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This is to certify that the
dissertation entitled

Neutrophils and Idiosyncratic Advese Drug Reactions

Resulting from Inflammation-Drug Interaction: Ranitidine and

Diclofenac as Examples

presented by

Xiaomin Deng

has been accepted towards fulfillment
of the requirements for the

PhD degree in Biochemistry and Molecular
Biology

 

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Major Professor’s Signature

3-1 1-2008

 

Date

MSU is an affirmative-action, equal-opportunity employer

PLACE IN RETURN BOX to remove this checkout from your record.
TO AVOID FINES return on or before date due.
MAY BE RECALLED with earlier due date if requested.

 

DATE DUE DATE DUE DATE DUE

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

5/08 K:lProi/Acc&Pres/CIRC/DateDua indd

 

Neutrophils and Idiosyncratic Advese Drug Reactions Resulting from Inflammation-Drug
Interaction: Ranitidine and Diclofenac as Examples

By

Xiaomin Deng

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

DOCTOR OF PHILOSOPHY
Department of Biochemistry and Molecular Biology

2008

ABSTRACT

NEUTROPHILS AND IDIOSYNRATIC ADVERSE DRUG REACTIONS
RESULTING FROM lNFLAMMATlON-DRUG INTERACTION

By

Xiaomin Deng

Idiosyncratic adverse drug reactions (IADRs) occur in a small fraction of people
taking a drug. The liver is a frequent target. For the vast majority of drugs associated with
these reactions, including the histamine2 (H2) receptor antagonist ranitidine (RAN) and
the nonsteroidal anti-inflammatory drug diclofenac (DCLF), the mechanism of toxicity is
unknown. Inflammatory stress might be a susceptibility factor for IADRs, which was
supported by the observation that mild inflammation renders nontoxic doses of RAN and
DCLF injurious to liver in rats. This dissertation tested the hypothesis that neutrophils
(PMNs) are activated in the livers of rats cotreated with LPS and an lADR-producing
drug (RAN, DCLF), and these cells contribute to liver injury by enhancing hemostasis
and/or releasing neutrophil proteases. In the LPS/RAN model, LPS induces the
accumulation of PMNs in the liver, and RAN causes the transmigration and the activation
of these PMNs. Activated PMNs release toxic proteases that kill hepatocytes or enhance
fibrin deposition in liver, both of which could lead to hepatocellular injury caused by
LPS/RAN treatment. PMNs enhance the fibrin deposition through plasminogen activator
inhibitor-1 G’AI-l), the negative regulator of the fibrinolytic system. A PAI-l inhibitor

reduced the hepatic fibrin deposition and hepatocellular injury caused by LPS/RAN

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cotreatment. The PAI-l inhibitor also reduced PMN activation independently of
neutrophil chemokines, suggesting a direct role for PAI-l in activating PMNs. The
enhanced PAI-l production by RAN treatment after LPS exposure occured through
tumor necrosis factor-alpha (TNFa). RAN augmented TNFa production after LPS
treatment, and the prolongation of LPS-induced TNFa production by RAN is crucial for
the liver injury caused by LPS/RAN cotreatment, since a TNFa-converting enzyme
(TACE) inhibitor, given immediately before RAN, reduced both serum TNFa protein and
the hepatocellular injury. The augmentation of TNF-a by RAN occured in a
post-transcriptional manner by enhanced p3 8-dependent TACE activation. In addition to
the RAN model, PMNs are also important in the liver injury caused by LPS/DCLF
cotreatment. These studies could aid in understanding mechanisms by which
inflammation acts as a susceptibility factor for IADRs and biomarker identification for

IADRs resulting from inflammation-drug interaction.

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ACKNOWLEDGMENTS

Of all the sections in this dissertation, this is perhaps the most difficult to write. A
very large number of people from Michigan State University, from the Department of
Biochemistry and Molecular Biology, the Department of Pharmacology and Toxicology
and the Center for Integrative Toxicology deserve tremendous thanks. In addition, my
interactions with other collaborators and colleagues from Abbott Biosrearch Center,
Abbott Laboratory, Bristol-Myers Squibb, Wyeth, University of Gratz and Society of
Toxicology have dramatically increased my excitement for toxicology. To paraphrase
these thanks would be insufficient.

Let’s start with the MSU folks. First and foremost I would like to thank my mentor,
Dr. Robert Roth. Deciding to join his laboratory was perhaps the most influential and best
decision I have made. I hope I can reflect Bob’s enthusiasm and perseverance in
toxicological research as I move onward in my career. He not only provided brilliant
scientific ideas in my thesis research but also gave advice on the mental development of
being a great scientist. Dr. Patricia Ganey also deserves my thanks. Patti played a very
significant role in my graduate career as another mentor in the lab. She even managed to
teach me lab techniques once after being a faculty for so many years. I perhaps own a big
thanks to Dr. James Lueyndyk. Jim is always ready to answer whenver I went to him for
questions and suprisingly he always knows the answers. I would like to thank Jim for the
significant amount of time he put in to training me in the lab as well as the teamwork on
tln's dissertation work especially in my early graduate study. Another big thank I shall

give to is Dr. Jane Maddox. She is my first mentor in lab during my rotation and it is my

iv

 

pleasure to work with her then and throughout the rest of my graduate study. She is the
expert on in vivo animal behavior and always knew the terms that I have never heard of
before. I would also like to thank Dr. John Lapres, Dr. Donald Jump, Dr.Kathy Gallo and
Dr. Dave Dewitt for serving on my graduate committee and for their continued interest in
my research and career goals.

I have had the pleasure of working with so many other incredible people in the
laboratory during my time at MSU and all of them deserve my thanks. Several post-docs
in the lab have helped me at times including Dr. Francis Tukov, Dr. Sachin Devi, and Dr.
Jesus Olivero-Verbel. In addition to these guys, I would like to thank all of the laboratory
technicians, graduate students, and undergraduates I have had the opportunity to work
with including Sandy Newport, Steve Bezdecny, Colin North, Pat Shaw, Rohan Pradhan,
Theresa Eagle, Wei Zou, Erica Sparkenbaugh, J ingtao Lu, Aaron Fullerton. In addition to
folks in the lab, I would also like to the administrative staff of the Center for Integrative
Toxicology, Department of Biochemistry and Molecular Biology and Department of
Pharmacology and Toxicology.

In addition to colleagues on campus I have offered the unique opportunity to
collaborate with others in industry and academia on the work presented herein as well as
other projects. Dr. Robert Stachelwitz has served as another mentor to me during my
internship at Abbott Bioresearch Center. Rob has been helpful in all aspects of my career
from experimental design to encouragement. His energetic personality and sense of
humor always excites me. We are able to forge a fantastic collaboration with Abbott
Laborotary. I would like to thank the Abbott group including Jeff Waring, Mike Liguori

and Eric Blomme. I would like to also thank Dr. Ernst Malle from University of Gratz,

Austria, Dr. Lois Lehman-Mckeeman from Bristol-Myers Squibb and Dr. Dave Drandell

for providing useful tools to my thesis research.

vi

TABLE OF CONTENTS

LIST OF TABLES ................................................................................. x
LIST OF FIGURES ............................................................................... xi
LIST OF ABBREVIATIONS .................................................................... xiii
CHAPTER 1
Introduction ........................................................................................ 1
1.1 Drug-induced idiosyncratic hepatotoxicity .................................... 2
1.1.1 Overview of drug induced idiosyncratic hepatotoxicity. 2
1 .1.2 Example: Troglitazone ................................................. 5
1.1.3 Inflammatory stress and IADRs 9
1.2 Inflammation ....................................................................... 12
1.2.1 Overview of inflammation as a contributor to tissue injury ...... 12
1.2.2 TNFa ..................................................................... 13
1.2.3 Neutrophils (PMNs). . . .. .............................................................. 17
1.2.4 The hemostatic system and hypoxia .................................. 20

1.3 Mechanisms of liver injury in models of inflammation-drug interaction:

ranitidine asexample ............................................................... 23
1 .3.1 Ranitidine-induced idiosyncratic hepatotoxicity in human
patients .................................................................... 23
1.3.2 Inflammation-ranitidine interaction .................................. 24
1.3.3 Involvement of hemostasis,PMN and TNFa ...................... 26
1.3.4 Interaction of TNFa, hemostasis and PMNs31
1.4 Diclofenac-induced idiosyncratic hepatotoxicity .............................. 38
1.5 Overview of the thesis ............................................................. 42
1.5.1 Knowledge gaps .......................................................... 42
1.5.2 Hypothesis and specific aims .......................................... 43
CHAPTER 2
Neutrophil interaction with the hemostatic system contributes to liver injury in rats
cotreated with lipopolysaccharide and ranitidine ............................................. 45
2.1 Abstract .............................................................................. 45
2.2 Introduction ......................................................................... 48
2.3 Methods ............................................................................. 50
2.3.1 Materials ................................................................. 50
2.3.2 Animals .................................................................. 50

vii

2.3.3 Experimental Protocol ................................................. 50
2.3.4 PMN Depletion, CD18 Neutralization and Eglin C Treatment.. 51
2.3.5 Hepatotoxicity Assessment ........................................... 52
2.3.6 Fibrin Immunohistochemistry and Quantification ................. 52
2.3.7 Evaluation of Coagulation System Activation and Plasma PAI-
1...................... ........................................................................ 1523
2.3.8 Evaluation of Liver Hypoxia ...................................................... 53
2.3.9 Evaluation of Circulating Leukocytes, PMNs, Liver PMNs and
PMN activation........... ............................................................ 54
2.3.10 Evaluation of PMN Protease Effects on Fibrinogcn and PAI-l in
tho.u.u..u.u..n.u..n.u..”.H..u.u.”..u.u.unnununuuuun.55
2.3.11 Statistical Analysis ........................................................... 56
2.4 Results ............................................................................... 58
2.4.1 PMN depletion studies in the LPS/RAN model .................... 58
2.4.2 HOCl-protein adduct staining ......................................... 66
2.4.3 Effects of CD18 neutralization on hepatotoxicity, PMNs and
hemostasis ............................................................... 75
2.4.4 Effects of eglin C on serum ALT activity and hemostasis ........ 82
2.5 Discussion ........................................................................... 87
CHAPTER3
p3 8-dependent TNF-a converting enzyme is important for liver injury in hepatotoxic
interaction between lipopolysaccharide and ranitidine ....................................... 96
3.1 Abstract .............................................................................. 97
3A Innoducfion .......................................................................... 99
3.3 Methods ............................................................................. 102
3.3.1 Materials ................................................................. 102
3.3.2 Animals .................................................................. 102
3.3.3 Experimental Protocol ................................................ 102
3.3.4 Treatment with inhibitors of p38, TACE or Plasminogen Activator
Inhibitor-1 (PAI-l) ...................................................... 103
3.3.5 Hepatotoxicity Assessment ............................................ 104
3.3.6 Evaluation of Hepatic Phospho-p38 and Phospho-MK-Z ....... 104
3.3.7 Evaluation of Coagulation System Activation and Plasma PAI-
1 ........................................................................... ‘l()fS
3.3.8 Serum TNF-o. and MlP-2 Evaluation ................................ 105
3.3.9 Fibrin Immunohistochemistry and Quantification .................. 105
3.3.10 Evaluation of Liver PMNs and PMN Activation ................... 106
3.3.11 Evaluation of Hepatic TNF-a mRNA and TACE activity ........ 107
3.3.12 Statistical Analysis ................................................... 108
3.4 Results ............................................................................... 117
3.4.1 p38 activation after LPS/RAN treatment ............................ 117
3.4.2 Effects of p38 inhibitor on hepatotoxicity ........................... 117
3.4.3 Effects of p38 inhibitor on biomarkers of hemostasis and liver
I’lvll\ls ..................................................................... 1 117

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3.4.4 Hepatic TNF-a mRNA after LPS/RAN treatment .................. 119
3.4.5 Effects of p38 inhibitor on serum TNF-a protein and hepatic TNF-
a mRN A after LPS/RAN treatment ................................... 119
3.4.6 Hepatic TACE activity after LPS/RAN treatment .................. 119
3.4.7 Effects of TACE inhibitor on serum TNFa and liver injury. . 128
3.4.8 Effects of TACE inhibitor on plasma PAI-l ......................... 128
3.4.9 Effects of PAH inhibitor on hepatotoxicity and markers of
hemostasis and PMNs ................................................... 129
3.5 Dicussion ............................................................................ 136
CHAPTER 4
Modest inflammation enhances diclofenac hepatotoxicity in rats: role of neutrophils and
bacterial translocation ................................... 144
4.1 Abstract .............................................................................. 145
4.2 Introduction ......................................................................... 147
4.3 Methods ............................................................................. 150
4.3.1 Experimental Design .................................................. 150
4.3.2 PMN depletion ......................................................... 151
4.3.3 Gut Sterilization ......................................................... 151
4.3.4 Bacterial Culture in Fecal and Liver Homogenates ................ 151
4.3.5 ALT Activity and Histopathology Assessmen ...................... 152
4.3.6 RNA Preparation ........................................................ 152
4.3.7 Gene Array Analysis ................................................... 153
4.3.8 Statistical Filtering of Genes Changed After LPS/DCLF
Treatment........... ...................................................................... 154
4.3.9 Serum MIP-2 Concentration and Blood/Liver Leukocyte
Evaluation ............................................................... 155
4.3.10 Statistical Analysis ................................................... 156
4.4 Results ............................................................................... 157
4.4.1 Hepatotoxicity from DCLF/LPS interaction ........................ 157
4.4.2 Microarray analysis of livers from rats treated with
LPS/DCLF ............................................................... 157
4.4.3 Neutrophil-related gene expression changes after LPS/DCLF
treatment and effects of neutrophil eepletion on LPS/DCLF-
induced hepatotoxicity ................................................ 171
4.4.4 Effects of gut sterilization on LPS/DCLF hepatotoxicity ....... 178
4.4.5 Effects of gut sterilization on liver injury in rats treated with a
hepatotoxic dose of DCLF ............................................ 179
4.5 Discussion ........................................................................... 186
CHAPTER 5
Summary and Conclusions ....................................................................... 191
5.1 Summary and conclusion .......................................................... 192
BIBLIOGRAPHY ................................................................................. 204

ix

 

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Table 4.1

Table 4.2

Table 4.3

Table 4.4

Table 4.5

LIST OF TABLES

Circulating Leukocytes after PMN Antiserum Treatment in LPS/RAN-treated
Rats ................................................................................... .57

Histopathological Evaluation of Livers from LPS/DCLF-treated Rats. . 162

Functional Classification of Genes Expressed to Greater Degree after
LPS/DCLF Treatment than after LPS or DCLF Given Alone ................. 168

Gene Ontology (GO) Biological Process Categories for Which Gene
Expression Was Affected to a Greater Degree after LPS/DCLF Treatment
than after Treatment with LPS or DCLF Alone ................................. 170
Effects of Anti-PMN Serum on Circulating Leukocytes and Liver PMNs... 180

Effects of Gut Sterilization on liver Bacterial Colony Growth after High Dose
DCLF Treatment ..................................................................... 185

LIST OF FIGURES

Images in this dissertation are presented in color

Figure 1.1
Figure 2.1
Figure 2.2

Figure 2.3

Figure 2.4

Figure 2.5

Figure 2.6
Figure 2.7
Figure 2.8

Figure 2.9

Possible pathogenic mechanism of LPS/RAN-induced liver injury ......... 37
Effects of PMN depletion on hepatotoxicity induced by LPS/RAN ......... 60
Effects of PMN depletion on liver fibrin after LPS/RAN treatment. . . . . 62
Effects of PMN depletion on coagulation system activation after LPS/RAN

treatment ............................................................................ 64
Effects of PMN proteases on plasma fibrinogen in vitro. . . . . . . . . . . . . 66

Effects of PMN depletion on plasma PAI- 1 concentration after LPS/RAN
treatment” . 70

Effects of PMN proteases on PAI-l concentration in vitro .................... 72
Effect of PMN depletion on liver hypoxia after LPS/RAN treatment ....... 74
PMN activation after LPS/RAN treatment ...................................... 77

Effects of CD1 8 neutralization on biomarkers of hemostasis, PMNs and
hepatotoxicity in LPS/RAN-cotreated rats ..................................... 79

Figure 2.10 Effects of CD18 neutralization on liver hypoxia after LPS/RAN

treatment ............................................................................ 8 1

Figure 2.11 Effects of PMN-protease inhibitor on hepatotoxicity after LPS/RAN

treatment ............................................................................. 84

Figure 2.12 Effects of PMN protease inhibitor on markers of hemostasis and liver injury

after LPS/RAN treatment ........................................................... 86
Figure 2.13 Interaction of PMNs with the hemostatic system in the LPS/RAN model of
liver injury: working hypothesis ................................................... 95
Figure 3.1 p38 activation after LPS/RAN treatment ........................................ 110
Figure 3.2 Effects of a p38 inhibitor on MK-2 phosphorylation .......................... 112
Figure 3.3 Effects of a p38 inhibitor on serum ALT acitivty .............................. 114
xi

