymatic inactivation Q: quinone Q : semiquinone radical
Figure 10.1 Mechanisms of quinone hepatotoxicity. Quinone cytotoxicity is elicited by two mechanisms: (i) the arylation/alkylation of nucleophiles among important biological constituents, for example, quinones react covalently with thiols, such as GSH or the cysteine residues of proteins, to form arylation/alkylation products that eventually cause cellular damage, and (ii) oxidative stress via redox cycling. Redox-cycling quinones accept one electron from intracellular reductases to form semiquinone radicals, which are auto-oxidized by donating one electron to molecular oxygen. In these processes, massive superoxide anion radicals are produced, leading to oxidative stress
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proteins, are often active sites of proteins such as proteases (Thornberry et al., 1992), phosphatases (Barrett et al., 1999) and antioxidative enzymes (Arner and Holmgren, 2000; Fujii and Ikeda, 2002), and the functional loss of protein thiols induces dysfunction of these proteins. Electrophilic moieties of quinones (e.g. 2, 3, 5 and 6 positions of 1,4-benzoquinone (BQ)) react covalently with these functional thiols to form arylation/alkylation products. This reaction causes protein dysfunction, leading to serious cellular damage, and often cell death (necrosis). Some groups have reported that when primary hepatocytes were treated with arylating quinone, BQ, intracellular levels of GSH and protein thiol decreased in a time-dependent manner, followed by cell death (Rossi et al., 1986; Tapper et al., 2000). However, intracellular levels of glutathione disulfide (GSSG) did not change, suggesting that the decreased levels of GSH might not be due to oxidative stress but is the result of chemical reaction of quinones with GSH (Rossi et al., 1986). The relationship between quinone toxicity and oxidative stress is described in Sections 10.2.3 and 10.2.4. 10.2.2
Reactive Oxygen Species and their Scavenging Systems
Reactive oxygen species (ROS) are the general term for highly reactive molecules derived from oxygen, such as the superoxide anion, hydrogen peroxide and hydroxyl radical. ROS are generated constitutively from cellular organelles, especially liver endoplasmic reticulum and mitochondria, and a variety of xenobiotics such as quinones induce ROS production. Cells have defense systems to cope with routinely generated ROS. The superoxide anion is dismutated to hydrogen peroxide and molecular oxygen by superoxide dismutase (SOD). Mitochondrial Mn–SOD efficiently eliminates the superoxide anion that arises from molecular oxygen in the respiratory chain. The intracellular Cu/Zn–SOD is present in the cytoplasm, nucleus and peroxisomes of all mammalian cells, where it scavenges superoxide anion (Crapo et al., 1992). Hydrogen peroxide produced by the dismutation of the superoxide anion radical is degraded to molecular oxygen and water by catalase, glutathione peroxidase (GPx) and peroxiredoxin (Prx). Catalases are present ubiquitously in aerobic organisms, and the highest level of enzyme activity is in the liver and erythrocytes. Within cells, catalases are located mostly in peroxisomes because of the presence of many hydrogen peroxide-producing enzymes. There are four types of GPx, i.e. classical GPx (GPx1), gastrointestinal GPx (GPx2), plasma GPx (GPx3) and phospholipid hydroperoxide GPx (GPx4) (Imai and Nakagawa, 2003). The former three GPxs reduce hydrogen peroxide and organic alkyl hydroperoxides to water and corresponding alcohols, at the expense of oxidation of GSH to GSSG. The last type of peroxidase, phospholipid hydroperoxide GPx, is different from the other three GPxs with regard to its substrate specificity and localization. This enzyme has the unique ability to reduce membrane lipid hydroperoxides such as phospholipid and cholesterol hydroperoxides directly, and is located mostly in the testis (Thomas et al. 1990). Prx was discovered more recently than the above-mentioned catalase and GPx, and is a hydrogen peroxide-scavenging enzyme. Prx enzymes contain conserved cysteine residues among Prx enzymes that undergo peroxide-dependent oxidation and thiol (thioredoxin)-dependent reduction during a catalytic cycle. Mammalian cells express six isoforms of Prx (Prx I to VI), which are classified into three subgroups (2-Cys, atypical 2-Cys and 1-Cys) on the basis of the number and position of cysteine residues as active sites. The relatively high abundance of Prx enzymes in mammalian cells appears to play a role for removing the low levels of peroxides produced during normal cellular metabolism
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(Rhee et al., 2005). Besides antioxidative enzymes as described above, cells have small molecules with antioxidative capacity, such as ascorbic acid, α-tocopherol and GSH (Mates, 2000). These molecules are able to scavenge ROS efficiently and specifically at relatively low concentrations. When ROS is not eliminated sufficiently by these anti-oxidative enzymes or small molecules, cells are damaged by oxidative ‘insults’, leading to cell death. 10.2.3
Redox Cycle
Quinones undergo one-electron reduction by various intracellular reductases to yield semiquinone radicals. The semiquinone radicals are then re-oxidized easily in co-operation with molecular oxygen to form the superoxide anion. This oxidation–reduction process between quinone and the semiquinone radical is called the quinone redox cycle. The superoxide anion generated via the redox cycle is dismutated to hydrogen peroxide and oxygen by superoxide dismutase. Then, hydrogen peroxide is converted into hydroxyl radicals in the presence of metal cations such as Fe2+ or Cu+ (Me(n−1)+ + H2 O2 → Men+ +OH− +HO· ). The hydroxyl radical is highly unstable and the most reactive oxygen radical among ROS, causing lipid peroxidation, protein oxidation and DNA damage, followed by cellular toxicity (Droge, 2002; Galli et al., 2005). There are some reports of quinone toxicity via redox cycling. Treatment with menadione, which has a redox cycling activity, increased oxygen consumption and superoxide anion production inside cells and decreased intracellular levels of GSH, leading to hepatotoxicity (Thor et al., 1982). When rat primary hepatocytes were treated with the redox cycling quinone, 2,3-dimethoxy-1,4-naphthoquinone (DMNQ), intracellular levels of GSH were decreased and levels of GSSG were increased, indicating oxidative stress induction, followed by cell death (Gant et al., 1988). In the liver, one-electron reduction between quinone and semiquinone radical is catalyzed mainly by flavoenzymes such as NADPH–cytochrome P450 reductase and NADH– cytochrome b5 reductase (Berlin and Haseltine, 1981; Kappus and Sies, 1981; Powis and Appel, 1980). These reductases use NAD(P)H as an electron donor to reduce quinones. The rate of one-electron reduction of quinones by these reductases is not considered to be associated with their structural profiles or lipid solubility but with the one-electron redox potential of these reductases. Two-electron reduction of quinones, which is mediated by NAD(P)H:quinone oxidoreductase 1 (DT-diaphorase), leads to hydroquinone formation (Danson et al., 2004; Ross and Siegel, 2004). Reduction of quinones by NAD(P)H:quinone oxidoreductase 1 may constitute a detoxification process since rapid reduction of the quinone to the hydroquinone would circumvent one-electron processes. Furthermore, hydroquinones may be conjugated with glucuronic acid or sulfate to form unreactive and water-soluble substances that are readily eliminated from the body. In fact, inhibition of NAD(P)H:quinone oxidoreductase 1 by dicoumarol enhanced hepatotoxicity induced by treatment with quinones (Keyes et al., 1985; Tsuda, 1990). 10.2.4
The Relative Contribution of Quinone-Induced Arylation/Alkylation or Redox Cycle to Hepatotoxicity
Quinone toxicity depends on two mechanisms, i.e. their arylating/alkylating or redoxcycling abilities. The contribution of arylating/alkylating and/or redox-cycling abilities
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of quinones to cytotoxicity was reported to be a function of the chemical structure or the redox potential of the quinone (Watanabe et al., 2004), and the contribution could be examined experimentally. The idea is as follows: arylation/alkylation products of thiols were the results of the reaction with quinones, and they could not be recovered to the original protein thiols by treatment with a reductant. ROS that resulted from redox cycling, especially hydrogen peroxide, react with the thiolate anion, which is in equilibrium with protein thiols to initially produce the sulfenic acid and then the disulfide. These oxidized products are rereduced to the original thiols in the presence of a reductant such as dithiothreitol. Therefore, we can discriminate decreases in protein thiols due to direct arylationl/alkylation from that due to oxidative insults via the redox cycle by measuring the total loss of protein thiols and recoverable protein thiols upon addition of a reductant. Thus, loss of total protein thiols minus recoverable protein thiols equals the loss of protein thiols by arylating/alkylating (Bellomo et al., 1990; d’Arcy Doherty et al., 1987). However, it is difficult to evaluate the extent to which the reaction with protein thiols contributes to the cytotoxicity of quinones. Efforts were made to develop model quinones that have either arylation/alkylation ability or redox cycling ability to induce cytotoxicity. One of the representative compounds of the former is considered to be BQ or 1,4-naphthoquinone, and that of the latter is thought to be DMNQ. Henry and Wallace showed that cytotoxicity of arylating quinones BQ and 1,4-naphthoquinone was stronger than that of the redox cycling quinone DMNQ in primary rat hepatocytes (Henry and Wallace, 1996). Similar results were obtained by other groups (Miller et al.,1986; Schmieder et al., 2003; Seung et al., 1998; Toxopeus et al., 1993), indicating that hepatotoxicity of quinone compounds depends mainly on their arylating activities, and that the redox cycle is less involved in hepatotoxicity. Our results suggest that the primary toxicity of BQ is stronger than that of DMNQ in rat primary hepatocytes in a short-term exposure (Ishihara et al., 2006). However, a long-term exposure of hepatocytes to DMNQ induced oxidative stress sufficient to damage cells, leading to necrosis. In this regard, the redox cycle is considered to contribute a great deal to hepatotoxicity (Ishihara et al., 2006). More importantly, under cytochrome P450 inhibition, the hepatotoxicity of DMNQ, a redox-cycling quinone, became stronger than that of BQ, an arylating quinone. This is described in the next section.
10.3
A Role of Cytochrome P450 in Quinone-Induced Hepatotoxicity
The membrane-bound microsomal monooxygenase system is localized in the endoplasmic reticulum of most animal tissues. Although the highest content of microsomal monooxygenase components is found in hepatocytes, the system is present also in the lung, kidney, brain, lymphocytes, vascular smooth muscle, olfactory and intestinal epithelium (McKinnon and McManus 1996; Kapitulnik and Strobel 1999). Cytochrome P450, which is a heme-thiolate protein, is a major enzyme in the microsomal monooxygenase system. The predominant function of cytochrome P450 appears to be the oxygenation of exogenous compounds as well as some endogenous substrates (Gonzalez, 1988). A variety of substrates can be oxidized because of the existence of multiple cytochrome P450 isozymes with broad substrate selectivity. Most drugs are metabolized by cytochrome P450, resulting in production of water-soluble metabolites, which are then excreted from the body. Mono-oxygenation reactions typically require the input of two electrons that are transferred to cytochrome P450
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by the flavoprotein NADPH–cytochrome P450 reductase and/or NADH–cytochrome b5 reductase. Because cytochrome P450 is closely related to the side-effects of drugs, interaction of cytochrome P450 with various drugs is an important subject in toxicology (Wienkers and Heath, 2005; Wilkinson, 2005). However, quinone compounds are metabolized via hydroquinone production by NAD(P)H:quinone oxidoreductase 1, followed by forming conjugates. Therefore, cytochrome P450 is not directly involved in quinone metabolism. In fact, the relationship between cytochrome P450 and metabolism of the quinone-based anticancer agent doxorubicin (adriamycin) was examined in NADPH–cytochrome P450 reductase-deleted mice in which cytochrome P450 cannot function. When doxorubicin was administered to both wild-type and NADPH–cytochrome P450 reductase-deleted mice, no change in the concentration of doxorubicin in the blood was observed between the two types of mice. Therefore, this result supports the idea that cytochrome P450 is not involved in quinone metabolism (Henderson et al., 2006). However, by considering two points – (1) cytochrome P450 accepts an electron from intracellular reductases, and (2) redox-cycling quinones also receive an electron from them – cytochrome P450 might be involved in the hepatotoxicity of redox-cycling quinone. Therefore, we examined the role of cytochrome P450 in the toxicity of the redox-cycling quinone DMNQ using rat primary hepatocytes. When rat primary hepatocytes were treated with 60 μM DMNQ and incubated for 5 h, cell viability was decreased to 65 %. Treatment with DMNQ in combination with cytochrome P450 inhibitors SKF or ketoconazole decreased the viability to 30 or 48 %, respectively, indicating that DMNQ-induced hepatotoxicity was enhanced under cytochrome P450 inhibition. On the other hand, the arylating quinone BQ-induced hepatotoxicity was not affected by pretreatment with a cytochrome P450 inhibitor (Ishihara et al., 2006). Therefore, enhancement of hepatotoxicity by cytochrome P450 inhibition occurs only when a redox-cycling quinone, not an arylating quinone, is treated. More importantly, both superoxide anion production and NADPH oxidation induced by treatment with DMNQ were potentiated by pretreatment with a P450 inhibitor (Table 10.1), indicating strongly that the DMNQ-induced redox cycle is enhanced under P450 inhibition. On the basis of these findings, we offer a putative mechanism for the enhancement of DMNQ-induced hepatotoxicity under cytochrome P450 inhibition, as shown in Figure 10.2. In the liver, an electron from NADPH–cytochrome P450 reductase is transferred to cytochrome P450, which metabolizes many endogenous substrates or xenobiotics (Gonzalez, 1988), and NADPH–cytochrome P450 reductase catalyzes the one-electron reduction of quinones in redox cycles (Powis et al., 1980; Berlin et al., 1981). Thus, when redox-cycling quinones exist in hepatocytes, the electron flows from NADPH–cytochrome P450 reductase are considered to be additionally transferred to redox-cycling quinones; this leads to the production of semiquinone radicals, which are re-oxidized by giving electrons to molecular oxygen to yield the superoxide anion. This explains how DMNQ increased the NADPH oxidation and production of the superoxide anion. More importantly, the inhibition of cytochrome P450 by SKF or ketoconazole enhanced NADPH oxidation and superoxide production, and NADPH-cytochrome P450 reductase inhibition suppressed DMNQinduced and DMNQ + SKF-induced or DMNQ + ketoconazole-induced NADPH oxidation (Table 10.1). Our scenario for these findings is as follows. The inhibition of cytochrome P450 suppresses the electron transfer from the reductase to cytochrome P450; thus, the reductase donates electrons only to quinones to produce semiquinone radicals, resulting in
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Table 10.1 Enhancement of DMNQ-induced redox cycling and cell death under cytochrome P450 inhibition (details of this procedure are described else where (Ishihara et al., 2006)). a,e
Conditions
Untreated DMNQ DMNQ + SKF DMNQ + ketoconazole
Cell viabilityb,e Superoxide production NADPH oxidation (%) (nmol/min/106 cells)c,e (nmol/min/mg protein)d,e 91.2 ± 1.2 64.6 ± 2.7 30.3 ± 4.2 f 47.6 ± 4.9g
14.4 ± 0.4 16.4 ± 0.5 18.3 ± 0.5g 18.1 ± 0.4g
6.8 ± 1.2 46.6 ± 3.2 77.4 ± 5.8 f 66.9 ± 4.2 f
a
Primary rat hepatocytes were pretreated with SKF (30 μM) or ketoconazole (30 μM) for 20 min and then incubated in the presence or in the absence of DMNQ (60 μ M). Cell viability was estimated as the percentage of LDH activity in the medium after incubation for 5 h. c The amount of superoxide anion radical produced by hepatocytes was quantified by a method using acetylated cytochrome c after 10 min incubation. d NADPH oxidation in rat liver microsomes was measured by changes in absorbance at 340 nm when DMNQ or DMNQ + cytochrome P450 inhibitors was added. e Values are given as the mean ± SE of five separate experiments. Data were analyzed by using the Student’s t-test, with Holm’s corrections for multiple comparisons. P values less than 0.05 were considered statistically significant. f P < 0.01 versus DMNQ-treated hepatocytes. g P < 0.05 versus DMNQ-treated hepatocytes. b
further increases in NADPH oxidation and production of superoxide anion (Figure 10.2). Namely, the quinone redox cycle is enhanced by cytochrome P450 inhibition. This scenario is supported by data showing that an NADPH–cytochrome P450 reductase inhibitor, diphenyleneiodonium chloride, suppressed NADPH oxidation induced by treatment with both DMNQ and DMNQ + SKF or DMNQ + ketoconazole (Ishihara et al., 2006). Many anticancer, antimalarial or antibacterial agents have quinone structures. These drugs have high therapeutic efficacy, but have the ability to induce cytotoxicity. To date, general considerations of quinone-induced cytotoxicity are as follows: (1) cytochrome P450 contributes less to quinone metabolism (detoxification), and (2) the toxicity of arylating quinones is greater than that of redox-cycling quinones. On the basis of these considerations, it is natural that quinone compounds having less arylating activity are required for developing less-toxic drugs. However, our results indicate that the toxicity of redox-cycling quinones is enhanced under cytochrome P450 inhibition. In addition, the hepatotoxicity of DMNQ under cytochrome P450 inhibition was much higher than that of BQ at the same concentration. In this regard, careful consideration of the redox-cycling ability in terms of hepatotoxicity must be taken when quinone-based drugs are used or developed.
10.4
Concluding Remarks
The hepatotoxicity of quinone compounds could be induced by two mechanisms, i.e. arylation/alkylation and the redox cycle. To understand the mechanisms of hepatotoxicity of quinones is to avoid or alleviate their hepatotoxicity. The hepatotoxicities of arylating quinones are generally recognized to be higher than that of redox-cycling quinones. However, the hepatotoxicity of redox-cycling quinones is potentially higher than that of arylating quinones under cytochrome P450 inhibition. Our proposal is that we should
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(i) Normal state Redox-cycling quinone NAD(P)H
NADPH–cytochrome P450 reductase NADH–cytochrome b5 reductase
Cytochrome P450
(ii) Under cytochrome P450 inhibition Redox-cycling quinone NAD(P)H
NADPH–cytochrome P450 reductase NADH–cytochrome b5 reductase
Cytochrome P450 SKF Ketoconzole Electron flow
Figure 10.2 Enhancement of quinone hepatotoxicity under cytochrome P450 inhibition: a putative mechanism. In the liver, cytochrome P450 accepts an electron from NADPH– cytochrome P450 reductase or NADH–cytochrome b 5 reductase. When redox-cycling quinones are used under conditions where cytochrome P450 is operating functionally, NADPH–cytochrome P450 reductase or NADH-cytochrome b 5 reductase supplies electrons to both cytochrome P450 and redox-cycling quinones. In other words, quinones compete with cytochrome P450 for electron flow from the reductases. However, under cytochrome P450 inhibition, reductases are not able to transfer an electron to cytochrome P450 and, thus, they donate an electron only to redox-cycling quinones, forming semiquinone radicals. As a result, the quinone redox cycle can be potentiated by cytochrome P450 inhibition
not underestimate the hepatotoxicity of redox cycling quinones if we use them together with cytochrome P450 inhibitors, regardless of whether they are drugs. Therefore, careful consideration for quinone-based drugs will help to avoid hepatotoxicity. In addition, the strong toxicity of quinone-based drugs might be alleviated by therapy in combination with other drugs if the toxic mechanism is known. One example of this is the cardiotoxicity of doxorubicin, which is an anticancer agent with a quinone structure, being attenuated by pretreatment with antioxidants (Lenzhofer et al., 1983). We stress the mechanism of the redox-cycling quinone under the condition of P450 inhibition, which is thought to be readily attainable, and thus arouses concern in the case of managing quinone compounds.
10.5
Acknowledgements
We thank Dr M. Iwasaki for critical reading of this manuscript.
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Abbreviations
BQ – 1,4-benzoquinone DMNQ – 2,3-dimethoxy-1,4-naphthoquinone GPx – glutathione peroxidase GSH – reduced glutathione GSSG – glutathione disulfide Prx – peroxiredoxin ROS – reactive oxygen species SOD – superoxide dismutase
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11 A Mechanistic View of Troglitazone Hepatotoxicity Rawiwan Maniratanachote and Tsuyoshi Yokoi
11.1
Introduction
Thiazolidinediones (Figure 11.1) are a class of oral antidiabetic agents and are the synthetic ligands for the peroxisome proliferator-activated receptor γ (PPAR γ ) (Lehmann et al., r , (±)-5-[[4-[(3,4-dihydro-6-hydroxy-2,5,7,8-tretramethyl1995). Troglitazone (Rezulin 2H-1-benzopyran-2-yl)methoxy]phenyl]methyl]-2,4-thiazolidinedione) was the first thiazolidinedione antidiabetic agent approved for clinical use by the US Food and Drug Administration in 1997. Troglitazone lowers the blood glucose levels through increased glucose uptake by skeletal muscle, decreased hepatic glucose production and increased insulin sensitivity of the target tissue in animal models of metabolic impairment (Ciaraldi et al., 1990; Fujiwara et al., 1988, 1995). These pharmacological effects are exerted through PPAR γ -dependent transcription of genes involved in glucose and lipid metabolism and energy homeostasis (Lehmann et al., 1995; Saltiel and Olefsky, 1996; Spiegelman, 1998). Based on the pharmacological advantages and the apparent absence of severe toxic effects, troglitazone was thought likely to become a promising treatment for type II diabetes mellitus in patients with insulin resistance. However, in the combined North American clinical trials, elevations of serum alanine aminotransferase more than three times the upper limit of normal were observed in 48 out of 2510 patients (1.9 %) treated with troglitazone. Liver biopsies from two patients confirmed the hepatocellular nature of the injury as an idiosyncratic drug reaction (Watkins and Whitcomb, 1998). Meanwhile, troglitazone had been concomitantly reported to be associated with idiosyncratic hepatotoxicity, with some patients showing severe or fatal
Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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H3C
CH3 O
H3C
O S NH
O S NH
HO3SO
ST1A3
CH3
HO CH3
CH3
CH3
Troglitazone sulfate (M1)
UGT
Troglitazone
CH3 O
H3C
O N
O NH
NH CH3
HO HO
Troglitazone glucuronide (M2)
O
Rosiglitazone N
S
O O OH
S
CH3
O
COOH
CYP3A4 CYP2C8
N
CH3
CH3
O
H3C
O
O
O OH
NH H 3C
H3C
S
S
NH
O O
Pioglitazone
Figure 11.1
CH3
Troglitazone quinone (M3)
Structures of thiazolidinediones and pathways of troglitazone metabolism
liver damage (Gitlin et al., 1998; Neuschwander-Tetri et al., 1998; Shibuya et al., 1998). Consequently, it was withdrawn from the market in the USA and Japan in March 2000. The hepatotoxic effects of troglitazone were not predicted from conventional animal models (Watanabe et al., 1999) or in cynomolgus monkeys (300–1200 mg/kg/day for 52 weeks), a primate model having similar metabolic profiles to humans (Rothwell et al., 2002). Two other thiazolidinediones, which are now on the market, rosiglitazone and pioglitazone, have been introduced in 1999 and are unlikely to share the hepatotoxic effects of troglitazone (Freid et al., 2000; Isley and Oki, 2000; Lebovitz et al., 2002). It should also be noted that the clinical dosage regimen for improvement of fasting glucose is distinguishable among these thiazolidinediones (Table 11.1). The recommended dose for troglitazone was 200 to 600 mg/day, for rosiglitazone 4 to 8 mg/day and for pioglitazone 15 to 45 mg/day (Hanefeld, 2001; Loi et al., 1999; PDR, 1999, 2005a,b). The dosage requirement for their efficacy might have reflected their hepatotoxic potential. This review summarizes the molecular mechanism of troglitazone hepatotoxicity from studies both in vivo and in vitro. Even though, there is no direct evidence to indicate the precise mechanism of the toxicity so far. Many points of view, however, have been proposed to contribute to the toxic effects of troglitazone.
11.2
Potential of Troglitazone Metabolites for Hepatotoxicity
In humans, troglitazone is predominantly metabolized by three pathways: sulfation, glucuronidation and oxidation to form a sulfate conjugate (M1), a glucuronide conjugate (M2) and a quinone metabolite (M3), respectively (Figure 11.1). M1 and M3 are the major metabolites in plasma, while M2 is a minor metabolite (Izumi et al., 1997a,b; Kawai et al.,
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Pharmacokinetic parameters of thiazolidinediones
Parameter
Troglitazone (1997–2000)a
Rosiglitazone (1999–present)a
Pioglitazone (1999–present)a
Oral dosage (mg/day) Plasma protein binding (%) Absolute bioavailability (%) Cmax (μg/ml) AUC (μg-h/ml) tmax (h) Plasma elimination half-life (h) Biliary excretion (%) Urinary excretion (%) Effects of food
200–600 >99 40–50 0.90–2.82 7.4–22.1 <2–3 16–34 85 3 Increases the extent of absorption by 30 to 80 %
1–8 99.8 99 0.076–0.598 0.358–2.971 1.75 3–4 23 64 Decrease in Cmax by 28 % and delay in tmax
15–45 >99 83 1.4 ± 0.2b 11.6 ± 2.2b <2 3–7 NAc 15–30 Slight delay in tmax to 3–4 h
a
Year on clinical application. R , Takeda Pharmaceutical Company Ltd, Japan). For 30 mg/day oral dosage (data from the product leaflet of Actos c Not applicable – most of the oral dose is excreted into the bile. b
1998; Loi et al., 1999). The main metabolite, troglitazone sulfate (M1), is formed by the action of phenol sulfotransferase, ST1A3 (Honma et al., 2002). It accounts for about 70 % of the metabolites detected in human plasma (Loi et al., 1999). Differing from other thiazolidinediones, troglitazone contains a 6-hydroxy-5,7,8trimethylchromane moiety (a chroman ring of vitamin E). This structure accounts for the effective antioxidant property of troglitazone and suggests an advantage in preventing diabetic vascular complications in addition to its hypoglycemic and hypolipidimic effects (Inoue et al., 1997). This structure, however, has the potential to undergo metabolic activation to form troglitazone quinone or metabolite M3 by cytochrome P450s (CYPs) 3A4 and 2C8 (Yamazaki et al., 1999). In humans, it is likely that CYP3A4 is primarily responsible for this reaction (He et al., 2001). In addition, troglitazone has been shown to induce CYP3A in human and rat hepatocytes, which stimulates the formation of the quinone (Ramachandran et al., 1999; Sahi et al., 2000). By the action of CYP3A, troglitazone yields several reactive intermediates in rats (Kassahun et al., 2001; Tettey et al., 2001) (Figure 11.2). In vitro, the formation of an epoxide of troglitazone quinone was also identified (Yamamoto et al., 2002). It is known that quinones represent a class of toxicological intermediates, which can result in acute cytotoxicity and immunotoxicity as well as carcinogenesis (Bolton et al., 2000). The maximum plasma concentrations in patients taking troglitazone at the dosage of 600 mg/day reached to only about 2.82 μg/ml or 6.3 μM (Loi et al., 1999). However, a study in rats demonstrated that the concentration of troglitazone in liver tissues was 10–12 fold higher than that in the plasma (Sahi et al., 2000). Therefore, the troglitazone levels in human liver might allow the formation of these putative reactive intermediates and their accumulation may lead to the hepatotoxicity (see next section). A relatively minor metabolite, troglitazone glucuronide (M2), is catalyzed by UGT (Yoshigae et al., 2000). The glucuronidation of troglitazone in human intestine is 3-fold higher than that in human liver. In the liver, the reaction is likely mediated by UGT1A1,
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H 3C
CH3 R1
.O
O R2
Phenoxy redical intermediate
NH
S O
CH3
O
Sulfoxide intermediate and radical
CH3 O
H 3C
CH3 R1
O R2
o-Quinone methide
. NH
S
O CH2
O
CH3
O
H 3C
O
H3C
R1 OH
R2 N C
Quinone epoxide HO
O H 3C
O
-Ketoisocyanate derivative
S
O
CH3
O O S
R1 =
O
H 3C NH
CH3 O
R2 = HO CH3
Figure 11.2
Reactive metabolites of troglitazone catalyzed by CYP3A
while in the intestine it is mediated by UGT1A8 and UGT1A10 (Watanabe et al., 2002). A polar, partially β-glucuronidase-sensitive metabolite with retention properties similar to M2 was found in the profiling of urine samples (Loi et al., 1999). There has been no reported evidence that M2 is responsible for the hepatotoxic effects.
11.3
Susceptible Genetic Factors Associated with Troglitazone Hepatotoxicity
As mentioned above, troglitazone can undergo metabolic biotransformation by CYP3A4 to form a quinone metabolite (M3) and epoxide specie (Izumi et al., 1997a,b; Kawai et al., 1998; Loi et al., 1999; Yamamoto et al., 2002). Quinones can react readily with sulfur nucleophiles such as glutathione (GSH) or cysteine residues on proteins (Bolton et al., 2000). The toxic effects of troglitazone have been thought to be mediated by the depletion of GSH, covalent binding to cellular macromolecules or oxidative stress. In cryopreserved-human hepatocytes, large variations in the sensitivity to troglitazone were observed and sensitive donors were demonstrated to form significantly lower amounts of GSH conjugates and glucuronides than resistant donors (Kostrubsky et al., 2000; Prabhu et al., 2002). It is known that the GSH conjugation is formed by the action of glutathione S-transferase (GST). A study in rats has shown that GSH adducts of troglitazone are formed and the reaction is enhanced by CYP3A (Tettey et al., 2001). An epoxide of troglitazone quinone catalyzed by CYP3A4 might also be eliminated by GSTs and epoxide hydrolase (Yamamoto et al.,
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2002). These findings indicate an association between metabolic activation by CYP and detoxification by GSTs. In a key report concerning this aspect, Watanabe et al. (2003) investigated the genetic factors responsible for troglitazone hepatotoxicity in vivo, in humans. Among one hundred and ten patients prescribed troglitazone, 25 case patients had an abnormal increase in ALT or AST levels to at least 9 times or 5 times the upper limit of the normal range, respectively, while 85 control patients showed no significant increase in the ALT levels during more than 6 months of treatment. Interestingly, they found that this abnormal elevation of liver enzymes caused by troglitazone treatment was highly associated with the double null genotype of GSTM1 and GSTT1 (Watanabe et al., 2003). Hence, interindividual differences in the detoxification ability might contribute to the susceptibility and individual risk for troglitazone hepatotoxicity. However, the complete mechanism is still largely unknown.
11.4
Implications of Canalicular Bile Salt Export Pump and Drug Transporters
The main metabolite of troglitazone, troglitazone sulfate, undergoes biliary excretion and accounts for up to 60 % of the dose in rats (Kawai et al., 1997). In patients with hepatic impairment, troglitazone sulfate was found to accumulate about 4-fold in plasma with a 3-fold increased half-life (Ott et al., 1998). This metabolite also showed an inhibition effect on the canalicular bile salt export pump (Bsep) as well as drug transporters, suggesting it contributes to the liver toxicity. Funk et al. (2001a,b) reported that troglitazone sulfate inhibits the ATP-dependent taurocholate transport mediated by Bsep in isolated canalicular rat liver plasma membrane (IC50 0.4–0.6 μM) about 10 times more strongly than the parent compound (IC50 3.9 μM). The inhibition of Bsep suggests it is one of the possible factors contributing to the hepatotoxicity since the subsequent accumulation of bile salts may lead to intrahepatic cholestasis in humans. Previously, cholestatic signs have also been described in a patient with troglitazone hepatotoxicity (Gitlin et al., 1998). Another group of researchers reported that troglitazone sulfate is transported by organic anion transporting polypeptide (OATP) transporters with higher affinity to OATP-C than OATP8, and showed a strong inhibitory effect on estrone-3-sulfate transport by these transporters (Nozawa et al., 2004). Both OATP-C and OATP8 are members of the organic anion transporting polypeptides, which are expressed in the basolateral membrane of hepatocytes (Hagenbuch and Meier, 2003; Kreb, 2006). They play important roles in the hepatic handling of endogenous compounds and xenobiotics. Therefore, any factors that affect or impair the OATP-C levels or activity, for example, genetic polymorphisms (Kreb, 2006), may cause an accumulation of M1, leading to troglitazone hepatotoxicity.
11.5
Hypersensitivity Reaction in Troglitazone Hepatotoxicity
Idiosyncratic toxicity is generally considered to be host-dependent, dose-independent, difficult to reproduce in experimental animals and relatively uncommon. Some idiosyncratic drug reactions are due to a metabolic abnormality of the host, but many have an
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immunological basis or result from immune-mediated hypersensitivity (Pohl et al., 1988). A well-characterized example is the idiosyncratic hepatitis induced by halothane. Sera of these patients contain autoantibodies directed against some trifluoroacetyl (TFA)–protein adducts, including protein disulfide isomerase, microsomal carboxyesterase, calreticulin, ERp-72, GRP-78 and GRP-94 (review in Gut et al., 1993), as well as CYP2E1 (Bourdi et al., 1996). Our recent report described that aldolase B, which is an enzyme predominantly localized in the liver and kidney (Penhoet et al., 1966; Rutter, 1964), was detected as an autoantigen that reacted with antibodies in the sera from two patients with type II diabetes mellitus with troglitazone-induced liver dysfunction (Maniratanachote et al., 2005b). The titer of anti-aldolase B remained high for several weeks after stopping troglitazone administration. This finding supported the idea that troglitazone hepatotoxicity may have an immunological basis. However, autoantibodies to aldolase B were also detected in the sera of patients with chronic hepatitis as well as liver cirrhosis (Brown et al., 1987; Maniratanachote et al., 2005b). At present, a definitive explanation for the occurrence of aldolase B autoantibodies, whether it is the cause or consequence of the progression of hepatotoxicity, is still lacking. There are several reactive metabolites generated by troglitazone (Figure 11.2) (Kassahun et al., 2001; Tettey et al., 2001; Yamamoto et al., 2002). Aldolase B, which is an enzyme predominantly localized in the liver (Penhoet et al., 1966; Rutter, 1964), may be one of the target proteins that interact with those reactive species and trigger the immune response. Further investigation may provide better understanding of this mechanism.
11.6
Molecular Mechanism of Troglitazone-Induced Liver Toxicity
Troglitazone has been shown to induce apoptosis in various hepatic (Bae and Song, 2003; Tirmenstein et al., 2002; Yamamoto et al., 2001) and non-hepatic (Shiau et al,. 2005) cell types depending on the concentration and time of exposure. Unlike its pharmacological effects, the toxicity of troglitazone seems to be a PPARγ -independent mechanism and the higher affinity PPARγ agonists such as rosiglitazone possess much lower toxic effects (Lehmann et al., 1995; Shiau et al,. 2005). In addition, Shiau et al. (2005) demonstrated that a synthetic counterpart of troglitazone, which lacks PPARγ activation activity, was also able to induce apoptosis in cultured cells. As mentioned above, troglitazone can generate the main metabolites M1 and M3 via the action of ST1 and CYP2C8/CYP3A4, respectively. In in vivo experiments in rats, M1 showed the potential to inhibit Bsep, suggesting it is one of the factors contributing to cholestasis in humans (Funk et al., 2001a,b). In addition, the troglitazone quinone metabolite M3 has been suggested to be associated with troglitazone hepatotoxicity in humans (Neuschwander-Tetri et al., 1998). However, these metabolites showed fewer toxic effects compared to the parent compound, troglitazone, when mammalian hepatocytes and hepatoma cell lines were treated directly (Honma et al., 2002; Kostrubsky et al., 2000; Tettey et al., 2001; Tirmenstain et al., 2002; Yamamoto et al., 2001, 2002). It is most likely that troglitazone causes hepatic cell death via apoptosis. Apoptosis is a normal physiologic form of cell death and plays a prominent role in liver pathogenesis such as autoimmune liver diseases, viral hepatitis, and drug-induced hepatitis. In this regard, we summarize below the molecular responses to troglitazone toxicity in the cells (Figure 11.3).
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305
P
PKR
eIF2 Protein translation
ER Lumen
P
Unfolded proteins
p-38
Ca 2+
Str
ess
BiP
P
Chaperone genes BiP
+
P0
P0
P
c-Jun
sig
Ribosome P
P
JNK
nal Troglitazone
P
Bid
tBid tBid
P
?
Bad Bak/Bax Mitochondrion
Protein translation
Bcl-2
Nucleus
Apoptosis
Figure 11.3
11.6.1
Caspase-3
Cytochrome c
Schematic representation of the effects of troglitazone in liver cells
MAPK-Mediated Cell Death Pathway
The three well-characterized subfamilies of mitogen-activated protein kinases (MAPKs), extracellular signal-regulated kinase (Erk), c-Jun N-terminal kinase (JNK) and p38, are regulated by phosphorylation and play important roles in a variety of cellular process including growth, differentiation and apoptosis (Johnson and Lapadat, 2002). Erk is generally activated by mitogens, while JNK and p38 are preferentially activated by stress and inflammatory cytokines. The most obvious effect of troglitazone on apoptosis is likely via the promotion of JNK, which in turn activates c-Jun by phosphorylation as well as by activation of p38 (Bae and Song, 2003). Gardner et al. (2005) reported that calcium/calmodulindependent kinase II (CaMKII) is a critical upstream activator of p38 phosphorylation in GN4 cells. In addition, troglitazone also causes the induction of Bax, Bad, the cleavage of Bid and release of cytochrome c. However, troglitazone showed a negligible effect on Erk (Bae and Song, 2003; Gardner et al., 2003, 2005). JNK is characterized as a stress-activated protein kinase based on its activation in response to the inhibition of protein synthesis. We will provide the additional information supporting this point in a later section.
11.6.2
Impairment of Mitochondrial Functions
Another obvious mechanism of troglitazone-induced toxicity in liver cells is by causing a reduction of the mitochondrial membrane potential with a concomitant depletion of the cellular ATP concentration (Bova et al., 2005; Tirmenstein et al., 2002). Subsequently, it increases the plasma membrane permeability and calcium ion (Ca2+ ) efflux. The result of these effects on mitochondria is the release of cytochrome c into the cytoplasm and activation of the caspases leading to apoptosis (Bova et al., 2005).
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Induction of Cell Cycle Arrest
Cyclin-dependent kinases (CDKs) are serine–threonine protein kinases that regulate the cell cycle progression. These kinases are activated by various cyclins, inhibited by natural inhibitors such as p21, p27 and p18 and are tightly controlled by transcriptional and posttranscriptional modifications (Sherr and Roberts, 1999). Bae et al. (2003) reported that troglitazone-induced cell cycle arrest by this pathway and that apoptosis of hepatoma cell lines was caused an elevation of the levels of p53 and its downstream proteins, Gadd45, p21 and p27, as well as by a reduction in the levels of cyclin D1 and phospho-Rb.