Figure 3.4 Effects ofa p38 inhibitor on biomarkers of hemostasis. . . . . . . . . . . . . 116
Figure 3.5 Effects of a p38 inhibitor on PMN biomarkers .................................. 121
Figure 3.6 Hepatic TNF-a mRNA after LPS/RAN treatment ............................. 123
Figure 3.7 Effects of a p38 inhibitor on serum TNF-a protein and hepatic TNF-a
mRNA ............................................................................... 125
Figure 3.8 Hepatic TACE activity after LPS/RAN treatment ............................. 127
Figure 3.9 Effects of a TACE inhibitor on serum TNF-a and liver injury ............... 131
Figure 3.10 Effects of TACE inhibitor on plasma active PAI-l ............................ 133
Figure 3.11 Effects of a PAI—l inhibitor on hepatotoxicity and markers of hemostasis and
PMN activation ..................................................................... 135
Figure 3.12 Diagram of pathogenic mechanism contributing to hepatocellular injury in
LPS/RAN model ................................................................... 143
Figure 4.1 Dose-responsive DCLF hepatotoxcity ........................................... 159
Figure 4.2 LPS potentiates DCLF hepatotoxicity ............................................ 161
Figure 4.3 Hierachical cluster analysis of hepatic gene expression after LPS/DCLF
treatment ............................................................................ 165
Figure 4.4 Venn diagram of gene expression changes caused by LPS/DCLF .......... 167
Figure 4.5 Neutrophil-related gene expression changes after LPS/DCLF treatment... 173
Figure 4.6 Serum MIP-2 protein concentration after LPS/DCLF treatment ............. 175
Figure 4.7 Effects of PMN depletion on LPS/DCLF-induced liver injury ............... 177
Figure 4.8 Effects of gut sterilization on LPS/DCLF-induced liver injury .............. 182
Figure 4.9 Effects of gut sterilization on hepatotoxicity induced by a Large Dose of

Figure 5.1

DCLF ................................................................................ l 84

Summary diagram of pathogenic mechanism for LPS/RAN-induced liver
injury ................................................................................. 200

xii

AN OVA
ALT
ALP
AST
COX-2
CINC- 1
CS
DCLF
EU
ELISA
FAM
fMLP
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GSH
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HA
HIF-l a
HPC
HPF
HOCL
IADR
ICAM-l
IL- 1

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LPS
MAPK
MIP-2
MIP- l a.
MK-2
NAS
NF-KB
CS
NSAID
PIM
PAI-l
PAR
PMN

LIST OF ABBREVIATIONS

analysis of variance

alanine aminotransferase

alkaline phosphatase

aspartate aminotransferase
cyclooxygenase-2
cytokine-induced chemoattractant-l
control serum

diclofenac

endotoxin unit

enzyme-linked immunosorbent assay
famotidine

f-Met-Leu-Phe

galactosarnine

gamma-glutamyl transferase
glutathione

histamine2-

hyaluronic acid

hypoxia inducible factor-1a

hepatic parenchymal cell

high power field

hypochlorous acid

idiosyncratic adverse drug reaction
intercellular adhesion molecule-1
interleukin 1

intravenous

intraperitoneal

c-Jun N-terminal kinase

Kupffer cell

Limulus arnoebocyte lysate
lipopolysaccharide

mitogen activated protein kinase
macrophage inflammatory protein 2
macrophage inflammatory protein-1a
MAPK-activated protein kinase 2
rabbit anti-rat PMN antiserum
nuclear factor kappaB

control serum

nonsteroidal anti-inflammatory drugs
pimonidazole

plasminogen activator inhibitor-1
protease activated receptor
neutrophil

xiii

RECA-l
SEC
TAT

TF

TGZ
TLR4
TNF
TVX

ranitidine

rat endothelial cell antigen-1
sinusoidal endothelial cell
thrombin-antithrombin dimmer
tissue factor

troglitazone

toll-like receptor 4

tumor necrosis factor-alpha
trovafloxacin

xiv

CHAPTER ONE

Introduction

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1.1 Drug-induced idiosyncratic hepatotoxicity
1.1.1 Overview of drug-induced idiosyncratic hepatotoxicity

Adverse drug reactions (ADRS) remain a challenging human health problem.
Although continued efforts are put forward to understanding and prevent these reactions,
ADRs still contribute significantly to hospitalization and mortality throughout the world.
Furthermore, due to under-reporting, the real incidence might be even higher than
reported by current methods (Bagheri et al., 2000;Sgro et al., 2002). An immediate
adverse outcome is the withdrawal or limited usage of certain efficacious drugs, leading
to deficits in successful treatment of diseases. One example is the anti-epileptic drug
felbamate (Pellock, 1999;Dieckhaus et al., 2002), an effective drug in treating severe
cases of epilepsy. Unfortunately, its usage was markedly restricted due to the aplastic
anemia and hepatotoxicity it caused in patients. In addition, ADRs cause a significant
loss of investment, commited effort and future profits and a possible financial loss from
lawsuits from the pharmaceutical industry view point.

A variety of drugs with different pharmacological targets are associated with ADRs.
One of the common target organs is the liver. Drug-induced liver injury is associated with
histopathologic and clinical features of acute hepatocellular necrosis, biliary injury, or a
combination of the two (Zimmerman, 1993;Zimmerman, 2000). The main culprits are

anti-infectious, central nervous system, musculoskeletal, and gastrointestinal drugs

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Adverse drug reactions can be divided into predictable, dose-related reactions (Type
A reaction) and idiosyncratic reactions (Type B reaction). Type A reactions are
dose-dependent and occur in a relatively defined time frame, and all individuals are
suscceptible. A typical example is acetaminophen-induced hepatotoxicity in people
(Amar and Schiff, 2007;Larson et al., 2005). In contrast, idiosyncratic adverse drug
reactions (IADRs) typically occur only in a small fraction of the patients taking a drug.
These reactions are usually unpredictable, not apparently dose-dependent and display a
variable onset time after the beginning of drug therapy (Kaplowitz, 2005c;Uetrecht,
2007;Waring and Anderson, 2005). Moreover, the toxicity resulting from most IADRs is
not related to the pharmacological target. All of those features of IADRs make these
reactions more insidious, more difficult to understand and, unfortunately, more difficult
to predict from current preclinical studies or clinical trials.

A well known example of drug-induced idiosyncratic hepatotoxicity is
troglitazone-induced liver injury. Troglitazone, which was marketed in 1997, is one of
the many thiazolidinediones for treatment of type 2 diabetes. It acts as a peroxisome
proliferator-activated receptor-gamma (PPARy) agonist and improves insulin resistance.
However, clinical trials indicated that 1.9% of patients taking the drug experienced
elevated serum alanine aminotransferase more that 3 times the upper limit of normal,
revealing significant frequency of mild and moderate hepatotoxicity (Watkins and
Whitcomb, 1998). Furthermore, it was reported that rare but severe liver injury also

occurred, and this serious idiosyncratic hepatotoxicity from troglitazone led to its

 

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withdrawal from the market in 2000 (Graham et al., 2003). Liver biopsy from affected
patients revealed predominately hepatic necrosis with occasional bridging fibrosis. Acute
and chronic inflammatory infiltrates were cited most often, with rare prominence of
eosinophils. The time frame of hepatotoxicity occcurance varied from within a month to
several years after initiation of maintenance therapy. IADRs induced by troglitazone
are most likely independent of its pharmacological target, since other thiazolidinediones
acting as PPARy agonists, such as rosiglitazone and pioglitazone, raise less concern of
hepatotoxicity for diabetic patients (Scheen, 2001a;Scheen, 2001b).

Better prediction of idiosyncratic liver injury will require a better understanding of
the mechanisms. It is noted that “no one model fits the characteristics of all idiosyncratic
drug reactions” (Seguin and Uetrecht, 2003a). Drug properties, genetic background and
environmental factors could all contribute to idiosyncratic liver injury (Boelsterli,
2003a;Kaplowitz, 2001). Two conventional hypotheses have arisen over the years to
explain the mechanisms of idiosyncratic drug reactions. One of them is that the reactions
are based on drug metabolism polymorphisms among patients, which result in different
levels of toxic drug metabolites (Williams and Park, 2003b). The other one argues that
the reactions arise from an adaptive immune response to proteins bound to the drug or its
metabolites (Park et al., 2001;Ju and Uetrecht, 2002). An extension of this hypothesis of
adaptive immune response is the “danger hypothesis” (Seguin and Uetrecht,
2003b;Pirmoharned et al., 2002b). This hypothesis suggests that a second “danger

signal,” in addition to the presense of antibodies, is necessary to mount a specific immune

 

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response and the consequent hepatotoxicity. This signal might be any number of factors
including some form of cellular stress, underlying disease or other environmental factors.
The following sections will use troglitazone, a well known drug associated with IADRs,
as an example to review the progress, knowledge gaps and alternative thinking in

understanding the mechanisms of idiosyncratic hepatotoxicity.

1.1.2 Example: Troglitazone

Since its withdrawal in 2000, a large amount of effort has been made to elucidate the
mechanism of troglitazone (TGZ)-induced hepatotoxicity. A variety of hypotheses were
proposed to explain TGZ-induced cell injury including reactive metabolites formation
and accumulation, mitochondrial dysfunction and oxidant stress, inhibition of bile salt
transporter, and apoptosis (Masubuchi, 2006;Chojkier, 2005b). For example, the major
TGZ metabolites (sulfate, glucuronide, quinone) have been identified in both cultured
cells, experimental animals and human patients (Loi et al., 1999;Kawai et al.,
1997;Yoshigae et al., 2000;Watanabe et al., 2002;Honma et al., 2002). However, TGZ
metabolite-protein adducts have only been demonstrated by incubation of liver
microsomes from rats with various P450 inducers or “Supersomes” (cDNA-expressed
human P450) (He et al., 2004a). Furthermore, although TGZ is cytotoxic to human
HepG2 cells and rat and human hepatocytes, inhibitors of drug metabolizing enzymes
responsible for TGZ metabolism have not succeeded in protecting against the

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al., 2002). In a recent study, HepG2 cells together with microsomes containing
cDNA-expressed CYP3A4 or HepG2 cells transfected with CYP3A4 were able to
metabolize TGZ, leading to increased cytotoxicity (Vignati et al., 2005). This finding has
been questioned since the TGZ quinone metabolite formed by CYP3A4 is less cytotoxic
than TGZ, both in rat hepatocytes and HepG2 cells (Tettey et al., 2001a). In addition, in
normal human hepatocytes, these CYP3A4-related metabolites are unlikely to be
generated and accumulate in these cells enough to exert toxic effects. In
summary, studies investigating TGZ-induced cytotoxicty were performed with cultured
cell lines using concentrations of TGZ 1—2 orders of magnitude above pharmacological
levels. These studies in vitro have not provided a relevant mechanistic explanation
for the infrequent troglitazone hepatotoxicity in patients. Thus, the role of the reactive
metabolite in TGZ cytoxicity remains a question.

Other evidence supporting the role of metabolic idiosyncrasy in TGZ-induced
hepatotoxicity is the finding of single-nucleotide polymorphisms (SNPs) for
TGZ-metabolizing enzymes in human populations. SNPs have been found in three
genes important to CYP3A activity (Kuehl et al., 2001b). This is relevant because the
hepatic gene expression of CYP3A4 varies about 50-fold, and the CYP3A4 overal
cnzymic activity in vivo varies at least 20-fold among individual people (Eichelbaurn and
Burk, 2001). SNPs of the CYP3A gene family members affect various ethnic groups ,
where it is expressed at high levels in a minority of Americans of European descent and

Europeans (Kuehl et al., 2001agHustert et al., 2001). However, the TGZ has not been

found 10 be 1“
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found to be toxic with doses up to 1200mg/kg after 52 treatment of monkeys, although
the TGZ treatment resulted in significant exposure to TGZ quinine (M3) (Rothwell et al.,
2002). In human patients, M3-derived reactive intermediates were found to bind
covalently to microsomal protein and glutathione (Tettey et al., 2001b;He et al., 2004b),
but the importance of these adducts to TGZ- associated hepatoxicity has not been
established. In a cohort study of 4,079 patients, combined genetic polymorphisms from
several genes were associated with increased susceptibility for TGZ-induced liver injury.
They included CYP1A1, NAD(P)H dehydrogenase, quinone 1 (NQOl), glucose
transporter type 1 (GLUT-l), PPARy-892, and PPARy-143l (de la Iglesia FA et al.,
2003). In another study, a strong correlation was also observed between TGZ-induced
liver injury and the combined glutathione S-transferase-theta l (GSTT1)- glutathione
S-transferase M1 (GSTM1) null genotype (Watanabe et al., 2003). However, none of
these studies tested the functional relationship of these SNPs with TGZ-induced liver
injury in human patients.

Some of the reported cases had histological evidence for an adaptive
immune-mediated reaction (Arioglu et al., 2000;Kohlroser et al., 2000;Murphy et al.,
2000) or the responses can be reduced by corticosteroids (Prendergast et al.,
2000;Bonkovsky et al., 2002). One of these patients developed a similar cholestatic
hepatitis when switched to rosiglitazone after TGZ treatment, consistent with an adaptive
immune-mediated reaction and suggesting a class effect related to the drug’s

pharmacological target. The occurrence of eosinophils or granulomatous inflammatory

 

irritates in t
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infiltrates in the liver is evidence, albeit inconclusive that hypersensitive immune
response plays a role in the injury induced by TGZ in some patients. Unfortunately, there
are no studies in experimental animals which could reproduce the liver injury,
particularly hypersensitivity reactions, caused by TGZ in human patients.

A recent study by Boelsterli and colleagues showed that TGZ could induce mild
hepatocellular injury after 4 weeks of treatment in SOD2+/-- heterozygous mice (Ong et
al., 2007). Hepatic mitochondria isolated from TGZ-treated mice exhibited enhanced
oxidative stress. Furthermore, in hepatocytes isolated from untreated SOD2+/--, but not
wild-type mice, TGZ caused a concentration- dependent increase in superoxide
anion production. This was the first demonstration in an animal model that TGZ
treatment induced mild liver toxicity. In agreement with previous studies, wild-type
animals tolerated the drug without adverse effects, however, mice with a deficiency
in mitochondrial antioxidant defense were found to be susceptible. The results support
the general concept that subclinical stresses from prolonged drug treatment superimposed
on a genetic deficiency in the same organism can lead to cell injury and organ
damage. However, a single genetic defect such as SOD in the human population might
not be responsible for all the cases of liver injury in patients after TGZ treatment.

The lack of evidence that TGZ is hepatotoxic in healthy experimental animals and
the findings of Boelsterli and colleages suggest that some undefined stress and/or

differences in certain human patients, either genetic or environmental, make them

susceptible to TGZ-induced hapatotoxicity. Stresses such as deficient mitochondrial

gfioxldmt dc

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antioxidant defense can be periodic and can go unnoticed. This could explain the erratic

temporal and dose relationships that characterize IADRs.

1.1.3 Inflammatory stress and IADRs

Results of numerous studies indicate that a modest inflammatory response can
enhance liver sensitivity to a variety of toxic chemicals in rats (Yee et al., 2000;Barton et
al., 2000a). Episodes of inflammation are commonplace in people, although most of them
cause no obvious injuries. Moreover, they occur irregularly and often go unnoticed.
These observations have led us to hypothesize that an episode of inflammation
during drug therapy might decrease the threshold for drug toxicity and thereby render an
individual susceptible to an adverse drug reaction. This could be an attractive concept to
explain the basis for IADRs. During drug therapy, concurrent, perhaps unnoticed
inflammatory or other environmental stresses could render an otherwise nontoxic drug
dose injurious to certain organs. This could explain the erratic temporal and dose
relationships of that characterize idiosyncratic reactions.

Small doses of lipopolysaccharide (LPS), the cell wall component of gram-negative
bacteria, precipitate modest inflammatory responses in mammals, resulting in increased
susceptibility to toxicitiy from numerous hepatotoxic chemicals (Roth et al., 1997;Ganey
and Roth, 2001). For example, exposing rats to a nonhepatotoxic dose of LPS results in a
several fold increase in sensitivity to liver injury from allyl alcohol (Sneed et al., 1997).

This increase in sensitivity depends on an LPS-stimulated inflammatory response

 

inciting Ru;
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involving Kupffer cell-derived eicosanoids (PGDz), neutrophils and the hemostatic
system, the system controls blood clotting in vessels (Kinser et al., 2002;Ganey et al.,
2001;Kinser et al., 2004).

In animal studies, evidence is growing that supports the ability of LPS to augment
hepatotoxicity induced by several classes of drugs that cause IADRs in people. For
example, coadministration to rats of nonhepatotoxic doses of LPS and the antipsychotic
drug chlorpromazine (CPZ) results in liver damage and other effects that resemble human
CPZ idiosyncrasy (Buchweitz et al., 2002). Trovafloxacin, a fluoroquinolone antibiotic
associated with idiosyncratic reactions, also interacts with LPS, resulting in hepatotoxcity
both in rats and mice (Waring et al., 2006c;Shaw et al., 2007). In contrast, levofloxacin,
another quinolone antibiotic without the tendency for idiosyncratic reactions, does not
share this interaction with inflammatory stress. Thus, this model of drug-inflammation
interaction is able to distinguish a drug that causes idiosyncratic adverse reactions from
one in the same pharmacologic class that does not. A recent study showed that sulindac, a
NSAID associated with IADRs, also could induce liver injury at an otherwise nontoxic
dose when cotreated with LPS (Zou et al., 2008). These animal models mimicking
idiosyncratic reactions in human patients could provide usefirl tools for mechanistic
study, which in turn could lead to novel biomarkers and methods to prevent or cure
adverse reactions. The models that will be explored by this thesis are ones in which

animals are treated with either ranitidine (RAN) or diclofenac (DCLF). Both of these

10

 

tags are as.»

it iiscttssed ::

 

drugs are associated with rare idiosyncratic hepatotoxicity in human patients, which will

be discussed in detail in the following sections.

11

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1.2 Inflammation
1.2.1 Overview of inflammation as a contributor to tissue injury

Inflammation is classically defined as a local protective reaction of tissue to
irritation, injury, or infection, characterized as “redness, swelling, pain and heat.”
However, inflammation is now viewed in terms of activation of cells of the innate
immune system, the coordination of the mediators they produce and altered gene
expression and cell signaling. Inflammation can both participate in host defense and have
the potential to injure tissues. Indeed, it has become clear that inflammation plays a role
in the pathogenesis of many diseases, can cause tissue injury by itself and can increase
sensitivity of tissues to the toxic effects of some other xenobiotics as mentioned above.

Inflammation comprises not only traditional inflammatory cells (e.g.,
neutrophils, macrophages) and the mediators they produce (e.g., cytokines/ chemokines,
coagulation factors, complement factors) but also endothelial cells and parenchymal cells
in the tissue (Ganey et al., 2004). The inflammatory cells can attack and damage the
tissues directly by releasing toxic factors such as reactive oxygen species, proteases, etc.,
or they can release cytokines, eicosanoids or other mediators that lead indirectly to other
damaging events. The hemostatic and complement systems are also activated in
inflammatory responses and can participate in tissue injury. Changes in gene expression
and the transcriptional regulators involved (e.g., nuclear factor kappa B, early growth
response gene-1) are integral players in the inflammatory response. The hemostatic and

complement systems are also activated in inflammatory responses and can participate in

12

 

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tissue injury. Transcriptional regulators not only control the production of inflammatory
mediators but also play an important role indirectly in the activation of various cells
involved.

Inflammation is known to be a feature of numerous diseases and may contribute to
disease pathogenesis. Several obvious examples are conditions for which inhibition of
inflammatory mediators have been successful in treatment (e.g, arthritis). However,
inflammation is now shown to be involved in several other conditions. The list includes
Parkinson’s disease (Barcia et al., 2003), diabetes (Tracy, 2003), obesity (Cottam et al.,
2004), cardiovascular diseases (Willerson and Ridker, 2004) and some types of cancer
(De Marzo et al., 2007;Whitcomb, 2004).

Exposure to large amounts of LPS, such as in sepsis, can result in damage to several
organs including the liver (Hewett and Roth, 1993c). LPS activates toll-like receptors on
macrophages, thus resulting in a cascade of inflammatory events. This renders LPS an
effective and commonly used model inflammagen in animal studies. The following
section will discuss the roles of some inflammatory mediators in the pathogenesis of liver
injury, with LPS-induced liver injury as a model. The involvement of such mediators in

liver toxicity from selected xenobiotic agents will also be mentioned.

1.2.2 TNFa
TNFa production is triggered by LPS in several cell types, in the liver especially by

Kupffer cells (the liver resident macrophage) (Hewett and Roth, 1993b). This cytokine

l3

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acts at two receptors (TNFR 1 and TNF R 2) present on many cell types to initiate cell
death signaling, promote inflammatory mediator release, increase expression of nitric
oxide sysnthesis 2, inducible (iNOS), activate the hemostatic system and induce cell
proliferation (Hehlgans and Pfeffer, 2005;Vassalli, 1992).

Expression of TNFa mRNA increases shortly after administration of LPS, and the
concentration of TNFu protein in blood rises within an hour (Hewett and Roth,
1993a). TNFa-neutralizing antibodies and inhibition of TNFu synthesis significantly
attenuate LPS—induced injury in rodents. In contrast, inhibition of TNFa biosynthesis or
neutralization of TNFa does not influence hepatic PMN‘ accumulation, suggesting either
that TNFa injures the liver after exposure to LPS by affecting activation or function of
PMNs (eg, release of cytotoxic proteases) or that it acts in a PMN-independent manner.

TNFa has been identified as the most important pro-inflammatory cytokine of the
acute inflammatory response as well as a major component in the pathogenesis of the
septic shock syndrome (Tartaglia et al., 1993;Rietschel et al., 1996). Indeed, TNFa
infusion into blood circulation leads to a sepsis-like syndrome in “rats (Tracey et al.,
1986). Administration of anti-TNFa antibodies in baboons protects from lethal
bacteraemia triggered by infusion of live Escherichia coli (Tracey et al., 1987). In these
models, TNFR 1 is essential in mediating TNF signaling, since TNFR l-deficient mice
are protected from LPS/D-galactasomine (D-GalN) and Staphylococcus aureus
superantigen/ D-GalN-induced sepsis shock (Pfeffer et al., 1993). However, in other

experimental models using concanavalin A (Con A) or Pseudomonas exotoxin A-induced

14

 

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hepatitis, TNF R 2 signalling seems to be important for the host damaging effects (Kusters
et al., 1997;Schumann et al., 1998).

In addition to its role in promoting tissue injury, TNFa can also have protective
influence. For example, the cecal ligation and puncture (CLP) model of abdominal sepsis
generates invasion of gut-derived bacteria into the blood stream and into organs, causing
septic-like syndromes. Studies employing this model suggested that TNFa is important
for recovery and survival from septic peritonitis independent of its affect on lymphocytes
(Echtenacher et al., 1995;Echtenacher et al., 1990). Furthermore, activation of the TNFR
II by endogenous TNFa constitutes an important interaction for the development of
LPS-induced resistance to bacterial infection (Echtenacher and Manuel, 2002). In
addition, a state of immunoparalysis characterized by a reduced production of TNFa
develops after CLP, which results in bacterial superinfection and leads to subsequent
lethality. Echtenacher and coworkers (Echtenacher et al., 2003) demonstrated in the CLP
model that TNFa administration during the phase of immunoparalysis can be beneficial
or deleterious, depending on the location of TNFa activity, timing of TNFa
administration and the type of infection. From these results, the prevalent concept
that TNFa is solely host-damaging needs to be re-evaluated, and additional studies are
required to analyze in more detail the role of TNFa in both pathogen defense and host
damage.

The signaling pathway for LPS-induced TNFa production has been extensively

studied (Kawai and Akira, 2007b;Takeda and Akira, 2005). Macrophages are the primary

15

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response immune cells to LPS. Toll-like receptor 4 (TLR4) on macrophages serves as a
pattern recognition receptor for LPS. After recognizing LPS, TLR4 causes the
recruitment of a set of intracellular Toll/IL-l receptor (TIR) domain-containing adaptors,
including myeloid differentiation factor 88 (MyD88), TIR-related adaptor inducing
interferon (TRIF) and TRIP-related adaptor molecule (TRAM). Among them, MyD88 is
a universal adaptor that activates inflammatory signaling pathways. The association of
TLR4 and MyD88 stimulates the recruitment of members of the IL-1 receptor-associated
kinase (IRAK) family, especially IRAK4. Once phosphorylated, IRAK dissociates from
MyD88 and interacts with TNF receptor-associated factor 6 (TRAF6). TRAF 6, an E3
ligase, forms a complex with ch 13 to promote the synthesis of polyubiquitin chains,
which in turn activate TGF-B-activated kinase 1 (TAKl), a MAPK kinase kinase. TAKl,
in combination with TAKl-binding protein (TAB), activates two downstream pathways
involving the IkappaB kinase (IKK) complex and the MAPK family (P38, JNK, ERK).
The phosphorylation of IKB by the IKK complex is necessary for the degradation of IKB
and the subsequent nuclear translocation of the transcription factor NF-KB, which
controls the expression of various inflammatory cytokine genes including TNFa. MAPK
can phosphorylate and activate the transcription factor AP-l, which has a central role in
inflammatory cytokine synthesis. In addition, MAPK (especially P38) can control
cytokine production in a post-transcriptional manner by increasing mRNA stability and
mRNA translation (Kotlyarov et al., 1999;Mavropoulos et al., 2005;Neininger et al.,

2002a;Neininger et al., 2002b).

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1.2.3 Neutrophils (PMNs)

PMNs are involved in producing liver injury induced from large, hepatotoxic doses
of LPS (Hewett et al., 1992). After exposure of rodents to LPS, various PMN
chemoattractants such as cytokine-induced neutrophil chemoattractant-l (CINC-l) and
macrophage inflammatory protein-2 (MIP-2) are elevated in plasma, and adhesion
molecules become upregulated on the surface of sinusoidal endothelial cells (SECS),
PMNs and hepatocytes (Jaeschke et al., 1996a). The upregulation of these chemokines
and adhesion molecules facilitates PMN transmigration across the endothelial cell barrier
and promotes subsequent localization of PMNs close to hepatocytes (Springer,
l994a;Butcher, 1991b). This transmigration and activation process involves rolling,
binding to and migrating across the endothelium. The migration of neutrophils from
the vasculature is regulated by at least three distinct molecular signals: selectin and its
receptor (ie., carbohydrate), integrin and its receptors (ie., immunoglobulin family) and
chemokines and their receptors. A key feature is that
selectin-carbohydrate, integrin-immunoglobulin family and chemoattractant-receptor,
interactions act in sequence, not in parallel (Butcher, 1991a;Ley, 1996). Selectins initiate
the rolling across the endothelium while integrins causes firm adhesion. Integrins and
chemokines comprise the driving force for the transmigration into parenchyma
(Kobayashi, 2008). This concept of sequential action has been confirmed by the
observation that inhibition of any one of these steps gives essentially complete, rather

than partial, inhibition of neutrophil emigration.

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When activated, PMNs release numerous cytotoxic factors, including reactive
oxygen species and PMN proteases (Jaeschke et al., 2002;Ganey et al., 1994c). Among
the latter, cathespsin G and elastase are important mediators of hepatic parenchymal cell
killing by activated PMNs in vitro (Ho et al., 1996a). Release of these factors is involved
in several pathological conditions such as acute lung injury, endotoxemia and
ischemia-reperfusion injury (Jaeschke et al., 1991;Jaeschke et al., 1990;Kawabata et al.,
2002). In addition, PMNs are critical mediators of injury in models of
endotoxin-potentiated hepatotoxicity, such as from aflatoxin Bl, monocrotaline and
trovoflaxacin (Barton et al., 2000b;Yee et al., 2003e;Waring et al., 2006b).

There exists controversy regarding the mechanisms of neutrophil-induced cell
injury in the liver. Most coculture experiments in vitro using activated neutrophils and
cultured hepatocytes showed that protease inhibitors but not antioxidant enzymes were
able to prevent the neutrophil-mediated cell injury, which develops over 15—20 h (Mavier
et al., 1988;Harbrecht et al., 1993;Ganey et al., 1994b). This time frame is much longer
than that by which hepatocellular injury develops in vivo in LPS-treated rats.
Furthermore, neutrophils may be less likely to attack and kill normal hepatocytes in vivo
since during inflammatory pathogenesis hepatocytes are exposed to other stress factors
(Gujral et al., 2004a) that are not reflected in most in vitro coculture systems. There are
studies in vivo showing that PMN protease inhibitors protect the liver from injury caused
by ischemia-reperfiision. However, this attenuation of liver injury was accompanied by a

decrease in chemokines and hepatic PMN accumulation, so that the protection effect

18

 

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might be due to inhibitions of actions other than direct hepatocyte killing by the proteases
(Soejima et al., 1999;Yamaguchi et al., 1997;Yamaguchi et al., 1999). Other studies
showed that PMNs can kill hepatocytes in vivo within 1 h, which correlates with the
appearance of an intracellular oxidant stress and the formation of hypochlorite-mediated
chlorotyrosine protein adducts (Hasegawa et al., 2005;Jaeschke et al., 1999a;Gujral et al.,
2004b). Furthermore, an inhibitor of PMN NADPH oxidase significantly delayed liver
injury in galactosoamine—sensitized mice given LPS (Gujral et al., 2004c). Mice deficient
in glutathione peroxidase-l showed enhanced susceptibility against
neutrophil-mediated cytotoxicity (Jaeschke et al., 1999b). These studies underline the
crucial role of the PMN-dependent oxidative burst in the pathogenesis of liver injury, but
they do not rule out the possibility of PMN protease killing of hepatocytes in vivo. The
longer time required for protease mediated killing of hepatocytes in vitro suggests that
PMN proteases might act in concert with other inflammatory mediators to damage
hepatocytes in vivo. These mediators could be either PMN-dependent or not. Previous
work showed that PMNs stimulated with fMLP plus PMA to release proteases and ROS,
respectively, kill hepatocytes with a timecourse similar with that of protease alone
(Ganey et al., l994a). This result suggests that an interaction of proteases either with
other, non ROS inflammatory mediators from other cell types or stresses not directly
related to PMNs that alter hepatocellular homeostasis are necessary for full manifestation

of LPS-induced liver injury.

19

 

hemOSlaSlS
factors in C0
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1.2.4 The hemostatic system and hypoxia

The coagulation and fibrinolytic systems are important controllers of vascular
hemostasis (Levi et al., 2003d). Tissue factor and factor VII are important initiating
factors in coagulation, and through a complex cascade they activate factor X. Factor X in
turn cleaves and activates thrombin. Thrombin is a protease that can cleave fibrinogen
into fibrin. Fibrin, upon cross-linking and polymerization, forms clots in blood vessels.
This activation of the coagulation system is involved in a variety of disease conditions
such as thrombosis and disseminated intravascular coagulation (DIC) (Levi, 2005b;Levi
et al., 2003c).

The formation of fibrin clots is tightly regulated by the interplay between the
coagulation and fibrinolytic systems. Plasminogen activators (PAS), including urokinase
PA and tissue-specific PA, are important activators of the fibrinolytic agent plasmin by
cleaving plasminogen into its active form. Plasmin can cleave and dissolve crosslinked
fibrin. The activity of plasminogen activator is controlled by endogenous plasminogen
activator inhibitor-1 (PAI- l ).

PAI-l is synthesized de novo by a variety of cells in vitro (e.g., hepatocytes,
adipocytes, endothelial cells, cardiac myocytes and platelets) (Westrick and Eitzman,
2007;Macfelda et al., 2002), but it is unclear which cell type is the major producer of
PAH in vivo. A PAI-l pool is also found in the granules of platelets, where it is stored
and could be released after vessel damage (Hoekstra et al., 2004b). Except in platelets,

transcriptional control of PAH is a pivotal mechanism in determining tissue and plasma

20

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PAI-l 0
induced

endotoxi

PAI-
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PAI-l content. Experiments in vitro have demonstrated that secretion of PAH can be
induced by a variety of agents, including TNF-a, IL-l, transforming growth factor-B,
endotoxin, very-low-density lipoprotein, glucose, glucocorticoid, and insulin. The
transcriptional induction is mediated through Spl element, hypoxia-responsive element
or SMAD 3/4 binding sites in the PAI-l-promoter (Fink et al., 2002;Hou et al., 2004).
Upregulation of PAH is associated with DIC and other thrombotic diseases (Padro et al.,
1997b;Padro et al., 1995).

PAI-l is present in 3 forms in the blood circulation: an active, an inactive, and a
latent form. The active form converts spontaneously into the latent form with an half-life
of 1 h (Hoekstra et al., 2004a). The latent form is more stable and can also be reconverted
into the active form. PAI-l is removed from the circulation by the liver and also is
inactivated by endothelium (Hekman and Loskutoff, 1985;0wensby et al., 1991).
Endothelium- or platelet-derived PAI-l is normally complexed to vitronectin, resulting in
a 2-4 fold increased half-life of PAI-l in the circulation.

Inhibition of thrombin activation significantly attenuates LPS-induced liver injury
(Hewett and Roth, l995b;Moulin et al., 2001a), but this protection is independent of
circulating fibrinogen (Hewett and Roth, l995a).This suggests that although thrombin
activation is important for liver injury from LPS, the formation of fibrin clots resulting
from fibrinogen processing is not required for the injury. Activation by thrombin of
protease activated receptor-l (PAR-1) influences inflammatory cell accumulation and/or

activation in the liver. Inhibition of thrombin reduced PMN activation as estimated by

21

 

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plasma elastase concentration (Copple et al., 2003b;Pearson et al., 1996). Furthermore,
perfusion of livers from LPS-treated rats with thrombin or the PAR-1 agonist TFLLR
caused hepatocellular injury (Moulin et al., 2001b;Copple et al., 2003a). This effect was
eliminated by prior depletion of PMNs. Thus, these results suggest that
thrombin-mediated activation of PAR-l can be important for PMN activation and that
these two events are sufficient for causing liver injury in LPS-treated rats.