11.7
Effect of Troglitazone on the Inhibition of Protein Translation
The endoplasmic reticulum (ER) is a major site of protein synthesis and its inside, or lumen, is a major site of protein folding (Gething and Sambrook, 1992). In mammalian cells, naturally the rate of protein synthesis is rapidly down-regulated following the induction of apoptosis. The phosphorylation of eukaryotic initiation factor 2α (eIF2 α) is important in the regulation of the selective translation during ER stress and the unfolded protein response (Holcik and Sonenberg, 2005). Troglitazone was shown to promote Ca2+ release from the ER leading to PERK and PKR activation, phosphorylation of eukaryotic initiation factor 2α(eIF2α), translation inhibition and growth arrest (Figure 11.3) (Fan et al., 2004; Gardner et al., 2005; Palakurthi et al., 2001). A study using PPARγ −/− and PPARγ +/+ mouse embryonic stem cells suggested that these effects were PPARγ -independent (Palakurthi et al., 2001). It is known that the ER is a major cellular storage site of Ca2+ in the cell, and ER chaperones play important roles in Ca2+ accumulation and release. Any disturbance in the ER homeostasis causes a release of Ca2+ , which in turn blocks ER protein processing, resulting in the accumulation of incompletely folded proteins, and activates the transcription of ER chaperone genes (Liu et al., 1998; Lodish and Kong, 1990). We found that the immunoglobulin heavy-chain binding protein (BiP), an abundant chaperone protein in the ER, was overexpressed in hepatoma cell lines by troglitazone treatment (Maniratanachote et al., 2005a). The important role of this chaperone protein was indicated by the phenotypic change in cell viability when BiP expression was inhibited by small interference RNA (Maniratanachote et al., 2005a). This condition rendered cells more susceptible to the toxic effects of troglitazone. Collectively, it might be postulated that troglitazone acts as a chemical stress signal that causes the release of Ca2+ from the ER, and that BiP expression is one of the cellular defense mechanisms of the ER in response to troglitazone-induced toxicity (Figure 11.3). With respect to the inhibition of translation by troglitazone toxicity, we recently found that the ribosomal protein P0 (P0) is also one of the targets of troglitazone cytotoxicity in HepG2 cells (Maniratanachote et al., r r r r). P0 is known as a phosphoprotein that functions in some processes of protein translation (Gonzalo et al., 2001). It was found that, rather than its overexpression, dephosphorylation of P0 occurred in troglitazone-induced cytotoxicity (Maniratanachote et al., r r r r). Therefore, dephosphorylation of P0 may play a role in the regulation of protein translation in response to the toxic effects of troglitazone (Figure 11.3).
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Mechanistic View of Troglitazone Hepatotoxicity
11.8
307
Conclusions and Perspectives
Troglitazone is a drug that can cause idiosyncratic hepatotoxicity in human. This kind of toxicity is usually unpredictable, pharmacologically independent, rare and not reproducible in experimental animal models, which makes it difficult to study (Lee, 2003). A number of toxicological tests, both in vivo and in vitro, have been performed. So far, no direct mechanism has been found that can explain why troglitazone hepatotoxicity occurred in only some individuals. The failure to develop such toxicity in animal models is still inexplicable. However, we have learned from previous reports that the mechanism of troglitazone hepatotoxicity is PPARγ -independent, the molecular mechanisms of apoptotic cell death are most likely involved in the hepatotoxicity and the idiosyncratic hepatotoxicity might be a consequence of a genetic basis in susceptible individuals. Recent findings concerning the miRNA functions in specific tissues has enabled better understanding of the molecular mechanisms of various pathologies and diseases (review in Bartel, 2004). Among several hundreds miRNAs, miR-122 is the most abundant and liverspecific (Lagos-Quintana et al., 2002; Baskerville and Bartel, 2005). It has been shown to have various roles, for example, in hepatitis C viral infection (Jopling et al., 2005) and in lipid metabolism (Esau et al., 2006). Therefore, studies on miRNAs and their targets might reveal clues concerning troglitazone idiosyncratic hepatotoxicity.
Acknowledgments We thank Mr Brent Bell for reviewing the manuscript.
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Honma, W., Shimada, M., Sasano, H., Ozawa, S., Miyata, M., Nagata, K., Ikeda, T. and Yamazoe, Y. (2002). Phenol sulfotransferase, ST1A3, as the main enzyme catalyzing sulfation of troglitazone in human liver. Drug Metab. Dispos. 30: 944–952. Inoue, I., Katayama, S., Takahashi, K., Negishi, K., Miyazaki, T., Sonoda, M. and Komoda, T. (1997). Troglitazone has a scavenging effect on reactive oxygen species. Biochem. Biophys. Res. Commu. 235: 113–116. Isley, W. L. and Oki, J. C. (2000). Rosiglitazone and liver failure. Ann. Intern. Med. 133: 393. Izumi, T., Enomoto, S., Hoshiyama, K., Sasahara, K. and Sugiyama, Y. (1997a). Pharmacokinetic stereoselectivity of troglitazone, an antidiabetic agent, in the KK mouse. Biopharm. Drug Dispos. 18: 305–324. Izumi, T., Hoshiyama, K., Enomoto, S., Sasahara, K. and Sugiyama, Y. (1997b). Pharmacokinetics of troglitazone, an antidiabetic agent: Prediction of in vivo stereoselective sulfation and glucuronidation from in vitro data. J. Pharmacol. Exp. Ther. 280: 1392–1400. Johnson, G. L. and Lapadat, R. (2002). Mitogen-activated protein kinase pathways mediated by ERK, JNK and p38 protein kinases. Science 298: 1911–1912. Jopling, C. L., Yi, M., Lancaster, A. M., Lemon, S. M. and Sarnow, P. (2005). Modulation of hepatitis C virus RNA abundance by a liver-specific microRNA. Science 309: 1577–1581. Kassahun, K., Pearson, P. G., Tang, W., McIntosh, I., Leung, K., Elmore, C., Dean, D., Wang, R., Doss, G. and Baille, T. A. (2001). Studies on the mechanism of troglitazone to reactive intermediates in vitro and in vivo. Evidence for novel biotransformation pathways involving quinone methide formation and thiazolidinedione ring scission. Chem. Res. Toxicol. 14: 62–70. Kawai, K., Kawasaki-Tokui, Y., Odaka, T., Tsuruta, F., Kazui, M., Iwabuchi, H., Nakamura, T., Kinoshita, T., Ikeda, T., Yoshioka, T., Komai, T. and Nakamura, K. (1997). Disposition and metabolism of the new oral antidiabetic drug troglitazone in rats, mice and dogs. Arzneimittelforschung 47: 356–368. Kawai, K., Odaka, T., Tsurata, F., Tokui, T., Ikeda, T. and Nakamura, K. (1998). Stereoselective metabolism of new oral anti-diabetic agent troglitazone stereoisomers in liver. Xenobio. Metab. Dispos. 13: 362–368. Kostrubsky, V. E., Sinclair, J. F., Ramachandran, V., Venkataramanan, Wen, Y. H., Kindt, E., Galchev, V., Rose, K., Sinz, M. and Strom, S. C. (2000). The role of conjugation in hepatotoxicity of troglitazone in human and porcine hepatocyte cultures. Drug Metab. Dispos. 28: 1192–1197. Kreb, R. (2006). Implications of genetic polymorphism in drug transporters for pharmacotherapy. Cancer Lett. 234: 4–33. Lagos-Quintana, M., Rauhut, R., Yalcin, A., Meyer, J., Lendeckel, W. and Tuschl, T. (2002). Identification of tissue-specific microRNAs from mouse. Current Biol. 12: 735–739. Lebovitz, H. E., Kreider, M. and Freed, M. I. (2002). Evaluation of liver function in type 2 diabetic patients during clinical trials. Diabetes Care 25: 815–821. Lee, W. M. (2003). Drug-induced hepatotoxicity. New Engl. J. Med. 349: 474–485. Lehmann, J. M., Moore, L. B., Smith-Oliver, T. A., Wilkison, W. O., Willson, T. and Kliewer, S. A. (1995). An antidiabetic thiazolidinedione is a high affinity ligand for peroxisome-activated receptor γ (PPARγ ). J. Biol. Chem. 270: 12953–12956. Liu, H., Miller, E., van de Water, B. and Stevens, J. L. (1998). Endoplasmic reticulum stress proteins block oxidant-induced Ca2+ increases and cell death. J. Biol. Chem. 273: 12858–12862. Lodish, H. F. and Kong, N. (1990). Perturbation of cellular calcium blocks exit of secretory proteins from the rough endoplamic reticulum. J. Biol. Chem. 265: 10893–10899. Loi, C. M., Young, M., Randinitis, E., Vassos, A. and Koup, J. R. (1999). Clinical pharmacokinetics of troglitazone. Clin. Pharmacokinet. 37: 91–104. Maniratanachote, R., Minami, K., Katoh, M., Nakajima, M. and Yokoi, T. (2005a). Chaperone proteins involved in troglitazone-induced toxicity in human hepatoma cell lines. Toxicol. Sci. 83: 293– 302.
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Maniratanachote, R., Shibata, A., Kaneko, S., Yamamori, I., Wakasugi, T., Sawazaki, T., Katoh, K., Tokudome, S., Nakajima, M. and Yokoi, T. (2005b). Detection of autoantibody to aldolase B in sera from patients with troglitazone-induced liver dysfunction. Toxicology 216: 15–23. Maniratanachote, R., Minami, K., Katoh, M., Nakajima, M. and Yokoi, T. (2006). Dephosphorylation of ribosomal protein P0 in response to troglitazone-induced cytotoxicity. Toxicol. Lett. 166: 189– 199. Neuschwander-Tetri, B. A., Isley, W. L., Oki, J. C., Ramrakhiani, S., Quiason, S. G., Phillips, N. J. and Brunt, E. M. (1998). Troglitazone-induced hepatic failure leading to liver transplantation. Ann. Intern. Med. 129: 38–41. Nozawa, T., Sugiura, S., Nakajima, M., Goto, A., Yokoi, T., Nezu, J., Tsuji, A. and Tamai, I. (2004). Involvement of organic anion transporting polypeptides in the transport of troglitazone sulfate: Implications for understanding troglitazone hepatotoxicity. Drug Metab. Dispos. 32: 291–294. Ott, P., Ranek, L. and Young, M. A. (1998). Pharmacokinetics of troglitazone, a PPAR-gamma agonist, in patients with hepatic insufficiency. Eur. J. Clin. Pharmacol. 54: 567–571. Palakurthi, S. S., Aktas, H., Grubissich, L. M., Mortensen, R. M. and Halperin, J. A. (2001). Anticancer effects of thiazolidinediones are independent of peroxisome proliferators-activated receptor γ and mediated by inhibition of translation initiation. Cancer Res. 61: 6213–6218. r , in Physician’s Desk Reference, 52nd Edition, Medical Economics Company, PDR (1999). Rezulin Inc., Montvale, NJ, USA, pp. 2310–2314. r , in Physician’s Desk Reference, 59th Edition, Thomson PDR, Montvale, NJ, PDR (2005a). Actos USA, pp. 3181–3185. r , in Physician’s Desk Reference, 59th Edition, Thomson PDR, Montvale, PDR (2005b). Avandia NJ, USA, pp. 1438–1443. Penhoet, E., Rajkumar, T. and Rutter, W. J. (1966). Multiple forms of fructose diphosphate aldolase in mammalian tissues. Proc. Natl. Acad. Sci. USA 56: 1275–1282. Pohl, L. R., Satoh, H., Christ, D. D. and Kenna, J. G. (1988). The immunologic and metabolic basis of drug hypersensitivity. Ann. Rev. Pharmacol. 28: 367–387. Prabhu, S., Fackett, A., Lloyd, S., McClellan, H. A., Terrell, C. M., Silber, P. M. and Li, A. P. (2002). Identification of glutathione conjugates of troglitazone in human hepatocytes. Chem. Biol. Interact. 142: 83–97. Ramachandran, V., Kostrubsky, V. E., Komoroski, B. J., Zhang, S., Dorko, K., Esplen, J. E., Strom, S. C. and Venkataramanan, R. (1999). Troglitazone increases cytochrome P-450 3A protein and activity in primary cultures of human hepatocytes. Drug Metab. Dispos. 27: 1194–1199. Rothwell, C., McGuire, E. J., Altrogge, D. M., Masuda, H. and de la Iglesia, F. A. (2002). Chronic toxicity in monkeys with the thiazolidinedione antidiabetic agent troglitazone. J. Toxicol. Sci. 27: 35–47. Rutter, W. J. (1964). Evolution of aldolase. Fed. Proc. 23: 1248–1257. Sahi, J., Hamilton, G., Sinz, M., Barros, S., Huang, S. M., Lesko, L. J. and LeCluyse, E. L. (2000). Effects of troglitazone on chromosome P450 enzymes in primary cultures of human and rat hepatocytes. Xenobiotica 30: 273–284. Saltiel, A. R. and Olefsky, J. M. (1996). Thiazolidinediones in the treatment of insulin resistance and type II diabetes. Diabetes 45: 1661–1669. Sherr, C. J. and Roberts, J. M. (1999). CDK inhibitors: Positive and negative regulators of G1-phase progression. Genes Dev. 13: 1501–1512. Shiau, C. W., Yang, C. C., Kulp, S. K., Chen, K. F., Chen, C.S., Huang, J. W. and Chen, C. H. (2005). Thiazolidinediones mediate apoptosis in prostate cancer cells in part through inhibition of Bcl-xl/Bcl-2 functions independently of PPARγ . Cancer Res. 65: 1561–1569. Shibuya, A., Watanabe, M., Fujita, Y., Saigenji, K., Kuwao, S., Takahashi, H. and Takeuchi, H. (1998). An autopsy case of troglitazone-induced fulminant hepatitis. Diabetes Care 21: 2140–2143.
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Spiegelman, B. M. (1998). PPAR-γ : Adipogenic regulator and thiazolidinedione receptor. Diabetes 47: 507–514. Tettey, J. N., Maggs, J. L., Rapeport, W. G., Pirmohamed, M. and Park, B. K. (2001). Enzyme induction dependent bioactivation of troglitazone and troglitazone quinone in vivo. Chem. Res. Toxicol. 14: 965–974. Tirmenstein, M. A., Hu, C. X., Gales, T. L., Maleeff, B. E., Narayanan, P. K., Kurali, E., Hart, T. K., Thomas, H. C. and Schwartz, L. W. (2002). Effects of troglitazone on HepG2 viability and mitochondrial function. Toxicol. Sci. 69: 131–138. Watanabe, T., Ohashi, Y., Yasuda, M., Takaoka, M., Furukawa, T., Yamoto, T., Sanbuissho, A. and Manabe, S. (1999). Was it possible to predict liver dysfunction caused by troglitazone during the nonclinical safety studies? Iyakuhin Kenkyu 30: 537–546. Watanabe, Y., Nakajima, M. and Yokoi, T. (2002). Troglitazone glucuronidation in human liver and intestine microsomes: high catalytic activity of UGT1A8 and UGT1A10. Drug Metab. Dispos. 30: 1462–1469. Watanabe, I., Tomita, A., Shimizu, M., Sugawara, M., Yasumo, H., Koishi, R., Takahashi, T., Miyoshi, K., Nakamura, K., Izumi, T., Matsushita, Y., Furukawa, H., Haruyama, H. and Koga, T. (2003). A study to survey susceptible genetic factors responsible for troglitazone-associated hepatotoxicity in Japanese patients with type 2 diabetes mellitus. Clin. Pharmacol. Ther. 73: 435–455. Watkins, P. B. and Whitcomb, R. W. (1998). Hepatic dysfunction associated with troglitazone. New Engl. J. Med. 338: 916–917. Yamamoto, Y., Nakajima, M., Yamazaki, H. and Yokoi, T. (2001). Cytotoxicity and apoptosis produced by troglitazone in human hepatoma cells. Life Sci. 70: 471–482. Yamamoto, Y., Yamazaki, H., Ikeda, T., Watanabe, T., Iwabuchi, H., Nakajima, M. and Yokoi, T. (2002). Formation of a quinone epoxide metabolite of troglitazone with cytotoxic to HepG2 cells. Drug Metab. Dispos. 30: 155–160. Yamazaki, H., Shibata, A., Suzuki, M., Nakajima, M., Shimada N., Guengerich, F. P. and Yokoi, T. (1999). Oxidation of troglitazone to a quinone-type metabolite catalyzed by cytochrome P-450 2C8 and P-450 3A4 in human liver microsomes. Drug Metab. Dispos. 27: 1260–1266. Yoshigae, Y., Konno, K., Takasaki, W. and Ikeda, T. (2000). Characterization of UDPglucuronosyltransferases (UGTS) involved in the metabolism of troglitazone in rats and humans. J. Toxicol. Sci. 25: 433–441.
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12 Role of the Kupffer Cell in Hepatotoxicity and Hepatocarcinogenesis James E. Klaunig, Stacy M. Corthals, Lisa M. Kamendulis, and Binu K. Philip
12.1
Introduction
The liver is composed of a variety of cell types, with the majority of the volume (∼ 90 %) and number (∼ 65 %) accounted for by parenchymal cells (hepatocytes), while nonparenchymal cells (endothelial, Kupffer, hepatic stellate and pit cells) comprise the remaining hepatic mass (Altin and Bygrave, 1988; Blouin, 1977; Widmann et al., 1972; Wisse, 1977). Kupffer cells, first identified by von Kupffer in 1876, originate from precursor cells in the bone marrow, which produce blood monocytes. These blood monocytes that are derived from the mononuclear phagocyte system, migrate into various tissues in the body and transform into tissue macrophages (van Furth, 1975, 1980, 1985, 1989, 1992; van Furth et al., 1972). Kupffer cells are macrophages that reside in the sinusoidal spaces of the liver and account for ∼ 15 % of all liver cells in number and ∼ 39 % of the sinusoidal nonparenchymal cell types (Wisse et al., 1996) (Figure 12.1). Kupffer cells are motile and are distributed throughout the liver lobule; however, they are about twice as abundant in periportal than the centrilobular regions of the liver lobule (Bouwens et al., 1992; Sleyster and Knook, 1986). Kupffer cells have an irregular stellate shape. Within the periportal region, they are frequently found either in gaps between adjacent sinusoidal endothelial cells or adhered to their surface (Rubin and Faber, 1988). They occasionally have extensions into the space of Disse through endothelial fenestrae and into the interior of the sinusoid, bringing them into direct contact with hepatic stellate cells and hepatocytes (Wisse, 1974). Evidence Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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Hepatocyte
Hepatic stellate cell (Ito cell)
Endothelial cell
Slnusoldal lumen
Kupffer cell
Space of Disse
Figure 12.1 Schematic representation of liver cell organization. Hepatocytes are arranged as cords radiating from the central vein. Hepatic sinusoids are channels between the cords of heapatocytes and are larger and more irregular than capillaries. Blood entering the hepatic sinusoid, via the portal vein and hepatic artery, travels toward the central vein along cords of parenchymal cells (hepatocytes). Endothelial cells, Kupffer cells, and stellate cells (Ito cells) are the three major types of cells in the sinusoids. Endothelial cells line the sinusoidal lumen. The space between the endothelium and hepatocytes is termed the space of Disse. Kupffer cells, the resident macrophage in the liver, are located in the lumen of the sinusoid. Ito cells are located between the endothelial cells and the hepatocytes.
also suggests that Kupffer cells in the periportal regions are larger, possess greater lysosomal enzyme activities and have more scavenging functions than those in the centrilobular regions (Sleyster and Knook, 1982; Bouwens et al., 1986; Mochida et al., 1989). However, the periportal Kupffer cells are less active in inflammatory reactions and generate less superoxide anion than the centrilobular Kupffer cells (Sleyster and Knook, 1982; Bouwens et al., 1986; Mochida et al., 1989; Naito et al., 2004). The sinusoidal positioning of the Kupffer cells makes them the first line of defense against many circulating endogenous and exogenous compounds as they pass through the liver (Mathison and Ulevitch, 1979). Their cytoplasm typically display an abundant number of lysozomes and fuzzy-coated vacuoles at the cell surface, components involved in endocytic mechanisms of the Kupffer cell (Wisse, 1972; Wisse et al., 1996; Naito et al., 2004). The phagocytic vesicles in the cytoplasm usually contain cellular fragments and hemosiderin resulting from phagocytosis of aged erythrocytes (Klika et al., 1986). Kupffer cells exhibit endogenous peroxidase patterns distinct from other macrophage populations (Wisse, 1974) and stain positively for macrophage markers ED1, ED2, anti-lysozyme, and Ki-M2R in rats and F4/80 in mice (Niki et al., 1995; Van Rooijen et al., 1990; Naito et al., 2004).
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Kupffer Cell Function
As a specialized macrophage, the Kupffer cell exhibits several important properties, including endocytosis, antigen presentation and secretion of biologically active products (Winwood and Arthur, 1993). These properties are integral to the physiological functions of the Kupffer cell and include phagocytosis of large foreign particles such as erythrocytes and bacteria, clearing of endotoxins, processing of ingested material, tumor cytotoxicity, immune and inflammatory responses, liver regeneration, metabolism of iron and effects on hepatocyte function (Winwood and Arthur, 1993). Kupffer cells contain Fc, C3 and scavenger receptors, as well as cytosol binding sites for specific ligands of particles and soluble substances that mediate phagocytosis. The destruction of these foreign bodies and blood cells can take place both in and around the Kupffer cell, often at the expense of the surrounding tissue. This destruction and clearance can occur through the release of reactive oxygen intermediates, nitric oxide, eicosanoids, cytokines and lysosomal enzymes, or other biologically active products, each of which may elicit effects on surrounding cell populations (Decker, 1990). Kupffer cells are activated through the contact between membrane receptors and xenobiotic particles. One important physiological substance that activates Kupffer cells is the gastrointestinal (GI)-derived bacterial endotoxin (lipopolysaccharides; LPS) which enters the liver via the portal vein. This has led to the hypothesis that Kupffer cells are always in a semiactivated state from continuous exposure to low-levels of GI-derived LPS (Lumsden et al., 1988). The various biologically active products secreted by activated Kupffer cells may have paracrine interactions between different liver cell populations. Lysosomal enzymes and sulfated proteoglycan secreted from activated Kupffer cells are involved in protein degradation and tissue remodeling (Knook et al., 1977; Laskin et al., 1991). Other secretory products of activated Kupffer cells such as eicosanoids (prostanoids and leukotrienes; Decker, 1990; Kmiec et al., 1997), nitric oxide (Gaillard et al., 1991), platelet-activating factor (Chao et al., 1989), IL-1α and IL-1β (Armbrust et al., 1995; Shirahama et al., 1988), TNF-α (Karck et al., 1988), TGF-α (Decker et al., 1995), TGF-β1 and TGF-β2 (Bissell et al., 1995; Meyer et al., 1990) act as intercellular mediators and modulators of cell function. Kupffer cells also undergo oxidative burst during phagocytosis, which leads to production of reactive oxygen species, including superoxide anions, hydroxyl radicals and their by-products (i.e. hydrogen peroxide; Li and Karin, 1999). The oxidative burst seen in Kupffer cells is dependent on the rapid uptake of molecular oxygen and subsequent transformation into reactive oxygen species. This is an early event in Kupffer cell activation, resulting in an enhanced production of reactive oxygen species within seconds of interaction with a stimulus. This oxidative burst and the accompanying oxidative stress is one of the stimuli involved in increased expression of cytokines (Vrba and Modriansky, 2002). Of the factors released from activated Kupffer cells, tumor necrosis factor alpha (TNF-α) is involved in the pathogenesis of septic shock and inflammation, altered gene expression, changes in cytochrome P450 activity, cellular proliferation and apoptosis (Arii and Imamura, 2000; Beutler and Cerami, 1987; Dinarello et al., 1986; Laskin, 1996; Milosevic et al., 1999). This molecule can also initiate the release of IL-1, IL-6, colonystimulating factors, platelet activating factor and prostaglandins from both parenchymal
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and non-parenchymal cells (Dinarello et al., 1986; Laskin, 1996). TNF-α has been shown to cause direct damage to endothelial cells and activate neutrophils and monocytes (Laskin, 1996; Sato et al., 1986). In addition, TNF-α decreases canalicular anion transport expression (Nakamura et al., 1999), induces IL-8 synthesis (Thornton et al., 1991) and iNOS mRNA and NO synthesis (Shiratori et al., 1998) within hepatocytes and at relatively high concentration can induce hepatocellular apoptosis (Hamada et al., 1999). Kupffer cell-derived IL-1 is a modulator of inflammation and fibrosis, and can induce both the proliferation and activation of macrophages, endothelial cells, B and T lymphocytes, and fibroblasts (Feder, et al., 1993; Laskin, 1996). It can also increase the cytotoxic response in Kupffer cells and natural killer cells, which may be of some importance in the antimetastatic properties of the Kupffer cell (Laskin, 1996). Kupffer cell-derived molecules have effects on growth regulation in the liver by inducing either cell proliferation and/or apoptosis. Proliferative factors include (1) transforming growth factor alpha (TGF-α), (2) hepatocyte growth factor, (3) TNF-α and (4) IL-6, while the antiproliferative apoptotic factors produced by Kupffer cells are TGF-β and IL-1 (Decker, 1990). TGF-α derived from Kupffer cells, exhibits a potent mitogenic response in hepatocyte cultures, suggesting that similar effects may occur from the release of this peptide in vivo. The antiproliferative factors TGF-β and IL-1 released from Kupffer cells are also known to affect the growth of surrounding cells (Decker, 1990). In addition to LPS, number of compounds (including the released molecules themselves) such as interferon-δ (macrophage activating factor), TNF-α, viruses, platelet-activating factor, phagocytosed particulates, ionophores, peroxisome proliferators and phenobarbital are capable of activating Kupffer cells (Decker, 1990; Kroll et al., 1999; Rose et al., 1997a.b). To assess the role of the Kupffer cell in hepatotoxicity and cell growth, investigators have utilized various techniques to deplete or inactivate Kupffer cells, such as methyl palmitate, gadolinium chloride, glycine and liposome encapsulated dichloromethylene diphosphonate (clodronate) (Brown et al., 1997; Ikejima et al., 1997; Ju and Pohl, 2001; Marzi et al., 1991; van Rooijen and Sanders, 1994; Rose et al., 1999a). However, there are several advantages and disadvantages associated with many of these inactivating or depletion approaches. Methyl palmitate inhibits phagocytosis in Kupffer cells (Cai et al., 2005) and prevents Wyeth-14 643-induced DNA synthesis (Rose et al., 1997b), however daily i.v. injections are required to achieve this effect (Rose et al., 1999). Further, this agent causes necrosis of Kupffer cells (Rentsch et al., 2005). Glycine also prevents Wyeth-14,643-induced DNA synthesis (Rose et al., 1997a) and Wyeth-14,643-induced superoxide production (Rose et al., 1999a); however, it lacks specificity and results have not been consistently observed among studies. Gadolinium chloride has been widely used to deplete Kupffer cells. Disadvantages of using gadolinium chloride include necrosis of Kupffer cells, thereby stimulating the release of cytokines including TNFα, decreased activity of cytochrome P450, hepatotoxicity at higher doses, selective depletion of a subset of Kupffer cells (ED2+ Kupffer cells), and stimulation of hepatocyte proliferation (Rentsch et al., 2005; Rose et al., 2001; Rai et al., 1995, 1996, Badger et al., 1997; Ju et al., 2002; Ding et al., 2003; Jarvelainen et al., 2000). Rai et al. (1996) showed that gadolinium chloride did not reduce the number of Kupffer cells, but rather altered their acinar distribution and switched the phenotype, resulting in the loss of KCR expression, a Kupffer cell-specific antigen. Gadolinium chloride is also reported to be systemically toxic and
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stimulate dying Kupffer cells to release cytokines (Rose et al., 2001) and that in turn affects hepatocytes (Naito et al., 1996). Thus, it is controversial whether gadolinium chloride eliminates Kupffer cells or alters their function (Lazar, 1973; Hardonk et al., 1995; Rai et al., 1996). Silicates and asbestos have also been used to deplete Kupffer cells, but these agents result in Kupffer cell activation and secretion of inflammatory cytokines (Kagan and Hartmann, 1984). Clodronate-encapsulated liposomes, have many advantages over other Kupffer cell modulating protocols. In this approach, clodronate is localized between the lipid bilayers of the liposomes and are readily phagocytized by macrophages. Once phagocytized by Kupffer cells, the phospholipid bilayers of the liposomes are digested by lysosomal phospholipases and clodronate is released into the cell. Clodronate accumulates intracellularly and depletes the ATP thereby triggering apoptotic cell death (van Rooijen, 1989; van Rooijen and Sanders, 1996; Frith et al., 1997). Since the cell dies by apoptosis, cell remnants/apoptotic bodies are cleared and the release of growth mediators seen with other Kupffer cell depletion protocols is avoided. However, repeated injections are necessary for continuous depletion of Kupffer cells with this approach. Kupffer cells begin to repopulate on day 4 after elimination as evidenced by the reappearance of ED1-positive cells, and basal levels are restored in the liver 16 days after clodronate liposome treatment in the rat (van Rooijen et al., 1990) and after 14 days in mouse (Yamamoto et al., 1996). Through the use of protocols to activate or deplete/inactivate Kupffer cells, a number of investigators have assessed the contribution of the Kupffer cells as mediators of hepatotoxicity and cell growth.
12.3
Role of the Kupffer cell in Hepatotoxicity
The Kupffer cell plays an important role in the normal physiology and homeostasis of the liver, as well as participating in acute and chronic responses of the liver to toxic compounds (Figure 12.2; Table 12.1). In addition, Kupffer cells have been shown to participate in the toxicity of several well-known hepatocarcinogens. Endogenous and exogenous factors that modulate Kupffer cell activities may affect the risk of individuals in developing liver injuries (Ju et al., 2002). Idiosyncratic drug responses are suggested to result from druginduced modulation of Kupffer cells. Understanding the role played by Kupffer cells in the induction of hepatocyte injury will help to further our understanding of the mechanisms of idiosyncratic drug induced liver diseases (Table 12.2). 12.3.1
Acetaminophen
Acetaminophen is a widely used nonprescription analgesic and antipyretic agent. At high doses and also in susceptible individuals it can cause liver toxicity and even hepatic failure. Acetaminophen hepatotoxicity has been attributed in part to the activation of Kupffer cells secondary to hepatocyte damage initiated by the formation of a reactive metabolite, N-acetyl-p-benzoquinone imine (Cohen and Khairallah, 1997; Cohen et al., 1997). This hypothesis is based on the fact that various proinflammatory cytokines, reactive oxygen and nitrogen species are released during acetaminophen toxicity and Kupffer cell activation
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Hepatotoxicity Cadmium
Ethanol
Acetaminophen
GI – O2
Carbon tetrachloride
LPS
CCl3
Activated Kupffer cell Eicosanoids
Cytotoxic cytokines
Reactive oxygen and
nitrogen species
Hepatocyte
Figure 12.2 Schematic representation of the role of Kupffer cell in hepatotoxicity-induced by various toxicants. Kupffer cells can be activated directly or indirectly by various toxicants resulting in the release of biologically active cytokines, growth regulators, as well as reactive oxygen and nitrogen species, all of which may contribute to liver damage.
is known to release these bioactive substances. Goldin et al. (1996) showed that mice pretreated with clodronate containing liposomes to knock out Kupffer cells exhibited lower acetaminophen-induced liver injury. However, liver injury was significantly decreased only at initial time points (0.5–2 h) and while no difference was seen at later time points as compared to control mice. Several investigators have shown that gadolinium chloride pretreatment decreased acetaminophen hepatotoxicity in rats and mice (Laskin et al., 1995; Laskin and Pendino, 1995; Michael et al., 1999). However, the efficiency of gadolinium chloride to completely deplete Kupffer cells remains controversial. Overall, these studies suggest a role for Kupffer cells in the development of acetaminophen hepatotoxicity. However, the contribution of Kupffer cells versus infiltrated macrophages in acetaminophen hepatotoxicity is not clear (Laskin and Pilaro, 1986; Laskin et al., 1986). Moreover, nitric oxide is also produced by hepatocytes and sinusoidal endothelial cells and the superoxide anion is produced during acetaminophen metabolism (Michael et al., 1999). Thus, the relative contribution of Kupffer cells in the production of reactive oxygen and nitrogen species has also not been clearly established.