Although fibrin deposition does not appear to be important for liver injury caused by
LPS treatment, other results suggest that fibrin deposition can to contribute to
hepatocellular injury (Luyendyk et al., 20040). One of the possible consequences of
intravascular fibrin deposition is tissue hypoxia caused by blood flow impaired by fibrin
clots. Hypoxia can deplete cellular ATP and interfere with intracellular pH and
homeostasis of ions such as Na+ and Ca2+, which could subsequently cause cell death
(Carini et al., 1997). In addition, hypoxia can induce the formation of reactive oxygen
species (Lluis et al., 2007;Lluis et al., 2005b). It can also activate numerous intracellular
signaling pathways related to cell stress, mostly mediated by hypoxia inducible factor-1a,
an important transcription factor responsive to hypoxia (Shibayama, 1987b).
Interestingly, breathing in a low 0; atmosphere enhanced the liver lesions in rats caused
by large, hepatotoxic dose of LPS (Shibayama, 1987a). This suggests that hypoxia might

interact with LPS-induced inflammatory mediators to cause hepatocellular injury in vivo.

22

 

L3 .‘Iec
ranitidin
1.3.1 Ran

A “i
tllZ'i-rece;
administrz
ulcers. g3:
RAN hep:

Elitfltilb I. .\l

Hutiation of
{EMU-”Tent“

ad'yersc re \

maniac

l‘ts‘.

 

 

I“) loot,

 

“Kid; 3.
h
RAN is E\

.‘\

 

1.3 Mechanisms of liver injury in models of inflammation-drug interaction:
ranitidine as example
1.3.1 Ranitidine-induced idiosyncratic hepatotoxicity in human patients

A widely used drug associated with idiosyncratic hepatotoxicity is the histamine 2
(H2)-receptor antagonist, ranitidine (RAN). RAN is available over the counter for oral
administration, or by prescription for parenteral administration for treatment of duodenal
ulcers, gastric hypersecretory diseases and gastroesophageal reflux disease. Idiosyncratic
RAN hepatotoxicity occurs in less than 0.1% of people taking the drug (Ribeiro et al.,
2000b). Most liver reactions are mild and reversible; however, extensive liver damage
and death have occurred in individuals undergoing RAN therapy (Ribeiro et al., 2000a).
Idiosyncratic hepatotoxicity to RAN typically manifests as elevations in serum markers
of hepatocellular injury with more modest increases in indicators of cholestatic injury.
These reactions are typical of IADRs, as the time of onset of hepatotoxicity relative to
initiation of therapy varies greatly. Rechallenge with RAN does not necessarily result in a
reoccurrence of toxicity (Halparin, 1984b;Hiesse et al., 1985b). Indeed, in some cases the
adverse response resolved despite continued therapy (Chung et al., 2000a). These
characteristics do not seem consistent with an adaptive immune response as the cause of
RAN IADRs.

RAN is minimally metabolized into three major metabolites in both humans and rats:
N-oxide, S-oxide and desmethyl metabolites (Chung et al., 2000b;Carey et al., 1981b).

RAN is excreted mostly unchanged in urine in humans as compared to through bile in

23

 

ratslfareyt

 

to llVCl ha
Accordingly
bioactit atie:
|

Althot:
idiosmcrati l
of PAN it

endotoxemi.

“3196a \ 0m

 

 

 

tnttamrnatio
Obsen Cd pr

PAN IADI

rats (Carey et al., l981a;Huang et al., 2005). However, no known RAN metabolite toxic
to liver has been reported either in human or rodent animals to our knowledge.
Accordingly, it seems unlikely that IADRs caused by RAN are due to metabolic
bioactivation of this drug.

Although direct evidence implicating inflammation as a contributing factor to these
idiosyncratic reactions is lacking, it is interesting that a majority of 34 human case reports
of RAN idiosyncratic hepatotoxicity mentioned prodromal signs consistent with
endotoxemia (i.e. increased LPS conc. in blood) or inflammation (e.g. diarrhea, fever,
nausea/vomiting, and/or abdominal pain) (Luyendyk et al., 2003a). The ability of modest
inflammation to potentiate the toxicity of numerous xenobiotic agents, along with
observed prodromal inflammatory signs in RAN idiosyncrasy, led us to hypothesize that
RAN IADRs can result from episodes of mild inflammation occurring during drug

therapy.

1.3.2 Inflammation-ranitidine interaction

Although RAN alone is not hepatotoxic in rats, studies in rats have demonstrated
that modest underlying inflammation induced by a nontoxic dose of LPS precipitates
IADR-like liver injury from RAN (Luyendyk et al., 2003b). This was not the case with
another histamine-2 receptor antagonist, famotidine (F AM), which does not share RAN’s
propensity to cause idiosyncratic adverse reactions in human patients. This suggested that

the pharmacological target is not the toxic target in terms of LPS/RAN-induced liver

24

 

3:?) O
b} HjJe'
hismmlne
H3468???
“15 n01 it
but insut‘l'.

PAN
mdzonal.
with a he?
might iner
The liver :
maximizes
comprised
liter injuq
expression
enhanced 11
One of the

negative re

 

 

 

injury. Other studies have shown that histamine attenuates LPS-induced TNFa synthesis
by H2-receptor-dependent mechanism in monocyte (Vannier et al., 1991). Furthermore,
histamine affords protection against P.acnes/LPS-and ethanol-induced liver injury by a
H2-receptor dependent mechanism (Yokoyama et al., 2004;Homyak et al., 2003). This
was not inconsistent with our findings since H2-receptor blockade might be a required
but insufficient factor for LPS/RAN-induced liver injury,

RAN given to rats 2hr after a nonhepatotoxic dose of LPS caused an acute,
midzonal, suppurative, necrotizing hepatitis that resembled lesions in animals treated
with a hepatotoxic dose of LPS (Luyendyk et al., 2003c). This result suggested that RAN
might increase hepatic parenchymal cell sensitivity to an LPS-like hepatotoxic response.
The liver injury in LPS/RAN cotreated rats begins at 2-3 hours after RAN treatment,
maximizes at 6hr, and is sustained for at least 24 hours. Hepatic inflammatory infiltrates
comprised predominately PMNs, suggesting the possibility of a role for these cells in the
liver injury (Luyendyk et al., 2005c). In addition, a gene array study showed that
expression of several genes involved in the hemostatic system and hypoxia were greatly
enhanced in LPS/RAN-cotreated rats compared to either LPS or RAN treatment alone.
One of these genes was PAI-l (Luyendyk et al., 2004p). Since PAl-l is an important
negative regulator of the fibrinolytic system, its selectively enhance expression suggested
the possibility of hemostatic system involvement. Furthermore, the crucial role of TNFa
and other cytokines in large-dose LPS-induced liver toxicity raised the possibility of the

importance of inflammatory cytokines in the LPS/RAN model. The following sections

25

 

will re\ 1:
discuss t

LPS RAF

thrombin-
the heme
Plaéma I
SIEUSQid-«i
I325 Hedi;
flit-tinehti.
PIEVemed
Slfiem in
ll'lle ”Sit I1 ‘5'
FIODQUDCQ
lilllSSi'm'h-I}~
Ron h

mgsered b

hepamclte

 

will review the evidence that supports the roles of these inflammatory factors and will
discuss how they interact with each other to contribute to the liver injury caused by

LPS/RAN cotreatment.

1.3.3 Involvement of hemostasis, PMNs and TNFa

In rats, LPS alone caused a mild and transient increase in plasma
thrombin-antithrombin dimer (TAT) and PAI-l concentration, suggesting activation of
the hemostatic system. RAN cotreatment augmented and sustained the increases in
plasma TAT and PAI-l (Luyendyk et al., 2004n). Consistent with this, significant
sinusoidal fibrin deposition occurred only in livers of LPS/RAN-treated rats relative to
rats treated with LPS or RAN alone. The anticoagulant agent heparin and the
fibrinolytic agent streptokinase, both of which reduced hepatic fibrin deposition,
prevented the liver injury induced by LPS/RAN, suggesting a role for the hemostatic
system in this model of idiosyncrasy-like liver injury (Luyendyk et al., 2004m).
Interestingly, the activation of the coagulation system in LPS/RAN cotreated rats is more
pronounced than that in LPS/F AM cotreated rats (Luyendyk et al., 2006a), suggesting the
possibility of a role for the coagulation system in the idiosyncratic liver injury caused by
RAN. It appears that RAN selectively augments the coagulation system activation
triggered by LPS, and this in turn causes subsequent fibrin deposition, tissue hypoxia and

hepatocyte death. Indeed, hepatic hypoxia occurs in LPS/RAN treated rats and heparin

26

u I
re: uceo

Inner OI“

On

LPS RA.

OCCUFI'CII

endothel.

COH‘CCII If.

addlm e ;

 

 

reduced the tissue hypoxia as well as fibrin deposition (Luyendyk et al., 2005d). The
latter observation suggested that fibrin deposition leads to liver hypoxia in this model.

One possible contributor to coagulation system activation ad PAI-l production in
LPS/RAN-treated rats could be endothelial cell activation. Previous results suggested the
occurrence of an early, transient effect of RAN and a prolonged LPS effect on sinusoidal
endothelial (SEC) function, reflected as an increase in serum hyaluronic acid (HA)
concentration (Luyendyk et al., 20041). The effects of the two agents appeared to be
additive at earlier times in LPS/RAN-cotreated rats. The early changes in serum HA
concentration were not accompanied by decreased rat endothelial cell immunostaining of
liver sections, suggesting the absence of overt SEC destruction. Overall, these results
suggested that SEC dysfunction in LPS/RAN-treated rats occurred to a greater degree
than in LPS/Veh-treated rats at a time prior to liver injury. Although its contribution is
not fully understood, a perturbation of SEC function might be important for the
coagulation system activation, PAI-l production and subsequent liver injury after
LPS/RAN treatment. For example, endothelial cells activated after LPS exposure could
express tissue factor and thereby initiate the coagulation cascade (Carnerer et al., 1996).
In addition, HA has been reported to enhance expression of PAH , thus causing sustained
fibrin deposition (Horton etal., 1999).

Data presented in this thesis will confirm and explore an essential role for PMNs in
liver injury caused by LPS/RAN. In the LPS/RAN model, PMN accumulation in liver is

caused largely by LPS, evidenced by the same degree of PMN accumulation in the livers

27

 

of rats U r:;'
:lllilSCl. l1;
night or SJ
hteractit t

laCl; 01‘ l v.
actit‘ated i."
fact that '_‘t
ll’S con p.
RAX [TC ‘1:
what: >‘,
lOltajimg ‘
through ‘ 1:
HIP-2) 2 r1N
1“ PM a.

PAH C tr.

 

of rats treated with LPS/RAN and as in those treated only with LPS (Luyendyk et al.,
2005c). The mechanism of initial recruitment of neutrophils into liver is unknown, but it
might occur through cytokines released from Kupffer cells activated by LPS, from
interaction of adhesion molecules (e.g., selectins) or from sinusoidal contraction. The
lack of liver injury after LPS treatment suggests that PMNs are not extravasated and
activated in liver after exposure to the noninjurious LPS dose used in these studies. The
fact that PMNs accumulate in the liver to about the same degree in rats treated only with
LPS compared to LPS/RAN indicates that PMNs need a secondary signal provided by
RAN treatment to be activated and cause their damaging effects. RAN itself does not
activate PMNs directly; in fact, it inhibits PMN activation both in vitro and in vivo
(Okajima et al., 2000d;Okajima et al., 2002d). This suggests that RAN acts indirectly
through other inflammatory mediators.These mediators might be PMN chemokines (i.e.,
MIP-2) and/or hemostatic factors like PAl-l since both have been shown to be involved
in PMN activation (Lentsch et al., 1998;Li et al., 2004;Maher et al., 1997). In addition,
PAI-l can potentiate LPS-induced PMN activation in vitro (Kwak et al., 2006c).
Interestingly, mRNA and serum protein concentration of both MIP-2 and PAI-l are more
upregulated after treatment with LPS/RAN than after LPS only at a time before liver
injury onset (Luyendyk et al., 2006h), and this increase is unique to RAN compared to
FAM, raising the possibility of MIP-2 and/or PAI-l as signals to activate PMNs in this

model.

28

decree;
LPS-in.
' epatoc
concentr
increase '
délelope‘
nor PrOd u
LPS'SllmL
one that d
ll-t’i. IL.1

 

 

 

Since TNFa is a critical cytokine involved in large-dose, LPS—induced liver injury,
the effects of LPS/RAN treatment on TNFu production was examined. At a nontoxic
dose, LPS rapidly induced serum TNFa production but did not cause liver injury (Tukov
et al., 20073). The serum TNFu concentration peaked at about 1.5 hours and rapidly
decreased after that, returning almost to normal by 8 hours. This indicates that an
LPS-induced increase in TNFa of this magnitude and duration is not sufficient to cause
hepatocellular damage. Despite not being damaging by itself, this large increase in TNFa
concentration could be critical for the pathogenesis of liver injury when combined with
other effects of RAN cotreatment. Indeed, RAN-cotreatment caused the serum TNFu
increase to last longer than in rats given LPS alone, and these RAN-cotreated rats
developed hepatotoxicity. In contrast, FAM neither enhanced TNFa production
nor produced liver injury when administered with LPS. Thus, the prolongation of
LPS-stimulated TNFa production distinguished a drug that causes human IADRs from
one that does not. Interestingly, some TNFa -dependent cytokines/chemokines, such as
IL-6, IL-lB and MIP-2 also had the same pattern of prolonged increase after RAN
cotreatment (Tukov et al., 2007r).

Kupffer cells (KCs) are a primary response cell to endotoxin challenge in liver. They
are also the major sources of many inflammatory cytokines/chemokines. Some of the
inflammatory mediator expression induced by xenobiotic/LPS coexposures in rats could
be recapitulated using a KC/HPC coculture system (Tukov et al., 2007q). Indeed, RAN

enhanced TNFa release in the presence of LPS in KC/HPC cocultures, but only at a large

29

 

‘4—
l
s .- “,5“ ' "h‘."n'.3‘3’ 9V! '

 

.1
I
‘-II

drug tre
that cau:
lllP-B p

here Ll

IPS PM

 

 

RAN concentration (0.5mM) that is unlikely to be achieved in the LPS/RAN animal
model. FAM had a similar effect on TNFa release in KC/HPC coculture (Tukov et al.,
2007p). However, RAN but not FAM caused a prolonged increase in serum TNFa after
drug treatment in vivo. Thus, this cell-based system in vitro failed to distinguish a drug
that causes IADRs in humans from one that does not. RAN had no effect on LPS-induced
MIP-2 production in KC/HPC coculture. This also differed from the response in vivo,
where LPS/RAN treatment caused a selective upregulation of serum MIP-2 compared to
LPS/FAM treatment (Luyendyk et al., 2006). The disparity between these responses in
vitro and in vivo suggests that the RAN-induced enhancement of TNFu and MIP-2
production seen in vivo likely occurs by mechanisms involving cell types other than or in
addition to KCs and the inflammatory mediators those cells produce. The exact
mechanism of RAN augmented TNFa production after LPS treatment in vivo remains
unknown.

To determine the role of TNFa in LPS/RAN-induced hepatotoxicity, both
pentoxyfylline (PTX) and etanercept (Etan) were used to reduce TNFa. PTX is a
methylxanthine that inhibits the synthesis of TNFa (Dezube et al., 1993;Barton et al.,
2001;Yee et al., 2003d). However, PTX also has several other pharmacological effects.
Accordingly, Etan was also used. It is a dimeric fusion protein that contains a soluble
TNFa receptor capable of selectively neutralizing serum TNFa. PTX and Etan
significantly reduced serum TNFa concentration and activity, respectively, and both were

effective in reducing hepatocellular injury in LPS/RAN cotreated rats (Tukov et al.,

30

 

20070). These results indicate that TNFa is critically involved in LPS/RAN-induced
liver injury. However, since PTX and Etan were given before the LPS challenge, those
results could not differentiate whether LPS-induced TNFa elevation or the RAN-induced

prolongation of the TNFu response was crucial for the pathogenesis.

1.3.4 Interaction of TNFa, hemostasis and PMNs

In LPS/RAN—cotreated rats, TNFu inhibition leads to decreases in PMN chemokines
such as MIP-2 and in plasma markers of hemostasis, such as TAT and PAI-l (Tukov et
al., 2007n). This suggests that TNFa is a more proximal inflammatory mediator in the
pathogeneic cascade relative to the hemostatic system and PMNs. Indeed, TNFa
inhibition reduced hepatic fibrin deposition, indicating that TNFa acts upstream of the
hemostatic system. How exactly TNFa contributes to hemostatic system activation
remains unknown in this model. One possibility is that TNFa could act on vascular
endothelial cells or on macrophages to induce tissue factor expression (Parry and
Mackman, 1995;Bierhaus et al., 1995;Schwager and Jungi, 1994). In addition, in the
presence of PMNs, TNFa exposure can cause sinusoidal endothelial cell (SEC) damage
in vitro (Smedly et al., 1986;Takei et al., 1995), which can activate the coagulation
system. TNFa and IL-1 can also stimulate the expression and release of PAH by
endothelial cells in vitro (Fan et al., 2000b). The contribution of TNFd to tissue factor
production and inhibition of fibrinolysis has also been demonstrated in vivo in a model of

lung hemorrhagic shock (Fan et al., 2000a). Thus, TNFa could cause the disruption of the

31

 

 

u-Ji

hemosuatic
PAl-l exp:
TNFu .
lBrad-ham
tSchleiffe r.
1993). In I
accumulat.
Klimt 1.
be dcpeng’j

heparin sir

 

0n hepatic
imitation
RCllVare p)
Inhib-
resulm‘ fl

pmucllor

hemostatic system and subsequent fibrin deposition by inducing either tissue factor or
PAI-l expression or both.