TGFβ1, platelet-derived growth factor (PGDF), thromboxane B2 (TxB2), leukotrienes, NO• , O•− 2 , TNFα ,TNFα, IL-1 NO• , O•− 2
Free radicals and endotoxins released from hepatocytes during CCl4 bioactivation activates Kupffer cells Might be directly activating Kupffer cells
Cadmium
TNFα, NO• , O•− 2
Nitric oxide and superoxide causes lipid peroxidation in hepatocytes. TNFα and IL-1 are involved in leucocyte infliltration
TNFα upregulates expression of adhesion molecules (ICAM-1) on endothelial cells leading to increased neutrophil infiltration in the liver and also stimulates mitochondrial oxidant production in hepatocytes by increasing respiration as well as inhibition of electron transport Nitric oxide and superoxide rapidly reacts to form peroxynitrite, which is a potent oxidant and nitrates protein tyrosine and also attack other macromolecules in the hepatocytes. The superoxide radical also causes lipid peroxidation in phospholipid membrane of hepatocytes and can also cause alteration in DNA or mitochondrial functions TGFβ1 and PGDF stimulates proliferation and collagen production by stellate cells leading to fibrosis. TxB2 upregulates TNFα and IL-1β production
Biological effect
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Endotoxins derived from GI tract following ethanol exposure activates Kupffer cells
Ethanol
Factors released from activated Kupffer cells
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Effect on Kupffer cells
Compound
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Kupffer cells and hepatotoxicity
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Table 12.2 Role of Kupffer cells in liver regeneration following partial hepatectomy Depletion of Kupffer cells
Mode of depletion
— Prior to partial hepatectomy
— Gadolinium chloride
Clodronate– liposomes
After partial hepatectomy
Clodronate– liposomes
Results
Reference
Increased TNF and IL-6 levels Increased TNFα mRNA levels and IL-6 levels Decreased TNF levels and increased IL-6 levels Increased hepatocyte proliferation and mitotic index, while decreasing TGFβ IL-1β and HGF mRNA Delayed hepatocyte DNA synthesis and decreased TNFα, IL-6, HGF, IL-10 and TGF-β1 levels Decreased HGF mRNA and hepatocyte proliferation Decreased TNFα mRNA levels
Iwai et al., 2001 Rai et al., 1996 Iwai et al., 2001 Boulton et al., 1998 Meijer et al., 2000 Takeishi et al., 1999 Prins et al., 2005
In addition to its pathologic role, the Kupffer cell has been postulated to have a protective role in acetaminophen toxicity. Ju et al. (2002) reported that Kupffer cell depletion with clodronate liposomes increased the susceptibility of mice to acetaminophen toxicity with a concomitant decrease in the hepatic expression of several regulatory cytokines released by the Kupffer cells. Raiford and Thigpen (1994) showed that immunostimulation of Kupffer cells with Corynebacterium parvum diminished acetaminophen-induced liver injury in rats. However, these authors attributed the decreased hepatic injury to decreased bioactivation of acetaminophen. Although the available literature suggest the involvement of the Kupffer cell in acetaminophen hepatotoxicity, additional research is needed to characterize the relative contribution of various factors released from the Kupffer cells in the toxicity process. 12.3.2
Ethanol
Acute and chronic consumption of ethanol alters the gastrointestinal bacteria and causes an overgrowth of gram negative bacteria leading to an increased influx of endotoxin from the gastrointestinal (GI) tract into the circulation, which is then cleared by Kupffer cells and hepatic sinusoidal endothelial cells (Bautista, 1998a,b; Enomoto et al., 1998, Thurman, 1998). Endotoxin activates Kupffer cells by interacting with CD14 receptors leading to increased production of reactive oxygen species via NADPH oxidase. These free radicals, in turn, activate NF-kB, leading to increased TNFα production and eventually liver damage (Dey and Cederbaum, 2006). Diphenyleneiodonium sulfate, an NADPH oxidase inhibitor, prevented early alcohol-induced liver injury in the rat suggesting that oxygen radicals derived from NADPH oxidase in Kupffer cells is a key step in the development of hepatotoxicity following ethanol exposure (Kono et al., 2001). Mice deficient in p47phox, a subunit of NADPH oxidase, exhibited decreased liver injury with a concomitant reduction in
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reactive oxygen species and TNFα levels following ethanol treatment (Kono et al., 2000). Acute ethanol challenge induced superoxide anion release into the hepatic sinusoid and subsequent liver damage was attenuated in rats by Kupffer cell depletion with gadolinium chloride (Yokoyama et al., 1999). In addition to reactive oxygen species, activated Kupffer cells release prostaglandins such as PGE2 that increase oxygen consumption causing hypoxia in hepatocytes (Qu et al., 1996). TNFα released from the activated Kupffer cells is believed to be responsible for increased inflammation, necrosis, and fatty accumulation in the injured liver (Iimuro et al., 1997; Wheeler, 2003; Wheeler and Thurman, 2003; Bautista, 2000). A glycine-rich diet expedited the recovery process from early ethanol-induced liver injury in rats by inhibiting TNFα production by activated Kupffer cells (Yin et al., 1998). Kupffer cells that are activated following ethanol exposure also produce macrophage inflammatory protein -2 (MIP-2) and intercellular adhesion molecule -1 (ICAM-1) which bind to polymorphonuclear neutrophils leading to neutrophil infiltration in the liver (Thurman, 1998). In addition to endotoxin, psychogenic stress is also shown to activate Kupffer cells and increase the susceptibility of mice to ethanol-induced liver injury (Panuganti et al., 2006). Calcium is a second messenger involved in signaling in Kupffer cells, which contain L-type voltage-dependent calcium channels that are easily activated following chronic ethanol treatment (Decker, 1990). Nimodipine, a calcium channel blocker, prevented alcoholic hepatitis caused by chronic intragastric ethanol exposure in rats, demonstrating that activation of Kupffer cells by intracellular calcium is involved in ethanol-induced liver injury (Iimuro et al., 1996). Treatment with gadolinium chloride (Adachi et al., 1994, 1995) or reduction of gastrointestinal gram negative bacteria by administration of antibiotics (Adachi et al., 1995) diminished early ethanol-induced liver injury in rats. Recently Ito et al. (2004) showed a protective role for Kupffer cells in rat liver against acetaminophen intoxication following ethanol binging. Taken together, these studies establish the involvement of Kupffer cell activation via gastrointestinal-derived endotoxin in the pathogenesis of alcoholic liver disease. 12.3.3
Carbon Tetrachloride
Carbon tetrachloride (CCl4 ) was widely used as a solvent, cleaning agent, fire extinguisher, grain fumigant and antihelminthic for humans. Its use has declined steadily since the 1970s, due mainly to its hepatic and renal toxicities. The hepatotoxicity produced by CCl4 is a complex multifactorial process. CCl4 is metabolized by CYP2E1 to the reactive trichloromethyl radical leading to cytotoxicity through lipid peroxidation (Alric et al., 2000). Lipid peroxidation causes changes in the permeability barrier of the plasma membrane leading to calcium influx resulting in increased intracellular calcium. Kupffer cell inhibition by gadolinium chloride pretreatment inhibited CCl4 -induced hepatic lipid peroxidation, injury, and liver cirrhosis (Muriel and Escobar, 2003) suggesting that Kupffer cells mediate the liver damage induced by CCl4 , and also that lipid peroxidation is produced or stimulated by Kupffer cells (Muriel et al., 2001). Kupffer cells are activated by high intracellular calcium levels resulting in the release of cytokines and other biologically active compounds that promote the pathogenic process (Bermelin and Decker, 1983). Expression of osteopontin, an extracellular matrix component, was found to be significantly higher in activated Kupffer cells in the hepatic necrotic areas of rats after CCl4 intoxication, suggesting that osteopontin can
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act as a chemokine that can induce the migration of Kupffer cells (Kawashima et al., 1999). Geerts et al. (1988) showed that the number of Kupffer cells in the liver increases during chronic CCl4 administration to rats, with a peak after nine weeks of exposure. Several cytokines, such as macrophage or granulocyte colony-stimulating factors, were postulated to play a role in the increased Kupffer cell proliferation (Hoedemakers et al., 1994). Kupffer cells isolated from rats exposed to CCl4 produced cytokines that stimulate proliferation and production of collagen in stellate cells (Edwards et al., 1993). Destruction of Kupffer cells with gadolinium chloride as well as inactivation with dietary glycine attenuated CCl4 -induced hepatic fibrosis in rats suggesting that collagen production by stellate cells is potentially regulated in vivo by early release of products from Kupffer cells (Rivera et al., 2001). Rats treated with polymyxin B to eliminate gram negative bacteria and endotoxin exhibited an acute decreases in CCl4 -induced hepatic injury (Nolan and Leibowitz, 1978). Increased levels of endotoxin stimulate the Kupffer cells to release mediators resulting in hepatic fibrosis. Treatment of rats with an inhibitor of 5-lipoxygenase activating protein partially depleted Kupffer cells and attenuated CCl4 -induced injury in rats suggesting that this is a critical protein for the survival of Kupffer cells (Titos et al., 2005). Administration of hepatocyte growth promoting factor attenuated CCl4 -induced acute liver injury in rats by acting as a mitogen to hepatocytes and Kupffer cells (Sato et al., 1999). Pretreatment of rats with large doses of vitamin A, an activator of Kupffer cells, potentiated CCl4 -induced lipid peroxidation and liver injury (elSisi et al., 1993). On the contrary, activation of Kupffer cells with Bacillus Calmette–Guerin (BCG), corynebacterium parvum or zymosan has been shown to protect rats from acute liver injury induced by CCl4 and acetaminophen (Pereira et al., 1997). However, this protection appears to be related to altered microsomal enzymes and increased production of glutathione (Raiford and Thigpen, 1994). Collectively, the available literature support the involvement of Kupffer cells in CCl4 -induced toxicity. 12.3.4
Cadmium
Cadmium is an environmental and industrial pollutant and is a known hepatotoxicant whose mode of action may be at least partially mediated by the Kupffer cell (Dudley et al., 1982; Sauer et al., 1997). This heavy metal accumulates in the livers of treated rodents, a distribution pattern which likely reflects the target specific toxicity of this compound (McKim et al., 1992). Time-course studies following cadmium exposure (i.v.) in rats demonstrated not only cadmium localization in the liver, but also the initial hepatocyte ultrastructural changes associated with exposure (Dudley et al., 1984). Within an hour of treatment, dilation of hepatocyte rough endoplasmic reticulum and the loss of membrane-associated ribosomes were observed, followed by mitochondrial swelling and the appearance of fibrillar material in the cytoplasm (Dudley et al., 1984). This cellular response can result in apoptosis and necrosis (Habeebu et al., 1998). While the development of hepatic injury following cadmium exposure is well-documented, the mechanisms that contribute to toxicity are not fully elucidated, and the contribution of the Kupffer cell remains a matter of contention. In a review by Rikans and Yamano (2000), the authors outlined some of the proposed mechanisms through which cadmium causes hepatotoxicity. These pathways implicate vascular endothelial cell damage (Nolan and Shaikh, 1986), sulfhydryl group inactivation
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(Nolan and Shaikh, 1986), oxidative stress (Bagchi et al., 1996), and Kupffer cell activation (Sauer et al., 1997) as possibly playing roles in cadmium hepatotoxicity. The Kupffer cell was focused on because of its potential to produce an biologically active molecules, including pro-inflammatory mediators such as TNFα and nitric oxide, which play prominent roles in the toxicity of compounds such as lipopolysaccharide (Bopst et al., 1998). Cadmium activates Kupffer cells, resulting in cytoplasmic vacuolization (Hoffmann et al., 1975), increased colloidal carbon clearance (Sauer et al., 1997), and alteration in cytokine production (Kayama et al., 1995). Interestingly, although TNFα has been shown to have primary involvement in other hepatotoxic events, TNFα-null mice are not resistant to cadmium chloride-induced hepatotoxicity (Harstad and Klaassen, 2002b). Similarly, inducible nitric oxide synthase (iNOS)-null mice are also not resistant to the effects of cadmium chloride-induced toxicity (Harstad and Klaassen, 2002c). This would suggest that neither TNFα nor nitric oxide is involved in the hepatotoxic effects of cadmium; however, these studies do not rule out the involvement of Kupffer cells. By depleting the Kupffer cell with gadolinium chloride, cadmium-induced liver damage was inhibited in rodents (Harstad and Klaassen, 2002a; Sauer et al., 1997; Kuester et al., 2002). Interestingly, low doses of gadolinium chloride that depleted Kupffer cells did not confer hepatic resistance to cadmium toxicity, which would indicate a protective mechanism of gadolinium chloride treatment beyond that of Kupffer cell (Harstad and Klaassen, 2002a). Of interest is that gadolinium chloride does not affect cadmium distribution in the liver, thereby removing the possibility of lessening the hepatic cadmium burden (Harstad and Klaassen, 2002a). Although gadolinium chloride can induce the formation of metallothionein, a cadmium-binding protein, the induction of this protein was not shown to be involved in the protective effect of gadolinium chloride against cadmium toxicity (Harstad and Klaassen, 2002a). Additionally, gadolinium chloride treatment in isolated hepatocytes did not protect against cadmium toxicity, suggesting that the presence of other cell types such as the Kupffer cell, are related to this effect (Badger et al., 1997). It is possible, for example, that oxidant production by activated Kupffer cells may exacerbate cadmium toxicity, although this has not been examined. Another mode of action may involve the nitric oxide-mediated release of cadmium from metallothionein, leading to enhanced toxicity in the liver (Misra et al., 1996). While the role of the Kupffer cell in cadmium toxicity is a matter of debate, the role of the change to the Kupffer cell in cadmium hepatotoxicity has not yet been ruled out. 12.3.5
Partial Hepatectomy
The liver has the ability to undergo compensatory hyperplasia after injury or partial hepatectomy until the full functional capacity of the liver is restored. The partial hepatectomy model (Higgins and Anderson, 1931) removes two-thirds of the liver, resulting in the remaining liver lobes to enlarge due to quiescent liver cells becoming stimulated to re-enter the cell cycle and proliferate to compensate for the removed mass of the liver within 7– 10 days. External stimuli and changes in gene expression, growth factor production, and morphological structure induce this response (Takeishi et al., 1999). Based upon the structure, function and activation of Kupffer cells which results in secretion of both positive (TNFα) and negative (TGFβ, IL-1) growth modulators, Kupffer cells
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may influence hepatocyte proliferation during liver regeneration (Table 12.2). TNFα has been suggested as an important early component of liver regeneration through activation of signaling pathways (Vandenabeele et al., 1995; Cressman et al., 1996; Taub et al., 1999). However, TNFα has also been identified as a cell-death mediator and inducer of apoptosis at higher dose levels (Beutler and Van Huffel, 1994; Bazzoni and Beutler, 1996; Wallach, 1997; Declercq et al., 1998). Several studies have investigated the role of Kupffer cells and the influence of TNFα on early events of liver regeneration after partial hepatectomy. However, the results are controversial. Depletion of Kupffer cells prior to partial hepatectomy enhances early hepatocellular proliferation in some reports (Rai et al., 1996; Boulton et al., 1998) but delays proliferation in other reports (Meijer et al., 2000; Zerbe and Gressner, 1988). Rai et al. (1996) hypothesized that if Kupffer cells released TNFα after partial hepatectomy, then Kupffer cell depletion by gadolinium chloride should inhibit liver regeneration. Rats treated with gadolinium chloride prior to partial hepatectomy resulted in the absence of Kupffer cell receptor (KCR) mRNA demonstrating the depletion of Kupffer cells. However, TNFα mRNA expression was increased in gadolinium chloride treated rats after partial hepatectomy, a finding in opposition to their hypothesis (Rai et al., 1996). These authors also reported increased IL-6 levels, possibly indicating that gadolinium chloride also amplified TNFα bioactivity after partial hepatectomy, since IL-6 is a TNFα-inducible cytokine. Their study, therefore, indicated that when Kupffer cells are eliminated, other cells in the liver produce TNFα to initiate regeneration of hepatocytes. Interestingly, bile ducts and blood vessels were shown to be a source of TNFα in regenerating livers (Loffreda et al., 1997). Callery et al. (1991) also demonstrated that after partial hepatectomy, TNFα production from isolated Kupffer cells was suppressed by regulation of eicosanoid metabolism. Iwai et al. (2001) have also shown that secretions of TNFα and IL-6 from perfused rat liver were increased after partial hepatectomy. Rats pretreated with gadolinium chloride suppressed TNFα but not IL-6 secretions, suggesting that Kupffer cells play an important role in hepatic TNFα secretion, whereas IL-6 production may be achieved by other hepatic cells (Iwai et al., 2001). In contrast to the results reported by Rai et al. (1996), plasma TNFα levels and hepatic, pulmonary and renal TNFα mRNA levels were decreased in Kupffer cell-depleted rats (via clodronate-encapsulated liposomes) after partial hepatectomy (Prins et al., 2005). This is also supported by findings showing that mitotic hepatocytes and PCNA-positive hepatocytes in TNFα knock out mice were similar to the wild type mice, suggesting that TNFα has little influence on liver regeneration after partial hepatectomy (Hayashi et al., 2005; Fujita et al., 2001). Boulton et al. (1998) studied the role of Kupffer cell modulation of early events after partial hepatectomy. Rats treated with clodronate-encapsulated liposomes prior to a partial hepatectomy depleted Kupffer cells, as evidenced by immunohistochemistry for ED1, which recognizes a cytoplasmic antigen in monocytes and macrophage populations, and ED2, recognizing membrane antigens of resident macrophages. The uptake of Indian ink by phagocytosis was also abolished in Kupffer cell-depleted rats. Hepatocyte proliferation and mitotic index were increased in the Kupffer cell-depleted rats compared to controls in both resting liver and after partial hepatectomy, indicating that the role of the Kupffer cell during this time is to inhibit proliferation. Expression of negative growth modulators, TGF-β and IL-1β were reduced in Kupffer cell-depleted rats (Boulton et al., 1998), which have been reported to increase in expression after partial hepatectomy (Evarts
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et al., 1992; Burr et al., 1993; Bissell et al., 1995; Boulton et al., 1997). This increase in expression of TGF-β and IL-1β may give rise to the increase in hepatocyte proliferation as seen after partial hepatectomy (Boulton et al., 1998). Takeishi et al. (1999) also examined the role of Kupffer cells in liver regeneration. Mice depleted of Kupffer cells by clodronate-encapsulated liposomes after a two-thirds partial hepatectomy exhibited reduced hepatocyte proliferation. This is in contrast to Boulton et al. (1998). However, both observed a reduction in HGF mRNA expression in Kupffer cell-depleted animals. This may also be responsible for the reduction in liver regeneration seen after Kupffer cell-depletion. Although HGF has been shown to be synthesized by Kupffer, Ito and endothelial cells (Maher, 1993), the study by Takeishi et al. observed HGF expression in Ito cells after partial hepatectomy while less was observed in Kupffer cell-depleted mice, indicating that Kupffer cells may control HGF production in Ito cells. Meijer et al. (2000) performed a splenectomy in rats one week prior to partial hepatectomy and administered clodronate-encapsulated liposomes prior to partial hepatectomy. Hepatocyte DNA synthesis was decreased compared to controls in Kupffer cell-depleted rats 48 h after partial hepatectomy but no difference was seen at 96 h after partial hepatectomy. This indicates a delayed hepatocyte proliferation, rather than decreased hepatocyte proliferation. Growth-stimulating cytokines (TNFα, IL-6, HGF, IL-10, TGF-β1) were also decreased in Kupffer cell-depleted rats (Meijer et al. 2000) which may be responsible for the delayed hepatocyte regeneration after partial hepatectomy. In regards to exploring the role of Kupffer cells in liver regeneration, one main difference between the studies is the Kupffer cell depletion technique. Gadolinium chloride causes necrosis of Kupffer cells (Rose et al., 2001), thus releasing cytokines (TNF-α), which may lead to the early proliferation response seen after partial hepatectomy (Rai et al., 1996). However, there are also conflicting reports when using clodronate liposomes to deplete Kupffer cells, depending on the time of administration, either before partial hepatectomy (Boulton et al., 1998) or after partial hepatectomy (Takeishi et al., 1999). Taken together, conflicting results remain concerning the roles of TNFα and Kupffer cells in liver regeneration. The mechanism of TNFα production and its influence on liver regeneration are also still under investigation. While some reports have shown that TNFα is induced after partial hepatectomy (Rai et al., 1996; Iwai et al., 2001), others have shown a suppression of TNFα after partial hepatectomy (Prins et al., 2005; Hayashi et al., 2005, Fujita et al., 2001).
12.4 12.4.1
Role of the Kupffer cell in Liver Carcinogenesis Hepatocellular Cancer
Hepatocarcinogenesis, a multistage process, involves the alteration of a normal cell to a preneoplastic intermediate (Dragan et al., 1993). The role of the Kupffer cell in hepatocarcinogenesis has not been clearly established; however, limited evidence suggests that the Kupffer cell may be involved in the carcinogenesis process (Table 12.3). The majority of evidence implicating the Kupffer cell as a mediator in hepatocarcinogenesis has been demonstrated with peroxisome proliferators activated receptor α (PPARα) agonists (Rose et al., 1997a, b; Rusyn et al., 1998). This class of compounds, that includes model
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Table 12.3 Role of Kupffer cells in liver carcinogenesis Compound Wyeth 14 643
Biological effect
Increased hepatocyte DNA synthesis and hepatic TNFα mRNA levels in vivo, suggesting TNFα, originated from Kupffer cells, is responsible for the mitogenic effects of Wyeth 14 643 Increased acyl CoA oxidase; however Kupffer cell inactivation did not prevent this increase, indicating that Kupffer cell inactivation prevents DNA synthesis without affecting peroxisome induction Increased NF-κB in rat hepatocytes and Kupffer cells in vivo. Activated superoxide production in isolated Kupffer cells Phenobarbital Elevated levels of cyclooxygenase-2, PGE2 and PGD2 in cultured Kupffer cells Lindane Elevated levels of cyclooxygenase-2, PGE2 and PGD2 in cultured Kupffer cells Nafenopin Increased hepatocyte DNA synthesis in the presence of nonparechymal cells Increased TNFα released from nonparechymal cells LPS Activated macrophages, thereby increasing endothelial cell DNA synthesis in vitro and increasing endothelial cell damage in vitro 2-Butoxyethanol Induced hemosiderin in Kupffer cells in vivo
Reference Rose et al., 1997a; Bojes et al., 1997; Holden et al., 2000 Rose et al., 1997b
Rusyn et al., 1998, 2000; Rusyn et al., 2000 Kroll et al., 1999 Kroll et al., 1999 Hasmall et al., 2000 Hasmall et al., 2000 Corthals et al., 2006 NTP, 2000; Siesky et al., 2002
hepatocarcinogens such as Wyeth 14 643, produces a variety of biological effects, including peroxisomal proliferation, liver hyperplasia, sustained elevation of cell turnover, altered growth in preneoplastic foci and the induction of hepatic tumors (Marsman et al., 1988; Butterworth, 1990; Roberts, 1999; Cattley and Roberts, 2000; Hasmall et al., 2000; Holden et al., 2000; Roberts-Thomson, 2000; Hasmall et al., 2001). The participatory role of the Kupffer cell in PPARα agonist-induced DNA synthesis has been demonstrated using both in vivo and in vitro models (Rose et al., 1997a, 1999a, b; Gill et al., 1998). In particular, TNF-α, released from activated Kupffer cells is believed to be involved in the mitogenic response of peroxisome proliferators. Following treatment with Wyeth 14 643, hepatocyte DNA synthesis and hepatic TNF-α mRNA transcripts were increased over control levels in rats. These results suggest that TNF-α of Kupffer cell origin is responsible for the mitogenic effects of Wyeth 14 643 (Rose et al., 1997a; Bojes et al., 1997). Treatment with peroxisome proliferators does not appear to mediate de novo TNF-α gene expression, suggesting that the response to this class of compounds may be mediated by bioactivation or release of pre-existing TNFα protein from Kupffer cells (Holden et al., 2000). A role for TNF-α in Wyeth 14 643-induced cell proliferation is further supported by results demonstrating a reduction in cell proliferation by Kupffer cell inhibition with glycine and methyl palmitate. Interestingly, Kupffer cell inactivation did not prevent the increase in acyl CoA oxidase
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activity by Wyeth 14 643, indicating that Kupffer cell inactivation prevents DNA synthesis caused by PPARα agonists without affecting induction of peroxisomes (Rose et al., 1997b). Furthermore, stimulation of cell proliferation caused by PPARα agonists was blocked by antibodies against TNF-α (Rusyn et al., 1998). However, in TNFα knock out mice, hepatocellular DNA synthesis induced by Wyeth was not inhibited (Lawrence et al., 2001). This suggests that cell proliferation induced by PPARα agonist treatment occurs independently of TNF-α signaling or does not exclusively involve TNF-α. In addition to the production of TNF-α, Kupffer cell activation results in the production of oxidants. In rat hepatocytes, an increase in nuclear factor-κB (NF-κB), an oxidative stressactivated transcription factor involved in the regulation of cell proliferation and apoptosis, activation and binding was seen shortly after Wyeth 14 643 administration. Pre-treatment with allopurinol, a xanthine oxidase inhibitor and free-radical scavenger, suppressed NFκB activation by Wyeth 14 643. It is concluded that NFκB is activated by reactive oxygen species and plays a central role in the mechanism of action of PPARα agonists (Rusyn et al., 1998). Wyeth 14 643 was also shown to produce a rapid oxidant-dependent activation of NFκB in Kupffer cells in vivo and activated superoxide production by isolated Kupffer cells (Rusyn et al., 2000). Addition of non-parenchymal cells or conditioned medium from non-parenchymal cell cultures, increased DNA synthesis. DNA synthesis induced by nafenopin- and TNF-α were prevented by the removal of non-parenchymal cells from normal hepatocyte cultures and was restored by returning non-parenchymal cells to the purified hepatocytes (Hasmall et al., 2001). This finding provides additional evidence showing that non-parenchymal cells are permissive for the growth response of hepatocytes in vitro to PPARα agonists and may be mediated, at least in part by TNFα. However, the levels of TNFα released spontaneously or in response to PPARα agonists were shown to be insufficient per se to induce a growth response (Hasmall et al., 2000). Thus, it is likely that oxidants play a role in signaling hepatocellular proliferation following exposure to peroxisome proliferators perhaps via activation of NFκB and increases in additional increases in mitogenic cytokines such as TNF-α. In addition, the activation of NFκB by peroxisome proliferators appear species-specific. This class of compounds activates NFκB in rat and mouse liver, while they do not induce any changes in hamsters. This may explain the differences seen between species in relationship to cell proliferation (Tharappel et al., 2001). Although experimental evidence exists that establishes a relationship between the Kupffer cells and hepatocarcinogenesis by peroxisome proliferators, specific studies examining the role of the Kupffer cell on the tumor-promotion stage of carcinogenesis are lacking. Many hepatic tumor promoters appear to elicit their carcinogenic effects through the induction of cell proliferation. Increased cell replication may increase the frequency of spontaneous mutations through increasing the frequency of errors in DNA repair or replication, or may alter methylation of the genome and facilitate clonal expansion of initiated cells via a change in gene expression that either silences tumor suppressor genes or increases the expression of oncogenes, either of which may result in the formation of hepatic focal lesions (Cattley et al., 1991, 1998; Huber et al., 1991). Tumor promoters, for the most part, produce a strong, albeit transient, increase in replicative DNA synthesis during the first few weeks of exposure that typically returns to baseline levels within 2–4 weeks of treatment (Marsman et al., 1988; Schulte-Hermann et al., 1990). Hepatic tumor promoters have been shown to promote the growth of chemically and spontaneously induced lesions through enhanced cell replication (Marsman et al., 1988; Cattley et al., 1991; Cattley
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and Popp, 1989; Isenberg et al., 1997). In addition, the development of hepatocellular neoplasms following treatment with tumor promoters is species-specific and appears to correlate with the induction of DNA synthesis (Huber et al., 1996; James and Roberts, 1996). Furthermore, the effects of tumor promoters on the induction of DNA synthesis are reversible in that removal of the compound returns DNA synthetic rates to that of control both in vivo and in vitro (Isenberg et al., 2001; Kolaja et al., 1996; Taketani and Oka, 1983). The precise mechanism(s) for the mitogenic response of liver tumor promoters is not clearly understood; however, limited studies have described results that suggest the involvement of Kupffer cells in the biological effects elicited by two well-established hepatic carcinogens. Kupffer cells isolated from either phenobarbital or lindane-treated rats were shown to have elevated levels of cyclooxygenase-2 (COX-2) protein. In addition, in cultured Kupffer cells, phenobarbital and lindane both caused an increase in prostaglandin expression, particularly PGE2 , PGD2 , as well as COX-2 (Kroll et al., 1999). Overexpression of COX-2, the principle enzyme in prostanoid synthesis, was shown to decrease cellular rates of apoptosis. Furthermore, many tumor cells have been shown to overexpress COX2 (Marnett, 1992; Ballif et al., 1996). An additional importance of this finding is that several prostaglandins, including PGF2α and PGE2 exhibit mitogenic activity in several cell systems, including primary cultured hepatocytes (Hashimoto et al., 1997). Additionally, prostaglandins appear to be involved in the signaling pathways that result from partial hepatectomy (Aono et al., 1993). It has also been reported, however, that phenobarbital does not produce increases in prostaglandins that correlate with increases in DNA synthesis (Leung and Glauert, 1997). Although strong evidence indicates that prostaglandins are involved in tumor promotion, contention remains in terms of direct effects of prostaglandins on hepatocytes – thus the role of the prostaglandin-producing Kupffer cell remains to be established (Lee and Edwards, 2001). 12.4.2
Hemangiosarcoma
Kupffer cells also participate in 2-butoxyethanol-induced hemangiosarcomas in the mouse liver. 2-Butoxyethanol induced an increase in liver hemangiosarcomas in B6C3F1 male mice following chronic inhalation exposure (NTP, 2000). The mechanism for 2butoxyethanol-induced hepatic carcinogenicity is not known, but is hypothesized to involve Kupffer cell-activation and the induction of oxidative damage, secondary to red blood cell hemolysis by 2-butoxyethanol metabolites. 2-Butoxyethanol-induced red blood cell hemolysis resulted in hemosiderin (iron deposition) in Kupffer cells following chronic 2-butoxyethanol exposure in rodents (NTP, 2000; Siesky et al., 2002). Iron is a pro-oxidant that, at high levels, can produce oxidative stress either directly or indirectly by Kupffer cell activation. It may produce oxidative stress directly by catalyzing the conversion of superoxide and hydrogen peroxide to more potent oxidants, such as hydroxyl radical or ferryl or perferryl species by the Fenton reaction (Aust et al., 1985; Halliwell and Gutteridge, 1984). Importantly, the products of activated Kupffer cells may participate in the carcinogenesis process through inducing oxidative stress and damage, altering signaling cascades, initiating lipid peroxidation, increasing cell proliferation and/or protein modifications (Siesky et al., 2002; Imlay and Linn, 1988; Park et al., 2002a,b; Arii and Imamura, 2000; Bacon and Britton, 1990; Klaunig and Kamendulis, 2004) which may ultimately lead to hemangiosarcomas in the liver.
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When Kupffer cells exceed their iron storage capacity, the metal accumulates in parenchymal cells producing a mild iron overload (Searle et al., 1987; Ley et al., 1982). Iron overload in humans, whether it occurs genetically or by dietary intake, has been associated with the induction of hepatic cancer (Mandishona et al., 1998; Niederau et al., 1985; Stevens et al., 1994). Iron overload also promotes free-radical generation, leading to oxidative damage of biomolecules, functional alterations of sub-cellular organelles and loss of cell viability (Britton et al., 1987; Aust et al., 1985). These features of iron overload are observed in several tissues including the liver (Britton et al., 1987). Activation of Kupffer cells via red blood cell hemolysis or iron overload in the Kupffer cell results in production of cytokines, such as vascular endothelial growth factor (Urata et al., 2002), whose importance has been implicated in the induction of hemangiosarcomas in rodents (Nyska et al., 2002). In vitro studies have been performed to examine the role of macrophages on endothelial cells. RAW 264.7 cells were used as a model for Kupffer cells and were treated with LPS and hemolyzed red blood cells. The macrophages were activated by both treatments as measured by TNFα release (Corthals et al., 2006). Co-culture experiments allowed for products released from activated macrophages to be directly assessed in target endothelial cell populations without the activating agent confounding the interpretation of the results. LPS-activated macrophages increased endothelial cell damage, as measured by the Comet assay, further supporting the role for macrophage activation in the induction of DNA damage (Corthals et al., 2006). LPS- or hemolyzed red blood cell-activated macrophages co-cultured with endothelial cells also resulted in an increase in endothelial cell DNA synthesis (Corthals et al., 2006). These results support the hypothesis that products released from activated Kupffer cells may participate in the carcinogenesis process through DNA damage and/or cell proliferation, which allows for clonal growth of mutated cells and/or preneoplastic cell populations.
12.5
Summary
The Kupffer cell, the macrophage of the liver, plays an important role in the control and maintenance of normal liver homeostasis. The Kupffer cell exhibits multiple physiological and pathological functions through the production of cytokines and reactive oxygen species. The Kupffer cell removes endotoxin from circulation, particiates in the regeneration of the liver (compensatory hyperplasia), following either physical (partial hepatectomy) or chemical injury to the liver. Recent evidence confirms an important role for the Kupffer cell in mediating liver damage from acute and chronic alcohol exposure. Similarly, experimental evidence has demonstrated a participatory role for the Kupffer cell for the formation and promotion of hepatocellular and endothelial neoplasms in the liver.
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Van Rooijen, N. and Sanders, A. (1996). Kupffer cell depletion by liposome-delivered drugs: comparative activity of intracellular clodronate, propamidine, and ethylenediaminetetraacetic acid. Hepatology, 23: 1239–1243. Vandenabeele, P., Declercq, W., Beyaert, R. and Fiers, W. (1995). Two tumour necrosis factor receptors: structure and function. Trends Cell Biol 5: 392–399. VEGF (• • •). • • • • • • • • • • • • • • ••. Vrba, J. and Modriansky, M. (2002). Oxidative burst of Kupffer cells: target for liver injury treatment. Biomed Pap Med Fac Univ Palacky Olomouc Czech Repub 146: 15–20. Wallach, D. (1997). Cell death induction by TNF: a matter of self control. Trends Biochem Sci 22: 107–109. Wheeler, M. D. (2003). Endotoxin and Kupffer cell activation in alcoholic liver disease. Alcohol Res Health 27: 300–306. Wheeler, M. D. and Thurman, R. G. (1999). Production of superoxide and TNF-alpha from alveolar macrophages is blunted by glycine. Am J Physiol 277: L952–L959. Wheeler, M. D. and Thurman, R. G. (2003). Up-regulation of CD14 in liver caused by acute ethanol involves oxidant-dependent AP-1 pathway. J Biol Chem 278: 8435–8441. Wheeler, M., Stachlewitz, R. F., Yamashina, S., Ikejima, K., Morrow, A. L. and Thurman, R. G. (2000). Glycine-gated chloride channels in neutrophils attenuate calcium influx and superoxide production. Faseb J 14: 476–484. Widmann, J. J., Cotran, R. S. and Fahimi, H. D. (1972). Mononuclear phagocytes (Kupffer cells) and endothelial cells. Identification of two functional cell types in rat liver sinusoids by endogenous peroxidase activity. J Cell Biol 52: 159–170. Winwood, P. J. and Arthur, M. J. (1993). Kupffer cells: their activation and role in animal models of liver injury and human liver disease. Semin Liver Dis 13: 50–59. Wisse, E. (1972). An ultrastructural characterization of the endothelial cell in the rat liver sinusoid under normal and various experimental conditions, as a contribution to the distinction between endothelial and Kupffer cells. J Ultrastruct Res 38: 528–562. Wisse, E. (1974). Observations on the fine structure and peroxidase cytochemistry of normal rat liver Kupffer cells. J Ultrastruct Res 46: 393–426. Wisse E (1977). Ultrastructure and function of Kupffer cells and other sinusoidal cells in the liver. In: Kupffer Cells and Other Liver Sinusodial Cells (eds Wisse E., Knook DL.) Amsterdam, Elsevier/North Holland pp 33–60. Wisse, E., Braet, F., Luo, D., De Zanger, R., Jans, D., Crabbe, E. and Vermoesen, A. (1996). Structure and function of sinusoidal lining cells in the liver. Toxicol Pathol 24: 100–111. Yamamoto, T., Naito, M., Moriyama, H., Umezu, H., Matsuo, H., Kiwada, H. and Arakawa, M. (1996). Repopulation of murine Kupffer cells after intravenous administration of liposome-encapsulated dichloromethylene diphosphonate. Am J Pathol 149: 1271–1286. Yin, M., Ikejima, K., Arteel, G. E., Seabra, V., Bradford, B. U., Kono, H., Rusyn, I. and Thurman, R. G. (1998). Glycine accelerates recovery from alcohol-induced liver injury. J Pharmacol Exp Ther 286: 1014–1019. Yokoyama, H., Fukuda, M., Okamura, Y., Mizukami, T., Ohgo, H., Kamegaya, Y., Kato, S. and Ishii, H. (1999). Superoxide anion release into the hepatic sinusoid after an acute ethanol challenge and its attenuation by Kupffer cell depletion. Alcohol Clin Exp Res 23: 71S–75S. Zerbe, O. and Gressner, A. M. (1988). Proliferation of fat-storing cells is stimulated by secretions of Kupffer cells from normal and injured liver. Exp Mol Pathol 49: 87–101.
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13 Sinusoidal Cells in Liver Injury and Repair Carol R. Gardner and Debra L. Laskin
13.1
Introduction
The liver is the major reservoir of microsomal and cytosolic, phase I and phase II drugmetabolizing enzymes [1, 2]. Thus, it is particularly sensitive to toxicity induced by reactive metabolites generated from cytochrome P450 (CypP450) mediated oxidation of drugs such as acetaminophen and carbon tetrachloride (CCl4 ). Hepatic injury initiated by these agents is associated with an inflammatory response characterized by increased numbers of phagocytic leukocytes in the tissue and activation of hepatic sinusoidal cells. These cells release inflammatory cytotoxic cytokines and mediators that function to destroy invading pathogens, rid the liver of debris and, ultimately, initiate tissue repair. However, many of the mediators released during the inflammatory response are nonspecific. Thus, over- or unregulated production of these mediators may, in fact, contribute to hepatotoxicity. The mediators involved in the cytotoxic process include reactive oxygen and nitrogen intermediates, inflammatory cytokines, eicosanoids and proteolytic enzymes. While some of the mediators induce cytotoxicity directly (e.g. reactive oxygen intermediates, nitric oxide, peroxynitrite), others degrade extracellular matrix (e.g. collagenase, matrix metalloproteinases) and/or promote inflammation (e.g. chemokines). There is also evidence that inflammatory mediators can modify hepatocyte protein and nucleic acid biosynthesis, as well as cytochrome P450-mediated metabolism, which may exacerbate hepatotoxicity [3, 4]. Under homeostatic conditions, the liver is in a mitotically quiescent state. However, following injury, it has the capacity to regenerate, ultimately leading to repair and restoration of normal liver architecture. Evidence suggests that cells of the hepatic sinusoids, in particular
Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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macrophages, endothelial cells and stellate cells, are involved in this process. It appears that regeneration is mediated by cytokines and growth factors released from these cells. Thus, studies have shown that tumor necrosis factor-α (TNFα) and interleukin-6 (IL-6) released from Kupffer cells [5], together with hepatocyte growth factor released by stellate cells, induce proliferation of hepatocytes and regeneration of the liver [2, 5]. TNFα also stimulates hepatocytes to produce IL-6 and transforming growth factor-α (TGFα) which also contribute to liver regeneration following injury [6, 7]. During regeneration, multiple growth factors, signal transduction and transcription pathways are activated and large numbers of genes are upregulated. The complex control of this process has yet to be fully elucidated. In this chapter, mechanisms of injury and repair induced by two model toxicants, acetaminophen and CCl4 , are reviewed. Both of these hepatotoxicants are metabolized to reactive intermediates by cytochrome P-4502E1 (Cyp2E1) and both induce centrilobular hepatic necrosis. However, the mechanisms by which they induce injury are distinct, as are the roles of nonparenchymal cells and inflammatory mediators in the pathogenic process (Table 13.1). These findings demonstrate that the response of hepatic nonparenchymal and infiltrating cells is toxicant-specific, suggesting that distinct approaches will be needed to develop effective therapies to limit or prevent hepatotoxicity.
13.2 13.2.1
Hepatic Nonparenchymal Cells Kupffer Cells and Inflammatory Macrophages
Kupffer cells constitute approximately 20 % of the hepatic sinusoidal cells and represent the largest population (80–90 %) of macrophages in the body. They are predominantly localized in periportal and central regions of the liver lobule and are anchored to the lumen of the endothelium by long cytoplasmic processes [8]. Thus, they are well positioned to phagocytize particulate and foreign materials from the portal circulation. Kupffer cells possess Fc, C3 and CRIg receptors, as well as scavenger receptors, carbohydrate receptors, pattern recognition receptors, such as toll-like receptor-4 (TLR-4), and cell adhesion molecules, which facilitate their ability to phagocytize opsonized and nonopsonized particles, apoptotic and damaged cells, and tumor cells [9–25]. Kupffer cells are among the most active secretory cells in the body, releasing an array of products with inflammatory, growth-promoting and regulatory activity. These include the superoxide anion, hydrogen peroxide, nitric oxide, peroxynitrite, proteolytic enzymes and eicosanoids which aid in antigen destruction [10, 26–32]. They also release a number of different cytokines with immunoregulatory and proinflammatory actions, including TNFα, interleukin-1 (IL-1), IL-6, interleukin-8 (IL-8), interleukin-10 (IL-10), interleukin-18 (IL-18), platelet activating factor (PAF), transforming growth factor-β (TGFβ), macrophage inhibitory protein (MIP-2) and interferon [10, 33–37]. Kupffer cells also elaborate growth factors involved in regulating the proliferation of hepatocytes, endothelial cells, stellate cells and inflammatory macrophages [32, 38–42]. The ability of liver macrophages to release mitogenic factors for hepatocytes suggests that these cells may also participate in the wound repair process [43–45]. The capacity of the liver to regenerate is critical for the survival of intoxicated humans and animals; however, the underlying mechanisms have only recently been explored. Activation of Kupffer cells
Increased number, chemotaxis, phagocytosis, cytotoxicity, ROI, RNI, IL-1, IL-6, IL-10, TNFα, HO-1, eicosanoids, KC/gro, MCP-1, MIP-1 Increased number, ROI, RNI, IL-1β, IL-10, TNFα, IL-6, IL-13, HGF, TGFβ, osteopontin, COX-2, HO-1, catalase
Acetaminophen
CCl4
Increased ROI
230, 235–237, 241, 246–251, 256–258, 270, 273, 276, 278, 279, 281, 290–300
ROI, reactive oxygen intermediates; RNI, reactive nitrogen intermediates; IL, interleukin; TNF, tumor necrosis factor; HO-1, heme oxygenase-1; HGF, hepatocyte growth factor; PAF, platelet activating factor; COX-2, cyclooxygenase 2; IFN, interferon; MIP-1, macrophage inhibitory protein-1; TGF, transforming growth factor; FGF, fibroblast growth factor; TIMP-tissue inhibitors of metalloproteinase; PDGF, platelet derived growth factor; MCP-1, macrophage chemoattractant protein-1; NGF, nerve growth factor; ET-1, endothelin-1; MMP, matrix metalloproteinase; KC, keratinocyte-derived chemokine; IP-10, IFNγ -inducible protein-10; RANTES regulated upon activation, normal T-cell expressed, and presumably secreted; Mig, monokine induced by gamma.
Increased number, TIMP-1, α2-macroglobulin, pleiotropin, osteopontin, nitric oxide, TGFβ, PDGF, FGF, HGF, PAF, ET-1, IL-10, NGF, TNFα, collagenases, MMPs, Type I and Type III collagens
114, 139–141, 143–145, 147, 150, 162, 163, 165, 199, 202, 207, 285–289
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Increased Increased IFNγ , number, osteopontin, RNI KC, MIP-1α, MIP-2, MCP-1, RANTES, IP-10, Mig
Increased nitric oxide
Neutrophils References
NK/NKT cells
Endothelial cells Stellate cells
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Macrophages
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Mediator response
Cells and mediators of acetaminophen- and CCl4 -induced hepatic injurya
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appears to be required for optimal regenerative activity in the liver, most likely due to the release of TNFα and IL-6 [5, 46, 47]. These cytokines initiate hepatocyte proliferation, at least in part, via transcription factors such as NF-κB and STAT-3. Interestingly, it appears that these events may be triggered by leukocyte–Kupffer cell interaction mediated by the intracellular adhesion molecule-1 (ICAM-1) [48]. 13.2.2
Endothelial Cells
Endothelial cells form the walls of the hepatic sinusoids and represent the major fraction of hepatic sinusoidal cells (approximately 50 %). Unlike endothelial cells in other vascular beds, hepatic sinusoidal endothelial cells are devoid of basement membrane [49]. Moreover, they possess pores or fenestrae, which provide an opportunity for direct contact between plasma and hepatocytes. Thus, sinusoidal endothelial cells function as a selective barrier between the blood and the liver parenchyma. Endothelial cells also possess unique ‘bristle-coated’ membrane invaginations and vesicles, and lysosome-like vacuoles, and are thought to play a role in the clearance of macromolecules from the circulation. Through Fc, carbohydrate, scavenger receptors and TLRs, endothelial cells endocytose a variety of particles in the portal circulation, including glycoproteins, lipoproteins, albumin, lactoferrin, hyaluronic acid, collagen and aldehyde-modified proteins [11, 16, 21, 23, 50–61]. It has been reported that endocytosis is upregulated in endothelial cells when Kupffer cell functioning is impaired [51, 52, 54, 62–64]. In response to inflammatory cytokines and bacterially derived lipopolysaccharide (LPS), hepatic endothelial cells, like Kupffer cells, are ‘activated’ to release mediators that regulate the function of parenchymal and nonparenchymal liver cells. These include chemokines, IL-1, IL-6, PAF, fibroblast growth factor, interferons, endothelin, eicosanoids, proteolytic enzymes, reactive oxygen and nitrogen intermediates [27, 34, 38, 65–70]. In normal liver, endothelial cells express various adhesion molecules, including ICAM-1, ICAM-2, lymphocyte function-associated antigen-3 (LFA-3), very late antigen-5 (VLA-5) and vascular adhesion protein-1 (VAP-1) [71]. Expression of these molecules, as well as P-selectin, E-selectin and vascular cell adhesion molecule-1 (VCAM-1), which facilitate inflammatory cell emigration into the liver, is upregulated in sinusoidal endothelial cells activated following hepatotoxicant exposure [9, 71–73]. These findings suggest that endothelial cells are important in inflammatory responses in the liver. The observation that endothelial cells express CD40, CD54, CD80, CD86 and MHC class I and II molecules, which are markers of antigen-presenting cells, indicate that they may also play a role in immune surveillance and potentially in the development of tolerance in the liver [49, 74, 75]. 13.2.3
Stellate Cells
Stellate cells, also referred to as Ito cells, fat-storing cells, perisinusoidal cells and lipocytes, constitute approximately 20 % of the hepatic sinusoidal cells. These cells normally reside in a quiescent state within the space of Disse between endothelial cells and hepatocytes or between hepatocytes. Morphologically, stellate cells resemble fibroblasts in that they possess numerous extensions, as well as dilated rough endoplasmic reticulum. Stellate cells store vitamin A, which is localized in intracellular lipid droplets in the form of retinyl esters [76, 77]. Stellate cells also have the capacity to synthesize large quantities of extracellular matrix
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proteins, including types I, III, and IV collagen, as well as matrix metalloproteinases (MMP) and tissue inhibitors of metalloproteinase (TIMP), and there is evidence that they play a major role in collagen synthesis in both normal and fibrotic liver [78–82]. It has been suggested that stellate cells can contribute to inflammatory responses in the liver. Following exposure of animals to toxicants, stellate cells undergo a process of activation [83–85]. This involves a loss of lipid droplets and vitamin A storage capacity, migration to sites of liver injury, and transformation into highly proliferative myofibroblast-like cells [84, 86, 87]. Cell adhesion molecules, such as ICAM-1 and VCAM-1, as well as receptors for C5a, endothelin, eicosanoids, TNFα, IL-1 and platelet-derived growth factor (PDGF), are also upregulated on activated stellate cells [88–94]. Stellate cells are also primed, following exposure to liver toxicants, to release cytotoxic and inflammatory mediators, including IL-1, IL-6, IL10, PAF, colony-stimulating factor-1, nitric oxide, hydrogen peroxide, superoxide anion, eicosanoids, gelatinase, fibronectin, TGFβ, endothelin, macrophage chemotactic protein-1 (MCP-1) and cytokine-induced neutrophil chemoattractants (CINC) [78, 95–102]. Activation and transformation of stellate cells during the pathogenesis of tissue injury and fibrosis, as well as collagen deposition, appear to be mediated by cytokines and growth factors elaborated by parenchymal and nonparenchymal liver cells. These are largely divided into mitogenic mediators (TGFα, PDGF, IL-1, TNFα and insulin-like growth factor) that stimulate proliferation and transformation of stellate cells, and fibrogenic mediators, including TGFβ and IL-6, that induce collagen gene expression [93]). Hepatic fibrosis represents an exaggerated form of the liver’s wound healing response to injury and is characterized by excessive accumulation of interstitial matrix components within the tissue. A number of factors have been proposed to initiate and perpetuate the fibrogenic process in stellate cells, including the accumulation of inflammatory cytokines and growth factors, alterations in the extracellular matrix and oxidative stress [103]. Activation of stellate cells can occur through the generation of lipid peroxides from damaged hepatocytes and/or oxidants and cytokines released from activated Kupffer cells and inflammatory macrophages [104, 105]. Recent studies have shown that stellate cells have the capacity to phagocytose apoptotic bodies derived from damaged hepatocytes, which markedly stimulates expression of TGFβ1 mRNA and collagens and induces NADPH oxidase [106, 107]. During the pathogenesis of fibrosis, stellate cells exhibit increased sensitivity to inflammatory mediators, such as TNFα [108]. This leads to the production of chemotactic and fibrogenic mediators by liver cells and may contribute to the maintenance of an inflammatory infiltrate dominated by macrophages [105, 109].
13.2.4
Natural Killer Cells
Natural killer cells (NK) and natural killer cells with T-cell receptors (NKT) are found in great abundance in the liver and comprise nearly 20 % of liver leukocytes [19, 110]. NK cells are important in innate immune responses; thus, they release cytokines and chemokines, and can lyse transformed cells [110, 111]. NKT cells consist of a subpopulation of lymphocytes that coexpress NK and T-cell receptors [110, 112]. Both NK and NKT cells are readily activated for cytotoxicity and generate large quantities of interferon-γ (IFNγ ) and have been implicated in toxicant-induced liver injury [113–115].
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Acetaminophen Hepatotoxicity
Acetaminophen is the most widely used analgesic and antipyretic agent in the United States and is the drug most commonly ingested in intentional and accidental overdoses. This is due to the fact that acetaminophen is frequently found in combination with many over-the-counter and prescription medications. Although considered to be safe and efficacious at therapeutic doses [116], when taken in excess, hepatotoxicity can develop and in fact, acetaminophen-induced acute hepatic failure is the leading cause of acute liver failure among patients referred for liver transplantation [117]. The majority of ingested acetaminophen is conjugated with sulfate or glucuronide and then excreted in the bile and urine [118, 119]. A smaller portion, however, is metabolized via CypP450, in particular, Cyp2E1, predominantly localized in centrilobular regions of the liver lobule, to a highly reactive intermediate, N -acetyl- p-benzoquinone-imine (NAPQI) [120, 121]. After conjugation to reduced glutathione, NAPQI is also eliminated in the bile [122]. At toxic doses of acetaminophen, the sulfation and glucuronidation pathways become saturated leading to increased generation of NAPQI, depletion of hepatic glutathione and tissue injury [123]. Hepatotoxicity is due to binding of NAPQI to critical cellular macromolecules, e.g. DNA and in mitochondria [124–128]. This leads to mitochondrial dysfunction and DNA damage, characterized by inhibition of mitochondrial respiration, declining ATP levels and excessive generation of reactive oxygen intermediates [129–135]. These perturbations initiate hepatocellular necrosis observed in centrilobular regions of the liver [136–138]. The importance of the Cyp2E1 pathway in hepatotoxicity is most clearly demonstrated by findings that mice lacking the gene for Cyp2E1 are protected from acetaminophen-induced hepatic necrosis [120]. 13.3.1
Role of Kupffer Cells and Infiltrating Macrophages in Acetaminophen-Induced Heptotoxicity
A growing body of literature generated over the past two decades has provided strong evidence implicating activated liver macrophages, including both resident Kupffer cells and infiltrating cells in hepatotoxicity induced by acetaminophen. The early observation that there were increased numbers of macrophages in centrilobular regions of the liver following administration of hepatotoxic doses of acetaminophen to rodents, and that they appeared prior to histological evidence of frank necrosis fueled this area of investigation [26, 139]. When these macrophages were isolated they were found to be larger and more stellate than cells from control animals, were highly vacuolated and displayed an increased cytoplasmic:nuclear ratio [26]. These are classic signs of activated macrophages. In addition, macrophages from animals treated with acetaminophen exhibit functional activation, including increased adherence, phagocytosis, chemotaxis and cytotoxicity, as well as excessive generation of inflammatory mediators, including reactive oxygen and nitrogen intermediates, IL-1, TNF α, MCP-1, KC/gro and macrophage inflammatory protein-1 (MIP-1) [3, 26, 139–145]. These data demonstrate that macrophages accumulating in the liver after acetaminophen are also functionally activated. Additional evidence for a role for activated macrophages in acetaminophen-induced hepatotoxicity is based on results of experiments in which animals were pretreated with agents that inhibit or deplete macrophages. In these studies, the accumulation of macrophages in
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the liver and subsequent toxicity of acetaminophen was prevented by pretreatment of animals with gadolinium chloride, dextran sulfate or liposome-encapsulated dichloromethylene diphosphonate, compounds known to depress macrophage activity [137, 146–148]. Hepatoprotection by macrophage inhibition was reported to be associated with decreased production of inflammatory mediators, including TNFα, reactive oxygen and nitrogen species, suggesting that these mediators are important in the pathogenic process [147, 148]. This is supported by findings that blocking production of these inflammatory mediators is also protective against acetaminophen-induced toxicity. Thus, acetaminophen-induced hepatotoxicity is suppressed in transgenic mice lacking inducible nitric oxide synthase (NOS-2), or animals pretreated with inducible nitric oxide synthase inhibitors, aminoguanidine, ONO-1714 or l-N6-(1-iminoethyl)-lysine (L-NIL) [140, 141, 149, 150]. Similarly, mice over expressing superoxide dismutase (SOD) or treated with antioxidants such at β-d-glucan, resveratrol, carvedilol, green tea polyphenols, S-adenosylmethionine, (RS)-npropylthiazolidine-4(R)-carboxylic acid or N -acetylcysteine, also exhibit hepatoprotection [143, 151–155]. Immunosuppressive agents, such as corticosteroids (e.g. prednisolone and pregnenolone-16 α-carbonitrile), as well as eicosanoids have also been found to protect against liver toxicity caused by acetaminophen [156–160], providing additional support for the role of inflammatory mediators in hepatotoxicity. IL-10 is an anti-inflammatory cytokine known to down-regulate macrophage activity [161]. A number of studies have shown that levels of IL-10 increase in the liver following acetaminophen administration [141–143, 162–164]. Moreover, knockout mice lacking IL-10 are hypersensitive to acetaminophen-induced liver injury [164]. These findings also indicate that macrophages and inflammatory mediators are important in acetaminopheninduced hepatotoxicity. This is supported by findings that NOS-2 knockout mice, which are less sensitive to acetaminophen-induced toxicity, express increased levels of IL-10 [141]. Recent studies have suggested that some populations of macrophages accumulating in the liver in response to hepatotoxic doses of acetaminophen play a role in the resolution of inflammation and in initiation of tissue repair. These infiltrating macrophage subpopulations are antigenically and phenotypically different from resident Kupffer cells and release a distinct spectrum of inflammatory and growth-promoting mediators [5, 144, 162]. Moreover, blocking the recruitment of these macrophage subpopulations into the liver augments tissue injury induced by acetaminophen [144, 162, 165]. These observations support a dual role of macrophages in hepatotoxicity, initially contributing to tissue injury and subsequently orchestrating tissue repair [43, 162]. A question arises as to the nature of the mediators released by macrophages that are important in tissue repair. One mediator of particular interest is TNFα. Although this cytokine has classically been identified as a proinflammatory mediator, more recent studies suggest that TNFα is key in the maintenance of homeostasis and in initiation of tissue repair [166]. Thus, TNFα is a potent mitogen, stimulating hepatocyte proliferation following acute injury [46, 167–169]. TNFα also stimulates macrophages and other nonparenchymal and parenchymal cells in the liver to produce mediators important in extracellular matrix remodeling, including transforming growth factor-α (TGFα), connective tissue growth factor (CTGF) and matrix metalloproteinase-9 (MMP-9). These findings, together with the observation that TNFα receptor knockout mice are markedly more sensitive to acetaminophen-induced hepatotoxicity, demonstrate the importance of TNFα in the repair of damaged tissue [142, 170, 171]. Another macrophage-derived cytokine that appears to be involved in tissue
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repair after acetaminophen-induced injury is IL-6. Mice lacking the gene for this cytokine have been reported to exhibit delayed hepatocyte regeneration following acetaminophen intoxication [172]. Recent studies have focused on analyzing mechanisms regulating macrophage activation following acetaminophen administration. It has been suggested that this process involves inappropriate or prolonged activation of biochemical signaling pathways in the cells leading to increased gene expression and inflammatory mediator production. For example, following acetaminophen administration to animals, a rapid increase in nuclear binding activity of the transcription factor, NF-κB has been observed in the liver [143, 170, 173, 174]. NF-κB is a ubiquitous transcription factor known to regulate the activity of numerous genes involved in inflammatory responses and implicated in toxicity, including NOS-2, cyclooxygenase-2 (COX-2), TNFα and ICAM-1 [175]. Interestingly, blockade of NF-κB activation or the loss of NF-κB p50 has been reported to have no effect on acetaminopheninduced hepatotoxicity suggesting that NF-κB-dependent responses do not play a major role in the pathogenesis of toxicity in this model [143, 174]. Of note, studies have also suggested that activation of the NF-κB pathway is also not required for hepatocyte proliferation [176]. Increased nuclear binding activity of the transcription factor AP-1 has also been described in the liver after treatment of animals with acetaminophen [170, 177]. The fact that this activity is prevented by pretreatment of animals with gadolinium chloride suggests that Kupffer cells are important in this response. Other signaling molecules implicated in macrophage responsiveness to acetaminophen, and in particular their role in wound repair, include the transcription factors c-jun and c-fos which regulate a battery of genes involved in cell proliferation, as well as STAT3 and early growth reponse-1 (Egr-1). These transcription factors have been reported to be activated in the liver following administration of acetaminophen [171, 178] and their activity may be important in determining the outcome of the hepatotoxic response. Another signaling pathway that is involved in the progression of acetaminophen-induced liver injury is Fas/FasL. Activated macrophages and other inflammatory cells can express FasL [114, 115, 179, 180]. These cells may directly kill Fas-expressing hepatocytes [115]. Alternatively, Fas–FasL crosslinking between inflammatory cells and hepatocytes or between the inflammatory cells themselves within the liver may result in continuous production of cytokines and chemokines [114, 181–183], which would further potentiate acetaminophen-induced injury. 13.3.2
Endothelial Cells and Acetaminophen-Induced Hepatotoxicity
Endothelial cells are targets, as well as effector cells, of acetaminophen-induced liver injury. As disruption in hepatic blood flow precedes acetaminophen-induced necrosis, it has been suggested that congestion contributes to toxicity [184, 185]. Following acetaminophen administration, large pores appear in endothelial cells and they partially detach from underlying hepatocytes [185]. In vivo microscopy has also revealed rounding up or swelling of the sinusoidal endothelial cells within 30 min of acetaminophen administration [186]. These alterations are thought to be due to the secretion of MMP-2 and MMP-9 by sinusoidal cells [187]. Gaps through the endothelial cells have been shown to be formed by the destruction and/or coalescence of fenestrae which allow red blood cells to penetrate into the space of Disse [188]. Sinusoidal endothelial cells also rapidly lose their ability
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to endocytose albumin following acetaminophen administration [188]. The toxicity of acetaminophen towards endothelial cells is due, in part, to the presence of Cyp2E1 in these cells and rapid depletion of intracellular glutathione levels [189]. Sinusoidal endothelial cells secrete a wide variety of inflammatory mediators that may contribute to hepatotoxicity [27, 34, 38, 65–70]. Of particular note is nitric oxide synthase which is generated in these cells by both inducible NOS-2 and constitutive or endothelial nitric oxide synthase (NOS-3) [188, 190] and appears to play a dual role in acetaminophen-induced hepatotoxicity. Thus, wild type mice treated with the NOS-3 inhibitor, NG -nitro-l-arginine methyl ester (L-NAME), or transgenic mice lacking the gene for NOS 3 are hypersensitive to acetaminophen-induced hepatotoxicity [150, 191]. In contast, mice treated with L-NIL, a specific NOS-2 inhibitor, as well as NOS-2 knockout mice, are protected from hepatocellular damage [141, 150, 191]. These data suggest that the role of endothelial cell-derived nitric oxide in acetaminophen-induced hepatotoxicity depends on the levels of this mediator generated and the enzyme regulating its synthesis. Angiogenesis is required for the repair of injured liver [192]. Mice treated with acetaminophen have been reported to express increased levels of vascular endothelial growth factor-A (VEGF-A), as well as receptors for this growth factor in the liver [193]. VEGF is a potent endothelial cell mitogen secreted by hepatocytes [194–196]. The angiogenic effects of VEGF are mediated by NOS-3 in sinusoidal endothelial cells [197]. It has also been suggested that VEGF induces the release of the hepatic mitogens, IL-6 and hepatocyte growth factor from endothelial cells [194, 198]. The ability of sinusoidal endothelial cells to regenerate and produce hepatocyte growth factors and cytokines may represent an important mechanism by which these cells contribute to repair. 13.3.3
Natural Killer Cells and Acetaminophen-Induced Hepatotoxicity
Recent studies have implicated NK and NKT cells in the progression of acetaminopheninduced hepatotoxicity. Liu et al. [114] reported that depletion of NK and NKT cells resulted in significant protection against acetaminophen-induced injury in mice. These investigators also identified NK and NKT cells as the source of hepatic IFNγ in acetaminophen-treated mice which suggests that this cytokine may be important in hepatotoxicity. In this regard, mice lacking the gene for IFNγ have been reported to exhibit decreased hepatotoxicity in response to acetaminophen [114, 145]. In addition to its antiviral and antibacterial activities, IFNγ activates macrophages and NK cells, and upregulates MHC class II expression on macrophages, and these activities may be important in its hepatotoxic actions in the liver. Activated NK and NKT cells also mediate the recruitment of neutrophils, macrophages and additional NK and NKT cells into the liver [114]. This is likely due to production of chemokines and other inflammatory proteins, including keratinocyte-derived chemokine (KC), MIP-1α, MIP-2, MCP-1, RANTES, IFNγ -inducible protein (IP-10) and Mig by these cells [114]. Welch et al. [199] recently showed that liver injury induced by acetaminophen is associated with increased expression of osteopontin, an inflammatory mediator, secreted by NKT cells, which regulates the severity of liver injury and neutrophil infiltration into the liver in other models [200]. Osteopontin knockout mice were also reported to be less susceptible to acetaminophen-induced hepatotoxicity when compared to wild type mice [199]. Taken together, these data provide evidence that hepatic NK and NKT cells participate in acetaminophen-induced liver injury.
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Infiltrating Neutrophils and Acetaminophen-Induced Hepatotoxicity
Although it is well documented that neutrophils accumulate in the liver after acetaminophen administration to rodents, their role in liver injury is controversial [114, 145, 201, 202]. The accumulation of neutrophils in areas of hepatic necrosis following acetaminophen administration was first reported by Mitchell et al. [203]. It was, however, subsequently shown that there was no evidence of neutrophil activation in the liver following acetaminophentreatment of mice, and that depletion of neutrophils using anti-CD-18 antibody or anti-GR-1 antibody did not protect against acetaminophen-induced liver injury [163, 202]. It has also been reported that blockade of neutrophil accumulation in acetaminophen-intoxicated rats had no effect on liver injury [174]. Similarly, ICAM-deficient mice, which exhibit decreased neutrophil adherence, were not protected from acetaminophen-induced injury [163]. High mobility group box 1 (HMGB1) protein is a potent mediator of inflammation that is secreted by activated monocytes and macrophages and induces neutrophil accumulation [204, 205]. Mice treated with anti-HMGB1 antibody were not protected from acetaminophen-induced liver injury [205]. Taken together, these data suggest that infiltrating neutrophils only remove cell debris and do not contribute to liver damage [202]. In contrast to these findings, Smith et al. reported protection against acetaminopheninduced liver injury in rats with anti-neutrophil antisera [201]. Hepatoprotection was also observed when animals were pretreated with anti-GR-1 antibody [206–208]. The protection was associated with reduced NOS-2 expression in the liver [207]. Ishida et al. [207] also reported that mice lacking CXCR2, a chemokine receptor that is key for neutrophil emigration, are protected against acetaminophen-induced hepatotoxicity. Moreover, ICAM-1deficient mice were resistant to acetaminophen-induced hepatotoxicity [208]. Thus, it is apparent that neutrophils are involved in acetaminophen-induced hepatotoxicity; however, their precise role remains to be determined.
13.4
Carbon Tetrachloride Hepatotoxicity
Carbon tetrachloride (CCl4 ) is a haloalkane that was widely used as a solvent, cleaner and degreaser [209]. Exposure to CCl4 causes severe liver injury, including inflammation, centrilobular necrosis, steatosis and cirrhosis [209, 210]. The toxicity of CCl4 , is due to loss of calcium sequestration, impairment of lipid homeostasis, lipid peroxidation, release of pro- or anti-inflammatory cytokines and apoptosis. Whereas, at low doses, toxicity is transient and reversible, at higher doses or after prolonged exposure, irreversible fatty degeneration, fibrosis and cirrhosis are observed [209]. The hepatotoxicity of CCl4 , like acetaminophen, is dependent on CypP450-mediated metabolism. Through the actions of Cyp2E1, and to a lesser extent, Cyp2B1 and Cyp2B2, a trichloromethyl free radical (CCl3 • ) is generated [209, 211–213]. Findings that Cyp2E1 knockout mice are less sensitive to the effects of CCl4 than wild type mice confirm the importance of this metabolic pathway and the generation of CCl3 • in hepatotoxicity [214]. CCl4 induced hepatotoxicity is thought to be due to the reactions of a CCl3 • radical with amino acids, nucleotides, fatty acids, proteins, nucleic acids and lipids. This involves haloalkylation and lipid peroxidation [209]. In the presence of oxygen, the CCl3 • radical can also
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be converted to the trichloromethyl peroxy radical, CCl3 OO• , which is more reactive and short-lived [215] and may, in fact, initiate the process of lipid peroxidation [216–218]. It has long been recognized that CCl4 affects intracellular calcium levels within hepatocytes [219]. Hepatocellular calcium is sequestered into three main compartments, the cytosol, endoplasmic reticulum and mitochondria, each with its own ion pumping system [209]. High or repeated CCl4 exposure results in depletion of the ATP supply, inactivation of plasma membrane Na+ /K+ and Ca2+ -ATPase activities, and uncontrolled activation of protein kinase C which suppresses lipid secretion [220, 221]. Uncontrolled levels of calcium destroy cytoskeletal structure and activate catabolic enzymes, such as proteases, endonucleases and phospholipases, resulting in necrotic cell death [222–224]. The extracellular matrix consists of a large number of macromolecules that comprise the scaffolding of normal and fibrotic liver and include collagens, glycoproteins and proteoglycans [225]. Fibronectin and laminin are among the glycoproteins present, while the proteoglycans include heparan, dermatan, chondroitin sulfate, perlecan and endoglin [226, 227]. The matrix of the normal liver is composed of non-fibril-forming collagens, including types IV, VI and XIV [228, 229]. Fibrosis is characterized by an increase in the synthesis and deposition of extracellular matrix proteins which interferes with hepatic functioning [230]. As fibrosis develops, the total content of collagens and noncollagenous components increases three- to five-fold [231]. In addition, there is a shift in the type of extracellular matrix from the low density membrane-like matrix to an interstitial type, consisting predominantly of type I collagen or fibrillar collagen [232, 233]. Initiation of the fibrotic process involves Kupffer cells, endothelial cells and stellate cells, as well as the various inflammatory mediators and growth factors produced by these cells [101, 230]. 13.4.1
Kupffer Cells and Infiltrating Macrophages in CCl4 -Induced Heptotoxicity
Following CCl4 -intoxication, there is a significant increase in the number of macrophages in the liver which appears to be due to proliferation of resident Kupffer cells and recruitment of peripheral monocytes/macrophages [234, 235]. In situ immunochemical localization studies indicate that there are increased numbers of cells in the liver expressing the macrophage-associated markers, ED1, ED2 and TNFα [235]. This was associated with the development of hepatic necrosis. Macrophages isolated from the livers of CCl4 -treated animals were found to be larger and more adherent than cells from control animals [236]. Expression of CD14, ED1, ED2, ICAM-1 and peroxisome proliferator-activated receptors (PPARs), as well as receptors for PAF and endothelin-1 (ET-1) were also upregulated on these cells [235–241]. These findings suggest that macrophages are activated during the pathogenesis of CCl4 -induced liver injury. Direct evidence for a role of Kupffer cells and infiltrating macrophages in CCl4 -induced toxicity has been obtained from studies in which animals have been selectively depleted of Kupffer cells. In these studies, mice were protected from CCl4 -induced steatosis, inflammation, necrosis and/or cirrhosis [242–245]. Following CCl4 administration, liver macrophages have been reported to produce a number of pro- and anti-inflammatory mediators, including TNFα, nitric oxide, superoxide anion, TGFβ, IL-1β, IL-6, IL-10, IL-13, hepatocyte growth factor (HGF), and osteopontin, and evidence implicates many of these mediators in the pathogenic process [236, 237, 241, 246–251]. For example, it has been reported that inhibition of NOS-2
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using aminoguanidine protects mice from CCl4 -induced liver injury [252]. In contrast, mice lacking the gene for NOS-2 were found to be more sensitive to the effects of CCl4 , thus suggesting that nitric oxide may be protective in some stages of the pathogenic process [246]. Increases in serum levels of TNFα have also been described following CCl4 intoxication [237, 246, 253]. The idea that this cytokine plays a role in the pathogenesis of liver injury is supported by studies demonstrating that soluble TNFα receptors, which block the actions of TNFα, or mice lacking the gene for TNFα or TNFα receptor I (TNFRI) are protected against CCl4 -induced hepatotoxicity [246, 254]. It has also been reported that NF-κB p50 knockout mice are hypersensitive to CCl4 and that this is associated with increased release of TNFα [255]. These findings suggest that inappropriate TNFα production in these mice is due to a lack of p50-dependent histone deacetylase 1-mediated repression of TNFα gene transcription [255]. TNFα also plays a significant role in repair and recovery from hepatic damage initiated by CCl4 . Thus TNFRI and TNFRI/RII knockout mice exhibit significantly impaired hepatocyte proliferation and exaggerated hepatotoxicity in response to CCl4 [253]. Taken together, these data indicate that TNFα is responsible for both potentiating liver damage during CCl4 intoxication and for initiating tissue repair. Other macrophage-derived proteins which are upregulated in the liver after CCl4 exposure include COX-2, which generates eicosanoids, and heme-oxygenase-1 (HO-1) and catalase which are involved in anti-oxidant defense [237, 256–258]. Whereas eicosanoids produced via COX-2 have been implicated in portal hypertension [256], HO-1 and catalase appear to be important in hepatoprotection. Thus, inhibition of HO-1 results in increased CCl4 induced liver injury [259]. Similarly, mice lacking the gene for catalase were found to be more sensitive to CCl4 -induced liver injury [260]. Liver macrophages from CCl4 -intoxicated animals also synthesize large quantities of fibronectin and produce factors that are fibrogenic for stellate cells (e.g. TGFβ), as well as factors which activate stellate cells [230, 261, 262]. This is important in tissue repair and also in fibrosis. That macrophages contribute to hepatic fibrosis is evidenced by findings that animals depleted of these cells exhibit a reduction in deposition of collagen III and elastin after chronic CCl4 administration [263]. In contrast, depletion of macrophages during recovery from fibrosis has been reported to lead to persistence of collagen III and elastin in the liver, suggesting that macrophages are also important in tissue repair [263]. Taken together, these data support the concept that liver macrophages contribute to both tissue injury/fibrosis and to tissue repair after CCl4 administration. 13.4.2
Stellate Cells and CCl4 -Induced Hepatotoxicity
Chronic CCl4 intoxication is associated with activation of normally quiescent stellate cells [264]. This is characterized by increased smooth muscle α-actin expression [265]. In addition, stellate cells begin to proliferate in centrilobular regions of the liver lobules and in bands of tissue between adjacent central veins. During this process, stellate cells take on a fibroblast-like appearance, release nitric oxide and produce increased amounts of collagens, including collagens I and III, thus promoting hepatic fibrosis [266–270]. Activated stellate cells also generate TIMP-1 which contibutes to a net deposition of fibril-forming collagens [270]. Expression of tropoelastin by stellate cells is also increased during the course of CCl4 -induced fibrogenesis [271]. These data, together with findings that mice
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overexpressing TIMP-1 exhibit a reduced ability to undergo spontaneous resolution of fibrosis following cessation of CCl4 , provide support for a role of stellate cells in the fibrogenic process [272]. Stellate cells also produce factors during the course of CCl4 intoxication which promote the inflammatory response, thus contributing to hepatic necrosis. Thus, expression of pleiotropin, a heparin-binding protein which induces growth, angiogenesis, differentiation and transformation of cells, and osteopontin, are upregulated in hepatic stellate cells following CCl4 treatment of rats [241, 273]. After CCl4 administration, the contractility of stellate cells increases, leading to increased portal resistance [274]. This is caused by stellate cell constriction of individual sinusoids and contraction the cirrhotic liver [275]. ET-1 is the major stimulus for contraction of stellate cells, whereas, nitric oxide is the physiologic antagonist of this mediator [275, 276]. Stellate cells produce nitric oxide after CCl4 -induced activation [276]; however, as liver disease progresses, ET-1 increases and nitric oxide production decreases leading to further disruption of blood flow [277]. Evidence suggests that stellate cell activation during CCl4 -induced injury is regulated by autocrine cytokines, including TGFβ1 , PDGF, fibroblast growth factor (FGF), HGF, PAF and ET-1 [278]. Anti-inflammatory cytokines, such as IL-10, are also generated by activated stellate cells [248, 279]. IL-10 reduces the expression of collagen I while increasing interstitial collagenase, thus exerting antifibrotic activity [230]. Moreover, IL-10 knockout mice exhibit more severe liver injury following CCl4 administration [248, 279]. Upregulation of stellate cell function during CCl4 intoxication involves activation of many intracellular signaling pathways. Recent studies have shown that treatment of mice with CCl4 results in decreased expression of RelB and weak induction of p52 (NF-κB2) mRNA in stellate cells [255]. Oakley et al. [255] have also reported that the p50 subunit of NF-κB1 is protective in that it limits TNFα production and neutrophilic infiltration. It has also been shown that stellate cells express both Fas and FasL [280] which may increase their susceptiblity to apoptosis. A key feature of recovery from liver fibrosis is hepatic stellate cell apoptosis, which effectively removes a major source of new matrix proteins and TIMPs from the liver, thereby facilitating matrix degradation. Recent evidence suggests that some growth factors can induce apoptosis in stellate cells leading to recovery from fibrosis. After CCl4 administration, hepatic stellate cells express receptors for nerve growth factor (NGF), an important mediator of tissue repair generated by hepatocytes. Moreover, maximal production of NGF by hepatocytes coincides with the peak of stellate cell apoptosis [281]. 13.4.3
Infiltrating Neutrophils and CCl4 -Induced Hepatotoxicity
Neutrophils have been strongly implicated in the development of liver damage and stellate cell activation during CCl4 -induced liver damage. This is thought to be due to their ability to produce reactive oxygen species and various hydrolytic enzymes [282, 283]. One of the most potent mediators of hepatic accumulation and activation of neutrophils is TNFα [284]. It has been reported that the livers of CCl4 -injured mice lacking the gene for the NF-κB p50 subunit express higher levels of TNFα and that these mice have significantly increased numbers of neutrophils in damaged areas when compared with wild type mice [255]. In addition, hepatic stellate cells from p50 knockout mice express increased TNFα
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and this is associated with excessive fibrogenesis, indicating that NF-κB controls TNFα production in these cells [255].
13.5
Conclusions
Evidence has accumulated over the past several years demonstrating that chemically induced toxicity is a multifactorial process involving direct tissue injury, as well as a cascade of protein and lipid mediators generated by cells in the liver. These include not only resident cells (hepatocytes, Kupffer cells, stellate cells, endothelial cells and NK cells), but also infiltrating leukocytes (macrophages and neutrophils). Proliferation and activation of these cells following exposure to toxicants like acetaminophen and CCl4 can augment injury of hepatocytes and potentate liver damage. Inflammatory mediators, including cytokines, growth factors and reactive mediators, such as nitric oxide, superoxide anion, hydrogen peroxide and eicosanoids, produced by ‘activated’ nonparenchymal cells and/or infiltrating leukocytes may be cytotoxic, pro-inflammatory and can compromise normal liver functioning or may help restore normal tissue structure and function. Defining the precise role of each of these mediators in tissue injury and tissue repair is essential for our understanding of the mechanism of action of hepatotoxic chemicals and for devising steps to prevent or abrogate toxicity.
Acknowledgements This work was supported by USPHS National Institutes of Health Grants GM034310, ES004738, ES005022 and AR055073.
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[284] M.L. Bajt, A. Farhood and H. Jaeschke, Effects of CXC chemokines on neutrophil activation and sequestration in hepatic vasculature. Am. J. Physiol. Gastrointest. Liver Physiol. 281, G1188–G1195 (2001). [285] D.L. Laskin, Nonparenchymal cells and hepatotoxicity. Semin. Liver Dis. 10, 293–304 (1990). [286] M.E. Blazka, M.R. Elwellk, S.D. Holladay, R.E. Wilson and M.I. Luster, Histopathology of acetaminophen-induced liver changes: Role of interleukin 1α and tumor necrosis factor α. Toxicol. Pathol. 24, 181–189 (1996). [287] M.E. Blazka, J.L. Wilmer, S.D. Holladay, R.E. Wilson and M.I. Luster, Role of proinflammatory cytokines in acetaminophen hepatotoxicity. Toxicol. Appl. Pharmacol. 133, 43–52 (1995). [288] H. Chiu, J.A. Brittingham and D.L. Laskin, Differential induction of heme oxygenase-1 in macrophages and hepatocytes during acetaminophen-induced hepatotoxicity in the rat: effects of hemin and biliverdin. Toxicol. Appl. Pharmacol. 181, 106–115 (2002). [289] D. Nakae, K. Yamamoto, H. Yoshiji, T. Kinugasa, H. Maruyama, J.L. Farber and Y. Konishi, Liposome-encapsulated superoxide dismutase prevents liver necrosis induced by acetaminophen. Am. J. Pathol. 136, 787–795 (1990). [290] W.D. Thompson, A.S. Jack and R.S. Patrick, The possible role of macrophages in transient hepatic fibrogenesis induced by acute carbon tetrachloride injury. J. Pathol. 130, 65–73 (1980). [291] S. Camandola, M. Aragno, J.C. Cutrin, E. Tamagno, O. Danni, E. Chiarpotto, M. Parola, G. Leonarduzzi, F. Biasi and G. Poli, Liver AP-1 activation due to carbon tetrachloride is potentiated by 1,2-dibromoethane but is inhibited by alpha-tocopherol or gadolinium chloride. Free Radic. Biol. Med. 26, 1108–1116 (1999). [292] A.E. elSisi, D.L. Earnest and I.G. Sipes, Vitamin A potentiation of carbon tetrachloride hepatotoxicity: role of liver macrophages and active oxygen species. Toxicol. Appl. Pharmacol. 119, 295–301 (1993). [293] W. Chamulitrat, M.E. Blazka, S.J. Jordan, M.I. Luster and R.P. Mason, Tumor necrosis factoralpha and nitric oxide production in endotoxin-primed rats administered carbon tetrachloride. Life Sci. 57, 2273–2280 (1995). [294] A. Katz, J. Chebath, J. Friedman and M. Revel, Increased sensitivity of IL-6-deficient mice to carbon tetrachloride hepatotoxicity and protection with an IL-6 receptor-IL-6 chimera. Cytokines Cell Mol. Ther. 4, 221–227 (1998). [295] I.G. Sipes, A.E. el Sisi, W.W. Sim, S.A. Mobley and D.L. Earnest, Reactive oxygen species in the progression of CCl4 -induced liver injury. Adv. Exp. Med. Biol. 283, 489–497 (1991). [296] W. Zhu and P.C. Fung, The roles played by crucial free radicals like lipid free radicals, nitric oxide and enzymes NOS and NADPH in CCl4 -induced acute liver injury of mice. Free Radic. Biol. Med. 29, 870–880 (2000). [297] C.A. Kawser, J.P. Iredale, P.J. Winwood and M.J. Arthur, Rat hepatic stellate cell expression of alpha2-macroglobulin is a feature of cellular activation: implications for matrix remodelling in hepatic fibrosis. Clin. Sci. (London) 95, 179–186 (1998). [298] J.P. Iredale, Matrix turnover in fibrogenesis. Hepatogastroenterology 43, 56–71 (1996). [299] J.P. Iredale, R.C. Benyon, M.J. Arthur, W.F. Ferris, R. Alcolado, P.J. Winwood, N. Clark and G. Murphy, Tissue inhibitor of metalloproteinase-1 messenger RNA expression is enhanced relative to interstitial collagenase messenger RNA in experimental liver injury and fibrosis. Hepatology 24, 176–184 (1996). [300] J.P. Iredale, R.C. Benyon, J. Pickering, M. McCullen, M. Northrop, S. Pawley, C. Hovell and M.J. Arthur, Mechanisms of spontaneous resolution of rat liver fibrosis. Hepatic stellate cell apoptosis and reduced hepatic expression of metalloproteinase inhibitors. J. Clin. Invest. 102, 538–549 (1998).
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14 Cytokines in Liver Diseases Pablo Muriel
14.1
Introduction
14.1.1
Cytokines
Cytokines are extracellular proteins involved in the regulation of innate and immunologically inflammatory reactions. These proteins are involved in various processes including cell differentiation, development, growth and repair processes that lead to the maintenance or recuperation of homeostasis (Gao, 2005). Production and secretion of most cytokines are of short duration, and basal production is absent or occurs at very low levels. Many cytokines influence different cell types by binding to specific receptors that transduce signals into the target cell. Cytokine receptors operate as tyrosine kinase and tyrosine-kinase-associated receptors or serine/threonine kinase associated receptors or G-protein-linked receptors. Cytokines are generally active over a short space by binding to the cell of their origin or to a neighboring cell. The function of cytokines consists in mediating interactions between inflammatory and immune systems. A specific immune response triggers a local inflammatory effect, and vice versa, that is, a local inflammatory response induces a specific immune response in the tissue. Interleukin (IL)-1 or tumor necrosis factor (TNF), for instance, produced by macrophages at the site of inflammation induces endothelial cells to produce cell adhesion molecules that in turn make contact with lymphocytes, inducing their extravasation. On the other hand, resident or stroma cells undergoing activation produce cytokines that attract lymphocytes and polymorphonuclear cells that when stimulated, produce chemokines recruiting monocytes to the tissue (Burger and Dayer, 2002). Due to the high impact of cytokines, their functions have to be tightly modulated. They may be regulated by inhibitory molecules which can be arbitrarily classified into two
Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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categories – cytokine inhibitors which interfere with the binding of the cytokine to its specific receptor, or cytokines which are active in affecting the production of other cytokines or their inhibitors. Inhibitors of cytokines are often generated by the shedding of specific cytokine receptors (Chizzolini and Dayer, 1998). This is the case for TNF-α, whose activity is inhibited by TNF-soluble receptors, and to IL-1, whose activity is inhibited by IL-1sRII (Burger et al., 1995). In addition to soluble receptors, IL-1 receptor antagonist (IL-1Ra) counterbalances the pro-inflammatory effects of IL-1 (Burger and Dayer, 2000), and IL1RII also functions as a decoy receptor at the cell surface (Mantovani et al., 2001). Cytokines with inhibitory activity such as IL-4, IL-10 and IL-13 are principally produced by Th2 cells (Rognani, 2000), while transforming growth factor-β (TGF-β) is mainly synthesized by Th3; they inhibit type 1 cytokine activity and stimulate IL-1Ra production in macrophages or Kupffer cells (KC). IL-1Ra production in macrophages is also induced by interferon-β (IFN-β) [Jungo et al., 2001]. Another example is IFN-γ , a type 1 cytokine that counteracts TGF-β stimulation of collagen synthesis in fibroblast (Korn, 1997). 14.1.2
The Liver
The liver has an important role in metabolic homeostasis, because it is responsible for the storage, synthesis, metabolism, and redistribution of carbohydrates, fats and vitamins. The liver produces large numbers of serum proteins, mainly albumin, enzymes, cofactors and acute-phase proteins. This organ is the main detoxifying of the body; it removes xenobiotics and wastes by metabolism or biliary excretion. The main cell type of the hepatic tissue that carries out most of these functions is the parenchymal cell, the hepatocyte, which constitutes around 80 % of liver cells. The remaining 20 % comprises the non-parenchymal cells, which include endothelial cells, KCs, lymphocytes and hepatic stellate cells (HSC) (Desmet, 2001). Endothelial cells line the intrahepatic circulatory vessels or sinusoids of the liver, which provide a large surface area for oxygen and nutrient. KC are found in the sinusoids and are in charge of phagocytosis of particles and infecting organisms; importantly, these cells are responsible for the synthesis of cytokines. Lymphocytes are also part of the immune system that resides within the hepatic tissue to help to avoid infection. HSC possess several functions, like the storage of vitamin A and, when activated, the production of the extracellular matrix. All of these cells are activated by hepatic damage and the HSC produce most of the factors that lead to liver fibrosis and cirrhosis (Desmet, 2001).