TNFa can prompt the accumulation of PMNs in tissues by activating endothelial cells
(Bradham et al., 1998;Vassalli et al., 1992) and can also prime PMNs for activation
(Schleiffenbaum and Fehr, 1990;Kushimoto et al., 1996;Nagaki et al., 1991;Vassalli,
1992). In LPS/RAN-cotreated rats, neutralization of TNFa did not impact hepatic PMN
accumulation but did reduce serum MIP-2 and PAI-l concentration (Tukov et al.,
2007m). These results suggest that the signal for PMN extravasation and activation might
be dependent on TNFa, whereas PMN accumulation is not. Interestingly, anticoagulant
heparin similarly reduced serum MIP-2 and PAI-l concentration while having no effect
on hepatic PMN accumulation (Luyendyk et al., 2006b). This suggests that TNFa
activation of coagulation induces the expression of MIP-2 and PAL], which could
activate PMNs accumulated in the liver.

Inhibition of coagulation did not reduce serum TNFa concentration (unpublished
results), firrther supporting the idea that coagulation activation lies downstream of TNFa
production. Coagulation activation is not the cause of the initial neutrophil accumulation,
since heparin treatment did not impact hepatic PMN accumulation, as mentioned above.
However, coagulation activation might elicit PMN activation in the LPS/RAN model, a
possibility suggested by the finding that heparin treatment reduced serum MIP-2
concentration. This could happen in several ways. One possibility is that coagulation

activation causes activation of PAR-l, the receptor for thrombin on Kupffer cells,

32

endotheli.

to contri

chemokir

occlusit e

at chemo

lPS R-IX

Call llltlllt't

P-selectin

 

:llllPLSl'lIL‘
I
Hansml‘lla

cliposure :

exposure .1

 

Min.
Hot
isms \i‘
lelOI 5.
rel’t'fred
lilo Pl

'llatada

   
   
 

l .3
h ”it???“ ‘\

Ulllfl mu.

endothelial cells and hepatic stellate cells (Copple et al., 2003c). PAR-1 has been shown
to contribute to PMN activation in an indirect manner through releasing PMN
chemokines (Copple et al., 2003d). Another possibility is that hypoxia caused by
occlusive fibrin clots in liver sinusoids promotes PMN activation by altering expression
of chemokines and adhesion molecules. As mentioned above, hypoxia occurs in
LPS/RAN treated rats before the onset of liver injury (Luyendyk et al., 2004k). Hypoxia
can induce chemokines such as MIP-2 or adhesion molecules such as ICAM-1 and
P-selectin in endothelial cells and/or hepatocytes (Pinsky et al., 1996;Laurens et al.,
2005;Shreeniwas et al., 1992;Xu et al., 1999). These actions would favor the adhesion,
transmigration and activation of PMNs. Indeed, PMNs isolated from humans after acute
exposure to a hypoxic atmosphere showed increased activation compared to normoxia
exposure as evidenced by superoxide anion production and elastase release (Harada et al.,
1999a)

How PMNs contribute to the hepatotoxicity in this model remains unknown. It
seems likely that PMNs are involved in TNFu production since Kupffer cells are the
major source of TNFa in liver. Both coagulation system activation and PMNs are
required for producing liver injury in this LPS/RAN model. In other models of liver
injury, PMN activation can enhance activation of the coagulation system and vice versa
(Harada et al., 1999b;Yee et al., 2003c). This raised the possibility of interplay between
the hemostatic system and PMNs in LPS/RAN-induced liver injury. In fact, PMNs in

other models are involved in coagulation system activation, fibrinolysis and platelet

33

 

actix’atior
:003d;Pi
plasma 1‘;
prozhmm
:DOBCLA}
C153)? an.
dulQij

In 3d
Such €15 re
Interesting
bipoxic e:
killing 0“
mm In I
and hm“
PMNS Cor
ant-1mm

As 8
INF“. \K

 

[TukOV e.

 

2% Rs}.

 

activation (Kimura and Yokoi-Hayashi, l996a;Wu et al., l995a;Goel and Diamond,
2003d;Pintucci et al., 1992a). For instance, the PMN protease cathepsin G can cleave
plasma factor V and factor X to induce thrombin activation, and it also can promote
prothrombinase and fibrin formation by activating platelets (Goel and Diamond,
2003c;Allen and Tracy, 1995;Sambrano et al., 2000). In addition, PMN proteases can
cleave and release active PAI-l from endothelial matrix to inhibit fibrinlysis (Pintucci et
al., 1992b;Kimura and Yokoi-Hayashi, 1996b).

In addition to their role in hemostasis, PMNs upon activation release toxic products
such as reactive oxygen species and proteases, which can directly damage hepatocytes.
Interestingly, killing of hepatocytes in vitro by neutrophil elastase is enhanced by a
hypoxic environment (Luyendyk et al., 2005b). The time course over which hepatocyte
killing occurred during hypoxia in vitro (ie, 2hr) is consistent with the onset of liver
injury in LPS/RAN-treated rats. Taken together with the evidence that fibrin deposition
and hypoxia are critical to liver injury in LPS/RAN-treated rats, it seems possible that
PMNs contribute to the pathogenesis by releasing proteases that kill hepatocytes in an
environment made hypoxic by fibrin deposition or other events.

As summarized in Fig 1.1, LPS causes an early and transient increase in serum
TNFa, which is prolonged by RAN treatment. TNFa causes coagulation system
activation and PAI-l production, leading to subsequent fibrin deposition and hypoxia
(Tukov et al., 20071;Luyendyk et al., 2004j). LPS also causes PMN accumulation in liver,

and RAN might indirectly activate these cells, which can release toxic ROS and

34

My“.

 

Gui-I

pI’O-ICESC:
activatio
lluyendj

mmnhm

 

 

proteases. TNFa dependent production of MIP—2 and PAI-l might contribute to PMN
activation. Hypoxia might act synergistically with PMN proteases to kill hepatocytes
(Luyendyk et al., 2005a). In addition to its direct killing of hepatocytes, PMNs might

contribute to fibrin deposition and hypoxia.

35

 

Figure 1.1 Possible pathogenic mechanism of LPS/RAN-induced liver injury. LPS
causes an early and transient increase in serum TNFa. which is prolonged by RAN
treatment. TNFa causes coagulation system activation and PAl-l production, leading to
subsequent fibrin deposition and hypoxia (Tukov et al., 2007k;Luyendyk et al., 2004i).
LPS also causes PMN accumulation in liver, and RAN might indirectly activate these
cells, which can release toxic ROS and proteases. TNFa dependent production of MIP-2
and PAI-l might contribute to PMN activation. Hypoxia might act synergistically with
PMN proteases to kill hepatocytes (Luyendyk et al., 2005f). In addition to its direct

killing of hepatocytes, PMNs might contribute to fibrin deposition and hypoxia.

   

TF \
LPS Thrombin
\*:: Fibrin
RAN \TNF\ /:;; as

 

PAI- 1

 
 
   

**

p rotease

37

 

l.-I Dic
.\'\
many a
hut sor
apparen
l to 2 c
ntxnber
lll'allter.
In a large
2003;. T]
Presentari
human ll\
JOil‘lllct'de
SCVer
mClUding

lCiirthnj c

 

 

1.4 Diclofenac-induced idiosyncratic hepatotoxicity

Non steroidal anti-inflammatory drugs (NSAle) comprise another class of drugs,
many of which cause hepatic IADRs. For example, diclofenac (DCLF) has caused rare
but sometimes serious hepatotoxicity in humans (Boelsterli, 2003f). Although the
apparent incidence of severe DCLF-induced hepatic adverse reactions is quite low (from
1 to 2 cases per million prescriptions or 6 to 18 cases/ 100,000 person-year), the large
number of patients treated with DCLF makes the absolute number of cases impressive
(Walker, 1997b). In addition, cases of severe injury leading to liver transplantation occur
in a large proportion of the reported cases of DCLF-induced hepatotoxicity (Lewis et al.,
2003). The scarceness of liver biopsies from patients and the diverse histopathlogical
presentations in available samples make it difficult to draw clues about mechanisms from
human liver pathology alone (Zimmerman et al., 1999). Thus, the pathogenesis of this
low-incidence/ high-severity DCLF hepatotoxicity is largely unknown.

Several mechanisms for diclofenac-induced hepatotoxicity have been proposed,
including formation of reactive drug metabolites (Lim et al., 2006), oxidative stress
(Cantoni et al., 2003a) and mitochondrial injury (Masubuchi et al., 2002a). However,
cytotoxocity of DCLF to hepatocytes is only observed in vitro with large concentrations
that are not achieved in vivo. In human patients developing hepatotoxicity after DCLF,
polymorphisms in enzymes that metabolize DCLF, such as UGTZB7 and CYP2C8, and
in the DCLF glucuronide transporter ABCC2 have been discovered (Daly et al., 2007).

These could lead to increased formation of hydroxylated and/or glucuronidated DCLF

38

metaboll
relations.

are repo

al- 300}
(KIIII 1:
cells i501
Plellousl

no animct

in the ll\

 

 

metabolites. However, no direct evidence has been found to support the a causal
relationship between these DCLF metabolites and hepatotoxicity in vivo, although there
are reports that DCLF acyl glucuronide is directly involved in small intestinal injury in
rats (Seitz and Boelsterli, 1998d).

There is some evidence suggesting immune-modulated hypersensitivity in
diclofenac-induced hepatotoxicity. For example, a 53-year-old female developed
fulminant hepatic failure caused by inadvertent rechallenge with diclofenac (Greaves et
al., 2001 a). DCLF-conjugated mouse serum albumin or keyhole limpet hemocyanin
(KLH) is immunogenic in mice (Kretz-Rommel and Boelsterli, 1995). Furthermore, T
cells isolated from DCLF-KLH conjugate-treated mice were able to kill hepatocytes
previously exposed in vitro to noncytotoxic concentrations of DCLF. However, there are
no animal models which have reproduced toxic DCLF-induced immunoallergic reactions
in the liver. In fact, the T cell popliteal lymph node reaction was attenuated if animals
were treated orally with diclofenac as compared to direct injection into the footpad
(Gutting et al., 2003;Gutting et al., 2002). Since DCLF is normally given orally to human
patients, this raises a question about the likelihood that an immunoallergic reaction
contributes to DCLF-induced IADRs. DCLF autoantibodies have been found in human
patients, but the relevance of those autoantibodies to idiosyncratic hepatotoxicity is
unknown (Aithal et al., 2004a). Thus, the lack of a demonstrable connection between an
immunoallergic response to DCLF and hepatotoxicity raises a question as to whether

DCLF IADRs occur by this mechanism.

39

Ba;
llDRs i
a modes
of DC Ll
mixed I
gastroim
extreme
laborato:
induce s
there's} j
KIOICOK‘
aI‘hritis.
0f lnflar

llIlllan.

 

dEVElUP;

 

 

Based on the observation that a number of drugs that have the ability to cause
IADRs in human patients also cause liver injury in LPS-treated rats, we hypothesized that
a modest inflammatory episode increases the sensitivity of the liver to hepatotoxic effects
of DCLF. This merits special attention since most selective cyclooxygenase (COX)-1 and
mixed COX-l/COX-2 inhibitors, including diclofenac, have been associated with
gastrointestinal injury, including increased permeability of the mucosa, ulceration, and, in
extreme cases, perforation of the small intestinal wall. This holds true for
laboratory rodents, in which a single, small dose of diclofenac (e.g., 1.5 mg/kg) can
induce small intestinal ulceration (Atchison et al., 2000;Seitz and Boelsterli, 1998c) and
thereby promote LPS and or bacterial translocation from the intestine into the circulation.
Moreover, DCLF is normally prescribed to patients with inflammatory diseases such as
arthritis, in which inflammatory mediators (ie.TNF, PMNs) are usually present. The role
of inflammatory stress in DCLF hepatotoxicity is further suggested by the clinical
findings that IL-4 and IL-10 polymorphisms have been found in human patients who
developed hepatotoxicity after DCLF treatment (Aithal et al., 2004b).

The potentiation of DCLF induced hepatotoxicity by LPS might appear paradoxical,
since DCLF acts as a NSAID to dampen inflammatory responses. COX catalyzes
production of prostaglandins (PGs) and thromboxanes (Tx) from arachidonic acid. PGs
and sz have diverse biological actions, either proinflarnmatory or anti-inflammatory.
However, as a nonselective COX inhibitor (i.e., inhibits both isoforrns of the enzyme,

COX-1 and COX-2), DCLF inhibits the synthesis of cytoprotective and

40

“ .lAIP

 

Emil-1111131

- J‘ 5' i
mouttble

DCLF xvi

 

 

anti-inflammatory PGs produced by constitutive COX activity as well as products of
inducible COX-2, which are mostly proinflammatory. Moreover, the toxic interaction of
DCLF with LPS might arise from inflammatory mediators (eg., cytokines) that are not

products of the COX pathway and are therefore not inhibited by NSAIDs.

41

  

matlum
be impor
30052)

i
4

their int:

model. I;
I0 cause
hepatoc;

POSshnt:

 

1.5 Overview of the thesis
1.5.1 Knowledge gaps

As mentioned above, LPS/RAN treatment caused an idiosyncrasy-like liver injury in
rats (Luyendyk et al., 2003d). Both PMNs and the hemostatic system have been shown to
be important in LPS/RAN-induced liver injury (Luyenkyk et al., 2004; (Luyendyk et al.,
2005g). However, how PMNs participate in the liver injury induced by LPS/RAN and
their interactions with the hemostatic system have not been elucidated. Moreover, how
PMNs are recruited and whether they become activated in the LPS/RAN model remains
unknown. Since fibrin deposition and hypoxia appear to be important in the LPS/RAN
model, PMNs might contribute to liver injury by interacting with the hemostatic system
to cause fibrin deposition. On the other hand, PMN-derived proteases might directly kill
hepatocytes in a hypoxic environment. The data presented in this thesis will explore these
possibilities. The results will contribute to the understanding of mechanisms by which
inflammation acts as a susceptibility factor for toxicity due to drugs and other xenobiotic
agents. In addition, the findings could lead to improved strategies for preventing or
treating conditions involving PMN-dependent tissue injury.

TNF-a has proved to be important in LPS/RAN-induced hepatotoxicity (Tukov et
al., 2007j). Activation of p38 and MAPKAPK-2 can be involved in the production of
this and other proinflarnmatory cytokines. Thus, p38 activation could be a potential
upstream signal that leads to production of cytokines/chemokines and to subsequent

cascades contributing to LPS/RAN-induced liver injury (Luyendyk et al., 2006g). Hence,

42

inflamr.
hepLIIOII

tux, a

 

 

we investigated the role of P38 activation in LPS/RAN-induced hepatotoxicity. These
studies will begin to explore the intracellular signaling crucial to the initiation of
LPS/RAN induced-liver injury.

There are no animal models that reproduce DCLF-induced idiosyncratic
hepatotoxicity in vivo. Based on the observation that a number of drugs that have the
ability to cause IADRs in human patients also cause liver injury in LPS-treated rats, we
tested the hypothesis that a modest inflammatory episode in rats increases the sensitivity
of the liver to hepatotoxic effects of DCLF. DCLF is often given to patients with
inflammatory disease such as arthritis, in which PMNs comprise at least some of the
inflammatory infiltrates. Furthermore, PMNs are involved in other idiosyncratic
hepatotoxicity models based on LPS-drug interactions. This leads us to hypothesize that

PMNs are important in the hepatotoxicity induced by LPS-DCLF interaction.

1.5.2 Hypothesis and specific aims

Overall Hypothesis: Neutrophils are activated in part by p38 dependent TNF
production in the livers of rats cotreated with LPS and an IADR-producing drug, and
these cells contribute to liver injury by enhancing hemostasis and/or releasing neutrophil
proteases. This general hypothesis will be addressed by specific hypotheses represented

in 3 specific aims:

43

lll'f".
and or:
{Che ter

lin
injur cu

\ir‘.‘

 

 

Aim 1 Hypothesis: Neutrophils are activated in the livers of LPS/RAN treated rats
and contribute to liver injury by enhancing hemostasis and/or releasing proteases.
(Chapter 2)

Aim 2 Hypothesis: p38 contributes to neutrophil activation and subsequent liver
injury caused by LPS/RAN cotreatment by inducing TNFu. (Chapter 3)

Aim 3 Hypothesis: LPS cotreament with a nontoxic dose of DCLF causes
hepatocellular injury in rats and this LPS/DCLF interaction is PMN dependent. (Chapter

4)

44

CHAPTER TWO
Xiaomin Deng, James P. Luyendyk, Wei Zou, Jingtao Lu, Ernst Malle, Patricia E.
Ganey, Robert A. Roth.(2007) Neutrophil interaction with the hemostatic system

contributes to liver injury in rats cotreated with lipopolysaccharide and ranitidine. J

Pharmacol Exp Ther. 322 (2): 852-61.