14.2 14.2.1
Cytokines and Liver Regeneration Liver Regeneration
In the normal adult liver cell division is rarely seen in hepatocytes, as these cells are in the G0 phase of the cell cycle (Michalopoulos and DeFrances, 1997). After removal of an important mass of the organ (50–70 %), around 95 % of hepatocytes quickly enter the S phase of the cell cycle. The rate of DNA synthesis in hepatocytes begins to increase after about 12 h and peaks around one day in the rat liver. The induction of DNA synthesis occurs latter in the non-parenchymal cells, at around 2 days for Kupffer and biliary epithelial cells, and around 4 days for endothelial cells (Taub, 2004). Most of the increase in liver mass
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occurred by 3 days after partial hepatectomy and mass restoration is complete in 5–7 days (Fausto, 2000; Michalopoulos and DeFrances, 2005). 14.2.2
Cytokines in Liver Regeneration
The hypothesis that cytokines might regulate the regenerative response comes from the observation that NF-κB and STAT3 transcription factors are activated by cytokines immediately after partial hepatectomy (Taub et al., 1999). IL-6−/− mouse studies have shown that after surgical partial hepatectomy, liver regeneration requires IL-6 cytokine (Cressman et al., 1996; Sakamoto et al., 1999). However, IL-6 is necessary but not enough stimuli for the induction of regeneration, since restitution of liver mass is only delayed in the absence of IL-6 (Cressman et al., 1996). IL-6 participates in various processes that are involved in liver pathophysiology, including hepatoprotection and mitogenesis (Yang et al., 2006). Some studies have questioned the proposed major role of IL-6 in the mitogenic response after partial hepatectomy (Wuestefeld et al., 2003; Blindenbacher et al., 2003b). IL-6 activates the Janus kinase JAK by binding to its receptor (Levy and Lee, 2002; Heinrich et al., 2003). Once activated, JAK phosphorylates the associated glycoprotein 130 (gp130) and STAT3 on a Tyr residue, which results in the dimerization of STAT3. After STAT3 is dimerized, it goes to the nucleus and activates the target genes. In addition, the stimulation of gp130 activates the MAPK (mitogen-activated protein kinase) cascade and JAK activates the SH2-domain-containing tyrosine phosphatase SHP2 and recruits growth-factor-receptorbound protein-2/son-of-sevenless (GRB2-SOS). Then, GRB2-SOS activates Ras, which leads to activation of Raf-MEK-ERK (where MEK stands for MAPK and ERK kinase). It is important to point out that MAPK is crucial for cellular proliferation (Talarmin et al., 1999), and evidence indicates that IL-6 can activate kinases that are involved in cell survival, including phosphatidylinositol 3-kinase (P13K) (Levy and Lee, 2002). Several studies have found the regulatory network that controls IL-6 signaling in hepatocytes. It was shown that TNF-α is also required for a normal proliferative response after partial hepatectomy by using TNF-receptor-1 (TNF-R1) knockout mice (Yamada et al., 1997). This effect seems to be performed by the ability of TNF-α to induce IL-6, because IL-6 treatment improves the defect in DNA synthesis that happens in TNF-R1 knockout mice subjected to partial hepatectomy. However, the absence of TNF-α does not impair liver regeneration (Fujita et al., 2001). KC produce most of the IL-6 in the liver (Aldeguer et al., 2002; Yang et al., 2006) and TNF-α induces the production of IL-6 by upregulating NF-κB, which, in turn, activates transcription of IL-6 (Taub, 2004). Recently it was reported that IL-6 functions acutely to stimulate hepatic regeneration, while chronic exposure not only abolishes the regenerative activity of IL-6, but actually sensitizes the liver to injury and death (Jin et al., 2006). 14.2.3
Pathways Mediated by Growth Factors: HGF and TGF-α
Some investigators suggest that IL-6 is not a powerful mitogen in the hepatocyte, as the stimulation of liver proliferation is minimal in the absence of partial hepatectomy, unless IL-6 is highly over expressed (Taub, 2004). Various cytokines function to stimulate cellular replication during hepatic regeneration (Yang et al., 2006). Based on in vivo and in vitro studies, tumor growth factor alpha (TGF-α) and hepatocyte growth factor (HGF) are considered to be two of the most important growth factors (Michalopoulos and DeFrances,
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1997; Taub et al., 1999; Fausto, 2000; Pediaditakis et al., 2001; Koniaris et al., 2003; Taub, 2004; Huh et al., 2004; Gherardi et al., 2006). Non-parenchymal cells, particularly stellate cells synthesize HGF, and therefore affect hepatocytes in a paracrine manner (Gherardi et al., 2006). Pro-HGF, the HGF precursor, is quickly activated by proteases, such as the urokinase-type plasminogen activator (uPA) and its downstream effector plasminogen, after liver injury (chemical hepatectomy) or surgical partial hepatectomy (Pediaditakis et al., 2001; Shimizu et al., 2001; Currier et al., 2003). Therefore, blocking uPA delays the appearance of HGF, and thus delays hepatic proliferation. On the contrary, blocking plasminogen-activator inhibitor (PAI) accelerates the release of HGF and therefore accelerates liver regeneration. Administration of antibodies to HGF at the same time as CCl4 intoxication (which produced liver necrosis and thus a chemical hepatectomy), blocks the regenerative response (Burr et al., 1998), and the infusion of HGF into rodents reduces the level of CCl4 -induced liver damage (Matsuda et al., 1997; Ueki et al., 1999). In the liver, HGF regulates various processes, and is a direct stimulant of liver regeneration (Taub, 2004; Gherardi et al., 2006). HGF and its receptor, Met, are important growth factors in several organs, and knockouts results in a lethal phenotype (Gherardi et al., 2006). Therefore, it was difficult to use genetic approaches to study the routes that are regulated by HGF in vivo (Burr et al., 1998; Pediaditakis et al., 2001; Huh et al., 2004). Huh et al. (2004), utilizing a liver-specific conditional knockout of Met, showed that the HGF-Met pathway is important for DNA synthesis after liver injury. Moreover, evidence from studies that use other gene-knockout mice, such as uPA, plasminogen and vascular endothelial growth factor (VEGF) gave supportive data (Taub, 2004). In addition, pharmacological approaches, like antibodies and adenoviruses, indicated that HGF and TGF-α, participate in the proliferative response of the hepatic cells (Burr et al., 1998; Pediaditakis et al., 2001; Huh et al., 2004; Yang et al., 2006). DNA synthesis is highly stimulated by HGF in cultured hepatocytes and also modulates cell motility and morphology. HGF has several effects on various mitogenic signaling pathways. HGF activates the receptor tyrosine kinase Met and several downstream pathways that include those that involve P13, ERK, and AKT (Okano et al., 2003; Gherardi et al., 2006). Studies in isolated hepatocytes indicate that the mitogenic effect of HGF is mediated, at least in part, through stimulation of another growth factor, TGF-α, because antibodies against TGF-α partially block the DNA-synthesis in the regenerating liver (Tomiya et al., 2000; Sheving et al., 2002; Okano et al., 2003). A large number of ligands and receptors belong to the TGF and EGF family; thus, the inability to block all of these ligands and receptors simultaneously made it difficult to demonstrate that the potent mitogens TGF-α and EGF are crucial for hepatic proliferation. Similarly, VEGF interacts specifically with endothelial cells within the liver and increases HGF production by non-parenchymal cells. This VEGF-mediated liver growth depends on the presence of endothelial cells, but the mechanism is not known, and blocked partially by treatment with antibodies to HGF (LeCouter et al., 2003). Liver regeneration, in humans, takes place most frequently after hepatic injury by ischemia or hepatitis, an inflammation of the liver that is induced by injury produced by xenobiotics, immune-mediated injury or viral infections. Then, understanding the fundamental aspects of the biology of liver regeneration will help to explain how the liver responds to toxic injury by alcohol, drug overdose or infections as viral hepatitis, which in turn, will provide the basic knowledge for rational treatments.
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Cytokines and Viral Hepatitis Interferon-α
Viral hepatitis has been treated with IFN-α as the primary medical choice for more than 15 years (Hofnagle, 1999; Liang et al., 2000; Gao, 2005; Wong and Lok, 2006). Thus, IFN-α induced activation of signal transduction and downstream antiviral genes in the liver has been extensively studied (Castet et al., 2002; Zhu et al., 2003; Ji et al., 2003; Radaeva et al., 2004). The effects of IFN-α are mediated via IFNAR1 and IFNAR2 complex. The IFNAR1 chain depicts only one form and is related to signal transduction, while IFNAR2 shows three forms, including hIFNAR2a (Pestka, 2000). The full length of hIFNAR2c is involved in signal transduction and ligand binding, while both the short-form hIFNAR2b and the soluble-form hIFNAR2a are associated to suppression of IFN-α signaling (Pfeffer et al., 1997; Pestka, 2000). After IFN-α binding, IFN-α receptor associated tyrosine kinases (JAK1 and Tyk2) become activated; then Y466 of the IFNAR1 is phosphorilated. Then, IFNAR1 phosphorilates STAT2 on Y690 and STAT1 on Y701. STAT1 and STAT2, once phosphorilated, constitutes heterodimers that translocate into the nucleus to induce the transcription of several genes, including many antiviral proteins, tumor suppressors and proapoptotic proteins (Darnell, 1998; Stark et al., 1998; Pestka, 2000). The action mechanism of IFN-α in the treatment of viral hepatitis is believed to be due by targeting hepatocytes and modulating the immune system via induction of the Jak-Stat signaling pathway (Gao et al., 2004). In vitro human hepatocytes express high levels of IFN-α receptor IFNAR1 and IFNAR2c, and respond well to IFN-α stimulation (Radaeva et al., 2002). IFN-α activates STAT1-3,5 in human hepatocytes and hepatoma cells. IFN-α upregulates expression of around 50 genes and downregulates expression of about 10 genes by 50 % as determined by micro array analyses (Gao, 2005). These include induction of four antiviral genes and a wide variety of proapoptotic/tumor suppressor genes (Radaeva et al., 2002), that may participate in the liver antiviral and antitumor activity of IFN-α (Wong and Lok, 2006). More than 60 % patients with viral hepatitis respond poorly to IFN-α therapy (Hofnagle, 1999; Liang et al., 2000; Wong and Lok, 2006). Inhibition of IFN-α antiviral activity by several viral factors may contribute to failure of IFN-α treatment in patients with viral hepatitis (Gao et al., 2004). These viral proteins include hepatitis C virus (HCV) E2 protein (Taylor et al, 1999), HCV core protein (Blindenbacher et al., 2003a; Basu et al., 2001) and HCV nonstructural 5A (NS5A) (Podevin et al., 2001). Various factors in the patient such as alcohol drinking (Nguyen et al., 2000), elevation of TNF-α (Hong et al., 2001), IL-1β (Tian et al., 2000) and IL-10 (Shen et al., 2000) have been observed to decrease IFN-α-activated STAT1 in the liver, which may contribute to failure of IFN-α treatment in viral hepatitis patients with alcohol drinking or high serum levels of TNF-α, IL-1β and IL-10 (Wong and Lok, 2006). These host factors could be potentially improved for IFN-α therapy. 14.3.2
Interferon-γ
IFN-γ is a cytokine that plays an important role in host defense due to its antiviral, proapoptotic, antiproliferative, pro-inflammatory and immunoregulatory activities (Gao, 2005). The IFN-γ action mechanism is mediated through activation of the JAK-STAT signaling (Pestka, 2000; Stark et al., 1998). Binding of IFN-γ to IFNGR1 produces the dimerization of IFNGR1 and IFNGR2; then, activation of the receptor-associated kinases activation,
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JAK1 and JAK2, takes place. Then, the complex of the receptor–kinase activates STAT that produces the expression of several genes including antiviral, proapoptotic and antiproliferative between others. STAT1−/− mice possess defective IFN-γ signaling and absent innate response to bacterial or viral infection (Meraz et al., 1996). Activation of STAT1 by IFN-γ has been associated with hepatic inflammation and injury, and suppression of liver regeneration (Brooling et al., 2005). Various models of liver injury activate STAT1, including Con A-induced T cell hepatitis and LPS/D-galactosamineinduced hepatic injury, and IFN-γ is the principal responsible for this stimulation (Kim et al., 2003). It is worth noting that high concentrations of the of mRNA (Abbate et al., 2003) and STAT1 protein (Radaeva et al., 2004; Duong et al., 2004) were observed in the livers of patients with chronic hepatitis C, suggesting the that IFN-γ /STAT1 play a causative role of chronic viral hepatitis. IFN-α, IFN-β and IFN-λ are the only reported activators of STAT2 (Pestka, 2000; Kotenko et al., 2003). STAT2 knockout mice show an increased sensibility to viral infection and the loss of a type I IFN autocrine/paracrine loop (Park et al., 2000). IFN-α treatment induces significant STAT2 activation in primary hepatocytes from humans (Radaeva et al., 2002), indicating that STAT2 activation is a key signal involved in the antiviral therapy of IFN-α in humans with viral hepatitis. There is evidence that STAT2 protein is diminished in the liver of humans with chronic alcoholism, which could be an important mechanism contributing to the IFN-α treatment failure (Nguyen and Gao, 2002).
14.4
Cytokine Participation in Alcoholic Liver Disease
The leading cause of death from hepatic disease at least in western countries is alcoholic liver disease (ALD), and abnormal cytokine metabolism is a major feature of the disease (Dey and Cederbaum, 2006). Several researchers reported increased serum TNF-α concentrations in alcoholic hepatitis (AH) (McClain et al., 2004), and TNF-α concentrations correlated well with injury severity and mortality (McClain et al., 1999). Polymorphism of TNF-α promoter is associated with a higher incidence to alcoholic steatohepatitis. This suggests that a subset of people who drink alcohol may be genetically predisposed to the development of ALD (Grove et al., 1997). Moreover, patients with AH or cirrhosis show elevated serum concentrations of TNF-α-inducible cytokines such as IL-6, IL-8 and IL-18 and levels correlated with markers of liver function and clinical outcome (McClain et al., 1999). Studies in animals and cells evaluated the role of cytokines (especially TNF-α) in experimental models of liver disease (McClain et al., 2004). Rats chronically fed alcohol had much higher LPS-stimulated plasma levels of TNF-α than controls, and hepatic damage can be attenuated by giving a prostaglandin analog that downregulates TNF-α production (Honchel et al., 1992). By utilizing the intragastric alcohol-feeding model, some researchers demonstrated that the development of liver injury was associated with an increase in TNF-α mRNA in the liver (Nanji et al., 1994; Kamimura and Tsukamoto, 1995). In addition to the increased levels of proinflammatory and cytotoxic cytokines in ALD, anti-inflammatory cytokines, like IL-10 production by KC and monocytes are diminished. The importance of this imbalance of cytokines was studied in IL-10 deficient mice; these mice were more susceptible to ethanol hepatotoxicity and exhibited increased levels of proinflammatory cytokines such as TNF-α. IL-10-plays an antifibrotic role in addition to
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its capacity to modulate TNF-α; therefore, a decrement in this cytokine probably plays a pathophysiological role in the establishment and progression of ALD (McClain et al., 1999, 2004). Cytokine production blockade has been utilized to counteract alcohol-induced hepatic injury. For example, antibiotics have been used to inhibit gut flora and LPS production, gadolinium chloride to destroy KC, and the glutathione precursor procysteine, to inhibit cytokine production. These strategies succeed in ameliorating alcohol hepatic damage in rats (McClain et al., 1999, 2004). Furthermore, the administration of anti-TNF-α to rats, effectively prevented alcohol induced liver injury (Iimuro et al., 1997) and TNF-α type I receptor knockout mice do not develop alcoholic hepatic damage (Yin et al., 1999). LPS and oxidative stress are thought to be to major stimuli for cytokine production in ALD (Dey and Cederbaum, 2006). In ALD, there is increased intestinal permeability and a high frequency of endotoxemia (McClain et al., 1999). The NF-κB factor is activated by LPS and sensible to oxidative stress; its activation results in the production of certain cytokines such as TNF-α (Karin et al., 2004). Apoptosis is an important mechanism of cell death in ALD and it has been demonstrated that apoptotic hepatocytes are taken up by KC and stimulate TNF-α production (Canbay et al., 2003). In addition, when hepatocytes die of proteasome inhibition-mediated apoptosis, the dying hepatocytes release IL-8 and IL-18 to produce important inflammation (Canbay et al., 2003). Several investigations show the role of impaired proteasome function in the hepatic damage and development of alcoholic hyaline in ALD. Cytokine liver alterations in ALD are related with functional changes in monocytes and KC. Neopterin, the most widely used marker of monocyte activation, is consistently elevated in ALD. Mechanisms involved in this activation are increased LPS binding protein, high levels of CD14, enhanced signaling through Toll-like receptors, direct effects of alcohol or acetaldehyde and oxidative stress. KC activation takes place in chronic alcohol intake (McClain et al., 2004). Acute alcohol ingestion has an inhibitory effect on cytokine induction. Rats given an acute dose of alcohol show decreased concentrations of TNF-α after LPS stimulation (Hill et al., 1997). Accordingly, short-term incubation of monocytes with alcohol leads to suppressed LPS-stimulated proinflammatory cytokine induction (Hill et al., 1997). On the contrary, when experimental rodents are fed alcohol during a prolonged time, for at least 4–6 weeks, they show augmented production of LPS-induced TNF-α (Honchel et al., 1992). Therefore, chronic alcohol exposure causes monocytes/KC to be activated to produce some cytokines in excess in response to stimuli such as LPS.
14.5 14.5.1
Cytokines and Non-Alcoholic Steatohepatitis Steatohepatitis
The major risk factor for non-alcoholic steatohepatitis (NASH) in humans is obesity, especially visceral adiposity (Clark et al., 2003; Farrell and Larter, 2006). Patients with NASH are frequently insulin-resistant even if they do not have an elevated body mass index (Marchesini et al., 2005). It is well known that adipose tissue is a source of free fatty acids that are sent to the liver, and then, triglycerides that are synthesized by hepatic cells are released into the blood.
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Fat produces hormones, such as adiponectin, resistin and leptin, that regulate fat itself and metabolism in other organs. Hypoxia and reactive oxygen species regulate adiponectin production in adipocytes (Chen et al., 2006). In addition, fat is a source of neurotransmitters like angiotensin II and noradrenalin (Diehl et al., 2005). Furthermore, fat is a component of the immune system since adipocytes produce immunomodulatory cytokines, such as TNF-α and IL-6 (Rajala and Scherer, 2003; Lyon et al., 2003; Diehl et al., 2005). Due to the immunomodulatory and neuroendocrine functions of fat, it is expected that increases in adipose mass influence immune function, energy homeostasis and metabolism. The situation is more complicated because there are both quantitative and qualitative differences in the factors (adipokines) that may be produced by different fat depots. Visceral adiposity, for instance, that appears to be less immature as compared with subcutaneous fat, induces less adiponectin but more TNF-α and free fatty acids than subcutaneous fat which produces large amounts of leptin (Lyon et al., 2003). Therefore, the final consequences of obesity may be different depending on which fat predominates. 14.5.2
Fat Depots and TNF-α
Fatty liver disease is promoted by visceral adiposity by various mechanisms. First, intraabdominal fat is a source of free fatty acids that are delivered into the portal vein and thus to the liver (Kelley et al., 2003; Farrell and Larter, 2006). The delivery of free fatty acids to organs, including the liver, induces localized insulin resistance in mice (Kim et al., 2001a). Furthermore, studies of gene disruption have demonstrated that interference with insulin signaling in hepatocytes activates fat-synthesizing enzymes in these cells and results in steatosis of the liver (Tobe et al., 2001). Secondly, patients with truncal obesity show very low levels of adiponectin and elevated levels of TNF-α (Misra and Garg, 2003). It is known that TNF-α activates intracellular signaling molecules, including the inhibitor of the kappa beta kinase and stress related kinases such as N-terminal kinase that make cells resistant to the actions of insulin (Kim et al., 2001a,b; Diehl et al., 2005). The production and activity of TNF-α are antagonized by adiponectin (Ruan and Lodish, 2003). Thus, TNF-α activity is increased when adiponectin is decreased. Moreover, TNF-α itself diminishes adiponectin (Bruun et al., 2003). Adiponectin inhibits fatty acid synthesis and uptake in hepatocytes and stimulates fatty acid oxidation (Yamauchi et al., 2002). In other words, adiponectin itself also augments hepatocyte sensitivity to insulin. The combination of high TNF-α and low adiponectin levels coexist in the context of increased hepatic exposure to free fatty acids and results in hepatic steatosis and severe liver insulin resistance (Diez and Iglesias, 2003). 14.5.3
Cytokines and Steatohepatitis
Similarities in the histopathology of NASH and alcohol-induced steatohepatitis suggest that similar mechanisms may play a role in both diseases. In experimental non-obese animals in which steatohepatitis was induced with chronic alcohol administration, inhibition of TNF-α prevented liver injury (Tsukamoto and Lu, 2001). Chronic ethanol consumption produces insulin resistance in rodents (Mohr et al., 1998). Since the cellular mechanisms for insulin resistance involves TNF-α production (Kim et al., 2001a,b), drugs that inhibit TNF-α (Yu et al., 2002) protect mice from alcohol-induced steatohepatitis (Enomoto et al., 2005). In a mouse model of NASH (leptin deficient ob/ob mice), Li et al. (2003) demonstrated that
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anti-TNF-α antibodies improved NASH and hepatic insulin sensitivity. Both non-obese ethanol fed mice and obese ob/ob mice show increased TNF-α and decreased adiponectin levels; administration of recombinant adiponectin prevented TNF-α, improved insulin sensitivity and reversed steatohepatitis in control ethanol-fed mice and in obese mice (Xu et al., 2003). Patients with an elevated body mass index very frequently show high serum levels of leptin (Lebovitz, 2003). Thus, it was argued that studies in leptin-deficient ob/ob mice may provide little knowledge to the mechanism of action of NASH and insulin resistance in humans (Green, 2003). However ob/ob mice and obese humans have in common low levels of adiponectin and relatively high levels of TNF-α (Lebovitz, 2003; Xu et al., 2003). Various treatments that improve the dysmetabolic syndrome (diet, exercise or thiazolidinediones for instance) increased adiponectin levels and reduce TNF-α in patients (Enomoto et al., 2003, 2005; Esposito et al., 2003). Therefore, it will be of interest to test if exogenous supplementation of adiponectin ameliorates resistance to insulin and NASH in humans.
14.6 14.6.1
Cytokines and Fibrosis Fibrosis
The process of hepatic fibrosis shares several characteristics with a wound-healing process: formation of a provisional clot influencing the invasion, inflammation and proliferation of inflammatory and connective tissue-producing cells, mainly HSC. These events may result in a complete repair or scar formation (Ramadory and Saile, 2004a,b). However, when the damage is chronic, matrix deposition occurs in excess of resorption; as a result of an imbalance between fibrogenesis and fibrolysis, a scarring process ensues. The scarring progress from bridging fibrosis to the formation of complete nodules results in an architectural distortion and blood-flow obstruction and finally in liver cirrhosis. Hepatic cirrhosis may be the result of a variety of repeated liver injuries, like chronic ethanol abuse, viral hepatitis, biliary obstruction, copper or iron accumulation. From the toxic animal models of liver fibrosis, we learned that fibrosis may result from recurrent liver injuries. Small but chronic liver injury usually associated with oxidative stress and lipid peroxidation processes (Muriel, 1997) induced in rodents leads to fibrosis and cirrhosis. However, different types of disease lead to different types of fibrosis as the injury progresses. The kinds of fibrosis can be divided into central and portal fibrosis. The major portal-based injuries that lead to cirrhosis are hemochromatosis, chronic cholestasis and chronic viral hepatitis. Steatohepatitis and chronic venous outflow obstruction include central-based diseases. Fibrosis can also be divided into porto–portal (e.g. following cholestatic liver injury), porto–central (e.g. in the case of viral hepatitis) or centro–portal (e.g. alcoholic liver disease) (Ramadori and Saile, 2004b). The etiology of the disease, with the exceptions of hemochromatosis, and in most of the cases of alcoholic liver diseases cannot be deduced from the inflammatory infiltrate (Ramadory and Saile, 2004a). 14.6.2
Cytokines and Matrix Synthesis
KC represent the resident macrophages in the liver and one of its functions is the production of cytokines. Interestingly, the inhibition of KC with gadolinium chloride prevents acute
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liver damage induced with CCl4 in the rat (Muriel et al., 2001). Furthermore, inactivation of KC prevents the fibrosis and cirrhosis experimentally induced by chronic CCl4 intoxication in rats (Muriel and Escobar, 2003). The most likely explanation for these results is that harmful cytokines produced by KC in response to CCl4 are involved in the signaling cascade that leads to the exacerbated production of scar tissue. Moreover, experimental cirrhosis induced by bile-duct obstruction induces changes in plasma and hepatic levels of cytokines (Fern´andez-Mart´ınez et al., 2006). HSC may be activated as a result of liver injury. The principal characteristic of activated HSC, and other extracellular matrix (ECM)-producing cells like fibroblast and myofibroblast, is the increased expression of smooth-muscle-α actin and an augmented production of ECM proteins (Issa et al., 2001). It is well known that fibroblast and myofibroblast actively participate in fibrogenesis. However, less is known about the factors that induce their proliferation and ECM synthesis when compared with activated HSC. It is known that myofibroblast proliferate in vitro, and that insulin growth factor-I (IGF-I) has an antiapoptotic effect (Issa et al., 2001), while TNF-α has a proapoptotic role (Ramadori and Saile, 2004a). On the contrary, activated HSC undergo apoptosis in vitro that may be inhibited by TNF-α and TGF-β and induced by IGF-I treatment (Ramadori and Saile, 2004a; Saile et al., 2004). HSC activation in vitro is regulated by various compounds, including cytokines, growth factors and products of oxidative stress, including nitric oxide (NO) (Muriel, 2006; Moreno and Muriel, 2006). The cytokine environment seems to be of crucial importance. For example, IFN-γ , one of the first molecules that is produced in the liver after intoxication, prevents ECM synthesis (Knittel et al., 1997; Batusic et al., 2004; Brooling et al., 2005) and leads to apoptosis of activated HSC (Saile et al., 2000), whereas the shortly later increased occurring TNF-α induces HSC activation, production of ECM synthesis (Knittel et al., 1997), and inhibits apoptosis in HSC (Ramadori and Saile, 2004b; Brooling et al., 2005). During liver cirrhosis, several hepatic and plasmatic cytokines are deregulated (Fern´andez-Mart´ınez et al., 2006). TGF-β appears to be a key cytokine/growth factor mediator in human fibrogenesis (Bataller and Brenner, 2005). TGF-β promotes the activation of HSC, stimulates the synthesis of ECM proteins and inhibits its degradation. One way to interfere with fibrosis is, therefore, by disrupting the TGF-β signaling pathway (Shek and Benyon, 2004). The most potent mitogen for HSC is platelet-derived growth factor (PDGF) and is upregulated in the fibrotic liver (Pinzani et al., 1989). As expected, its inhibition attenuates experimental liver fibrogenesis (Borkham-Kamphorst et al., 2004). Several investigations have shown relationships between increased levels of IL-10 and decreased stage of fibrosis (Nelson et al., 2000; Ramadori and Saile, 2004b). Recently, a mechanism for IL-10-dependent fibrosis inhibition in vivo was suggested (Safadi et al., 2004). The antifibrotic effect of IL-10 was confirmed, and then the relative increase of intrahepatic T lymphocytes during fibrogenesis was eliminated in IL-10 knockout mice as compared with wild type leading to the conclusion that T lymphocytes may be important for activation of cells that produce ECM. A possible role of T lymphocytes by reducing their migration into the hepatic tissue through total body irradiation was suggested. However, other investigations suggest a direct beneficial effect of irradiation on hepatocytes regarding cell death (Kojima et al., 2000) associated with an antifibrotic effect. As most of the factors that are known to affect HSC activation are produced by natural killer (NK) cells and KC, research was aimed to find factors inhibiting their cytokine
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production (Muriel et al., 2001, 2003; Muriel and Escobar, 2003; Fern´andez-Mart´ınez et al., 2004). Thalidomide, for example, has been reported to modulate various cytokines in the liver (Fern´andez-Mart´ınez et al., 2004) and to ameliorate CCl4 -induced cirrhosis in the rat (Muriel et al., 2003).
14.7
Conclusions
Many cytokines and other related molecules act in concert to contribute to liver physiology and pathophysiology of diverse etiologies. It is clear that liver regeneration requires several coordinated functions and signals to proceed normally. Pathways are very redundant and thus the elimination of a single gene delays but not impairs the recuperation of liver mass. It is not completely elucidated how the size of the liver is determined, why hepatic mitogen cytokines or factors are hepatoprotective in models of liver damage and how the complex structure of the hepatic parenchyma is re-established after cellular proliferation. Understanding the molecular basis for human hepatic regeneration will not only help to define the pathology but will ultimately provide new treatment options for patients with liver damage. IFN-α is used to treat viral hepatitis. STAT1, principally activated by IFN-α and IFN-γ , not only plays a fundamental role in the antiviral defense in hepatitis virus infection but also participates in liver injury and suppresses hepatocyte proliferation. Activated by IFN-α and IFN-γ , STAT2 also contributes importantly to the antiviral properties of interferons. Modulation of these STATs may offer a new approach in the treatment of viral hepatitis. Abnormal cytokine metabolism is a major feature of ALD. In patients with ALD, limited pilot studies using a TNF-α-soluble receptor and an antibody suggest that these compounds are generably safe and tolerable (Menon et al., 2004). A multicenter study, however, using both high-dose infliximab (anti-IFN-α antibody) and prednisone had to be discontinued due to increased infections (Naveau et al., 2004). The strategy employing inhibitors of cytokine synthesis or secretion, such as pentoxifyline, has generated promising results. The complementary and alternative medicine S-adenosylmethionine (SAM) (Muriel et al., 1994) have efficacy in ALD via inhibition of TNF-α and stimulation of IL-10. SAM was shown to improve survival and decreased need for liver transplantation in patients with alcoholic cirrhosis (Mato et al., 1999). Modulation of cytokine therapy for ALD is in its infancy, but, on the basis of its success in experimental models of liver damage (Muriel et al., 2003; Muriel and Escobar, 2003), rapid progress in this area is predicted. NASH is characterized by necrosis and inflammation, factors that seem to be modulated by cytokines, hormones and neurotransmitters that regulate the biological activity of TNF-α and other proinflammatory cytokines. Liver necroinflammation is necessary but not enough, for progression to cirrhosis. Factors like leptin that regulate the activity of profibrogenic cytokines, such as IL-6 and TGF-β, determine the extent of fibrosis that occurs during hepatic damage. Liver fibrosis is a wound-healing process in which the accumulation of ECM proteins is the predominant feature. Monocytes and KC participates by producing inflammatory cytokines after chronic liver injury that stimulates HSC and other ECM-producing cells to be activated. Activation is characterized by proliferation, motility, contractility and synthesis of ECM. TGF-β and IL-6 are the two main fibrogenic cytokines. Potential regulatory drugs
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of the production of cytokines and/or the activation of HSC are important targets for future antifibrogenic treatments. A better understanding of how cytokines and other factors regulate the function of the different types of liver cells should help us to develop rational treatments for hepatic human diseases.
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Pediaditakis P, Lopez-Talabera JC, Petersen B, Monga SP and Michalopoulos GK (2001). The processing and utilization of hepatocyte growth factor/scatter factor following partial hepatectomy in the rat. Hepatology 34: 688–693. Pestka S (2000). The human interferon α species and receptors. Biopolimers 55: 254–287. Pfeffer LM, Basu L, Pfeffer SR, Yang CH, Murti A, Garcia JV, Croze E, Constantinescu SN, Mullersman JE and Pfeffer LM (1997). The short form of the interferon α/β receptor chain 2 acts as a dominant negative for type I interferon action. J Biol Chem 272: 11002–11005. Pinzani M, Gesualdo L, Sabbah GM and Abboud HE (1989). Effects of platelet-derived growth factor and other polypeptide mitogens on DNA synthesis and growth of cultured rat liver fat storing cells. J Clin Invest 84: 1786–1793. Podevin P, Sabile A, Gajardo R, Delhem N, Abadie A, Lozach PY, Beretta L and Brechot C (2001). Expression of hepatitis C virus NS5A natural mutants in a hepatocytic cell line inhibits the antiviral effect of interferon in a PKR-independent manner. Hepatology 33: 1503–1511. Radaeva S, Jaruga B, Hong F, Kim WH, Fan S, Cai H, Strom S, Liu Y, El-Assal O and Gao B (2002). Interferon-α activates multiple STAT signals and down regulates c-Met in primary human hepatocytes. Gastroenterology 122: 1020–1034. Radaeva S, Jaruga B, Kim WH, Heller T, Liang TJ and Gao B (2004). Interferon-γ inhibits interferonα signaling in hepatic cells: evidence for the involvement of STAT1 induction and hyperexpression of STAT1 in chronic hepatitis C. Biochem J 379: 199–208. Rajala MW and Sherer PE (2003). Minireview: The adipocytes – at the crossroads of energy homeostasis, inflammation and atherosclerosis. Endocrinology 144: 3765–3773. Ramadory G and Saile B (2004a). Inflammation, damage repair, immune cells and liver fibrosis: Specific or nonspecific, this is the question. Gastroenterology 127: 997–1000. Ramadory G and Saile B (2004b). Portal tract fibrogenesis in the liver. Lab Invest 84: 153–159. Rognani S. 2000. T-cell subsets (Th versus Th2). Ann Allergy Asthma Immunol 85: 9–18. Ruan H and Lodish HF (2003). Insulin resistance in adipose tissue: direct and indirect effects of tumor necrosis factor-alpha. Cytokine Growth Factor Rev 14: 447–455. Safadi R, Ohta M, Alvarez CE, Fiel MI, Bansal M, Mehal WZ and Friedman SL (2004). Immune stimulation of hepatic fibrogenesis by CD8 cells and attenuation by transgenic interleukin-10 from hepatocytes. Gastroenterology 127: 870–882. Saile B, Eisenbach C, Neubauer K, Stoeckigt C and Ramadori G (2000). Antiapoptotic effect of interferon-A versus proapoptotic effect of interferon-gamma on hepatic stellate cells (RSC) – a novel pathway of IFN-A signal transduction via janus kinase 2 (JAK2) and caspase 8. Hepatology 32: 191A. Saile B, DiRocco P, Dudas J, El-Armouche H, Sebb H, Eisenbach C, Newbauer K and Ramadori G (2004). IGF-I induces DNA synthesis and apoptosis in rat liver hepatic stellate cells (HSC) but DNA synthesis and proliferation in rat liver myofibroblasts (rMF). Lab Invest 84: 1037–1049. Sakamoto T, Liu Z, Murase N, Ezure T, Yokomuro S, Poli V and Demetris AJ (1999). Mitosis and apoptosis in the liver of interleukin-6-deficient mice after partial hepatectomy. Hepatology 29: 403–411. Shek FW and Benyon RC (2004). How can transforming growth factor beta be targeted usefully to combat liver fibrosis? Eur J Gastroenterol Hepatol 16: 123–126. Shen X, Hong F, Nguyen VA and Gao B (2000). IL-10 attenuates IFN-α-activated STAT1 in the liver: involvement of SOCS2 and SOCS3. FEBS Lett 480: 132–136. Sheving LA, Stevenson MC, Taylormoore JM, Traxter P and Russell WE (2002). Integral role of the EGF receptor in HGF-mediated hepatocyte proliferation. Biochem Biophys Res Commun 290: 197–203. Shimizu M, Hara A, Okuno M, Matsuno H, Okada K, Ueshima S, Matsuo O, Niwa M, Akita K, Yamada Y, Yoshimi N, Uematsu T, Kojima S, Friedman SL, Moriwaki H and Mori H (2001).
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15 Bile Acids as Modulators of Apoptosis Rui E. Castro, Susana Sol´a, Clifford J. Steer and Cec´ılia M.P. Rodrigues
15.1
Introduction
Bile acids are derivatives of cholesterol that play an important role in digestion and absorption of fat. They are also signaling molecules that activate nuclear receptors and regulate many physiological pathways, including cholesterol homeostasis. When bile acids accumulate to high levels, a negative feedback mechanism is activated and synthesis is repressed. In contrast, bile acid synthesis increases when cholesterol accumulates. Furthermore, the expression of genes that synthesize cholesterol, fatty acids and bile acids are regulated by intermediates and/or end-products of the bile acid pathway itself. Bile acids are also important in providing the driving force for the formation and secretion of bile, thereby maintaining an efficient enterohepatic circulation. The different biological activities of bile acids have been related to chemical structure. In fact, hydrophobic bile acids are generally cytotoxic compounds, capable of inducing apoptosis, whereas hydrophilic bile acids are less cytotoxic and, in some cases, inhibitors of apoptosis. However, recent studies suggest that the regulation of apoptosis by bile acids is not entirely related to their inherent structure. Rather, it is the differential ability of bile acids to modulate specific cell survival and death pathways that may determine their ultimate role in apoptosis. Interestingly, the activation of survival and antiapoptotic signaling limits programmed cell death in hepatocytes, which are constantly exposed to a variety of proapoptotic stimuli. In this regard, ursodeoxycholic acid prevents cell death by blocking classic pathways of apoptosis and activating certain survival pathways. Further, ursodeoxycholic acid and its derivatives may prove to be important therapeutic agents in the treatment of neurodegenerative and other disorders in which increased levels of apoptosis contribute to the pathogenesis of the disease.
Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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Bile Acid Metabolism and Physiology
Bile acids, the major constituents of bile, are detergent molecules synthesized in the liver from neutral sterols by a complex series of chemical reactions (Russell and Setchell, 1992). In humans and most animal species, bile acids are produced primarily from the cholesterol metabolic pathway (Rodrigues, 2000a). The expression of enzymes involved in bile acid synthesis is tightly regulated by nuclear hormone receptors and several other transcription factors, which ensure a steady supply of bile acids to a highly demanding metabolic environment. The initial and rate-limiting step in bile acid biosynthesis is the 7α-hydroxylation of cholesterol, catalyzed by the cytochrome P450 enzyme, cholesterol 7α-hydroxylase (CYP7A1); CYP7A1 gene transcription is repressed by primary bile acids but activated in a feed-forward manner by dietary cholesterol (Li et al., 1990). Subsequently, a soluble end product is obtained through a sequence of enzymatic reactions, including the addition of hydroxyl groups and the oxidation of the cholesterol side chain. Although the final product is always an amphipathic molecule, different bile acid species have diverse degrees of hydrophobicity, as determined by their biochemical and physicochemical properties. Further, the amphipathic structure of bile acids allows these water-soluble compounds to interact with proteins and insert into lipid bilayers. These effects will have profound influences on cell function and structure, particularly when intracellular concentrations of bile acids exceed certain limits, as it is in the case of cholestasis. Primary bile acids, such as cholic and chenodeoxycholic acids, are synthesized in the liver, conjugated with the amino acids glycine or taurine, and then secreted via the bile ducts and gallbladder into the lumen of the small intestine (Russell and Setchell, 1992). Due to their unique biochemical characteristics, bile acids act as detergents to emulsify dietary lipids and fat-soluble vitamins. Nevertheless, they also act within the liver for solubilization of bilirubin and other catabolites. Bile acids re-enter the liver via the portal vein, together with digested nutrients, and are recirculated back into the gallbladder for use in the next feeding cycle (Rodrigues, 2000a). The biliary bile acid pool also includes secondary bile acids, such as deoxycholic and lithocholic acids, which are generated by hydroxylation of primary bile acids in intestinal bacteria. An important example of this metabolism is the formation of ursodeoxycholic acid, which occurs by oxidation of chenodeoxycholic acid to 7-oxolithocholic acid, followed by bacterial reduction yielding the 7β-isomer. Nevertheless, ursodeoxycholic acid represents only a small fraction of the bile acid pool in humans. Microbial biotransformations may also include deconjugation and 7α-dehydroxylations.
15.3
Cell Death in Liver Diseases
The liver is a multifunctional organ with essential roles in metabolism, biosynthesis, excretion, secretion and drug detoxification. The complexity and energy requirements of these processes expose the liver to anoxia and toxic insults that, in normal situations, do not damage liver cells. This can perhaps be attributed to the activation of protective mechanisms, as well as to the large repair capacity of the liver. In fact, the controlled balance between
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cell division and cell death is an essential feature for the development and maintenance of liver homeostasis; any disturbance impairs liver function and contributes to liver disease. In chronic liver diseases and after acute or chronic exposure to insults such as alcohol, cytokines or bile acids, the rate of cell death may exceed the ability to regenerate. In contrast, insufficient cell death is a prerequisite for the development of hepatocellular carcinoma. A better understanding of how hepatocytes die and how cell death can be prevented has profound clinical implications. Hepatocyte death occurs in both acute and chronic liver diseases. Acute liver failure usually develops as a consequence of viral hepatitis, drug- and toxin-induced toxicity or liver transplantation. Chronic liver failure results from viral hepatitis and metabolic or autoimmune diseases, and is often accompanied by cholestasis and liver fibrosis (Schoemaker and Moshage, 2004). In both cases, however, endothelial cells, hepatocytes and cholangiocytes may be damaged due to the accumulation of toxic metabolites, reactive oxygen species (ROS) and bile acids. This ultimately leads to extensive cell death, which may compromise liver function and result in hepatic failure. In hepatocytes, cell death occurs through either apoptosis or necrosis (Majno and Joris, 1995), as well as a combination of the two. Necrosis usually results from metabolic disruptions, which quickly activate degradative enzymes and lead to cell lysis. It is often seen during ischemia/reperfusion or acute drug-induced hepatotoxicity. Apoptosis develops in a more orderly manner, usually as a result of very specific stimuli (Kerr et al., 1972). Unlike necrosis, cells break into small membrane-bound fragments, commonly known as apoptotic bodies. These are eliminated by circulating as well as neighboring cells, thus minimizing inflammatory responses and leakage of cellular components into the extracellular space (Reed, 2000). Apoptosis is particularly characteristic of acute liver injuries (Strand et al., 1996; Kasahara et al., 2000). Nevertheless, features characteristic of both apoptosis and necrosis may coexist in the same tissue, and may even represent the edges of a cell death continuum (Lemasters, 2005). In addition, there is growing evidence that hepatocytes can die via programmed cell death processes without typical features of apoptosis, such as caspase activation. Indeed, several other forms of cell death have been described, including paraptosis, autophagy, mitotic catastrophe and others (Broker et al., 2005).
15.3.1
Morphology and Physiological Implications of Apoptosis
Apoptosis represents a key feature of several acute and chronic liver diseases, such as viral hepatitis, cholestasis, iron/copper overload states, alcohol abuse, nonalcoholic fatty liver disease and ischemia–reperfusion liver injury (Guicciardi and Gores, 2005). This tightly controlled process can be characterized by its specific morphological and nuclear changes, such as cell shrinkage, chromatin condensation, internucleosomal DNA degradation and nuclear fragmentation. Ultimately, hepatocytes fragment into apoptotic bodies, which are removed by circulating macrophages and neighbor cells, mainly Kupffer and hepatic stellate cells, with little inflammatory response (Kerr et al., 1972). However, in certain pathological conditions, liver cell apoptosis may be accompanied by an inflammatory response, such as infiltration of neutrophils and the development of fibrosis (Canbay et al., 2004). Moreover, apoptotic cells may produce chemokines, and their engulfment by macrophages may also
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generate cytokines (Savill and Fadok, 2000; Faouzi et al., 2001) with numerous downstream effects. 15.3.2
Biochemical Pathways of Apoptosis
Apoptosis in the liver is initiated by two distinct pathways: the extrinsic pathway, activated by death receptors located at the cellular membrane, and the intrinsic pathway, involving the mitochondria organelle (Malhi et al., 2006). 15.3.2.1
Death Receptor Signaling
Death receptors are type 1 transmembrane proteins with three domains, belonging to the tumor necrosis factor/nerve growth factor receptor superfamily. The most largely expressed death receptors in the liver are Fas (CD95/Apo-1), tumor necrosis factor receptor 1 (TNFR1), tumor necrosis factor-related apoptosis inducing ligand (TRAIL) receptor 1 (TRAILR1) and TRAIL receptor 2 (TRAIL-R2) (Yoon and Gores, 2002). The triggering point in death receptor signaling is the engagement of transmembrane proteins by their cognate ligands, followed by recruitment of intracellular adaptor molecules forming the deathinducing signaling complex (DISC) (Kischkel et al., 1995) (Figure 15.1). Once activated, death receptors initiate cleavage of procaspase-8 to active caspase-8. In type I cells, the apoptotic cascade continues as caspase-8 directly activates downstream caspases, such as caspase-3, perpetuating the apoptotic process (Peter and Krammer, 2003). In type II signaling cells, such as hepatocytes, the progression of the apoptotic cascade depends entirely on its amplification by mitochondria (Li et al., 2002). In this case, caspase-8 cleaves Bid which is then translocated to the mitochondria, causing mitochondrial permeabilization and release of apoptogenic effectors, such as cytochrome c, which ultimately results in caspase-3 activation (Scaffidi et al., 1998). Toxic stimuli, such as bile acids, induce translocation of Fas to the plasma membrane in hepatocytes (Sodeman et al., 2000). In fact, hepatocytes constitutively express Fas on their cell surface and, therefore, are very susceptible to Fas-mediated signaling. In addition, bile acids trigger epidermal growth factor receptor (EGFR)-mediated tyrosine phosphorylation of Fas, enabling the formation of the DISC at the cell membrane (Reinehr and Haussinger, 2004). Ligand-independent activation of Fas may also occur if high levels of Fas are expressed at the cell membrane and the Fas ligand (FasL) itself induces translocation of Fas to the plasma membrane (Reinehr et al., 2004). In both cases, there is recruitment of Fas-associated death domain (FADD) protein to the oligomerized receptors. FADD contains a death effector domain (DED) that enables the activation of initiator caspases. Of note, apoptosis initiated by FasL is responsible for hepatocyte loss in diseases such as nonalcoholic steatohepatitis (Ribeiro et al., 2004) and fulminant hepatic failure, characterized by an uncontrolled massive death of hepatocytes (Ryo et al., 2000). In addition, it has a significant role in the removal of virus-infected hepatocytes and cancer cells by T lymphocytes (Berke, 1995). TNF-α overlaps with Fas signaling in many aspects, such as the activation of caspase-8 and targeting of the mitochondria. In fact, TNF-R1 activation is followed by recruitment of the adaptor TNF-R-associated protein with death domain (TRADD), which in turn may also recruit FADD as well as other adaptor proteins (Hsu et al., 1995). TNF-α signaling plays a pivotal role in liver diseases, including alcoholic hepatitis and fulminant hepatic failure (Bird et al., 1990; Streetz et al., 2000). However, TNF-R1 is uniquely different from
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FasL TRAIL Fas TRAIL-R1/R2
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Bcl-2 Bcl-x L
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DED
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DED
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Apaf-1 unfolding oligomerization
Caspase 3 Apoptosome
APOPTOSIS Figure 15.1 Schematic overview of receptor- and mitochondria-mediated apoptosis. In the death receptor pathway, and following interaction with its cognate ligand, the receptors recruit adaptor proteins such as FADD and initiator caspase-8 and or -10. Active initiator caspases then act on procaspase-3 that once activated cleaves key substrates in the cell to orchestrate apoptosis. In the mitochondrial pathway, death stimuli target mitochondria, either directly or through transduction by proapoptotic members of the Bcl-2 family, such as Bax. The mitochondria then release cytochrome c, Smac/DIABLO and other apoptogenic proteins. Cytochrome c induces the oligomerization of Apaf-1, which recruits and activates procaspase-9. Caspase-9 then activates procaspase-3 to induce apoptosis. The crosstalk between both pathways, mediated by Bid, is characteristic of type II cells, such as hepatocytes (see text for a more complete description) DED, death effector domain; FADD, Fas-associated death domain; IAPs, inhibitor of apoptosis proteins; TRAIL, tumor necrosis factor-related apoptosis inducing ligand’ TRAILR1/R2, TRAIL-receptor 1/receptor 2
Fas and TRAIL receptors, since it activates both apoptotic and survival signals such as the nuclear factor κB (NF-κB) (Barnhart and Peter, 2003). The role of TRAIL and TRAIL receptors during hepatocyte apoptosis is still controversial. Nevertheless, recent studies suggest that TRAIL may play a crucial role in hepatic cell death and hepatic inflammation (Zheng et al., 2004).
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15.3.2.2
Mitochondrial Pathway
In hepatocytes, apoptosis amplified by the mitochondrial pathway is necessary to carry out the apoptotic process (Bradham et al., 1998a). In addition, intracellular stress such as ROS or increased calcium levels, and endoplasmic reticulum (ER) stress may directly target mitochondria (Lemasters, 2005). Therefore, mitochondria represent key players in the apoptotic process, acting as integrating sensors of various death signals. Following an apoptotic stimulus, the mitochondrial membrane is rapidly permeabilized, releasing apoptogenic factors from the intermembrane space to the cytosol, whereas the mitochondrial membrane potential is disrupted (Ricci et al., 2003). This phenomenon is generally classified as the mitochondrial permeability transition (MPT). Furthermore, ATP production is impaired and, consequently, generation of ROS occurs. ROS production, frequent in hepatocytes during acute and chronic liver injuries, and a result of Kupffer and inflammatory cell activation (Kurose et al., 1997; Loguercio and Federico, 2003), can be enhanced by a variety of stimuli, including bile acids. In these situations, and with compromise of cell detoxification mechanisms, oxidative stress may reach levels capable of inducing mitochondrial dysfunction and apoptosis. However, with depletion of ATP or in the presence of exacerbate levels of ROS, necrosis usually overlaps apoptosis as the predominant type of cell death (Lemasters et al., 1998). Members of the Bcl-2 family of proteins control the regulation of mitochondrial membrane integrity and cytochrome c release (Table 15.1). Antiapoptotic Bcl-2 family members such as Bcl-2 and Bcl-XL are integral membrane proteins, which are usually localized in the outer mitochondrial membrane in healthy cells. On the other hand, proapoptotic Bax, Bid and Bad are sequestered in the cytosol as inert forms or inactive precursors, prior to Table 15.1 Classification of Bcl-2 mammalian family members according to their function. Antiapoptotic members have four Bcl-2 homology (BH) domains, BH1-4. Proapoptotic members have either three BH domains, BH1-3, and are classified as multidomain proteins, or only the BH3 domain, and are designated BH3-only proteins Bcl-2 family members Proapoptotic Antiapoptotic
Multidomain
BH3-only
Bcl-2 Bcl-xL Bcl-W Mcl-1 A1 Boo/DIVA
Bax Bak Bok/Mtd
Bid Bad Bim/Bod Bmf Bik/Nbk Krk/DP5 Blk Nip3 Bnip3/Nix Puma Noxa
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induction of cell death (Gross et al., 1999). In addition to onset of the MPT, proapoptotic Bcl-2 family members, such as Bax and Bak, may directly form specific pores in the mitochondrial outer membrane, contributing to the release of apoptogenic factors. However, the structure and composition of such pores is still not completely understood. In addition, proapoptotic Bcl-2 family members may also interact with and antagonize antiapoptotic Bcl-2 family members. Antiapoptotic Bcl-2 family proteins bind to and inhibit proapoptotic Bax and Bak, and sequester truncated Bid and apoptotic protease-activating factor 1 (Apaf-1), thus preventing the release of cytochrome c and other apoptogenic factors (Vander Heiden and Thompson, 1999). Once released, cytochrome c oligomerizes with Apaf-1, a cytosolic protein that normally exists as an inactive monomer, dATP and cytosolic procaspase-9 to form the apoptosome (Hengartner, 2000). This complex formation results in cleavage and activation of caspase-9, which in turn cleaves and activates executioner caspases such as caspase-3. Active downstream caspases will then cleave many important intracellular substrates that ultimately lead to the morphological changes characteristic of apoptosis. Other mitochondrial factors released from mitochondria include Smac/DIABLO and Omi/HtrA2, which act as inhibitors of members of the inhibitor of apoptosis proteins (IAPs) family. Ultimately, they influence both genomic stability and nuclear integrity through apoptotic pathways. Finally, the ER is capable of releasing proapoptotic signals that target mitochondria. Oxidative stress and other stimuli can inhibit ER calcium pumps and activate calcium release pathways. Cytosolic calcium can either promote mitochondrial signaling, or be taken up by the mitochondria itself and induce a calcium-dependent MPT. The release of apoptogenic factors is thought to trigger caspase-12 and Jun N -terminal kinase (JNK) activation, which are both mediators of caspase-3 activity. Interestingly, Bcl-2 family members may also associate with the ER. It has been suggested that pro-apoptotic Bcl-2 family members, such as Bax and Bak, can enlarge the size of ER calcium stores, thus increasing the proapoptotic potential of ER calcium release (Lemasters, 2005).
15.4 15.4.1
Bile Acid Induction of Apoptosis Bile Acid Effects in Cholestasis
Cholestasis is a common feature of several chronic human liver diseases. High levels of toxic bile acids accumulate to promote liver cell death and development of liver injury that can lead to cirrhosis (Hofmann, 2002). In fact, progression of disease appears to result from the effects of toxic bile acids, such as glycochenodeoxycholic and taurochenodeoxycholic acids, that accumulate within hepatocytes, and not as a direct result on the surface plasma membrane (Schmucker et al., 1990). Cholestasis is characterized by morphological, biochemical, physiological and clinical changes, such as alterations of the cytoskeletal and intracellular organelles, as well as increased concentrations of canalicular marker enzymes and solutes secreted into bile. Further, cell membrane damage, disruption of cellular integrity and leakage of intracellular components may also occur from solubilization of membrane-bound lipid by hydrophobic bile acids. This detergent action of bile acids occurs essentially at the canalicular membrane, where bile acid concentrations allow mixed micelle formation. Nevertheless, serum bile
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acid levels in cholestatic patients are insufficient to produce a significant detergent effect and, therefore, bile acid cytotoxicity appears to result from distinct processes. Furthermore, in order to reduce the intrahepatic load, transcriptional regulation of bile acid transporters decreases the uptake and increases secretion of bile acids (Guo et al., 2003). Survival genes are also activated. These mechanisms are usually not sufficient to impair the progress of hepatic injury and the occurrence of bile acid-induced cell death, from apoptosis and necrosis, is characteristic of cholestatic liver diseases. The presence of classic Councilman bodies and cell dropout in the liver of cholestatic patients suggests that apoptosis may play a prominent role in cholestasis. However, the predominant type of liver cell injury may depend on factors such as cell type, magnitude of the toxic stimulus and metabolic status of the cells. 15.4.2
Bile Acid-induced Apoptosis
The mechanisms by which bile acids induce apoptosis in hepatocytes are still not fully defined. Bile acids may initiate both ligand-dependent and -independent death-receptor oligomerization, as well as modulate downstream signaling pathways, a combination that strongly sensitizes cells to apoptosis (Figure 15.2). It has been shown that toxic bile acids induce apoptosis in a Fas- and TRAIL-dependent fashion (Faubion et al., 1999; Higuchi et al., 2003). Studies using a Fas-deficient cell line have demonstrated that glycochenodeoxycholate induces TRAIL-receptor 2/DR5 expression and oligomerization, followed by increased apoptosis (Higuchi et al., 2001). However, Fas-mediated signaling appears to be the most important mechanism behind bile-acid induced liver injury (Higuchi and Gores, 2003). Finally, deoxycholic acid (DCA) induces apoptosis of primary hepatocytes through ligand-independent activation of Fas and EGFR (Qiao et al., 2001). Activation of Fas by bile acids may result from modulation of the cellular trafficking of death receptors, which exhibit very distinct post-transcriptional regulation. They are usually sequestered in intracellular vesicles associated with the Golgi complex, and transported to the cytoplasmic membrane upon a specific stimulus. Toxic bile acids induce death receptor transport, by a Golgi-associated and microtubule-dependent pathway, a phenomenon that also involves protein kinase C (PKC) and JNK (Sodeman et al., 2000; Graf et al., 2002). Once at the cell membrane, spontaneous receptor oligomerization occurs. Alternatively, Fas activation by FasL or other agonists may also be increased. Curiously, this process is stimulated by the p53 tumor suppressor protein. In fact, p53 was shown to transiently increase Fas expression at the cell surface via transport from the Golgi complex, and to induce Fas-FADD binding, sensitizing human vascular smooth muscle cells to Fas-induced apoptosis (Bennett et al., 1998). Following Fas signaling, apoptosis proceeds with DISC assembly, caspase-8 activation and induction of the mitochondrial pathway of apoptosis. It is interesting to note that bile acids may directly inhibit apoptosis inhibitor cellular FLICE inhibitory protein (cFLIP) assembly into DISC. By phosphorylating cFLIP through a PKC-dependent mechanism, bile acids decrease cFLIP ability to bind FADD, resulting in increased activation of caspase8/10 (Higuchi et al., 2003). In addition, PKC function may be associated with activation of the lysosomal cysteine protease cathepsin B, a key effector contributing to the characteristic morphological features of apoptosis (Roberts et al., 1999). Nevertheless, cathepsin B may also play a role in liver necrosis (Jaeschke and Lemasters, 2003).
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Death Receptor
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FADD MAPKKK
PI3K Bid Caspase 8 Akt NF-κB
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p38 MPT ERK1/2
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Figure 15.2 Schematic representation of death and survival transduction pathways modulated by bile acids. Toxic hydrophobic bile acids induce apoptosis by activating both liganddependent and -independent death-receptor oligomerization. The activation of death receptors by bile acids invariably signals the mitochondrial pathway of apoptosis in type II cells, such as hepatocytes. Furthermore, toxic bile acids may directly target mitochondria, either through induction of the MPT and ROS or activation of proapoptotic Bcl-2 members. Finally, hydrophobic bile acids partially activate death receptor-dependent survival pathways, such as the NF-κB. Hydrophilic bile acids do not induce apoptosis as they simultaneously activate survival signaling pathways, such as MAPK and PI3K, antagonizing Bcl-2 proapoptotic members and preventing mitochondrial dysfunction and apoptosis (see text for a more complete discussion). MAPKK, MAPK kinase; MAPKKK, MAPKK kinase. cIAP1, inhibitor of apoptosis protein 1; ERK1/2, extracellular-regulated kinase 1/2; FADD, Fas-associated death domain; IκB, inhibitor of κB; MAPKK, MAPK kinase; MAPKKK, MAPKK kinase; MPT, mitochondrial permeability transition; NF-κB, nuclear factor κB; P13K, phosphoinositol 3-kinase; TRAF1, tumor necrosis factor receptor-associated factor 1
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The activation of death receptors by bile acids invariably signals the mitochondrial pathway of apoptosis in hepatocytes. Some of the mechanisms implicated in bile acid toxicity include stimulation of lipid peroxidation and induction of mitochondrial dysfunction. In fact, DCA induces significant hepatocyte apoptosis in vivo and in vitro, as well as the MPT in isolated mitochondria (Rodrigues et al., 1998a). Furthermore, bile acid-induced MPT is prevented by a known inhibitor of the megapore channel, cyclosporine A, which in turn inhibits bile acid-induced hepatocyte cytotoxicity (Botla et al., 1995; Rodrigues et al., 1998b). Curiously, apoptosis triggered by toxic bile acids in colon cancer cell lines appears to result solely from the direct induction of the MPT (Wachs et al., 2005). Nevertheless, DCA-induced apoptosis may also result from disruption of mitochondrial transmembrane potential, increased ROS production and translocation of proapoptotic Bax to the mitochondria and release of cytochrome c (Rodrigues et al., 2003a). The mechanisms by which toxic bile acids promote oxidative stress and trigger the Fas signaling pathway may involve the activation of NADPH oxidase isoforms (Reinehr et al., 2005). DCA-induced generation of ROS from the mitochondria may also result in inhibition of PTPases, with subsequent activation of multiple growth factor receptors and downstream signaling pathways in hepatocytes, including the extracellular-regulated kinase (ERK) pathway (Fang et al., 2004). Curiously, for conjugated bile acids, the activation of G-protein coupled receptors is necessary for activation of receptor tyrosine kinases (Dent et al., 2005a). The generation of ROS is also required for activation of the insulin receptor/phosphoinositol 3-kinase (PI3K)/Akt pathway by DCA in primary rat hepatocytes (Han et al., 2004). In fact, DCA toxicity may be either reduced, if Akt and ERK1/2 are activated, or enhanced, if ROS levels affect and disrupt the mitochondrial membrane potential, leading to the release of cytochrome c into the cytosol. Ultimately, the production of ROS by hepatocytes may result in pro- and/or antiapoptotic events, depending on the specificity and level of the stimulus (Tirosh et al., 2003). In this respect, the type of liver injury observed during cholestasis may depend on the concentration of the toxic bile acid pool within cells. In fact, oxidative stress induced by moderate levels of toxic bile acids, that are typically observed during cholestasis, usually triggers cell death through apoptosis (Rodrigues et al., 1998a,b). On the other hand, bile acid-induced oxidative stress may reach levels in which mitochondrial function is severely compromised, leading to a rapid loss of ATP synthesis, coupled with substantial cytochrome c release. In this case, hepatocyte typically die from necrosis (Sokol et al., 2001).
15.5 15.5.1
Bile Acid Inhibition of Apoptosis Cell Survival Signaling
The liver is a unique organ that exhibits a large number of protective mechanisms against cytotoxic insults, as well as a great capacity for repair. In fact, liver cells are typically rich in antioxidants such as glutathione (Xu et al., 1998). Bile acids transporters also exhibit adaptive responses in the presence of high intracellular levels of toxic bile acids to protect liver cells (Trauner et al., 1999). Nevertheless, it is the activation of survival signaling pathways that characterizes the unique antiapoptotic response of hepatocytes to a variety of toxic stimuli. In this regard, bile acids specifically prevent and counteract their inherent cytotoxicity by activating survival pathways. These include the NF-κB, PKC,
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PI3K and mitogen-activated protein kinase (MAPK) pathways, calcium-dependent signal transduction cascades and cyclic adenosine monophosphate (cAMP). In the liver, the NF-κB signaling pathway acts as a critical sensor and integrator of both death and survival signals. In normal situations, NF-κB is tightly bound to an inhibitory protein, termed IκB, in the cytoplasm, which in turn inhibits its translocation to the nucleus and subsequent activation of target genes. Following cellular stresses, NF-κB can be activated by several molecules, including inflammatory cytokines (Schuchmann and Galle, 2002). In fact, TNF-α and interleukin-1β were found to be overexpressed during acute and chronic liver failure (Bradham et al., 1998b). Despite its involvement in apoptosis, TNF-α can also stimulate survival pathways via NF-κB-dependent gene expression. In addition, by recruitment of adaptor proteins, it induces IκB kinase (IKK) and proteosomal IκB degradation, thus leading to NF-κB nuclear translocation and activation (Barnhart and Peter, 2003). The inhibition of NF-κB was shown to induce apoptosis in hepatocytes, supporting the role of this transcription factor in modulating antiapoptotic genes (Bellas et al., 1997). Some candidate genes, which represent prime targets for novel therapies in liver diseases, are TNF receptor-associated factor 1 and 2 (TRAF1 and TRAF2), cIAP1 and cIAP2, interfering with caspase-8 activation (Wang et al., 1998), as well as members of the Bcl-2 family. Thus, the activation of NF-κB may explain, in part, the resistance of hepatocytes to TNF-α-induced cytotoxicity. The PI3K family of enzymes has also been implicated in controlling hepatocyte survival in response to bile acids. Furthermore, this pathway is involved in the regulation of transporters during bile formation. Downstream effectors of PI3K signaling include several PKC isoforms, such as PKCζ , and Akt, a serine/threonine kinase with multiple roles. Akt may inhibit apoptosis by modulating the expression and/or activity of caspases and Bcl-2 family members, including Bad. It is thought that Akt phosphorylates Bad, which will no longer associate with and inhibit antiapoptotic Bcl-XL . In addition, Akt phosphorylates and inactivates caspase-9, suppresses translocation of proapoptotic Bax to the mitochondria and activates the forkhead family of transcription factors (Datta et al., 1999). Interestingly, the PI3K signaling cascade may communicate with the NF-κB pathway. In fact, much like Akt, some PKC isoforms activate IKK, which phosphorylates IκB or the RelA subunit of NF-κB, increasing its transcriptional activity. In that sense, PKCζ has been described as a critical component of the taurochenodeoxycholate-induced PI3K survival cascade and its activation resulted in NF-κB-dependent transcriptional activity (Rust et al., 2000). Nevertheless, activation of PKCs have also been implicated in bile-acid induced cholestasis and apoptosis. For example, glycochenodeoxycholic acid-induced apoptosis requires activation of two PKC isoforms (Jones et al., 1997). Therefore, different PKC proteins, as well as their downstream targets, may have opposite effects in regulating apoptosis, although their precise role has not yet been clearly established. Finally, the MAPK pathway is another important survival pathway highly modulated by bile acids in hepatocytes. MAPKs are a superfamily of homologous enzymes that integrate and transduce intracellular and extracellular signals into the nucleus and are usually classified in three distinctly regulated groups, the ERK1/2, p38 kinase and JNK pathways. Each MAPK cascade is characterized by a cassette of three enzymes, which sequentially phosphorylate and activate a downstream kinase and, finally, transcription factors. While the ERK pathway transduces growth factor signals involved in differentiation, proliferation and secretion, the others function as stress signal transducers. Interestingly, signaling
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pathways involving ERK1/2 and p38 have also been implicated in biliary excretion of bile acids. DCA-induced activation of EGFR signaling, via the MAPK pathway, counteracts its proapoptotic effects (Qiao et al., 2001). Additionally, DCA activates the PI3K signaling pathway. Recent studies have suggested that the activation of the insulin receptor and Akt by DCA provides a greater cytoprotective effect than DCA-induced activation of EGFR and ERK1/2 (Dent et al., 2005). Depending on the conjugation state of the bile acid, either one or another survival pathway is more robustly activated to preserve cell viability. JNK signaling may either have a facilitating or inhibitory role in apoptosis, depending on the cell type and level of activation. In hepatocytes, it has been more extensively studied with respect to its proapoptotic role. In fact, JNK activation occurs during Fas-mediated apoptosis and is responsible for modulation of apoptosis-related proteins, such as p53 and c-myc, or activation of activator protein 1 (AP-1) (Davis, 2000). AP-1 may then transcriptionally up-regulate proapoptotic proteins; it has been shown to act upstream of the mitochondrial changes that result in cytochrome c release (Liu et al., 2002). Glycochenodeoxycholateinduced apoptosis is associated with modulation of AP-1 in a human hepatoma cell line by increasing transcription of c-Fos and JunB, up-regulating AP-1 activity and phosphorylating JNK and p38 (Bernt et al., 2006). The protective effect of NF-κB in hepatocytes may also result from the inhibition of JNK signaling by an NF-κB responsive gene product (Tang et al., 2001). In this regard, JNK inhibition was shown to reduce liver damage in a model of ischemia reperfusion injury (Uehara et al., 2005). Nevertheless, it is conceivable that JNK1 and JNK2 may differentially regulate the apoptotic response of cells exposed to toxic stimuli, depending on the status of parallel survival pathways (Dent et al., 2005). In fact, the JNK1/2 pathway plays a protective role in TNF-α-induced apoptosis in rat hepatocytes (Liedtke et al., 2002). 15.5.2 15.5.2.1
Antiapoptotic Pathways Role of Ursodeoxycholic Acid
Unlike several toxic hydrophobic bile acids, ursodeoxycholic acid (UDCA) displays a hepatoprotective effect. Its beneficial effects in primary biliary cirrhosis provided the first clues on the different biological effects of bile acids in health and disease. In fact, UDCA is currently used as a therapeutic agent for several hepatobiliary disorders, especially cholestatic liver diseases (Beuers et al., 1998; Lazaridis et al., 2001). Both the unconjugated form and its amidated conjugates, tauroursodeoxycholic acid (TUDCA) and glycoursodeoxycholic acid (GUDCA), are effective at protecting against the toxicity of more hydrophobic bile acid species (Rodrigues and Steer, 2000). Some major mechanisms of action include protection of cholangiocytes against the cytotoxicity of hydrophobic bile acids, stimulation of hepatobiliary secretion and protection of hepatocytes against bile acid-induced apoptosis (Paumgartner and Beuers, 2002). Indeed, in contrast to the effects of hydrophobic bile acids, cells exposed to UDCA alone up to certain levels exhibit no significant apoptotic changes. More importantly, while toxic bile acids fed to rats induce apoptosis in the liver, UDCA inhibits this effect in vivo, by preventing formation of ROS and translocation of the proapoptotic protein Bax from the cytosol to the mitochondria (Rodrigues et al., 1998a, 1999) (Figure 15.3). Furthermore, the ability of UDCA to modulate apoptosis is relevant to a variety of agents acting through different apoptotic pathways, and in different cell
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Figure 15.3 Modulation of DCA-induced apoptotic features by UDCA in isolated mitochondria and primary rat hepatocytes. (a) Incubation of mitochondria with UDCA does not affect organelle morphology; however, exposure to DCA results in mitochondrial swelling, rupture of the outer membrane and reduced number of intact mitochondria. Coincubation with UDCA significantly prevents mitochondrial alterations in structure. (b) As predicted, DCA-induced cytochrome c release from isolated mitochondria is significantly inhibited by UDCA. (c) DCA induces apoptosis in primary rat hepatocytes as evidenced by Hoechst staining showing nuclear condensation and fragmentation. UDCA is not significantly different from controls and reduces DCA-induced morphological changes. (d) Increased ROS production (HE → ethidium) and reduced mitochondrial transmembrane potential [DiOC6(3)] during DCA-induced apoptosis are inhibited when hepatocytes are preincubated with UDCA. As analyzed by cytofluorometry, the percentages reflect mitochondrial depolarization and ROS production. (e) The final steps of the apoptotic pathway are also strongly modulated by bile acids. When coincubated with DCA, UDCA significantly reduces cytochrome c release from the mitochondria to the cytosol, as well as PARP cleavage. DiOC6 (3), 3,3 -dihexyloxacarbocyanine iodide; FSC, forward scatter; HE, dihydroethidium. Reprinted by permission from Macmillan Publishers Ltd: CELL DEATH DIFFER, Rodrigues CMP, Ma X, Lineham-Stieers C, Fan G, Kren BT and Steer CJ, Ursodeoxycholic acid prevents cytochrome c release in apoptosis by inhibiting mitochondrial membrane depolarization and channel formation, 6, 842–854, copyright 1999
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types (Rodrigues et al., 1998b). UDCA almost completely inhibited DCA-induced apoptosis in primary rat hepatocytes and HuH-7 cells, as well as apoptosis induced by ethanol, transforming growth factor β1 (TGF-β1), anti-Fas antibody and okadaic acid. Perhaps somewhat unexpected, UDCA was also protective against apoptosis in nonhepatic cells, such as Saos-2, HeLa and Cos-7 cells. One mechanism for the reported antiapoptotic function of UDCA involves the mitochondrial membrane, either directly or indirectly (Figure 15.4). UDCA, and its taurine and glycine conjugates, prevent the release of cytochrome c, caspases activation, cleavage of the nuclear enzyme Poly(ADP-ribose) polymerase (PARP) and alterations in transmembrane potential induced by apoptotic triggers (Rodrigues et al., 1999). Furthermore, TUDCA has a profound effect on Bax-mediated structural changes in the lipid and protein environment of the mitochondrial membrane (Rodrigues et al., 2003a). In fact, Bax modifies the polarity and fluidity of the membrane lipid core, thus promoting the formation of an apoptotic pore and allowing the subsequent efflux of proteins. This was assessed by spectroscopic analyses of isolated mitochondria exposed to recombinant Bax. In addition, Bax insertion into the mitochondrial membrane(s) markedly disturbs the order structure of several proteins. TUDCA was proven to be a potent suppressor of Bax-induced alterations, in part, by preventing the binding of Bax to the mitochondria and inhibiting oxidative lipid injury. UDCA was also shown to partially prevent apoptosis, via the death receptor pathway, in primary mouse hepatocytes co-cultured with fibroblasts that express the Fas ligand (Azzaroli et al., 2002). However, a nitric oxide derivative of UDCA, but not UDCA alone, was shown to protect against liver damage in murine models of autoimmune hepatitis and to prevent caspase activation in HepG2 cells treated with a Fas agonistic antibody (Fiorucci et al., 2001). In this case, it was suggested that cysteine S-nitrosylation by intracellular nitroxide formation was the primary mechanism responsible for caspase inhibition, rather than an effect of UDCA. Although not fully understood, the endoplasmic reticulum stress-mediated pathway, which involves disruption of calcium homeostasis and caspase-12 activation, may also be regulated by TUDCA (Xie et al., 2002). Recent studies have suggested that UDCA may interfere with alternate molecular targets upstream of the mitochondria in hepatocytes. UDCA inhibited TGF-β1-induced E2F-1 transcriptional activation, p53 stabilization and p53-associated Bax expression, independently of its effect on mitochondria and/or caspases (Sol´a et al., 2003a). Furthermore, UDCA modulated the down-regulation of antiapoptotic Bcl-2 induced by TGF-β1, which correlates with decreased stabilization and/or degradation of p53 and NF-κB, respectively. Finally, it is becoming increasingly evident that activation of survival pathways may represent an important additional mechanism by which UDCA inhibits apoptosis. For example, it can stimulate the intracellular MAPK signaling pathway, via activation of the EGFR (Qiao et al., 2002). Indeed, abolishment of UDCA-induced MAPK signaling potentiated bile-acid induced apoptosis. In addition, UDCA was shown to inhibit DCA-induced apoptosis in human colon cancer cells, also through modulation of EGFR/Raf-1/ERK signaling (Im and Martinez, 2004). It appears that TUDCA is able to regulate cell death through similar mechanisms. Moreover, it was recently shown that the p38/ERK/MAPK and PI3K pathways are involved in the protection of TUDCA against glycochenodeoxycholic acid-induced apoptosis in hepatocytes (Schoemaker et al., 2004), probably representing a mechanism that converges on Bax. The protective effect of TUDCA occurs, in part, at a post-transcriptional
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Figure 15.4 Proposed mechanisms for the antiapoptotic actions of UDCA and TUDCA. UDCA protects hepatocytes from FasL-induced apoptosis, probably through interactions involving mitochondria. In fact, in addition to the activation of survival pathways, UDCA has been shown to negatively modulate the mitochondrial pathway of apoptosis by inhibiting Bax translocation to the mitochondrial membrane, ROS formation, cytochrome c release, and caspase-3 activation. Furthermore, UDCA and TUDCA also inhibit apoptosis associated with ER stress. TUDCA, specifically does so through modulating intracellular calcium levels, thus inhibiting calpain activation and procaspase-12 processing (see text for a more complete discussion). FasL, Fas ligand; TUDCA, tauroursodeoxycholic acid
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level, and is independent of inhibition of caspases and competition at the cell membrane with the toxic bile acid. Interestingly, TUDCA inhibits apoptosis in neuronal cells through the activation of survival pathways such as the PI3K pathway. In fact, glutamate-induced neurotoxicity in primary rat cortical neurons is associated with down-regulation of antiapoptotic members of the Bcl-2 family and dephosphorylation of Akt (Castro et al., 2004). Pretreatment with TUDCA significantly reduced apoptosis induced by glutamate, in part through phosphorylation and translocation of Bad from mitochondria to the cytosol. Importantly, inhibition of the PI3K pathway abrogated the protective effects of TUDCA, including the phosphorylation and translocation of Bad. Furthermore, amyloid-β (Aβ)-induced neuronal apoptosis is also associated with translocation of Bax to the mitochondria, cytochrome c release, caspase activation and DNA and nuclear fragmentation (Sol´a et al., 2003b). Remarkably, coincubation with TUDCA significantly modulates each of the Aβ-induced apoptotic events. In addition, while PI3K inhibitor, wortmannin, does not markedly affect Aβ-induced cell death, it decreases TUDCA protection against Aβ-induced apoptosis, reduces Akt phosphorylation and increases Bax translocation to mitochondria, supporting a strong role for the PI3K signaling cascade in regulating the antiapoptotic effects of TUDCA in several models. Recent studies have provided a further insight into the mechanisms by which cytoprotective bile acids may induce survival signaling pathways in cholangiocytes (Marzioni et al., 2006). In contrast to the direct activation of MAPK and PI3K signaling on hepatocytes, bile acids mostly dependent on the calcium signaling for the same activation in cholangiocytes. In this model, in vivo administration of UDCA and TUDCA to rats subjected to bile duct ligation and vagotomy, a model of cholestasis, prevented cholangiocyte apoptosis and the inhibitory effect on the phosphorylation of ERK1/2 and Akt. In contrast, these effects were blunted by blockage of calcium signaling, confirming the importance of the calcium-dependent PKC-α signaling induced by UDCA and TUDCA. It is interesting to note that other bile acids may activate survival signaling pathways, thus counteracting their inherent cytotoxicity. Taurochenodeoxycholic acid stimulates PI3K and, subsequently, PKCζ activity; the inhibition of this survival pathway renders taurochenodeoxycholic acid cytotoxic (Rust et al., 2000). Finally, it is conceivable that UDCA may act differentially on death/survival pathways, depending on the physiological conditions of the cell and/or type of stimulus. In fact, UDCA displays a chemopreventive role in a rodent model of colonic carcinogenesis (Earnest et al., 1994) and protects against the development of human colon cancer in several premalignant conditions (Tung et al., 2001; Serfaty et al., 2003). In addition, UDCA was demonstrated to inhibit the growth of tumors when associated with photodynamic therapy (Garbo et al., 2003). Recent studies suggest that cyclin D1 may be an important regulatory factor implicated in these processes (Wali et al., 2002). In that sense, apoptosis and cell cycle related genes, including cyclin D1, were recently shown to be regulated by UDCA (Castro et al., 2005). More importantly, targeting of cyclin D1 by UDCA and TUDCA may modulate their ability to prevent apoptosis (Castro et al., 2007). 15.5.2.2
Role of Nuclear Steroid Receptors
Many natural ligands of nuclear steroid receptors (NSR) are products of the cholesterol biosynthetic pathway (Whitfield et al., 1999). Thus, as cholesterol-derived molecules, bile
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acids have recently been identified as activators of NSR. Indeed, the structure of most bile acids is similar to that of steroid hormones (Mendoza et al., 2002). NSR regulate genes involved in bile acid biosynthetic pathways and cholesterol metabolism. For example, the farnesoid X receptor (FXR) modulates bile acid synthesis, transport and absorption, as well as reverse cholesterol transport (Makishima et al., 1999; Parks et al., 1999; Wang et al., 1999). It has been shown that a number of bile acids, including chenodeoxycholic, cholic, deoxycholic and lithocholic acids, bind and inactivate FXR in cultured cells. In addition, the pregnane X receptor (PXR), or its human ortholog steroid and xenobiotic receptor (SXR), regulates lithocholic acid and drug metabolism, exhibiting an essential role during detoxification of cholestatic bile acids (Staudinger et al., 2001; Xie et al., 2001). Interestingly, UDCA is a modulator of NSR activity. Although UDCA itself does not bind FXR (Makishima et al., 1999), this bile acid inhibits receptor activation by more hydrophobic bile acids (Howard et al., 2000). Other studies have shown that UDCA activates PXR to induce CYP3A4, a bile acid-metabolizing enzyme in human hepatocytes (Schuetz et al., 2001). Furthermore, UDCA interacts with the glucocorticoid receptor (GR) in the absence of specific ligands, suggesting it to be a novel and selective GR modifier (Tanaka and Makino, 1992; Tanaka et al., 1996; Mitsuyoshi et al., 1997; Miura et al., 2001). Under certain conditions, UDCA may also enhance GR-responsive gene expression. It has been described that in the presence of the classical GR agonist, dexamethasone, UDCA increases GR-induced tyrosine aminotransferase, a hepatocyte-specific marker of GR (Mitsuyoshi et al., 1997). Nevertheless, the interaction between UDCA and GR also results in nuclear translocation of GR without receptor transactivation. UDCA appears to act at the level of a distinct region of the LBD when compared with dexamethasone, attenuating the interaction between GR and coactivators (Miura et al., 2001). These data may explain the anti-inflammatory properties of UDCA. In fact, UDCA suppressed NF-κBdependent transcription and subsequent cytokine expression via the interaction of GR and NF-κB. It is assumed that transrepression of transcription factors, such as AP-1 and NF-κB, is the primary mechanism by which glucocorticoids mediate their anti-inflammatory activity. Different ligands induce various conformations of NSR, resulting in unique regulatory properties of the receptor (Bourguet et al., 2000). In addition, UDCA pretreatment was shown to suppress DNA binding activity of AP-1 during DCA incubation of human colon cancer cells (Im and Martinez, 2004). It is, therefore, possible that UDCA may modulate several apoptotic-related genes by functionally modulating GR (Figure 15.5). The mineralocorticoid receptor (MR) blocks apoptosis by increasing the ratio of antito proapoptotic proteins (Almeida et al., 2000) and inhibiting proteolytic processing of procaspase-3, -8 and -9 and PARP (Planey et al., 2002). However, in contrast to GR, there is no evidence that MR and UDCA interact directly. Nevertheless, it has been shown that MR and GR can bind similar ligands and target the same genes (Lim-Tio and Fuller, 1998). MR and GR are heteromeric complexes found in dynamic association with a growing number of chaperone proteins and other factors mediating their actions. Thus, it is conceivable that UDCA-mediated modulation of GR may also affect MR. In addition, bile acid-induced nuclear translocation of NSR may not always result in transactivation or transrepression of NSR. In fact, it is possible that bile acids may only require NSR to reach the nucleus, where they themselves regulate genes through modulation of other transcription factors. In this regard, it has been shown that hormone-bound NSR
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Figure 15.5 Role of nuclear steroid receptors in the antiapoptotic function of hydrophilic UDCA. UDCA appears to interact with a specific region of the NSR ligand binding domain, leading to NSR/hsp90 dissociation and nuclear translocation of the receptors. The nuclear trafficking of UDCA allows it to modulate the E2F1/p53/Bax apoptotic pathway, thus preventing apoptosis. UDCA may further inhibit the apoptotic cascade by regulating other apoptoticrelated genes through modulation of other transcription factors (see text for a more complete discussion). hsp90, heat shock protein 90; NSR, nuclear steroid receptor; UDCA, ursodeoxycholic acid
are not permanently localized in the nucleus. Rather, they rapidly and continuously shuttle between the nucleus and cytosol (Madan and DeFranco, 1993; Sackey et al., 1996). Similar to other bile acids, UDCA has been detected in the nucleus of hepatocytes (Setchell et al., 1997; Monte et al., 2002). During liver regeneration, an increased amount of UDCA was shown to inhibit bile acid-FXR mediated repression of several genes (Chiang et al., 2000; Repa et al., 2000). Moreover, in human liver tissue obtained from patients on oral bile acid therapy, while only a small portion of the bile acid pool resides within the liver, there appears to be selective uptake by the hepatocyte nucleus. In fact, UDCA was shown to be the most abundant bile acid detected in the nucleus of hepatocytes (Setchell et al., 1997; Monte et al., 2002). Increased levels of UDCA were also observed in nuclei during hepatocarcinogenesis
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(Mendoza et al., 2002). Therefore, it has been suggested that UDCA may bind to cytosolic NSR and translocate to the nucleus as part of the ligand–receptor complex. It was recently demonstrated that UDCA modulates the E2F-1/Mdm-2/p53 apoptotic pathway in hepatocytes via a NSR-dependent mechanism (Sol´a et al., 2004). UDCA pretreatment of hepatocytes during TGF-β1-induced apoptosis up-regulated both GR and MR expression. This effect was interesting in light of previous studies showing that activation of GR represses the effects of TGF-β1 in rat hepatoma cell lines (Iida et al., 1998; Yamamoto et al., 1998; Li et al., 2003). Moreover, these results suggested a mechanistic function for MR during hepatocyte apoptosis. Although the antiapoptotic action of MR has been extensively described in neuronal cells, less was known about its role during hepatocyte apoptosis. It was subsequently shown that UDCA promotes GR/heat shock protein 90 (hsp90) dissociation, thus inducing subsequent NSR translocation (Sol´a et al., 2005). However, with deletion of the C-terminal region of GR, UDCA no longer induced GR/hsp90 dissociation and GR nuclear translocation, nor protected against apoptosis, indicating that GR legend binding domain (LBD) is required for the antiapoptotic function of UDCA. Additional studies suggested that UDCA either directly binds to the COOH-terminal region of GR LBD, or somehow interacts via a second messenger signal. However, the nuclear translocation of UDCA in association with GR strongly suggested that they were bound to each other as an UDCA–GR complex. Nevertheless, a physical interaction between the classical GR and the UDCA molecule has not been absolutely confirmed. Surprisingly, the bile acid does not appear to require NSR transactivation to prevent apoptosis in hepatocytes (Sol´a et al., 2005). Other studies have shown that UDCA attenuates the interaction between GR and its co-activators (Tanaka et al., 1996; Hong et al., 1997; Miura et al., 2001). Nevertheless, UDCA alone increases the activation of glucocorticoid response elements in hepatocytes that overexpress GR, but does not protect them against TGF-β1-induced apoptosis by further increasing GR transactivation. Therefore, GR appears to function as a transporter of UDCA to the nucleus where it then acts to modulate apoptosis-related genes.