45

2.1 .-‘ bstr

.otr
lll’f l CL:
idios net
the l em
l1}p( he~
prod tcir;
anti- {\t‘

redu ed

 

2.1 Abstract

Cotreatment of rats with nontoxic doses of ranitidine (RAN) and lipopolysaccharide
(LPS) causes liver injury, and this drug-inflammation interaction might be a model for
idiosyncratic adverse drug responses (IADRs) in humans. Both neutrophils (PMNs) and
the hemostatic system have been shown to be important in the injury. We tested the
hypothesis that PMNs cause liver injury by interacting with the hemostatic system and
producing subsequent hypoxia. In rats cotreated with LPS/RAN, PMN depletion by
anti-PMN serum reduced fibrin deposition and hypoxia in the liver. PMN depletion also
reduced the plasma concentration of active plasminogen activator inhibitor-1 (PAI-l), a
major downregulator of the fibrinolytic system. This suggests that PMNs promote fibrin
deposition by increasing PAI-l concentration. PMNs were activated in the livers of
LPS/RAN-cotreated rats as evidenced by increased staining for hypochlorous acid
(HOCl)-modified proteins generated by the myeloperoxidase-hydrogen peroxide-chloride
system of activated phagocytes. Antiserum against the PMN adhesion molecule CD18
protected against LPS/RAN-induced liver injury. Since CD18 is important for PMN
transmigration and activation, these results suggest that PMN activation is required for
the liver injury. Furthermore, anti-CD18 serum reduced biomarkers of hemostasis and
hypoxia, suggesting the necessity for PMN activation in the interaction between PMNs
and the hemostatic system/hypoxia. Liver injury, liver fibrin and plasma PAI-l
concentration were also reduced by eglin C, an inhibitor of proteases released by

activated PMNs. In summary, PMNs are activated in LPS/RAN-cotreated rats and

46

parti ipu'

 

 

participate in the liver injury in part by contributing to hemostasis and hypoxia.

47

2.2 Intrr

In r.
injury re
causes ir
and the 3
al“ :0")

contribt'

 

 

 

2.2 Introduction

In rats, cotreatment with lipopolysaccharide (LPS) and ranitidine (RAN) causes liver
injury resembling hepatotoxic idiosyncratic adverse drug responses (IADRs) that RAN
causes in humans (Luyendyk et al., 2003c). Both polymorphonuclear neutrophils (PMNs)
and the hemostatic system are important in LPS/RAN-induced liver injury (Luyendyk et
al., 2005h;Luyendyk et al., 2004b). Hemostasis-associated fibrin deposition likely
contributes to injury in this model by causing liver hypoxia (Luyendyk et al., 2005i).

The hemostatic system is tightly regulated by the interplay between the coagulation
and fibrinolytic systems (Lasne et al., 2006). Tissue factor is the principal initiator of the
coagulation system, a complex cascade that ultimately generates active thrombin.
Thrombin cleaves circulating fibrinogen into fibrin monomers, which upon cross-linking
and polymerization can form obstructive clots in blood vessels. Plasminogen activators
(PAS), including urokinase and tissue-specific PA, are important proteolytic activators
of plasmin, which cleaves and dissolves crosslinked fibrin. The activity of plasminogen
activators is inhibited by plasminogen activator inhibitor-1 (PAI-l) (Keller et al.,
2006;Padro et al., 1997a).

PMNs usually require transmigration across the endothelial barrier and subsequent
activation to kill pathogens or injure tissues (Springer, 1995). These cytotoxic effects are
mediated in part by release of reactive oxygen species and/or granular proteases
(Jaeschke et al., l996b). PMN derived-proteases such as elastase and cathepsin G kill

hepatocytes directly in vitro (Hill and Roth, 1998;Ho et al., 1996b). Moreover, the killing

48

the hem
are 3Cll\

releasing

 

 

of hepatocytes by PMN-derived proteases is potentiated by hypoxia (Luyendyk et al.,
2005j).

In the LPS/RAN model of hepatotoxic drug-inflammation interaction, PMNs
accumulate in liver, but how they participate in the pathogenesis and their relationship to
the hemostatic system are unknown. Here we tested the hypothesis that hepatic PMNs
are activated in the livers of LPS/RAN-cotreated rats and contribute to liver injury by

releasing proteases and interacting with the hemostatic system.

49

2.3 )Ietl

2.3.1 Mu

louis. .\
number
El' mg

determirl

Bio Sclc

23.2 ml

Ma
Laborat.
Chou T;

T36) Vt k

 

 

hepi-‘IIOC ,\

 

 

2.3 Methods
2.3.1 Materials

Unless otherwise noted, all chemicals were purchased from Sigrna-Aldrich (St.
Louis, MO). Two lots of LPS derived from Escherichia Coli serotype 055:85 (catalog
number L-2880) with activities of 6.6 x 106 EU/mg (lot number 51K4115) and 13 x 106
EU/mg (lot number 43K4l 12) were used for these studies. These activities were
determined using a QCL Chromogenic LAL endpoint assay purchased from Cambrex

Bio Science, Inc. (Baltimore, MD).

2.3.2 Animals

Male, Sprague-Dawley rats [CrlzCD (SD)IGS BR; Charles River Breeding
Laboratories, Portage, MI] weighing 250 to 350 g were fed standard chow (rodent
chow/Tek 8640; Harlan Teklad, Madison, WI) and allowed access to water ad libitum.

They were allowed to acclimate for 1 week in a 12h light/dark cycle prior to use.

2.3.3. Experimental Protocol

Rats fasted for 24hr were given 2.5 x 106 EU/kg LPS (lot 43K4112) or its saline
vehicle (Veh) i.v. at 5ml/kg, and food was then returned. For the 6hr CD18 antiserum
study, 44.4 x 106 EU/kg LPS (lot 51K4115) or its vehicle at 2ml/kg was given. These
two LPS doses from different lots render approximately the same degree of

hepatocellular injury in terms of serum alanine aminotransferase (ALT) activity. Two

50

hours later, 30 mg/kg RAN or its vehicle [sterile phosphate-buffered saline (PBS)] was
administered (i.v.). RAN solution was administered at 2 kag at a rate of approximately
0.15 ml/min. Two, three or six hr after RAN administration, rats were anesthetized with
sodium pentobarbital (75 mg/kg i.p.). Plasma was collected by drawing 2 ml of blood
from the vena cava into a syringe containing sodium citrate (final concentration, 0.38%).
Another portion of blood was collected and allowed to clot at room temperature; serum
was prepared from this fraction and stored at -20°C until use. Representative slices
(3—4-mm thick) of the ventral portion of the left lateral liver lobe were collected and
fixed in 10% (v/v) neutral buffered formalin. For immunohistochemical analysis, a
portion of the left medial lobe of the liver was flash-frozen in isopentane cooled by liquid

nitrogen.

2.3.4 PMN Depletion, CD18 Neutralization and Eglin C Treatment

PMN depletion was accomplished by administration of a PMN antiserum before
treatment with LPS/RAN. Rats were given 0.25 ml of either normal rabbit serum (CS) or
rabbit anti-rat PMN serum (NAS) (Intercell Technologies, Jupiter, FL) diluted 1:1 in
sterile saline (0.5 ml per rat, iv.) 18 hr before administration of LPS. A previous study in
which anti-PMN serum was administered to rats demonstrated a selective and complete
depletion of PMNs (Snipes et al., 1995). Rats were killed 2 and 6 hr after treatment with
RAN, and citrated plasma was collected. Total blood leukocytes were quantified using a

Unopette white blood cell determination kit (BD Biosciences, San Jose, CA) and a

51

hemt :}1
min? 1g
perfc rr.1

or
‘peci 1c
Pept: le
lVVl ir.
injec ior
plasr ta
alter R“.

"Pro: ido

imm or.

 

 

hemocytometer. Slides were prepared from whole blood and stained using the Hema 3
staining system (Fisher Diagnostics, Middletown, VA), and differential counting was
performed.

For the CD18 neutralization study, rabbit anti-CD18 serum raised against rat CD18
specific peptide (Ac-CLKFDKGPFQKN-amide) was obtained from New England
Peptide (Gardner, MA). The anti-CD18 serum or normal rabbit serum was diluted 1:1
(v/v) in sterile saline and administered (0.5ml per rat, i.v.) immediately after LPS
injection. Rats were killed 6hr after RAN treatment for evaluation of serum ALT activity,
plasma PAI-l concentration, liver PMN accumulation and fibrin. Rats were killed 3hr
after RAN for hypoxia and PMN activation evaluation. For the eglin C study, eglin C
(provided by Novartis Pharm AG, Basel, Switzerland) was given at 8mg/kg (i.v.)

immediately after RAN, and 2hr and 4hr later. Rats were killed 6hr after RAN treatment.

2.3.5 Hepatotoxicity Assessment

Hepatic parenchymal cell injury was estimated as an increase in serum ALT activity.
ALT activity was determined spectrophotometrically using Infinity-ALT reagent from
Thermo Electron Corporation (Louisville, CO). Previous studies in this LPS/RAN model
have shown that serum ALT activity reflects histopathologic evidence of hepatocellular

necrosis (Luyendyk et al., 2003f).

2.3.6 Fibrin Immunohistochemistry and Quantification

52

pret

6X0

stai: .

posi

usir

l D61

lCflL

CORT ~

CT)
com

lLsin

{SOL 1:

it»

V6

EI

0';
\

 

 

 

Fibrin immunohistochemistry and quantification were performed as described
previously (Copple et al., 2002a). This protocol solubilizes all fibrinogen and fibrin
except for cross-linked fibrin (Schnitt et al., 1993a). Therefore, only cross-linked fibrin
stains in sections of liver. The positive area fraction refers to the percentage of area with

positive staining in the total microscopic field.

2.3.7 Evaluation of Coagulation System Activation and Plasma PAl-l

Plasma fibrinogen was determined from thrombin clotting time of diluted samples
using a fibrometer and a commercially available kit (B4233) from Dade Behring Inc.
(Deerfield, IL). Plasma thrombin-antithrombin (TAT) concentration was determined by
an enzyme-linked immunosorbent assay (ELISA) using a kit from Dade Behring Inc.
(catalog number OWMG15). Total plasma PAI-l concentration was evaluated using a
commercially available ELISA purchased from American Diagnostica Inc. (Greenwich,
CT). This ELISA measures PAI-l in active, inactive, and plasminogen activator/PAI-l
complexed forms. The concentration of functionally active PAI-l in plasma was assessed
using a commercially available ELISA kit purchased from Molecular Innovations Inc.

(Southfield, MI).

2.3.8 Evaluation of Liver Hypoxia
Liver hypoxia was evaluated by immunohistochemical staining for pimonidazole

(PIM) adducts. PIM is a 2-nitroimidazole marker of hypoxia and has been used to

53

 

identif;
other“
C hemi.

intm un

lSClon
intensit
occurs;

protein

 

 

 

2Dl ljl

gene!

..
‘L.

identify regions of hypoxia in liver (Arteel et al., 1995;Arteel et al., 1998b). Unless
otherwise noted, rats were given 120 mg/kg Hypoxyprobe-l (PIM hydrochloride;
Chemicon International, Temecula, CA) i.p. 2 hr before they were killed. PIM-adduct
immunostaining was performed as described previously (Copple et al., 2004).
Quantification of immunostaining was performed using Scion Image Beta 4.0.2 software
(Scion Corporation, Frederick, MD). Background was set to be the average pixel
intensity in periportal regions of VehNeh-treated livers (i.e., an area where no hypoxia
occurs) (Arteel et al., 1998a). An increase in positive immunostaining for PIM-modified

proteins indicates hypoxia in the liver tissue.

2.3.9 Evaluation of Circulating Leukocytes, PMNs, Liver PMNs and PMN
Activation.

Total blood leukocyte count was quantified using a Unopette white blood cell
determination kit (BD Biosciences, San Jose, CA) and a hemocytometer. Slides were
prepared from whole blood and stained using the Hema 3 staining system (Fisher
Diagnostics, Middletown, VA), and differential counting was performed. PMNs
accumulated in liver were visualized by immunohistochemical staining and quantified as
described previously (Yee et al., 2003b). PMN activation was measured by staining of
hypochlorous acid (HOCl)—protein adducts in liver. The monoclonal antibody (clone
2D10G9, subtype IgG2bk) is specific for hypochlorous acid (HOCl)—modified epitopes

generated in vivo (Malle et al., 1997a) and in vitro (Malle et al., l995a) and does not

54

CIOSSICS
in 4° 0 l‘
l

W61? \\

blocked

Molecu
hr at rt
examin
and pr:
Staining

0le PC f

 

 

crossreact with other oxidative protein modifications. Frozen sections of liver were fixed
in 4% (v/v) formalin for 10 minutes at room temperature with gentle rocking. The slides
were washed 3 times, 5 minutes each, with PBS (phosphate-buffered saline), then
blocked for 1 hour at room temperature with 3% goat serum in PBS. Antibody (diluted
1:1 in 3% goat serum) was added and incubated for 2 hr at room temperature with gentle
rocking. The slides were washed 3 times, 5 minutes each, with PBS. Alexa Fluro
488-labeled goat anti-mouse secondary antibody (diluted 1:500 in 3% goat serum,
Molecular Probes, Carlsbad, California) was applied, and the slides were incubated for 3
hr at room temperature. After washing 3 times, 5 minutes each, with PBS, they were
examined microscopically. Staining was quantified as for fibrin staining described above
and presented as positive area fraction. The integrated density within areas of positive
staining was calculated as the sum of the products of pixel gray intensity and pixel

numbers with that intensity.

2.3.10 Evaluation of PMN Protease Effects on F ibrinogen and PAI-l in vitro

Normal rat plasma was incubated with a mixture of elastase and cathepsin G or
vehicle (PBS) at two different conditions: elastase (0.04U/mL)/cathepsin G (4ug/ml) and
elastase (8U/ml)/ cathepsinG (4ug/ml). Elastase activity was determined using the
colorometric PMN elastase substrate, MeOSuc-Ala-Ala—Pro-Val-pNA (Calbiochem, San
Diego, CA). One unit of PMN elastase activity was defined as the amount of enzyme

required to cause a change of absorbance of 1.0 at 410 nm in 10 min at 37°C. After 2hr

55

PAl-l n
elastase
adding

ELISA
“as in.
elastase
and It

on 4-1‘

1
CAL

2.3.11 y
Ft
time 31‘.

Vla'v Ax

 

latilOrg
I651 “it

lOF all

\

 

and 6hr incubation, fibrinogen was measured by fibrometer as described above. Purified
PAI-l purchased from American Diagnostica Inc. (Greenwich, CT) was incubated with
elastase and cathepsin G under the same conditions, and the reaction was stopped by
adding anti-trypsin (40nM) after 2 or 6hr. The total PAI-l level was measured by PAI-l
ELISA as described above. For the SDS-PAGE analysis, purified fibrinogen (250ug/ml)
was incubated with PBS, thrombin (0.1U/ml), elastase (8U/ml)/cathepsin G (4ug/ml) or
elastase (0.04U/ml)/cathepsin G (4ug/ml). The reaction was stopped by addingl% SDS
and 2% 2-mercaptoethanol after 2 or 6hr. The resulting protein products were separated
on 4-12% gradient SDS-PAGE and visualized with silver staining (Biorad, Hercules,

CA).

2.3.11 Statistical Analysis

For the PMN depletion study and in vitro study, two way ANOVA was applied with
time and treatment as the two factors. For the CD18 neutralization and eglin C study, two
way ANOVA was used with LPS/RAN and drug (anti-CD18 serum or eglin C) as the two
factors. For the ALT data from the eglin C study, a student’s t-test was applied. Tukey
test was used as post-hoe analysis for ANOVA. The criterion for statistical significance

for all studies was p<0.05.

56

 

6h

Tal

LP! Kl

in )
LPS L
i
eml
l

17‘
.IT]: P

 

 

 

 

 

 

 

 

 

 

 

 

 

Time after RAN Treatment Leukocytes(#/ul) PMNs(#/ul)
CS/Veh/Veh 4012+457 776+277

2hr CS/LPS/RAN 4640+628 2482+661 *
NAS/LPS/RAN 1560+181* # 75+17* #
CS/Veh/Veh 4138+819 497+153

6hr CS/LPS/RAN 4236+591 2594+559
NAS/LPS/RAN 1812+368* # 344+121#

 

Table 2.1. Circulating leukocytes

LPS/RAN-treated rats. PMN depletion of LPS/RAN-treated rats was accomplished as
in Methods. Circulating total leukocytes and PMNs were measured at 2hr and 6hr after
LPS/RAN treatment as described in Methods. *significantly different from CS/Veh/Veh

group at the same time. #significantly different from CS/LPS/RAN group at the same

time. P<0.05, n=4-7.

57

after PMN antiserum

 

treatment

 

uerer
its co:
Circul.
RIM
reiuce.
or 6hr
lPS R.
IllOtlUCt
act‘llmu
All ac

lPS RA

36mm

 

(lulu.
CS

 

QEIWSII f

Coat‘rl.