15.6
Bile Acids for the Treatment of Neurodegenerative Disorders
As for many liver diseases, mitochondrial impairment appears to play a major role in a variety of brain disorders, including acute stroke and chronic neurodegenerative diseases. Mitochondrial compromise in Huntington’s disease (HD), for example, is characterized by abnormal energy metabolite concentrations and utilizations, impaired striatal mitochondrial respiratory chain complex II/III activity and increased stress-induced mitochondrial depolarization, free radical production and associated oxidative damage. A consequence of mitochondrial dysfunction in neuronal cells may be the activation of apoptotic pathways. In that sense, caspases, crucial for the initiation and execution of apoptosis are elevated and activated in HD brain. In acute types of injury, such as in the ischemia–reperfusion injury, neurons also exhibit several features of apoptosis, including chromatin condensation, transferase mediated dUTP nick-end labeling (TUNEL) and caspase activation. Furthermore, Bax protein is increased in the core, as well as in cells at the periphery of the ischemic infarct that exhibit DNA fragmentation. In contrast, Bcl-2 and Bcl-xL proteins are increased only in neurons that are adjacent to an infarct and appear to survive ischemic brain injury. The therapeutic role of UDCA has been established in the treatment of certain liver diseases and may have an even greater application for non-liver diseases in which increased
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levels of apoptosis contribute to their pathogenesis. In fact, the ability of UDCA to modulate apoptosis at several levels suggests a basic, essential function of this bile acid in regulating cell survival, independently of cell type. After conjugation with taurine, UDCA administrated in high doses can be delivered to other tissues, including the brain (Keene et al., 2001). In this regard, it has been shown that TUDCA inhibits apoptosis induced by several stimuli in neuronal cells (Rodrigues et al., 2000b; Sol´a et al., 2003; Castro et al., 2004). More significantly, the antiapoptotic properties of TUDCA have been explored in various animal models of neurological disorders associated with increased levels of apoptosis, including HD, Parkinson’s disease (PD) and acute ischemic and hemorrhagic stroke. Chronic administration of 3-nitropropionic acid (3-NP), an irreversible inhibitor of succinate dehydrogenase, provides a relatively accurate animal model of HD, where metabolic compromise, followed by activation of excitatory amino acid receptors, and free radical production appears to play a major role in the pathogenesis of the lesion. The systemic administration of TUDCA in the 3-NP rat model of HD was shown to reduce the associated morphologic striatal lesions (Keene et al., 2001). Moreover, behavioral studies correlated well with histological findings, since significant neuroproptection resulted in almost complete prevention of hyperactive behavior associated with 3-NP administration and maintained neophobia characteristic of cognitively intact animals. Similar conclusions were obtained in vitro, where TUDCA markedly reduced mitochondrial perturbations of apoptosis induced by 3-NP in cultured neuronal cells (Rodrigues et al., 2000b). These data suggested that TUDCA could possibly be used as a therapeutic agent for HD. Additional studies were performed in the R6/2 transgenic mouse model of HD, which expresses exon 1 of the human huntingtin gene with ∼ 150 CAG repeats, as well as the human huntingtin promoter. Because it results from genetic rather than chemical alterations, and involves chronic versus acute pathophysiology, the R6/2 transgenic mouse may more accurately reflect the true pathophysiology of HD. TUDCA administration led to a marked reduction in striatal cell apoptosis and degeneration of R6/2 transgenics (Keene et al., 2002). Control R6/2 mice exhibited prominent atrophy associated with reduced striatal volumes compared with control and TUDCA-treated wild-type mice. More importantly, TUDCA-treated R6/2 mice showed significantly less cerebral atrophy when compared with untreated transgenic animals. Furthermore, TUDCA decreased the average size and number of individual intracellular huntingtin inclusions, significantly improving HD pathology. Finally, it was not surprising that both locomotor and sensorimotor abilities improved substantially in HD mice treated with TUDCA. Based on its antiapoptotic properties, TUDCA was added to cell suspensions prior to transplantation to improve survival of nigral grafts in a rat model of PD (Duan et al., 2002). In fact, the results showed that TUDCA reduced apoptosis of cultured cells and, specifically, of dopamine positive cells. In transplantation studies, where TUDCA treatedcell suspensions were injected into the striatum of 6-hydroxydopamine lesioned rats, there was a significant improvement of the amphetamine-induced rotation scores. Furthermore, the number of apoptotic cells was much smaller in the graft areas of TUDCA-treated groups, whereas tyrosine–hydroxylase positive cells were more than 3-fold increased. More recently, TUDCA was shown to partially rescue a Parkinson’s disease model of C. elegans from mitochondrial dysfunction (Ved et al., 2005). Prompted by these studies, a well-characterized rat model of transient focal cerebral ischemia was studied to assess the protective effect of TUDCA (Rodrigues et al., 2002).
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Intravenous TUDCA administration resulted in improved neurologic function and reduced infarct volumes by ∼ 50 %. In addition, TUDCA significantly reduced the number of TUNEL-positive cells, prevented mitochondrial swelling and membrane disruption, and partially inhibited downstream caspase activation and endogenous substrate cleavage associated with apoptosis. TUDCA also reduced the appearance of TUNEL-positive cells, activation of caspase-3-like proteases and histological damage of the peri-hematoma region, in a collagenase-induced hemorrhagic model of stroke (Rodrigues et al., 2003b). In addition, NF-κB activation was significantly decreased, Bcl-2 levels remained elevated and rats showed improved neurological function after TUDCA administration. The activation of survival pathways may also be playing a prominent role in this model, as TUDCA also induced Bad phosphorylation through the activation of the PI3K survival pathway.
15.7
Summary
Bile acids are important signaling molecules, capable of modulating distinct celltransduction pathways. The hepatic accumulation of toxic bile acids is thought to play a key role in cholestatic liver injury, which is caused in part by hepatocyte apoptosis. In fact, toxic hydrophobic bile acids induce hepatocyte apoptosis through both the Fas receptor and classical mitochondrial pathways. In contrast, the antiapoptotic properties of certain bile acids, such as UDCA, against different cytotoxic stimuli have been demonstrated in several cell types. In addition to its effects at the mitochondrial level, UDCA prevents apoptosis through modulation of key cellular survival pathways, including the MAPK and the PI3K pathways. Interestingly, the antiapoptotic function of UDCA appears to involve NSR as key factors, which are specifically required for the nuclear traffic of this bile acid. Finally, the taurine-conjugated form of UDCA, TUDCA, also inhibits apoptosis. Furthermore, it is a hydrophilic molecule, readily water-soluble, that can be administered orally or intravenously, and is taken up by the liver with a high first-pass clearance. Given their lack of toxicity and striking antiapoptotic properties, UDCA and TUDCA may prove to have therapeutic benefit over other drugs in treating disorders where the metabolic compromise renders cells susceptible to apoptosis.
Acknowledgments Supported, in part, by grant POCI/SAU-FCT/62479/2004 from Funda¸ca˜ o para a Ciˆencia e a Tecnologia (FCT), Lisbon, Portugal (to CMPR). REC and SS are recipients of Ph.D. (SFRH/BD/12655/2003) and postdoctoral (SFRH/BPD/20834/2004) fellowships, respectively, from the FCT. The authors thank all members of their laboratory for help during the course of these studies.
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Russell DW and Setchell KD (1992). Bile acid biosynthesis. Biochemistry 31: 4737–4749. Rust C, Karnitz LM, Paya CV, Moscat J, Simari RD and Gores GJ (2000). The bile acid taurochenodeoxycholate activates a phosphatidylinositol 3-kinase-dependent survival signaling cascade. J Biol Chem 275: 20210–20216. Ryo K, Kamogawa Y, Ikeda I, Yamauchi K, Yonehara S, Nagata S and Hayashi N (2000). Significance of Fas antigen-mediated apoptosis in human fulminant hepatic failure. Am J Gastroenterol 95: 2047–2055. Sackey FN, Hache RJ, Reich T, Kwast-Welfeld J and Lefebvre YA (1996). Determinants of subcellular distribution of the glucocorticoid receptor. Mol Endocrinol 10: 1191–1205. Savill J and Fadok V (2000). Corpse clearance defines the meaning of cell death. Nature 407: 784–788. Scaffidi C, Fulda S, Srinivasan A, Friesen C, Li F, Tomaselli KJ, Debatin KM, Krammer PH and Peter ME (1998). Two CD95 (APO-1/Fas) signaling pathways. EMBO J 17: 1675–1687. Schmucker DL, Ohta M, Kanai S, Sato Y and Kitani K (1990). Hepatic injury induced by bile salts: correlation between biochemical and morphological events. Hepatology 12: 1216–1221. Schoemaker MH and Moshage H (2004). Defying death: the hepatocyte’s survival kit. Clin Sci (London) 107: 13–25. Schoemaker MH, Conde de la Rosa L, Buist-Homan M, Vrenken TE, Havinga R, Poelstra K, Haisma HJ, Jansen PL and Moshage H (2004). Tauroursodeoxycholic acid protects rat hepatocytes from bile acid-induced apoptosis via activation of survival pathways. Hepatology 39: 1563– 1573. Schuchmann M and Galle PR (2002). Dead or alive – NF-κB, the guardian which tips the balance. J Hepatol 36: 827–828. Schuetz EG, Strom S, Yasuda K, Lecureur V, Assem M, Brimer C, Lamba J, Kim RB, Ramachandran V, Komoroski BJ, Venkataramanan R, Cai H, Sinal CJ, Gonzalez FJ and Schuetz JD (2001). Disrupted bile acid homeostasis reveals an unexpected interaction among nuclear hormone receptors, transporters and cytochrome P450. J Biol Chem 276: 39411–39418. Serfaty L, De Leusse A, Rosmorduc O, Desaint B, Flejou JF, Chazouilleres O, Poupon RE and Poupon R (2003). Ursodeoxycholic acid therapy and the risk of colorectal adenoma in patients with primary biliary cirrhosis: an observational study. Hepatology 38: 203–209. Setchell KD, Rodrigues CMP, Clerici C, Solinas A, Morelli A, Gartung C and Boyer J (1997). Bile acid concentrations in human and rat liver tissue and in hepatocyte nuclei. Gastroenterology 112: 226–235. Sodeman T, Bronk SF, Roberts PJ, Miyoshi H and Gores GJ (2000). Bile salts mediate hepatocyte apoptosis by increasing cell surface trafficking of Fas. Am J Physiol Gastrointest Liver Physiol 278: G992–G999. Sokol RJ, Straka MS, Dahl R, Devereaux MW, Yerushalmi B, Gumpricht E, Elkins N and Everson G (2001). Role of oxidant stress in the permeability transition induced in rat hepatic mitochondria by hydrophobic bile acids. Pediatr Res 49: 519–531. Sol´a S, Ma X, Castro RE, Kren BT, Steer CJ and Rodrigues CMP (2003a). Ursodeoxycholic acid modulates E2F-1 and p53 expression through a caspase-independent mechanism in transforming growth factor β1-induced apoptosis of rat hepatocytes. J Biol Chem 278: 48831–48838. Sol´a S, Castro RE, Laires PA, Steer CJ and Rodrigues CMP (2003b). Tauroursodeoxycholic acid prevents amyloid-β peptide-induced neuronal death via a phosphatidylinositol 3-kinase-dependent signaling pathway. Mol Med 9: 226–234. Sol´a S, Castro RE, Kren BT, Steer CJ and Rodrigues CMP (2004). Modulation of nuclear steroid receptors by ursodeoxycholic acid inhibits TGF-β1-induced E2F-1/p53-mediated apoptosis of rat hepatocytes. Biochemistry 43: 8429–8438. Sol´a S, Amaral JD, Castro RE, Ramalho RM, Borralho PM, Kren BT, Tanaka H, Steer CJ and Rodrigues CMP (2005). Nuclear translocation of UDCA by the glucocorticoid receptor is required to reduce TGF-β1-induced apoptosis in rat hepatocytes. Hepatology 42: 925–934.
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Staudinger JL, Goodwin B, Jones SA, Hawkins-Brown D, MacKenzie KI, LaTour A, Liu Y, Klaassen CD, Brown KK, Reinhard J, Willson TM, Koller BH and Kliewer SA (2001). The nuclear receptor PXR is a lithocholic acid sensor that protects against liver toxicity. Proc Natl Acad Sci USA 98: 3369–3374. Strand S, Hofmann WJ, Hug H, Muller M, Otto G, Strand D, Mariani SM, Stremmel W, Krammer PH and Galle PR (1996). Lymphocyte apoptosis induced by CD95 (APO-1/Fas) ligand-expressing tumor cells – a mechanism of immune evasion? Nat Med 2: 1361–1366. Streetz K, Leifeld L, Grundmann D, Ramakers J, Eckert K, Spengler U, Brenner D, Manns M and Trautwein C (2000). Tumor necrosis factor alpha in the pathogenesis of human and murine fulminant hepatic failure. Gastroenterology 119: 446–460. Tanaka H and Makino I (1992). Ursodeoxycholic acid-dependent activation of the glucocorticoid receptor. Biochem Biophys Res Commun 188: 942–948. Tanaka H, Makino Y, Miura T, Hirano F, Okamoto K, Komura K, Sato Y and Makino I (1996). Ligand-independent activation of the glucocorticoid receptor by ursodeoxycholic acid. Repression of IFN-gamma-induced MHC class II gene expression via a glucocorticoid receptor-dependent pathway. J Immunol 156: 1601–1608. Tang G, Minemoto Y, Dibling B, Purcell NH, Li Z, Karin M and Lin A (2001). Inhibition of JNK activation through NF-κB target genes. Nature 414: 313–317. Tirosh O, Aronis A and Melendez JA (2003). Mitochondrial state 3 to 4 respiration transition during Fas-mediated apoptosis controls cellular redox balance and rate of cell death. Biochem Pharmacol 66: 1331–1334. Trauner M, Meier PJ and Boyer JL (1999). Molecular regulation of hepatocellular transport systems in cholestasis. J Hepatol 31: 165–178. Tung BY, Emond MJ, Haggitt RC, Bronner MP, Kimmey MB, Kowdley KV and Brentnall TA (2001). Ursodiol use is associated with lower prevalence of colonic neoplasia in patients with ulcerative colitis and primary sclerosing cholangitis. Ann Intern Med 134: 89–95. Uehara T, Bennett B, Sakata ST, Satoh Y, Bilter GK, Westwick JK and Brenner DA (2005). JNK mediates hepatic ischemia reperfusion injury. J Hepatol 42: 850–859. Vander Heiden MG and Thompson CB (1999). Bcl-2 proteins: regulators of apoptosis or of mitochondrial homeostasis?. Nat Cell Biol 1: E209–E216. Ved R, Saha S, Westlund B, Perier C, Burnam L, Sluder A, Hoener M, Rodrigues CMP, Alfonso A, Steer CJ, Liu L, Przedborski S and Wolozin B (2005). Similar patterns of mitochondrial vulnerability and rescue induced by genetic modification of α-synuclein, parkin and DJ-1 in Caenorhabditis elegans. J Biol Chem 280: 42655–42668. Wachs FP, Krieg RC, Rodrigues CMP, Messmann H, Kullmann F, Knuchel-Clarke R, Scholmerich J, Rogler G and Schlottmann K (2005). Bile salt-induced apoptosis in human colon cancer cell lines involves the mitochondrial transmembrane potential but not the CD95 (Fas/Apo-1) receptor. Int J Colorectal Dis 20: 103–113. Wali RK, Khare S, Tretiakova M, Cohen G, Nguyen L, Hart J, Wang J, Wen M, Ramaswamy A, Joseph L, Sitrin M, Brasitus T and Bissonnette M (2002). Ursodeoxycholic acid and F(6)-D(3) inhibit aberrant crypt proliferation in the rat azoxymethane model of colon cancer: roles of cyclin D1 and E-cadherin. Cancer Epidemiol Biomarkers Prev 11: 1653–1662. Wang CY, Mayo MW, Korneluk RG, Goeddel DV and Baldwin AS Jr (1998). NF-κB antiapoptosis: induction of TRAF1 and TRAF2 and c-IAP1 and c-IAP2 to suppress caspase-8 activation. Science 281: 1680–1683. Wang H, Chen J, Hollister K, Sowers LC and Forman BM (1999). Endogenous bile acids are ligands for the nuclear receptor FXR/BAR. Mol Cell 3: 543–553. Whitfield GK, Jurutka PW, Haussler CA and Haussler MR (1999). Steroid hormone receptors: evolution, ligands, and molecular basis of biologic function. J Cell Biochem 32–33(Suppl): 110– 122.
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Xie W, Radominska-Pandya A, Shi Y, Simon CM, Nelson MC, Ong ES, Waxman DJ and Evans RM (2001). An essential role for nuclear receptors SXR/PXR in detoxification of cholestatic bile acids. Proc Natl Acad Sci USA 98: 3375–3380. Xie Q, Khaoustov VI, Chung CC, Sohn J, Krishnan B, Lewis DE and Yoffe B (2002). Effect of tauroursodeoxycholic acid on endoplasmic reticulum stress-induced caspase-12 activation. Hepatology 36: 592–601. Xu Y, Jones BE, Neufeld DS and Czaja MJ (1998). Glutathione modulates rat and mouse hepatocyte sensitivity to tumor necrosis factor toxicity. Gastroenterology 115: 1229–1237. Yamamoto M, Fukuda K, Miura N, Suzuki R, Kido T and Komatsu Y (1998). Inhibition by dexamethasone of transforming growth factor beta1-induced apoptosis in rat hepatoma cells: a possible association with Bcl-xL induction. Hepatology 27: 959–966. Yoon JH and Gores GJ (2002). Death receptor-mediated apoptosis and the liver. J Hepatol 37: 400– 410. Zheng SJ, Wang P, Tsabary G and Chen YH (2004). Critical roles of TRAIL in hepatic cell death and hepatic inflammation. J Clin Invest 113: 58–64.
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16 Drug-Induced Intrahepatic Cholestasis by Interaction with the Hepatic Bile Salt Export Pump (BSEP) Christoph Funk, Johannes No´e, Ren´ee Portmann, Rub´en Alvarez-S´anchez, Florian Klammers, Christiane Lamy, Axel Paehler and Michael Pantze
16.1
Introduction
Drug-induced liver disease is one of the major challenges for drug development and drug safety. Many drugs and herbal remedies have been implicated in causing hepatic injuries, which can be divided into a group comprising predictable, dose- and time-dependent type I lesions occurring in most subjects exposed to the causative substance and which are usually readily reproducible in animals. On the other hand the group of type II lesions are considered ‘unpredictable’, dose- and time-independent, occurring only occasionally in patients and are not reproducible in animals (Kaplowitz, 2005). It is the latter type of liver injury which becomes overt only late in drug development when many individuals have been exposed. Often an interplay between different mechanisms in connection with polymorphic expression and induction/inhibition of key enzymes/transporters lead to such multifactorial toxicities. Nowadays, several of the individual mechanisms potentially contributing to such toxicities can be studied. The drug–drug interaction potential on the level of active (drug) transporters (summarized in this review) and the formation of reactive intermediates associated with covalent binding and immune-mediated responses are two major processes
Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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being investigated lately in more detail in this context (Uetrecht, 1999; Alvarez-S´anchez et al., 2006). Drug–drug interactions are widely described for drug metabolizing enzymes, such as cytochrome P450 monooxygenases and glucuronosyl transferases and they are increasingly recognized to occur also at the level of active drug transport, both the cellular uptake carriers and cellular exporters (Zhang et al., 2005). The interaction might be among two co-medicated drugs or with endogenous products of metabolism competing for the same transport/export process. Such an interference with the major hepatobiliary transport functions might pose an important contribution to the formation of intrahepatic cholestasis (Pauli-Magnus and Meier, 2005). In addition, several mutations leading to dysfunctional BSEP protein have been reported to pose a predisposition for cholestasis (No´e et al., 2005) and these patients are likely to exhibit a higher sensitivity towards cholestatic drugs. 16.1.1
Intrahepatic Cholestasis
Interference with the vectorial transport of biliary constituents from blood to bile can result in an accumulation of bile salts in the hepatocytes, leading to intrahepatic cholestasis (Trauner et al., 1998; Kullak-Ublick and Meier, 2000). High intracellular bile salt levels have been reported to induce cellular necrosis and mitochondrial dysfunction due to their intrinsic detergent activity (Gores et al., 1998). The more lipophilic bile acids, deoxycholate and chenodeoxycholate exert higher mitochondrial in vitro toxicity as compared to cholate and dehydrocholate (Figure 16.1). These hydrophobic bile acids are known to induce intrahepatic cholestasis in rats. The vectorial transport of both endobiotics (bile acids, steroids, bilirubin glucuronide and other metabolic products) and xenobiotics, along with their metabolites from plasma to bile, is facilitated by several transport systems (Zimniak et al., 1999). The export across the canalicular membrane, where the greatest uphill concentration gradient has to be overcome, often represents the rate-limiting step in this excretion process (Kadmon et al., 1993; Arrese et al., 1998). For bile acids, this step is catalyzed by a primary active, ATP-dependent transporter of the ABC protein family, the canalicular bile salt export pump (Abcb11/Bsep), localized in the canalicular liver plasma membrane (Gerloff et al., 1997). The cloning of the human bile salt export pump (BSEP) has greatly facilitated the studies of transport and inhibition properties of this ABC transporter in corresponding in vitro systems (Byrne et al., 2002; No´e et al., 2002). 16.1.2
Drug-Induced Intrahepatic Cholestasis
For several cholestatic compounds, interactions with the export of bile acids were found at the level of the canalicular ATP-dependent Bsep/BSEP. The immune suppressive agent cyclosporin A was observed to inhibit the canalicular Bsep in vitro (Kadmon et al., 1993), accompanied by an increase in liver taurocholate levels in the rat (Boehme et al., 1994). A similar mechanism was postulated for lovastatin (Spreckelsen et al., 1991), the NSAID sulindac (Bolder et al., 1999), for rifamycin, rifampicin (Stieger et al., 2000), glibenclamide (Stieger et al., 2000; Funk et al., 2001a), bosentan (Fattinger et al., 2001) and for fusidate (Bode et al., 2002). In addition, for troglitazone (Funk et al., 2001a,b), nefazodone
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140 120 MTT (% of control)
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1000
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(b)
120 TPP (% of control)
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100 80 60
Cholate Dehydrocholate
40
Deoxycholate 20
Chenodeoxycholate
0 0.1
1
10
100
1000
Concentration (µM)
Figure 16.1 Cytotoxic effects of major bile acids the cholate, dehydrocholate, deoxycholate and chenodeoxycholate, assessed by the effects on mitochondrial toxicity markers: (a) the reduction of MTT (methylthiazocyediphenyl-tettazolium-bromid) and (b) the retention of TPP (tetra phenylphosphonium bromide).
(Kostrubsky et al., 2006), ritonavir, saquinavir and efavirenz (McRae et al., 2006), inhibition of Bsep/BSEP was proposed as the mechanism leading to intrahepatic cholestasis and thereby contributing to liver toxicity. The observed in vivo effects, typical dose ranges and in vitro Bsep/BSEP inhibition potentials are summarized for a number of cholestatic compounds in Table 16.1. The cholestatic potential in vivo is not only dependent on the affinity towards the Bsep/BSEP transporter but depends on numerous additional factors, such as dose, liver tissue exposure, metabolization, co-medications and genetic predisposition.
2.2b 5.7
1500 10 400–600 200–400 100-500
Rifampicin Nefazodone
Bosentan
Cholestasis Jaundice, bile duct proliferation, cholestasis 18 % liver enzyme increase in patients
b
Determined by competitive inhibition of bile acid transport in rat liver plasma membrane vesicles. In addition to Bsep, fusidate also inhibits Mrp2 (IC50 = 5.5−7.6 μM). c IC50 value determined at one taurocholate concentration using Sf9 vesicles expressing BSEP. d Inhibition measured in sf9 vesicles expressing Bsep.
12d
6.4 9c
Fattinger et al., 2001
Bode et al., 2002 Krivoy et al., 1996; Stieger et al., 2000 Stieger et al., 2000 Kostrubsky et al., 2006
Funk et al., 2001a,b
Kadmon et al., 1993; Boehme et al., 1994
Reference
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a
1.3 (0.23)
400–600
Troglitazone (TGZ sulfate) Fusidate Glibenclamide
Multiple drug interactions, interference, with hepatic bile salt elimination Liver injury, cholestatic DDI potential with glibenclamide Hyperbilirubinemia Intrahepatic cholestasis
Clinical effect described
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0.2
50–200
Cyclosporin A
Bsep inhibition potential K I (μM)a
Typical dose range (mg per day)
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Table 16.1 Compounds for which a cholestatic/liver toxic effect was recently described as an interference with hepatic bile salt elimination by an inhibition of the Bsep/BSEP
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425
Cholestatic Potential of Selected Drugs Cyclosporin A
Systemic treatment with cyclosporin A (CsA) can lead to an interference with the hepatic elimination of bile acids and finally cholestasis, a common side effect of CsA therapy (Kadmon et al., 1993). The cholestatic effect could be attributed to an inhibition of the canalicular bile salt exporter Bsep by in vitro inhibition of the ATP-dependent taurocholate transport into liver plasma membrane vesicles (Boehme et al., 1994). The bile acid transporter was inhibited with an IC50 value of about 0.8 μM (Figure 16.2). The cholestatic effect could also be induced (Boehme et al., 1994) in vivo in rats. CsA elicited a very rapid decrease of bile secretion, with a maximum at 10–15 min after intravenous administration (Roman et al., 1989). The observed inhibition was transient, with almost complete recovery within 40 to 50 min post-administration (pa). The concomitant increase of radioactivity in the liver after injection of a tracer amount of radio-labeled taurocholic acid (Boehme et al., 1994) measured at 30 min after CsA administration gave additional evidence for an interaction at the bile canalicular membrane and a resulting intrahepatic accumulation of bile acids. The acute cholestatic effect of CsA in rats was used to establish an in vivo model to assess the cholestatic potential of test compounds under in vivo conditions (Figure 16.3) (Fattinger et al., 2001; Funk et al., 2001b). 16.2.2
Glibenclamide and other Sulfonylurea Antidiabetics
Glibenclamide has been reported to induce intrahepatic cholestasis in a case of a fatal hepatotoxicity (Krivoy et al., 1996). In a number of cases, glibenclamide was co-medicated with
ATP-dependent taurocholate transport (%)
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80
Cyclosporin A, IC50 = 0.8 ± 0.23 µM
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40
Glibenclamide, IC50 = 8.6 ± 1.9 µM
Troglitazone sulfate, IC50 = 0.4 ± 0.06 µM
20
0 0.01
0.1
1
10
100
Inhibitor concentration (µM)
Figure 16.2 In vitro inhibition effects of troglitazone, troglitazone sulfate and the two cholestatic model compounds, cyclosporin A and glibenclamide, on the ATP-dependent bile acid transport assessed by using cLPMV.
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Hepatotoxicity 180 160 Absolute change in plasma bile acids, 10 min, pa (µM)
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140 [ ] Cyclosporin A
120 100 80
[ ] Troglitazone
60 40 20
[ ] Glibenclamide
0 –20 0
10
20 30 Drug (mg/kg iv dose)
40
50
Figure 16.3 Acute in vivo effects of troglitazone and two cholestatic model compounds on the plasma bile acid levels in rats. The changes in total bile acid levels (difference pre-dose and 10 min. post-dose) are shown as the mean of three to four rats per concentration.
other cholestatic drugs, such as bosentan (Fattinger et al., 2001) or troglitazone (Shibuya et al., 1998; Funk et al., 2001b), increasing the incidence for developing intrahepatic cholestasis. Glibenclamide was shown in vitro to inhibit both the rat and human bile salt transporters (Bsep, BSEP) with similar potencies (IC50 ∼ 6 μM; Figure 16.2 (Stieger et al., 2000)). The cholestatic effect of glibenclamide was also observed in an acute rat model for cholestatic compounds (Figure 16.3) and by co-administration of both compounds, glibenclamide and bosentan, the additive cholestatic effect could be reproduced in this in vivo model (Fattinger et al., 2001). Similar to glibenclamide, other sulfonylurea antidiabetic compounds could also be associated with a small increased risk for liver injury. Due to a low overall number of cases detected in the patient population evaluated, the risk of individual sulfonylurea compounds could, however, not be estimated conclusively (Huerta et al., 2002). When comparing these clinical data with the in vitro Bsep inhibition potential determined using rat liver plasma membrane vesicles, it is evident that two compounds, tolbutamide and chlorpropamide, do not inhibit Bsep and did not increase the in vivo incidence for liver injury in spite of the high dose used (Table 16.2). On the other hand, the two compounds inhibiting Bsep in vitro, glipizide and glibenclamide/glyburide, were associated with an increase of the observed cases of liver injury (Table 16.2). 16.2.3
Cholestatic Potential of Bosentan
Bosentan caused dose-dependent and reversible liver injury in 2 to 18 % of patients, linked with a significant increase of serum bile salt levels (Fattinger et al., 2001). Concomitant administration of glibenclamide enhanced the cholestatic potency of bosentan. Similar
N
O
S
O
H N
S
N H
O
N H
O
O
H N
O
S
O
N H
O
N H
>500
>500
1.4
0
0
>500
80–320
500
500–1000
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Cl
O
Typical dose range (mg per day)
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Tolbutamide
Chemical structure
Liver injuries (cases per 100 000 prescriptionsa )
Bsep inhibition potential IC50 (μM)
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Table 16.2 Cholestatic effects of several sulfonylurea compounds. Cases of in vivo liver injury (as number of liver injury cases per 100 000 prescriptions) and in vitro inhibition potential on Bsep using taurocholate transport interference in liver plasma membrane vesicles
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O
H N
O
Data on incidence of liver injury in diabetic patients (Huerta et al., 2002).
O
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H N
S O O
H N
H N
2.7 ± 0.4
115 ± 20
3.4
4.3
10
5–60
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N
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cholestatic effects were seen in rats, serum bile acid levels were increased by both, glibenclamide and bosentan, with an additive effect if both compounds were co-administered. In vitro, bosentan inhibited the Bsep-mediated taurocholate transport (K I , ∼ 12 μM; Fattinger et al., 2001). The three major metabolites of bosentan showed a Bsep inhibition similar to bosentan, suggesting that they contribute to the cholestatic effect in vivo (Fattinger et al., 2001). The in vitro and in vivo cholestatic potential measured in rats supported the relevance of this mechanism for the liver injury observed in man. 16.2.4
Cholestasis in Troglitazone-Induced Hepatotoxicity
Troglitazone is an insulin sensitizer of the thiazolidinedione class, compounds used for the treatment of type 2 non-insulin-dependent diabetes mellitus (Chen, 1998). Elevations in liver enzyme levels observed during clinical trials and several cases of fulminant hepatic failure were reported, leading to the withdrawal of this compound from the market (Gitlin et al., 1998; Shibuya et al., 1998; Herrine and Choudhary, 1999). Different hypotheses towards the mechanism(s) underlying the troglitazone-associated hepatotoxicity have been published. Several reports suggested a cholestatic mechanism to be involved in this liver injury (Gitlin et al., 1998) and a strong reduction of the bile flow has been observed in isolated perfused rat liver (Preininger et al., 1999). Several cases of liver injury following concurrent troglitazone and glibenclamide treatment lead to the speculation that this latter drug, known to induce cholestasis in some patients (Krivoy et al., 1996), might enhance the probability for troglitazone-induced liver injury (Shibuya et al., 1998; Fukano et al., 2000). This cholestatic potential of troglitazone is further supported by in vitro and in vivo mechanistic studies (Funk et al., 2001b). Alternatively, reactive metabolite formation, associated with covalent protein binding (haptenization) and induction of an immune-mediated idiosyncratic response, has also been shown to occur for troglitazone (Kassahun et al., 2001; Tettey et al., 2001; Alvarez-S´anchez et al., 2006). Cellular toxicity studies pointed towards a direct toxicity of troglitazone or troglitazone sulfate, while the metabolization by cytochrome P450 3A4 (which is also responsible for the formation of the reactive intermediate metabolites) rather represents a detoxification pathway (Hewitt et al., 2002; Masubuchi et al., 2006). In other studies, the conjugated metabolites troglitazone sulfate and glucuronide showed a lower potential for direct cellular toxicity (Kostrubsky et al., 2000; Honma et al., 2002). A combination of multiple mechanisms, namely cholestasis leading to increased drug levels in hepatocytes, mitochondrial toxicity induced by troglitazone and/or bile acids, reactive metabolite formation and covalent protein binding associated with antigen presentation and stimulation of an immune-response might be triggered by troglitazone. This could fit with the proposal of haptenization-associated low-grade toxicity (danger signal) needed for a significant immune reaction (Uetrecht, 1999; Kaplowitz, 2005). The different mechanisms of troglitazone hepatotoxicity have been recently reviewed (Smith, 2003). 16.2.4.1
The cholestatic Potential of Troglitazone
Troglitazone and to a greater extent its main metabolite, troglitazone sulfate, inhibited the canalicular Bsep (Figure 16.2) in a competitive manner with apparent K I values of 1.3 and 0.23 μM, respectively (Table 16.1; Funk et al., 2001b). The higher inhibition potential
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of troglitazone sulfate in these rat liver membrane vesicles indicated the importance of this metabolite in contributing to the overall cholestatic potential of troglitazone. This was supported by studies in the isolated perfused rat liver (IPRL), where troglitazone triggered a strong decline in the bile flow in livers from male rats (Preininger et al., 1999). In this model, troglitazone was completely cleared from the perfusate, metabolized in the liver mainly to troglitazone sulfate and glucuronide and eliminated to a great extent into bile (Funk et al., 2001b). Troglitazone elicited a rapid decrease of both bile flow and biliary excretion of a radio-labeled taurocholic acid tracer added to the perfusion medium, while more of the labeled bile acid was recovered from the perfusion medium (Figure 16.4). In equilibration 10 µM troglitazone --------------------------------- ++++++++++++++++++++++++++++++++++ 110 male rat liver 100
(a)
120
Taurocholate excreted (%)
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equilibration 10 µM troglitazone --------------------------------- ++++++++++++++++++++++++++++++++++ female rat liver Excretion into perfusate 100 120
Taurocholate excreted (%)
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Time (min)
Figure 16.4 Cholestatic effect of troglitazone in the isolated perfused liver from (a) male and (b) female rats. Interference with the biliary export of a radio-labeled taurocholate tracer (3 nM) is shown as the mean of four-individual liver perfusions.