\“l;

 

2.4 Results
2.4.1 PMN depletion studies in the LPS/RAN model.

To assess the role of PMNs in hemostasis and hepatocellular injury, PMN numbers
were reduced prior to LPS/RAN-cotreatment. Rabbit anti-rat PMN antiserum (NAS) or
its control normal rabbit serum (CS) was administered 16 h before LPS treatment.
Circulating total leukocytes and PMN and liver PMNs were evaluated at 2hr and 6hr after
RAN treatment. As shown previously (Luyendyk et al., 2005k), the antiserum markedly
reduced blood PMNs but did not significantly affect other blood leukocytes at either 2hr
or 6hr (Table. 2.1). It also significantly reduced liver PMN accumulation caused by
LPS/RAN treatment at both 2hr and 6hr (data not shown). In addition, the antiserum
produced a qualitative change in liver PMNs, causing a shift toward the hepatic
accumulation of immature PMNs (i.e. band cells). Antiserum alone did not cause serum
ALT activity to increase either at 2hr or 6hr. Serum ALT activity was unaffected by
LPS/RAN treatment at 2hr after RAN administration but was significantly elevated by
6hr (Fig. 2.1). Treatment with the antiserum significantly decreased ALT activity in the
serum of LPS/RAN-treated rats at 6 hr after RAN, confirming previous results
(Luyendyk et al., 20051).

CS/LPS/RAN treatment caused a significant increase in liver fibrin compared to
CS/Veh/Veh control both at 2hr and 6hr (Fig. 2.2). NAS pretreatment did not affect fibrin
deposition at 2hr after RAN but significantly reduced it at 6hr. Activation of the

coagulation system leads to a decrease in plasma fibrinogen, as this protein is cleaved to

58

 

Figure 2.1. Effects of PMN depletion on hepatotoxicity induced by LPS/RAN. PMN
antiserum (NAS) or control normal serum (CS) was administered 16 hr before LPS or its
vehicle. Two hours after LPS treatment, RAN or its vehicle was administered. Serum
ALT activity was measured 2hr and 6hr after administration of RAN or its vehicle. *
significantly different from the respective group without LPS/RAN at the same time. #
significantly different from the respective group without CS at the same time. a,

significantly different from the same treatment group at 2hr. P< 0.05, n= 4-7.

59

IAN. I’ll.)

 

800 w

 

 

 

 

 

 

 

rights — CSNehNeh 3*
A :3 NASNehNeh
ed. Serum g — CSILPSIRAN
v 600 [:1 NAS/LPSIRAN
\ehicle.‘ E
..>.
. ‘6
jcltmtlt N 400 _
5
[11115-3- < a*#
E
a 200 -
d)
"’ ll
, jui
2hr 6hr

60

Figure 2.2. Effects of PMN depletion on liver fibrin after LPS/RAN treatment. PMN
depletion of LPS/RAN-treated rats was accomplished as in Fig 2.1. Fibrin staining was
performed 2hr and 6hr after treatment with RAN or its vehicle treatment. * significantly

different from CS/Veh/Veh group at the same time. # significantly different from

CS/LPS/RAN group at the same time. P< 0.05, n= 4-7.

61

Fibrin (Positive Area Fraction)

.030 -

.025 -

.020 ~

.015 -

.010 -

.005 -

 

 

_ CSNehNeh
m CSILPSIRAN
— NASILPSIRAN

 

 

 

0.000

 

2hr

62

 

6hr

 

Figure 2.3. Effects of PMN depletion on coagulation system activation after
LPS/RAN treatment. PMN depletion of LPS/RAN-treated rats was accomplished as in
Fig 2.1. Plasma fibrinogen and TAT concentration were measured 2hr and 6hr after RAN
or vehicle treatment. * significantly different from CS/Veh/Veh group at the same time. #

significantly different from CS/LPS/RAN group at the same time. P< 0.05, n= 4-7.

63

 

— CSNehNeh
It: CSILPSIRAN

300
l — NAS/LPSIRAN

 

 

 

250 -

N

O

O
1

Fibrinogcn (mg/L)
'3‘

 

 

 

 

 

 

 

 

 

 

 

 

 

 

100 ‘
50 -
Ition after
0
lished 35311 2hr 6hr
alierllll B
. e, — CSNehNeh
”W“ so * 2::3 CSILPSIRAN
l — NAS/LPSIRAN
.7, *
*
*
60 a
Q
3’
v 40 -‘
I'-
<
I—
20 t -.
0
2hr 6hr

64

Figure 2.4. Effects of PMN proteases on plasma fibrinogen in vitro. A: elastase and
cathepsin G or their vehicle (PBS) was incubated with plasma fibrinogen as described in
Methods. Fibrinogcn was measured using a fibrometer 2hr and 6hr later. *. significantly
different from PBS group. #. significantly different from elastase (0.04U/ml)/cathepsin G
treatment. P< 0.05, n= 4. B: thrombin, elastase/cathepsin G or their vehicle (PBS) was
incubated with purified fibrinogen, and the protein products were analyzed by
SDS-PAGE. Lane 1,2: fibrinogen+PBS. 2hr and 6hr respectively; lane 3.4.5: fibrinogen +
thrombin, 2hr; lane 6,7: fibrinogen + elastase(8U/ml) /cathepsin G(4ug/ml), 2hr and 6hr;
lane 8,9: fibrinogen + elastase(0.04U/ml)/cathepsin G(4ug/ml), 2hr and 6hr. Molecular

weight was marked at the right margin (kDa).

65

Issue in!
:scn'ool lfl
znii‘ieantl)‘
:hepsinli
1351 no
zed l‘.‘
Hagen '
IlCl f‘l‘tfi

.jecular

Fibrinogen(mgldL)

300

250

200 -

150

100 -

50-

 

— PBS
:21 Elastase(0.04U/mI)ICathepsin G
- EIastase(8U/ml)lCathepsin G

 

 

 

 

 

2hr 6hr

 

66

form insoluble fibrin clots. At both 2hr and 6hr, LPS/RAN treatment decreased plasma
fibrinogen and increased plasma TAT, a biomarker of coagulation system activation.
This suggests early and persistent activation of the coagulation system (Fig. 2.3). In
PMN-depleted rats, there was a more modest decrease in plasma fibrinogen (Fig. 3A), but
the elevation in TAT was unaffected (Fig. 2.38).

In vitro, incubation of PMN elastase/cathepsinG with fibrinogen reduced
fibrinogen concentration after 2 or 6hr of incubation (Fig. 2.4A). These results suggest
that PMN proteases cleave and inactivate fibrinogen. SDS-PAGE of fibrinogen produced
three predominant bands identified by silver staining, with molecular weights of
approximately 62 kD, 53 kD, 47 kD (Fig. 2.4B). These are likely the Au (62Kd), BB
(53Kd) and y chains (47Kd) of fibrinogen as described by Gorkun et al (Gorkun et al.,
1997). There was also a minor band, which is likely the smaller Aa chain lacking a
C-terminal fragment. Thrombin treatment reduced the size of the smaller Au and BB
chains, probably due to the release of fibrin peptides A and B, respectively (Gorkun et al.,
1994;Weisel et al., 1993). Elastase (0.04U/ml) and cathepsin G (4ug/ml) treatment
caused the degradation of the smaller Aa chain and the formation of degradation products
of smaller size. Increasing the concentration of elastase in the mixture resulted in the
degradation of BB, 7 and the smaller size Au chain and the formation of degradation
products of smaller size.

PAI-l is the major enodogenous downregulator of the fibrinolytic system, which

lyses insoluble fibrin clots and thereby controls the degree of fibrin deposition. Total

67

PAI-l includes active PAI-l, inactive PAH and PAI-l complexed with plasminogen
activator. LPS/RAN treatment caused a significant increase in the plasma concentrations
of both total and active PAI—l at 2hr and 6hr after RAN (Fig. 2.5). PMN depletion did not
affect total plasma PAI-l concentration at 6hr but increased it at 2hr (Fig. 2.5A). The
increase in total plasma PAI-l concentration with PMN depletion suggested that
PMN-derived proteases might degrade PAI-l. Indeed, incubation of purified PAI-l with
elastase/cathepsinG reduced total PAI-l concentration (Fig. 2.6). On the other hand,
PMN depletion did not significantly affect active PAI-l concentration at 2hr but
decreased it at 6hr (Fig. 2.58).

Since fibrin deposition and hemostasis can lead to tissue hypoxia,
immunohistochemistry of pimonidazole (PIM)-bound protein was used as a marker for
liver hypoxia. LPS/RAN treatment caused a significant increase in PIM-protein adduct
staining at 2hr after RAN, and this was prevented by NAS treatment (Fig. 2.7). Hypoxia
staining in LPS/RAN treated livers tended to be panlobular, as compared to the necrotic

foci which were predominately midzonal.

2.4.2 HOCl-protein adduct staining.

Activation of PMNs is required for many of their damaging effects on host tissue.
The potent oxidant HOCI, generated in vivo by the myeloperoxidase-hydrogen
peroxide-chloride system of activated phagocytes, reacts with proteins to form

chloramines that may be used as a biomarker for PMN activation. Neither LPS alone nor

68

Figure 2.5 Effects of PMN depletion on plasma PAI-l concentration after LPS/RAN

treatment. PMN depletion of LPS/RAN-treated rats was accomplished as in Fig 2.1.

 

Plasma active PAI-l (A) and total PAI-l (B) was measured 2hr and 6hr after LPS/RAN
treatment as described in Methods. * significantly different from CS/Veh/Veh group at
the same time. # significantly different from CS/LPS/RAN group at the same time. a,

significantly different from the same treatment group at 2hr. P< 0.05, n= 4-7.

69

 

— CSNehNeh
CSILPSIRAN

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2500 r — NASILPSIRAN
*

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Bi
5
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<
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Ta

5 *
,_ coo -

I

<

n.

m 400 ‘

I2

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0

< zoor

 

 

 

 

CD

 

2hr 6hr

70

 

Figure 2.6. Effects of PMN proteases on PAI-l concentration in vitro. Elastase and
cathepsin G or their vehicle was incubated with purified PAI-l as described in Methods.
Total PAl-l concentration was measured by ELISA 2hr and 6hr later. * significantly
different from vehicle group. # significantly different from elastase (0.04U/ml)/Cathepsin

G treatment. P< 0.05, n= 4.

7l

Total PAl-1 (nglml)

 

 

- PBS
E23 Elastase (0.04UImI)/Cathespin G(4uglml)
— Elastase (BU/ml) ICathespin G (4uglml)

 

 

 

 

 

 

72

Figure 2.7. Effect of PMN depletion on liver hypoxia after LPS/RAN treatment.
PMN depletion of LPS/RAN-treated rats was accomplished as in Fig 2.1. PIM was
administered to rats 1hr after RAN, and the rats were killed at 2hr after RAN. *
significantly different from CS/Veh/Veh group. # significantly different from

CS/LPS/RAN group. P< 0.05, n= 4-7.

73

 

 

— CSIVehNeh
CSILPSIRAN
— NASILPSIRAN

 

 

 

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T . . 4 a n

50505050
332211

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74

RAN alone affected HOCl-protein adduct staining (Fig. 2.SB,C,E). LPS/RAN
cotreatment increased the staining, indicating the activation of PMNs in
LPS/RAN-treated livers (Fig. 2.8D,E). The HOCl-protein epitopes were predominatly
within the midzonal areas of liver lobules, which was where the necrotic foci occurred in
livers of LPS/RAN-cotreated rats. This increase was apparent 3-6hr after RAN but not at

211: (Fig. 2.8F).

2.4.3 Effects of CD18 neutralization on hepatotoxicity, PMNs and hemostasis.

CD18 is an adhesion molecule expressed on PMN plasma membranes that is
important for the transmigration and subsequent activation of these cells (Springer,
l994b). Based on staining for HOCl-modified epitopes (Fig. 2.8), it was clear that PMNs
are activated in livers of LPS/RAN-treated rats. To investigate the role of PMN activation
in the LPS/RAN model, anti-CD18 serum was administered immediately after LPS, and
various biomarkers were evaluated 6hr after RAN. The anti-CD18 serum did not reduce
the number of circulating PMNs after LPS/RAN treatment (data not shown). LPS/RAN
caused an increase in PMNs accumulated in liver (Fig. 2.9A). Anti-CD18 serum alone
caused an increase in the number of PMNs in livers of Vetheh-treated rats but did not
affect hepatic PMN accumulation after LPS/RAN treatment. Anti-CD18 serum reduced
staining for HOCl-modified epitopes at 3hr after RAN (Fig. 2.9B) and protected from

LPS/RAN-induced hepatotoxicity, as reflected by reduction of serum ALT activity (Fig.

75

Figure 2.8. PMN activation after LPS/RAN treatment. Rats were treated with LPS
or vehicle 2hr before RAN or its vehicle treatment. HOCl-protein protein staining was
performed 3hrs after treatment with RAN or vehicle. The representative photomigraphs
for individual treatment group are presented in gray scale. A: Veh/Veh; B: Veh/RAN; C:
LPSNeh; D: LPS/RAN; E: Quantification of staining for all four treatment groups. *
significantly different from the respective group without LPS. # significantly different
from the respective group without RAN. F: Rats were treated with LPS/RAN or Veh/Veh
as described in Fig 2.1. HOCl-modifed protein staining was performed 2hr, 3hrs, 4hrs
and 6hr after LPS/RAN treatment. The Veh/Veh-treated animals were killed at 2hr after
RAN for HOCl-protein adduct staining. * significantly different from Veh/Veh group.

P<0.05, n=5-7.

76

 

S
* P
L
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9:53» 6:25 522.. Go:

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77

 

Figure 2.9. Effects of CD18 neutralization on biomarkers of hemostasis, PMNs and
hepatotoxicity in LPS/RAN-cotreated rats. Rabbit anti-C D18 serum or normal serum
was administered at the same time as LPS or its vehicle. RAN or its vehicle was
administered 2hr after LPS treatment. A: Liver PMN accumulation; B: HOCl-protein
adduct staining; C: Serum ALT activity; D: Liver fibrin; E: Plasma total PAI-l and F:
Plasma active PAI-l were evaluated 6hr after RAN or its vehicle treatment except the
HOCl-protein adduct staining was evaluated 3hrs after RAN treatment. * significantly
different from the respective group without LPS/RAN. # significantly different from. the

respective group without anti-CD18 serum. P<0.05, n=4-7.

78

fibrin I Fraction Tot! Area)

9 . . . .
8=ss s s

 

  

 

  

 

 

 

 

 

 

 

 

 

 

 

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02
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Figure 2.10. Effects of CD18 neutralization on liver hypoxia after LPS/RAN
treatment. CD18 neutralization in LPS/RAN-treated rats was accomplished as described
in Fig 2.9. PIM probe was administered 1hr after RAN treatment, and PIM-protein adduct
staining was performed 3hrs after RAN treatment. * significantly different from the

respective group without LPS/RAN. # significantly different from the respective group

without anti-CD18 serum. P<0.05, n=4-7.

80

PIM staining (positive area fraction)

.10 r

.08 J

.06 n

.04 -

.02 ~

 

 

— Normal Serum
:3 Anti-CD18 serum

 

 

 

0.00

 

 

VehNeh

81

 

 

LPS/RAN

2.9C). In addition, anti-CD18 serum decreased liver fibrin at 6hr after RAN treatment
(Fig. 2.9D).

As reported above (Fig. 2.9E and 9F), LPS/RAN treatment increased both total PAH
and active PAI-l concentration 6hr after RAN. Anti-CD18 serum did not affect the
increase in total PAl-l in plasma but reduced active PAl-l concentration. As reported
previously, liver hypoxia occurred 3hr after LPS/RAN treatment, as marked by increased
PIM protein adduct staining (Luyendyk et al., 2005 and Fig. 2.10). Cotreatment with

anti-CD18 serum prevented the liver hypoxia.

2.4.4 Effects of eglin C on serum ALT activity and hemostasis

Eglin C is a peptide inhibitor of both PMN elastase and cathepsin G (Braun et al.,
1987). It was used to investigate the contribution of PMN proteases to LPS/RAN-induced
hepatotoxicity and biomarkers of hemostasis. Eglin C administration significantly
reduced serum ALT activity, indicating attenuation of hepatocellular injury (Fig. 2.11).
Eglin C also reduced plasma active PAI-l concentration and liver fibrin after LPS/RAN
treatment (Fig. 2.12 A and B). In addition, it partially restored the fibrinogen
consumption caused by LPS/RAN treatment but did not affect plasma TAT concentration

(Fig. 2.12 c and D).

82

Figure 2.11. Effects of PMN-protease inhibitor on hepatotoxicity after LPS/RAN
treatment. Rats were treated with LPS/RAN as described in Fig 2.1. Eglin C or its
vehicle was administered 0hr. 2hr and 4hrs after RAN treatment. Serum ALT activity
was evaluated 6hr after RAN treatment. # significantly different from Veh group. P<0.05,

n=ll-12.

83

 

Eglin C

Veh

 

 

 

3:: S<

84

Figure 2.12. Effects of PMN protease inhibitor on markers for hemostasis and liver
fibrin after LPS/RAN treatment. Rats were treated with LPS/RAN or Veh/Veh as
described in Fig. 2.1. Eglin C or its vehicle was administered 0hr, 2hr, 4hr after RAN
treatment. A: Plasma active PAl-l . B: liver fibrin, C: plasma TAT and D: fibrinogen
were evaluated 6hr after RAN treatment. * significantly different from the respective
group without LPS/RAN. # significantly different from the respective group without

eglin c. P<0.05, n=5-12.