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female rat livers, the effect was milder and delayed as compared to the livers from male rats. This gender difference could be attributed to quantitative differences in the metabolism of troglitazone, mainly to a rapid formation of troglitazone sulfate in male liver due to the presence of a high affinity male-specific sulfotransferase (Funk et al., 2001a). In vivo, in rats, troglitazone similarly interfered with the hepatic elimination of bile acids and induced a transient increase in plasma bile acid levels (Figure 16.3.), while an accumulation of an intravenously administered radio-labeled taurocholate tracer was observed in liver tissue (Funk et al., 2001b). These results support the cholestatic mechanism of troglitazone and its metabolite troglitazonesulfate at the canalicular pole of the hepatocytes, where Bsep is localized, as the site of interference with the hepatobiliary excretion of bile acids. 16.2.4.2
Interrelation of Drug Metabolism and Hepatobiliary Export Processes
Several lines of evidence suggested that the hepatobiliary export of troglitazone and related metabolites might represent a rate-limiting step in the overall elimination of troglitazone. In patients with hepatic impairment, troglitazone sulfate was found to accumulate about 4-fold in plasma, with a 3-fold increased half-life (Ott et al., 1998). In rats, troglitazone sulfate accumulated as the major drug-related metabolite in liver tissue, reaching high intracellular concentrations (Funk et al., 2001b). The higher cholestatic potential of troglitazone in isolated perfused male rat livers as compared to the livers from female rats (Figure 16.4) can be mainly attributed to differences in metabolism rather than differences in bile excretion or in Bsep inhibition. Liver plasma membrane vesicles isolated from either male or female rat liver tissue exhibited a very similar capacity for ATP-dependent taurocholate transport, based on the intrinsic clearance calculated from the respective apparent K M and Vmax parameters (Table 16.3 (a)). In addition, the potential of troglitazone (Table 16.3(b)) and troglitazone sulfate (Table 16.3(c)) to inhibit this ATP-dependent transport was very similar among the two genders. The conversion of troglitazone to troglitazone sulfate was studied using liver cytosolic fractions prepared from male and female rat liver tissue. The respective kinetic evaluation indicated the presence of a high-affinity sulfotransferase in male rat liver in addition to a low-affinity isoenzyme similarly present in both genders (Funk et al., 2001a). The calculated intrinsic clearance values are more than 5-fold higher in male rats (Table 16.3(d)) and consistently explain the higher levels of the troglitazone-sulfate metabolite in male rats (Table 16.3(e)). These higher liver tissue levels, together with the 5-fold-lower IC50 value for Bsep of troglitazone sulfate, can explain the higher sensitivity of male animals towards troglitazone as observed in the in vivo acute rat cholestasis model. At a dose of 6.25 mg/kg, no effect was observed in female animals, while in male rats a marked increase in the total plasma bile acid level of more than 40 μM was observed (Table 16.3(f)). These in vitro and in vivo studies clearly underline the importance of both drug metabolism and drug export processes in interference of a drug molecule with the hepatic bile formation. 16.2.4.3
Reactive Troglitazone Metabolite Formation as an Additional Trigger for Hepatotoxicity
In addition to the formation of conjugated phase II metabolites, troglitazone has been shown to undergo bioactivation processes in rat and human liver microsomes, leading to the formation of reactive intermediates capable of covalent binding to protein thiol groups.
(f)
In vitro transport capacity of taurocholate into cLPMV In vitro inhibition of Bsep by TGZ in rat cLPMV In vitro inhibition of Bsep by TGZ-S in rat cLPMV Metabolism of TGZ by rat cytosolic sulfotransferases Liver tissue levels of TGZ-S (30 min after 6.25 mg/kg iv dose) In vivo cholestatic effect of intravenous TGZ, increase in plasma bile acid levels 10 min after an intravenous dose (6.25 mg/kg)
(a) (b) (c) (d) (e)
34.9
34 25 90 159 30
μl min−1 mg−1 % inhibition at 1 μM % inhibition at 1 μM μl min−1 · mg−1 nmol g−1 μM
Male
Unit
>−2.5
∼ 24.5 ∼ 20 ∼ 88 >35 > 8.2
Female
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Table 16.3 Gender differences in metabolism and cholestatic potential of troglitazone in rats. The metabolism of troglitazone (TGZ) to the main metabolite troglitazone sulfate (TGZ-S), liver tissue levels of TGZ-S and in vitro and in vivo cholestatic effects of TGZ and TGZ-S are shown. Data from Funk et al., 2001a,b
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R'
P450 P450, O
all TZDs
O
P450 P450, TGZ, RGZ
C * * N* *
S
* * * NH S * RO
O
R'
O
* S * XH
X=O, 15NH
Non TZD-related GSH conjugates
O
N R=
O
GS
OH
433
N
N
PGZ
RGZ
HO TGZ
H N R' =
N desmethyl-RGZ
Figure 16.5 Bioactivation of thiazolidinedione (TZD) derivatives troglitazone (TGZ), rosiglitazone (RGZ) and pioglitazone (PGZ). For TGZ, the non-TZD-related glutathione adduct appears to be the predominant adduct formed in human liver microsomes (∗ indicates the positions of 13 C or 15 N).
Glutathione adducts of these reactive intermediates have been characterized to elucidate the structure of the reactive intermediates formed during phase I metabolism of troglitazone predominantly catalyzed by P450 3A4 (Figure 16.5). P450-mediated S-oxidation is suggested to lead to an unstable thiazolidinedione (TZD)-sulfoxide which undergoes spontaneous ring opening to form a reactive sulfenic acid-α-keto-isocyanate (Kassahun et al., 2001). This TZD-dependent bioactivation was seen for all three TZD drugs in rat and human liver microsomes by a characteristic change of the isotopic difference between the non-labeled and stable isotope-labeled analogues of the TZD-drugs (Alvarez-S´anchez et al., 2006). In addition, troglitazone, rosiglitazone and pioglitazone are also prone to form reactive intermediates at higher substrate concentrations, suggesting that covalent binding to liver proteins may also be expected for these two compounds. In human liver microsomes, the major glutathione adduct derived from troglitazone showed, however, the conservation of all four labeled positions of the TZD moiety. This was the predominant adduct detected and appears to be more abundant than the GSH adducts derived from TZD ring scission. Its structure was consistent with the previously reported adduct derived from the quinone methide activated troglitazone metabolite (Kassahun et al., 2001; Alvarez-S´anchez et al., 2006). This observation suggests that TGZ bioactivation via TZD ring scission does not contribute significantly to the extent of GSH adduct formation. Although the relative contribution of less-abundant reactive intermediates to hepatotoxic events remains unknown,
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these findings suggest that the formation of these reactive intermediates not related to the TZD-moiety might be associated with the hepatotoxic potential of TGZ. Formation of quinone and quinone methide-type reactive intermediates has been linked to the prooxidant activity of TGZ in rat primary hepatocytes on cumene hydroperoxide-induced lipid peroxidation and cytotoxicity. This effect was significantly higher for TGZ as compared to other vitamin E analogues (Tafazoli et al., 2005). Cytotoxicity in hepatic cells and oxidative stress-inducing properties for thiazolidinedione derivatives were dependent on the presence of the 6-hydroxychromane moiety, suggesting a link to this specific molecular structure of TGZ. In addition, N1S1 rat hepatoma cells were more sensitive to TGZ and TGZ quinone than to other TZD derivatives lacking the 6-hydroxychromane moiety (Narayanan et al., 2003). Reactive metabolite formation and covalent binding to proteins might represent one mechanism contributing to the hepatic toxicity observed with troglitazone in some patients, in which drug-induced intrahepatic cholestasis might increase the individual susceptibility.
16.2.4.4
Active Hepatobiliary Export of Troglitazone Sulfate
Only recently, the human canalicular bile salt export pump, BSEP, has been cloned and therefore has enabled more mechanistic in vitro studies (Byrne et al., 2002; No´e et al., 2002). For the rat Bsep, the available data indicate that this transporter is involved in the excretion of taurocholate and related non-glucuronidated (except acyl glucuronide) nonsulfated bile salts (Keppler et al., 1992). It is mainly responsible for the bile salt-dependent portion of the overall bile flow. An involvement of Bsep in the transport of xenobiotics was suggested for non-conjugated organic anions (Hofmann, 1992). Based on its expression pattern, a broader substrate specificity of Bsep has been postulated in analogy to other multidrug resistance proteins (T¨or¨ok et al., 1999). Recently, pravastatin was reported as being a substrate for both the rat Bsep and the human BSEP transport proteins (Hirano et al., 2005). In addition, for troglitazone and troglitazone sulfate we have studied the ATP-dependent transport properties using liver plasma membrane vesicles from Sprague–Dawley rats (wt) and Mrp2-deficient rats (TR−), respectively (Figure 16.6). The model substrates, taurocholate and LTC4 , for the two ABC transporters, Bsep and Mrp2, were transported as expected, the former one in both vesicle preparations, while the latter one was transported only in wt vesicles. The radio-labeled troglitazone did not show any detectable ATP-dependent transport in this experimental in vitro system (data not shown), in spite of its Bsep inhibition potential. However, troglitazone sulfate showed ATP-dependent transport in both vesicle preparations, although considerable unspecific binding was seen, indicating the involvement of Bsep in the hepatobiliary transport of this compound. This was further verified using membrane vesicles recombinantly expressing either one of the two human canalicular ABC transporters, MRP2 or BSEP. Both transporters were able to mediate an ATP-dependent uptake of troglitazone sulfate into the respective vesicles (Figure 16.7). The functional activity of both transporters was verified by transport of the corresponding model substrate, LTC4 or taurocholate, respectively. Biliary excretion of troglitazone sulfate has also been described as one step of an enterohepatic circulation of both troglitazone sulfate and glucuronide metabolites (Kawai et al., 2000).
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+ ATP
Taurocholate
45 40 35 30 25 15 0.6
LTC4
–1
20
–1
wt
TR–
wt
TR–
0.4
0.2
0.0
Troglitazone sulfate
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Figure 16.6 Hepatobiliary transport of troglitazonesulfate using rat liver plasma membrane vesicles (cLPMV) prepared from Sprague–Dawley rats (wt) and Mrp2-deficient rats (TR− ), respectively. Taurocholate and LTC4 were used as model substrates testing Bsep and Mrp2 functional activities.
16.2.4.5
Dose and Co-mediations as Important Factors Contributing to Troglitazone-Induced Cholestasis
Typical daily troglitazone doses in man are in the range of 400 to 600 mg, producing troglitazone plasma concentrations of 1 to 2 μg/ml (Table 16.1; Loi et al., 1999). A quantitative extrapolation of the cholestatic effect observed in rats to man is not possible, due to species-specific differences in the process of bile formation and in the overall disposition of troglitazone. However, the processes involved in the hepatobiliary metabolism and export of the absorbed fraction of troglitazone represent potential targets for drug interactions and regulative changes. Therefore, troglitazone might induce intrahepatic cholestasis in combination with other cholestatic drugs, other diseases or pharmacogenetic liabilities, thereby contributing to the hepatotoxicity observed in some patients treated with troglitazone. Simvastatin (Herrine and Choudhary, 1999) and lisinopril (Gitlin et al., 1998), both drugs known to induce cholestatic side effects, were co-medicated with troglitazone in patients developing signs of liver toxicity. Another report described three out of four cases of
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MRP2
BSEP
Membrane vesicles used Figure 16.7 ATP-dependent transport of troglitazone sulfate into vesicles prepared from insect cells expressing either MRP2 or BSEP, or mock transfected as control, respectively. ATPdependent transport is shown as the mean of three independent experiments.
troglitazone-induced fulminant hepatitis where glibenclamide has been co-medicated with troglitazone. An interaction of the two drugs was suspected without further elaborating on a possible mechanism (Shibuya et al., 1998). Co-medication of potentially cholestatic drugs with troglitazone might increase the incidence in patients to develop liver toxicity. 16.2.5
Cholestatic Potential of Nefazodone
The antidepressant nefazodone (serzone) was withdrawn from the market due to drugmediated hepatic injury. Based on the clinical symptoms of the idiosyncratic hepatotoxicity, including jaundice, ALT and AST increase, high biliary elimination of nefazodone seen in animals and the relative high therapeutic doses of nefazodone in the range 200–400 mg/day, the potential for a cholestatic mechanism was studied in more details (Kostrubsky et al., 2006). In vitro, nefazodone caused a strong inhibition of the human bile salt transporter BSEP with an IC50 value of 9 μM, using membrane vesicles expressing BSEP. Nefazodone was furthermore found to interfere with the bile acid elimination in sandwich-cultured human hepatocytes. In addition, cytotoxicity was observed in human hepatocytes at nefazodone concentrations in the range of 10 μM. By inhibition of the metabolism, this could be associated with the concentrations of intact nefazodone rather than its metabolites. In rats, a transient increase in the serum bile acid level was observed at 1 h after oral drug administration. Based on these data, it is likely that the clinical hepatotoxicity of nefazodone is linked to the ability of the drug to interfere with the bile acid transport, primarily by an inhibition
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of the canalicular bile acid export protein BSEP. Interestingly, also for nefazodone the formation of reactive metabolites and covalent binding to liver proteins, including P450 3A4, have been suggested as a factor contributing to the hepatotoxicity of this compound (Kalgutkar et al., 2005). 16.2.6
Estrogen and Progesterone Steroid Hormones
Intrahepatic cholestasis of pregnancy (ICP) appears as transient cholestasis in pregnant women. The mechanism underlying this cholestasis has been investigated in detail (Vallejo et al., 2005). Certain progesterone metabolites, such as 5α-pregnan-3α-ol-20-one (PM4), are elevated in serum of women with ICP. PM4 is conjugated with sulfate in the liver and then excreted into bile and this PM4–sulfate metabolite was found to have a strong transinhibition effect on BSEP in an experimental system where BSEP was expressed in oocytes preloaded with bile acids. This trans-inhibition of BSEP by the PM4–sulfate metabolite is likely involved in the formation of ICP and a genetic deficiency in BSEP might be a contributing factor (Pauli-Magnus and Meier, 2005). Natural and synthetic estrogenic compounds and their metabolites are well known to interfere with the hepatic bile formation (Bossard et al., 1993). For 17α-ethinylestradiol, a decreased bile excretion capacity, due to multiple transporter defects in the canalicular membrane, has been shown. After a five-day treatment with 17α-ethinylestradiol rats showed a marked increase in liver weight, plasma bile acid levels (over 10-fold) and alkaline phosphatase (Bossard et al., 1993). In parallel, a decrease of both the ATP-dependent taurocholate transport and dinitrophenyl glutathione transport ratios were found in the canalicular membranes, indicating a down-regulation of Bsep and Mrp2. For a physiological estrogen metabolite, estradiol-17β-glucuronide (E2 17G), which can induce cholestasis at higher concentrations in rats, a dose-dependent inhibition of the ATPdependent taurocholate transport has been observed using rat liver membrane vesicles (Stieger et al., 2000). The effect was not seen in Bsep-expressing membrane vesicles, indicating a trans-inhibition of Bsep after export by Mrp2, which is only present in the vesicles, prepared from rat liver tissue. This in vitro finding was consistent with the fact that E2 17G induced a cholestasis in normal rats but not in TR− rats deficient of the canalicular Mrp2 exporter (Stieger et al., 2000). In conclusion, for several estrogenic compounds and progesterones, a trans-inhibition of Bsep/BSEP by a (conjugated) metabolite after being excreted into bile has been shown. This mechanism is likely to be involved in the induction of intrahepatic cholestasis of these steroid hormones by the accumulation of bile acids within the hepatocyte. 16.2.7
HIV Protease Inhibitors
Hepatotoxicity as a significant adverse effect in antiretroviral therapy of HIV-infected patients often forces withdrawal of the respective therapy (McRae et al., 2006). The in vitro cholestatic potential was studied for four compounds, ritonavir, saquinavir, nevirapine and efavirenz (McRae et al., 2006). Both the interference with hepatic bile acid uptake and biliary excretion were addressed in different experimental systems. The NTCP- and OATPmediated bile acid uptake into hepatocytes was studied in sandwich-cultured rat and human
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hepatocytes, while the BSEP-mediated biliary excretion of bile acids was studied using BSEP-expressing membrane vesicles. Ritonavir and saquinavir inhibited both NTCP- and OATP-mediated bile acid uptake and the BSEP-mediated biliary excretion of bile acids. Efavirenz only modestly inhibited the two processes, while nevirapine did not alter bile acid transport in any of the experimental systems. This rank order was not in agreement with the reported clinical hepatotoxic potential. Ritonavir-associated hepatotoxicity has been reported in up to 9 % of patients while hepatic toxicity is not generally associated with saquinavir or efavirenz, whereas nevirapine is commonly associated with the development of cholestatic and hepatocellular hepatotoxicity (McRae et al., 2006). The data suggest that inhibition of BSEP by the selected antiretroviral drugs is unlikely to be cause of antiretroviral-associated hepatotoxicity. The BSEP inhibition observed for some compounds appeared to be countered by an inhibition of the bile acid uptake transport systems, NTCP and OATP, potentially reducing the liver bile acid level. To the probably multifactorial hepatotoxicity different/other mechanism(s), such as interplay with metabolism, co-medications, genetic predisposition to name just a few, might contribute. 16.2.8
Fusidate
The steroid antibiotic fusidate has been shown to induce conjugated hyperbilirubinemia as early as two days after initiating the treatment in up to 48 % of the patients given the compound intravenously (Bode et al., 2002). The molecular mechanism leading to this conjugated hyperbilirubinemia was investigated in rats, in vitro using liver membrane vesicles and in vivo by investigating the effects on the bile flow and biliary elimination of different tracers (Bode et al., 2002). Fusidate inhibited both the ATP-dependent transport of the Mrp2 substrates estradiol17β-glucuronide and leukotriene C4 with K I values of 7.6 and 5.5 μM, as well as the ATP-dependent transport of the Bsep substrate cholyltaurine with a K I value of 2.2 μM. In vivo, in rats, fusidate reduced the bile flow and the excretion of substrates of both transporters (estradiol-17β-glucuronide and cholyltaurine), infused as tracers by 75 and 80 %, respectively. Based on these in vitro and in vivo data, fusidate competitively interacts with both the Bsep and Mrp2 canalicular ABC transporters, both involved in the elimination of biliary constituents. 16.2.9
Sulindac
In patients, hepatotoxicity, often of cholestatic nature, is an adverse effect described for long-term treatment with the nonsteroidal anti-inflammatory drug sulindac (Bolder et al., 1999). The effects of sulindac on the bile flow and bicarbonate secretion were studied in bile fistula rats (Bolder et al., 1999). A marked choleretic effect was observed, along with elimination of the unconjugated compound into bile. Studies with isolated perfused rat livers indicated a prolonged choleretic effect indicative of cholehepatic recirculation of sulindac. In the same model, an interference with the hepatic elimination of a cholyltaurine tracer was observed and upon co-infusion of cholyltaurine an acute cholestasis was induced. A competitive inhibition of bile acid transport was observed at both the basolateral and the canalicular pole of the hepatocyte. Overall, in the lower dose range sulindac is thought
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to undergo biliary excretion and cholehepatic recirculation inducing choleresis, while at higher concentrations sulindac is likely to competitively interfere with the canalicular bile salt transporter, thereby contributing to the cholestatic effect observed in patients (Bolder et al., 1999). 16.2.10
Rifampicin and Rifamycin SV
Rifampicin and rifamycin SV, macrolide antibiotic drugs, frequently induce cholestasis as a side-effect (Stieger et al., 2000; Mita et al., 2006). Both compounds interfered with the ATP-dependent taurocholate transport into liver membrane vesicles and membrane vesicles expressing rat Bsep, with K I values in the range of 1 to 12 μM (Stieger et al., 2000). In addition, in the polarized LLC-PK1 cells, expressing both human NTCP and BSEP, rifamycin SV and rifampicin interfered with the bidirectional transport of fluorescently labeled bile acids (Mita et al., 2006). Rifampicin interfered mainly with the bile acid exporter BSEP, leading to an increased intracellular bile acid concentration, while rifamycin SV interfered with both the bile acid uptake carrier NTCP and the exporter BSEP, resulting in a reduction of the intracellular bile acid level and a reduced overall bidirectional bile acid transport (Mita et al., 2006). These studies clearly indicate the complexity of in vitro interaction studies with the active transport processes involved in hepatic bile acid elimination.
16.3
Species Differences in Bsep/BSEP Inhibition
Orthologues of human BSEP have been identified in mouse, rat and rabbit, sharing 81, 82 and 87 % amino acid identity, respectively, with the human protein (No´e et al., 2002). A similar transport of individual bile acids was found for the mouse, rat and human Bsep/BSEP (taurochenodeoxycholate > taurocholate > tauroursodeoxycholate ≥ glycocholate) (No´e et al., 2002). In patients with BSEP deficiency, bile salts are nearly absent in bile, while mice appear to have alternative bile secretory pathways (e.g. Mdr1 and, to a lesser extent, Mdr2, Mrp2 and Mrp3), based on studies with Bsep knockout mice (Lam et al., 2005). This might explain the fact that for many cholestatic compounds no signals of impaired bile secretion and associated hepatotoxicity were seen in rodent toxicity studies. Species differences were also observed in the affinity of rat Bsep and human BSEP for cholestatic compounds. While the inhibition of Bsep/BSEP by midecamycin and cyclosporin A was similar between rat and human cLPMV, cloxacillin and glibenclamide showed a more marked inhibition of the human BSEP as compared to the rat Bsep (Horikawa et al., 2003). The inhibition potential of glibenclamide and three thiazolidinedione antidiabetics, troglitazone, rosiglitazone and pioglitazone, has been analyzed for the rat (Bsep) and human bile salt (BSEP) transporters expressed in membrane vesicles (Table 16. 4). The rank-order of inhibition was similar for the two transporters studied. However, the human BSEP appeared to be more sensitive for the compounds tested, especially for rosiglitazone, for which a 10-fold lower IC50 was observed when using the human BSEP. In addition, the low apparent inhibition of pioglitazone on the human BSEP is not in agreement with the clinical observations, as this compound is not associated with significant liver enzyme elevations. The dose, tissue distribution, metabolism and other factors have to be considered to explain this apparent discrepancy.
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Table 16.4 Species differences in the inhibitory potential of different glitazones on the rat and human bile salt transporters (Bsep, BSEP). The inhibitory effects were determined on the ATP-dependent taurocholate transporters expressed in membrane vesicles
Inhibitor Rosiglitazone Glibenclamide Troglitazone Troglitazone sulfate Pioglitazone
16.4
Inhibition of ATP-dependent taurocholate transport in membrane vesicles, IC50 (μM)
Typical dose range used in clinics (mg/day)
Rat Bsep
Human BSEP
8 10 400–600 — 45
54.4 9.6 10.3 9.3 5.5
6.5 6.3 2.3 4.2 0.4
Regulation of BSEP Expression
ABC transporters, which are involved in hepatic bile formation (BSEP, MRP2, MDR3), are regulated by different transcriptional cascades: ligand-activated nuclear receptors FXR, PXR and CAR, ligand-independent nuclear receptors and transcriptional cofactors (Eloranta and Kullak-Ublick, 2005). For bile acid homeostasis, the nuclear farnesoid X receptor (FXR) plays a pivotal role regulating key enzymes involved in bile acid synthesis and metabolism (CYP7A1, UGT2B4), as well as transport (NTCP, BSEP) (Cai and Boyer, 2006). Mrp3, an ABC transporter located at the basolateral membrane exporting compounds from the hepatocyte into plasma, is significantly up-regulated under cholestatic conditions (Donner and Keppler, 2001), reducing the intracellular bile acid concentration and potentially protecting the liver from the toxic effects of high bile acid levels. A similar regulation can be observed in primary cultured hepatocytes, indicating that these cells acquire a cholestatic phenotype (Rippin et al., 2001). While the bile acid uptake carriers Ntcp and Oatp1 were down-regulated, the exporters involved in bile acid excretion, Bsep, Mrp2 and Mrp1 (into bile or plasma) were up-regulated. Regulation of transporters in disease states and by nutrients and drugs might significantly alter the bile acid transport capacity in the hepatocytes and contribute to the inter-individual variability and sensitivity towards potentially cholestatic drugs.
16.5
Genetic Origin of Intrahepatic Cholestasis
One form of progressive familial intrahepatic cholestasis (PFIC) is characterized by high serum and low biliary bile acid levels and normal serum γ -GT levels (Trauner et al., 1998; Jansen et al., 1999). In one form, a mutation in FIC1, a P-type adenosine triphosphatase, is responsible for the cholestatic disease called ‘Byler disease’ according to the Amish population descending from Jacob Byler where this form was discovered. Linked with a slightly different form of PFIC, PFIC2, mutations in BSEP were found to be responsible for this progressive liver disease (Strautnieks et al., 1998). An impaired activity of the
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hepatobiliary bile salt export pump leads to increased intracellular bile acid levels and intrahepatic cholestasis. Numerous mutations have been described in BSEP, leading to different degrees of intrahepatic cholestasis. In PFIC2 patients, a functional expression of BSEP is typically missing, while in milder forms (benign recurrent intrahepatic cholestasis, BRIC) different mutations in BSEP lead to proteins with lower functional activities. Such a mutation was described in a patient with BRIC and the BSEP protein bearing the respective mutation, showing in vitro a significantly reduced taurocholate transport activity (No´e et al., 2005). Such Bsep transport proteins with reduced activity, as found in BRIC, might render a patient more sensitive and in combination with cholestatic drugs lead to a significant intrahepatic cholestasis.
16.6
Addressing BSEP Inhibition in Drug Discovery and Development to Reduce the Cholestatic Properties of New Molecular Entities
The potential of certain drugs to induce intrahepatic cholestasis has been outlined above and the mechanism by which these compounds interfere with the hepatobiliary elimination of bile acids, mainly by inhibition of BSEP by the drug or major drug metabolites, is well documented for many of these cholestatic compounds. Drugs, which are used at relatively high doses and for which biliary excretion of parent drug or metabolites represents a major elimination pathway, might be particularly prone to interfere with bile acid elimination. The in vitro inhibition of Bsep/BSEP can be easily determined using appropriate in vitro systems, such as assessing the ATP-dependent taurocholate transport in liver plasma membrane vesicles or membrane vesicles expressing the rat/human transporter (Stieger et al., 2000; Funk et al., 2001b; Kostrubsky et al., 2006). Special attention has to be given to include the relevant major drug metabolites in these studies. A high-speed screening system has been reported recently; ATP-dependent uptake was measured using BSEP expressing membrane vesicles and 96-well filtration plates (MultiScreen, Millipore) (Hirano et al., 2006). The vectorial transport of taurocholate was also assessed using double-transfected cells (Mita et al., 2006). The polarized LLC-PK1 cells expressed NTCP and BSEP at the respective membrane and were grown as monolayers on membrane inserts and fluorescent bile acids were used as substrates. The inhibition of the bidirectional transport by a group of cholestatic model compounds was assessed. The drawback of this system is that for the evaluation of the quantitative inhibition potential of the exporter BSEP, the intracellular concentration of the test compound is unknown. For the evaluation of the cholestatic potential of metabolized compounds, the relevant metabolites have to be assessed separately, as these cells are metabolically not competent (Mita et al., 2006). In addition to the in vitro inhibition studies with Bsep/BSEP, it is important to consider the in vivo effect, including tissue (liver) distribution kinetics and metabolism of the test drug. Animal models are used for this purpose with the obvious limitations of species differences in transport and metabolism. An acute cholestasis model in rats was used to characterize cholestatic compounds assessing the effects of intravenously administered compounds on the plasma bile acid level (Funk et al., 2001a,b). A transient increase in bile acids can be observed for cholestatic compounds, allowing a ranking of the in vivo
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Rel. in vivo effect on plasma bile acids –1 (ED50· (Δ10+ Δ30) )
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6
8
10
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In vitro IC50 ( M) Figure 16.8 In vitro and in vivo cholestatic potential of reference compounds (cyclosporin and glibenclamide) and representatives of one chemical series. The in vitro cholestatic potential was determined as the IC50 of Bsep inhibition in cLPMV, while the in vivo effect represents the increment of plasma bile acids increase in rats. Compounds with both low in vitro (high IC50 values) and in vivo cholestatic potentials (*) were selected for further characterization.
cholestatic potential. Results of the in vitro Bsep inhibition are shown in correlation to the in vivo cholestatic potential in rats for a series of molecules within one chemical class along with the corresponding results of the cholestatic reference compounds cyclosporin A and glibenclamide (Figure 16.8). This approach allows for the selection of compounds with a low cholestatic potential and can be systematically applied early on in the drug research and development process.
Abbreviations ALT Alanine transaminase AST Aspartate transaminase Bsep (Abb11) Rat canalicular bile salt export pump BSEP (ABCB11) Human canalicular bile salt export pump cLPMV Canalicular liver plasma membrane vesicles LTC4 Leukotriene C4 , a leukotriene–glutathione conjugate Mrp2 (Abcc2) Rat multidrug resistance related protein 2 MRP2 (ABCC2) Human multidrug resistance related protein 2 MV Membrane vesicles isolated from insect cells (Sf9) recombinantly expressing selected ABC transporters TGZ Troglitazone TZD(s) Thiazolidinedione(s) wt Wild type
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Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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17 Application of Toxicogenomics in Predicting Hepatotoxicity – Potentials and Challenges Wen Lin, Guoxiang Shen, Tin Oo Khor and Ah-Ng Tony Kong
17.1
Introduction
The liver is an important organ not only because it is the largest organ in the body, but also because it is the major organ to metabolize numerous xenobiotics and facilitates the detoxification of toxicants. Therefore, the liver itself is the main target of many potential adverse events generated from its exposure to xenobiotics. Drug-induced hepatotoxicity is a common cause of liver injury which mimics all forms of acute and chronic liver diseases and accounts for a large percentage of the cases of acute liver failure in drug therapy. It is not surprising that drug-induced idiosyncratic hepatotoxicity is the most frequent cause of drug withdrawal post-marketing. Hence, it is critical to be able to identify potential signals or ‘tags’ of hepatotoxicity in pre-clinical or early clinical trials during drug development processes. Traditional toxicological studies usually focus on the short-term 14-week and long-term 2-year rodent (such as rats) animal experiments to evaluate toxicological endpoints, including body weight, organ weight, death rate, tumor incidence and level of serum toxicity biomarkers. Therefore, traditional toxicology studies have the obvious disadvantages, such as being time-consuming and the generation of phenomenological results, plus at times, it’s almost impossible to predict the toxicity of new drug candidates based on currently existing traditional toxicology databases with these phenomenological endpoints. All of these shortcomings form the bottleneck in drug discovery and development processes in the pharmaceutical industry. The rapid development of genomic science provides
Hepatotoxicity: From Genomics to in vitro and in vivo Models Edited by S. C. Sahu C 2007 John Wiley & Sons, Ltd. ISBN: 978-0-470-05716-2
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many opportunities in drug research and development. One of these is the application of genomics in toxicology, or toxicogenomics. Toxicogenomic studies integrate knowledge of genomics, bioinformatics and toxicology to measure the gene expression changes provoked by exposure to a known toxicant or unknown drug candidates (Nuwaysir et al., 1999; Chin and Kong, 2002; Khor et al., 2006). It is believed that toxicants could elicit unique gene expression profiles which could be used to classify them from other non-toxicants. Therefore, databases constructed based on gene expression profiles elicited by known classes of toxicants could be used to predict phenotypes of toxicity of unknown drug candidates, such as potential hepatotoxicity. However, the application of toxicogenomics to hepatotoxicity studies requires an awareness of the limitation of gene expression profiling in the context of mechanistic and predictive toxicology. For example, many known toxicants induce toxicity by directly binding to proteins and DNA, not by directly altering genes involved in toxicity. Gene expression profiles also depend on time, dose and duration of the exposure of the chemical. For many genes, their expression changes reflect secondary outcomes due to primary upstream events rather than a direct response to toxicant exposure. Nevertheless, microarray-based toxicogenomics is still a very young and promising tool to address the mechanisms of hepatotoxicity of known toxicants or predict potential liver toxicity of new drug candidates (Lord et al., 2006).
17.2
Methodology
Because of the availability of genome-scale DNA sequences and functional information, microarray-based gene expression profiling has become an unprecedented powerful tool in toxicogenomic studies. During the past decades, numerous systems were developed inhouse or commercially, to construct large-scale DNA arrays. However, in general, all of these platforms are either cDNA- or oligonucleotide-based arrays (Butte, 2002). For all microarray analysis, the total RNA from samples is amplified to produce antisense cRNA (Figure 17.1). For spotted cDNA microarrays, different fluorescence-labeled cDNAs are prepared from cRNA amplified from treated and control samples; the labeled cDNAs are then purified and hybridized to the cDNA arrays. A cDNA microarray uses a single probe to detect a target gene, and the two biological (control vs. treated) samples are labeled with different colors and mixed at 1:1 ratio. After hybridization, the two colors are scanned separately using a laser and the relative expression level of each gene is determined by comparing the intensities after overlapping the two images. For oligonucleotide-based genomic arrays such as ‘Affymetrix GeneChips’, the cRNAs amplified from biological samples are further subjected to biotin-labeling and fragmentation. Each biological sample is then hybridized with an individual microarray (Figure 17.1). Affymetrix arrays use 11 to 20 pairs of oligonuleotides (containing a pair of ‘mis-matched’ oligonucleotides) to probe a target gene; after hybridization and laser scanning, the intensity differences between ‘perfect-match’ and ‘mis-match’ probes are calculated by appropriate image analysis software to give both quantitative (signal intensity) and qualitative (statistical significance of signals) measurements. The relative expression levels of each gene between different biological samples are obtained after normalization and comparison using software such as ‘Affymetrix GCOS’ and ‘GeneSpring’. Whole genome-based microarrays contain probes representing expressed sequence tags (ESTs). Many ESTs represent transcribed genes with
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Figure 17.1 Schematic illustration of using a microarray-based toxicogenomics approach to classify toxicants or predict the classes of unknown compounds
unknown functions. Therefore, genome-scale microarrays are not ideal platforms for toxicity molecular mechanisms studies due to the lack of functional information of these ESTs. However, gene expression profiles generated from these platforms after toxicants exposure have been successfully used to identify genes differentially regulated by different classes of toxicants or unknown compounds. The identification of genes, whose expression changes, are either directly linked to toxicity or related to the perturbation of cellular signaling pathways related to toxicity, appears to be the most important step to classify toxicants or predict toxicity. On the other hand, genes whose expression changes are not related to toxicity may interfere or confound the above applications in genomic array assays. For arrays designed specifically for toxicogenomic studies, especially these spotted cDNA arrays, sets of genes related to toxicity are selected to construct the array, such as the ‘NIEHS’ in-house printed ‘Human ToxChip V1.0’ cDNA array. The advantages of using these specific toxicologyfocused microarray platforms include the generation of less data points for further analysis and interpretation, and almost all the alternations of genes on the array could be related to toxicity induced by toxicants exposure. With respect to the technical aspect, each type of microarray has employed appropriate data normalization techniques. For spotted cDNA microarrays, simple normalization could be done by adjusting the overall brightness of each scanned microarray image, assuming
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that the quantities of RNA used are equal, or by normalizing the expression levels of housekeeping genes, assuming their expression levels are constant across the experimental conditions. More advanced normalization methods include the use of a locally weighted scatterplot-smoothing regression algorithm, followed by standard deviation regularization between array sub-grids. Normalization to the expression levels of housekeeping genes, and RMA and GCRMA normalization methods, are used in ‘Affymetrix GeneChip’ array analysis. These approaches have substantially improved the reliability and reproducibility of microarray data, even across different platforms or laboratories (Larkin et al., 2005). Therefore, under the same experiment design and similar RNA handling protocols, biological differences between samples have more significant effects on gene expression profiles than platform-specific effects, suggesting that gene expression profiles generated from different microarray platforms are comparable to each other. However, the gene expression levels of both platforms should be transformed to the log2 ratio of gene expression in response to treatments, to the mean value of control, which appear to be more biologically relevant values than the intensity measurements among different platforms (Larkin et al., 2005). To further improve the reliability of microarray data in toxicogenomic applications, a proper design of experiments should be at least equally emphasized. For example, the transcription of many genes is regulated by hormones-related signaling pathways; therefore, the gender of animal or time of sampling should be consistent cross all the biological samples. Another important factor needing to be considered is the heterogenecity of the organ or tissue to be used for RNA isolation, since different parts of an organ may have different subpopulations of cells which might respond differently to the same toxicants exposure. Failure to minimize these variations during experimental design or RNA preparation may generate biological differences that could lead to the misinterpretation of microarray data.
17.3
Data Analysis
The successful application of toxicogenomics in predicting hepatotoxicity relies on the appropriate analysis of microarray data. Experiments using high-density array platforms could easily generate thousands to millions of data points in one experiment, which represents a big challenge for further downstream data analysis. Currently, there are at least two types of mathematical methods to identify genes or expression patterns of interest from microarray data based on the different mathematical algorithms used. The supervised methods are used to identify genes that fit a pre-defined pattern, while the unsupervised methods are used to characterize the signature of a data set without a priori input or knowledge of a training set (Butte, 2002; Maggioli et al., 2006). Supervised methods use statistical approaches such as the ANOVA F-test to identify genes whose expression levels are significantly different between different biological samples; more importantly, supervised methods could also detect a discriminatory gene set among expression profiles generated from studies of toxicants with known toxicological properties. Therefore, supervised methods not only can detect genes related to toxicantsinduced toxicity, but could also perform class comparisons based on gene expression profiles generated from different classes of toxicants. Overall, these methods could be used for discriminant analysis, class prediction and supervised pattern recognition. All of these
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functions are important for toxicogenomic studies to predict the potential hepatotoxicity of new drug candidates or their toxicity classification, using the training databases constructed from gene expression profiles of known liver toxicants. The most popular supervised technique is the nearest-neighbor analysis. Nearest-neighbor methods are based on measuring the distance between observations, such as the Euclidean distance between two gene expression profiles. This method is commonly used to identify a gene expression pattern that best matches a designated query pattern. In toxicogenomics, a query pattern or training set can be generated by microarray analysis of tissues exposed to various toxicants with known types of toxicity compared with that in untreated control tissues. New compounds or drug candidates can then be tested in the