85

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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86

Discussion

In LPS/RAN-treated rats, hepatocellular injury begins 2-3hr after RAN
administration and progresses through 6hr, at which time serum ALT activity is maximal
(Luyendyk et al., 2003g). Confirming previous results (Luyendyk et al., 2005m), PMN
depletion with NAS dramatically reduced serum ALT activity 6hr after RAN, suggesting
an important role for PMNs in the liver injury. PMNs could contribute to this response by
a variety of mechanisms. One possibility is that PMNs promote coagulation and fibrin
deposition in liver, leading to hypoxia. Fibrin deposition occurs prior to injury (2-3 hr) in
liver sinusoids of LPS/RAN-cotreated rats (Luyendyk et al., 2004g). PMN depletion
decreased fibrin deposition at 6 hr after LPS/RAN treatment (Fig. 2.2) but not at 2 hr.
This result suggests that PMN—dependent fibrin deposition contributes to the progression
but not to the initiation of hepatocellular injury in LPS/RAN-treated rats. To investigate
how PMNs contribute to fibrin deposition in the LPS/RAN model, coagulation system
activation and PAI-l were evaluated. Interestingly, PMN depletion attenuated the
decrease in plasma fibrinogen caused by LPS/RAN (Fig. 2.3A) but did not influence the
elevation in plasma TAT concentration (Fig. 2.3B). Since TAT concentration reflects
generation of active thrombin, this result suggests that PMNs do not participate in
activation of the coagulation system but are capable of thrombin-independent cleavage of
fibrinogen. The latter might be due to the action of PMN-derived proteases such as
elastase/cathepsin G. Indeed, a combination of PMN elastase and cathepsin G cleaved

fibrinogen in vitro in a manner different from thrombin (Fig. 2.4). Furthermore, eglin C

87

reduced the consumption of fibrinogen caused by LPS/RAN treatment but did not affect
plasma TAT concentration (Fig. 2.12 C and D). Taken together, these results suggest that
PMNs contribute to fibrin deposition but not via activating the coagulation cascade.

An alternative mechanism by which PMNs could contribute to deposition of fibrin is
through inhibiting fibrinolysis by increasing active PAI-l. PMN lysosomal proteases
cathepsin B, D and G increased PAI-l activity in the medium of human umbilical vein
endothelial cells by cleaving PAI-l from extracellular matrix (Kimura and
Yokoi-Hayashi, 1996c;Pintucci et al., l993a;Pintucci et al., 19920). Consistent with this,
PMN depletion reduced active PAl-l at 6hr after RAN, although it did not affect it at 2hr
(Fig. 2.5B). This suggests that PMNs contribute to fibrin deposition after 2hr by
activating PAH and thereby inhibiting fibrinolysis. In contrast, PMN depletion did not
affect total plasma PAI-l concentration at 6hr, but it did increase total plasma PAI-l
concentration at 2hr. The mechanism by which PMN depletion increases total PAI-l
concentration is unknown. PMN derived proteases such as elastase can cleave and
inactivate PAI-l (Wu et al., 1995b), which has also been shown here (Fig. 2.6). However,
this is unlikely in vivo at 2hr in the LPS/RAN model since PMNs are not activated. to
release granular products at that time (Fig. 2.8). Interestingly, internalization of
UPA-PAI-l complexes by immune cells can occur through uPA receptors (Conese et al.,
1995;Nagatomi et al., 1997a), this might explain why PMN depletion increased total

PAI-l but had no effect on active PAI-l at the earlier time (ie., 2hr).

88

One explanation for the dual effects of PMNs on active and total PAI-l is that
PMNs are not activated to release proteases during earlier times but begin to degranulate
thereafter. In this regard, the shedding of PAI-l from endothelial matrix by PMNs
proteases might play a more dominate role at later times (ie. 6hr). This is consistent with
the results showing that formation of HOCl-modified epitopes, a marker of PMN
degranulation/activation, did not increase until 3hr after LPS/RAN treatment (Fig. 2.8),
suggesting that PMNs did not start the transmigration and activation process until after
2hr.

In the LPS/RAN model, PMN accumulation in liver is caused largely by LPS .
However, LPS exposure alone did not lead to activation of the accumulated PMNs (Fig.
2.8A). RAN did cause activation of the PMNs that accumulated in liver after LPS
treatment, as reflected by an increase in HOCl—protein adduct formation. Anti-CD18
serum did not alter PMN accumulation but reduced PMN activation in liver (Fig. 2.9A,
9B), suggesting that the early, LPS-induced sinusoidal accumulation of PMNs does not
require CD18 whereas their activation is CD18-dependent. In addition, anti-CD18
serum markedly reduced hepatocellular injury (Fig. 2.9A), suggesting that PMN
transmigration/activation is required for LPS/RAN-induced liver injury.

The anti-CD18 serum and NAS had similar effects on the hemostatic system.

Anti-CD18 serum also reduced fibrin deposition 6hr after LPS/RAN treatment (Fig.
2.9D), suggesting that activation of PMNs is required for their effect on hemostasis.

Moreover, PMN proteases are important in this regard since eglin C also reduced the

89

deposition of fibrin (Fig. 2.12B). This promotion of fibrin deposition by activated PMNs
is most likely mediated by PAI-l since both anti-CD18 serum and eglin C reduced the
elevation in active PAI-l. In addition, the reduction of active PAI-l by eglin C treatment
supports the findings of others (Kimura and Yokoi-Hayashi, 1996;Pintucci et al., 1993)
that PMN proteases can cleave PAI-l from endothelial matrix and increase the
concentration of active PAI-l in plasma.

There are several reports showing that hypoxia can cause hepatocyte death in vitro
(Nagatomi et al., 1997b;Lluis et al., 2005a). In the LPS/RAN model, agents that reduce
hypoxia also reduce hepatocellular injury. These include heparin (Luyendyk et al.,
2005m), PMN antiserum (Fig. 2.7) and CD18 antiserum (Fig. 2.8). Accordingly, it seems
likely that hypoxia caused in part by sinusoidal fibrin deposition that reduces blood flow
plays a role in inducing hepatocellular injury in this model. PMNs could also contribute
to tissue hypoxia. In LPS/RAN-treated rats, PMN depletion reduced PIM staining at 2hr
after LPS/RAN treatment, suggesting that PMNs contribute to early development of liver
hypoxia. This could occur by several mechanisms. One possibility is that accumulated
PMNs could clog sinusoidal vessels and impair microvascular blood flow. Furthermore,
PMN-derived factors could impair the balance between vasodilators and vasoconstrictors,
causing constriction of sinusoids and subsequent hypoxia. For example, PMN-derived
proteases reduce vasodilatory prostacyclin production both in vitro and in vivo (Harada et
al., 1997;Okajima_ et al., 2004b;Weksler et al., 1989). Myeloperoxidase, abundantly

Present in PMNs (5% of total cell protein content) and released by PMNs during the

90

oxidative burst, could also oxidize NO and thereby impair vascular relaxation (Eiserich et
al., 2002). This early contribution of PMNs to hypoxia did not depend on sinusoidal
fibrin deposition, since PMN deletion in LPS/RAN-treated rats did not affect liver fibrin
at 2hr after RAN (Fig. 2.2). Furthermore, PMNs accumulated in livers are not activated at
2hr (Fig. 8), suggesting that enhancement of hypoxia by PMNs at this time does not
require their activation and might be mediated directly by plugging sinusoids.

LPS-RAN cotreatment caused a greater degree of tissue hypoxia than LPS given
alone (Luyendyk et al., 2004f), although these two treatments had a similar effect on
PMN accumulation (Luyendyk et al., 20050). It is possible that RAN cotreatment causes
PMNs to adhere more firmly to sinusoidal endothelium and/or to undergo a shape change
that results in reduced sinusoidal blood flow and consequent hypoxia. Interestingly, the
early tissue hypoxia and PMN accumulation were both panlobular in distribution,
whereas the hepatocellular necrosis that followed was midzonal. The HOCl adduct
staining also tended to be midzonal. These results suggest that the early hypoxia might
be necessary but insufficient to cause the liver injury without additional factors such as
PMN activation.

The contribution of PMNs to liver hypoxia at later times might occur by a different
mechanism. Anti-CD18 serum reduced liver hypoxia at 3hr, suggesting that the
contribution of PMNs to liver hypoxia after 2hr requires PMN transmigration and/0r

activation. Since sinusoidal fibrin deposition appears to be important for liver hypoxia

9]

occurring at 3hr after LPS/RAN treatment (Luyendyk et al., 2005p), PMNs might
contribute to liver hypoxia after 2hr by promoting sinusoidal fibrin deposition.

Among the products released by activated PMNs, cathepsin G and elastase are
important mediators of hepatic parenchymal cell killing caused by PMNs in vitro (Ho et
al., 1996c). These proteases also have been implicated in models of liver injury in vivo
such as ischemia-reperfusion (Okajima et al., 2004a) and LPS-induced liver dysfunction
(Luyendyk et al., 2005q). Interestingly, killing of hepatocytes in vitro by neutrophil
elastase is enhanced by a hypoxic environment (Luyendyk et al., 2005r). The time course
over which hepatocyte killing occurred during hypoxia in vitro is consistent with the
development of liver injury in LPS/RAN-treated rats. Taken together with the evidence
that fibrin deposition and hypoxia are critical to liver injury in LPS/RAN-treated rats, it is
likely that PMNs contribute to the pathogenesis by releasing proteases that kill
hepatocytes in an environment made hypoxic by fibrin deposition. Indeed, eglin C
attenuated the hepatocellular injury caused by LPS/RAN treatment. Thus, PMN proteases
could contribute to liver injury by direct killing hepatocytes and indirectly by promoting
fibrin deposition and hypoxia, thereby increasing hepatocellular susceptibility to injury.

In conclusion, PMNs and the hemostatic system are key players in the pathogenesis
of liver injury in LPS/RAN-cotreated rats. PMN accumulation in liver in this model is
mediated by LPS and is insufficient to damage hepatocytes. Activation of hepatic PMNs
by RAN occurs around the time liver injury begins and is critical for the progression of

hepatocellular injury. Morever, the protection afforded by eglin C suggests that proteases

92

released during PMN activation are important. These proteases may injure hepatocytes
directly, but they also participate in liver injury progression by promoting hemostasis, in
part by activating antifibrinolytic PAI-l. PMNs also contribute to the liver hypoxia at a
time before PMN activation is evident. In addition, during injury progression, PMNs
likely enhance tissue hypoxia by promoting hemostasis in this model of idiosyncratic

adverse drug reaction.

93

Figure 2.13. Interaction of PMNs with the hemostatic system in the LPS/RAN
model of liver injury: working hypothesis. LPS causes modest coagulation system
activation and PAI-l production, leading to fibrin deposition and hypoxia (Luyendyk et
al., 2004e). RAN augments fibrin deposition by enhancing coagulation system activation
and PAI-l. LPS also causes PMN accumulation in liver. and RAN activates these cells,
which release toxic PMN proteases (Fig 2.8). The early hypoxia (2hr) is PMN-dependent
(Fig 2.7), whereas later hypoxia is enhanced by hemostasis. Hypoxia acts synergistically
with PMN proteases to kill hepatocytes (Luyendyk et al., 20055). In addition to its direct
killing of hepatocytes, PMN proteases increase active PAl-l production and subsequent

fibrin deposition (Fig. 2.12).

94

 

 

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CHAPTER THREE
Xiaomin Deng, Jingtao Lu, Lois D.Lehman-McKeeman, Ernst Malle, David Crandall,
Patricia E.Ganey, Robert A. Roth. (2008) p38-dependent TNF-a converting enzyme is
important for liver injury in hepatotoxic interaction between lipopolysaccharide and

ranitidine. J Pharmacol Exp Ther. Submitted

96

3.1 Abstl

[dio
health an
is ranitidi
and RAX
in humai
factor-31p
and that t
hEPOthesi
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more p35
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POSI-[ram
(TACE).
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Funhem;

 

afier Lpt

3.1 Abstract

Idiosyncratic adverse drug reactions (IADRs) remain an important issue in human
health and drug development. One of the drugs associated with IADRs in human patients
is ranitidine (RAN). In rats, cotreatment with nontoxic doses of lipopolysaccharide (LPS)
and RAN causes liver injury. This is a potential animal model for RAN-induced IADRs
in humans. Previous studies showed that RAN augmented serum tumor necrosis
factor-alpha (TNF-u) production and hepatic neutrophil activation after LPS treatment
and that both TNF-a and neutrophils are crucial for the liver pathogenesis. We tested the
hypothesis that p38 is necessary for TNF-a production, neutrophil activation and
subsequent liver injury caused by LPS/RAN cotreatment. LPS/RAN cotreatment caused
more p38 activation compared with treatment only with LPS. 83239063, a p38 kinase
inhibitor, reduced liver injury in rats cotreated with LPS/RAN. This inhibitor also
reduced neutrophil activation in livers and attenuated the hemostatic system. SB239063
decreased serum TNF-a concentration afier LPS/RAN treatment to the same level as LPS
treatment. However, the inhibitor did not reduce TNF-a mRNA in liver, suggesting a
post-transcriptional mode of action. This might occur through TNF-a converting enzyme
(TACE), which cleaves pro-TNF-a into its active form. Indeed, a TACE inhibitor
administered just before RAN treatment reduced serum TNF-a protein. The TACE
inhibitor also reduced liver injury and active plasminogen activator inhibitor-1 (PAI-l).
Furthermore, a PAI-l inhibitor reduced neutrophil activation and subsequent liver injury

afier LPS/RAN treatment. In summary, RAN enhanced TNF-a production after LPS

97

treatmen

actix atio

RAN cot

 

 

treatment through augmented p38 activation, and this appears to occur through the
activation of TACE. The prolonged TNF-a production enhanced PAI-l production after

RAN cotreatment and this is important for the hepatotoxicity.

98

seemir
ofieni
rare id
(RAN)
parent:
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them 0.
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3.2 Introduction

Idiosyncratic adverse drug reactions (IADRs) occur during treatment with numerous
drugs, typically in a small fraction of people taking the drug. These responses are
seemingly unrelated to dose, and the time of onset relative to beginning of drug therapy is
often variable (Kaplowitz, 2005b;Uetrecht, 2007). A widely used drug associated with
rare idiosyncratic hepatotoxicity is the histamine 2 (H2)-receptor antagonist, ranitidine
(RAN). RAN is available over the counter for oral administration or by prescription for
parenteral administration for treatment of duodenal ulcers, gastric hypersecretory diseases
and gastroesophageal reflux disease. Idiosyncratic RAN hepatotoxicity occurs in less
than 0.1% of people taking the drug (Fisher and Le Couteur, 2001a;Fisher and Le
Couteur, 2001b). Most liver reactions are mild and reversible; however, extensive liver
damage and death have occurred in individuals undergoing RAN therapy (Cherqui et al.,
1989;Ribeiro et al., 2000c). Rechallenge with RAN does not necessarily result in a
reoccurrence of toxicity (Halparin, 1984a;Hiesse et al., 1985a).

In rats, cotreatment with nontoxic doses of lipopolysaccharide (LPS) and ranitidine
(RAN) causes liver injury. This was not the case with another histamine-2 receptor
antagonist, famotidine (FAM), which is not associated with IADRs in human patients
(Fisher and Le Couteur, 2001c). Thus, this LPS-drug interaction model in rodents could
differentiate a drug that causes IADRs from a drug that does not. Previous mechanistic
studies showed that RAN augmented serum tumor necrosis factor-alpha (TNF-a)

production and hepatic neutrophil activation after LPS treatment, and both TNF-a and

99

.\1
.21..

I161.

pro]
trans
TNF-
TNF-
protei
Come]
genera
This cl
311d in -
that inl
1994),

INN

 

 

neutrophils are crucial for the liver pathogenesis (Deng et al., 2007g;Tukov et al., 2007i).
Moreover, TNF-a is likely to be a proximal signal in the pathogenic cascade (Tukov et
al., 2007b). The mechanism behind RAN augmentation of TNF-u production and
neutrophil activation is unknown.

TNF-a production involves gene expression of proTNF-a mRNA, translation of
proTNF-a protein and its cleavage and release of active TNF-u. LPS-induced TNF-u
transcriptional activation has been well studied (Kawai and Akira, 2007a). However,
TNF-a production can also been regulated at a post-transcriptional level. For example,
TNF-a mRNA stabilization and translation are regulated by p38 mitogen activated
protein kinase (MAPK) (Hitti et al., 2006d;Neininger et al., 2002c). Furthermore, TNF-a
converting enzyme (TACE) cleaves the 26 kDa membrane-bound proTNF-a protein to
generate secreted 17 kDa mature TNF-u (Aggarwal et al., l985b;Mullberg et al., 2000a).
This cleavage occurs at the Ala76-Val77 bond. The release of TNF-u from cells in vitro
and in vivo can be selectively blocked by hydroxamate-based metalloprotease inhibitors
that inhibit TACE activity (Mohler et al., 1994b;Gearing et al., l994;McGeehan et al.,
1994). These TACE inhibitors protect against endotoxin-mediated lethality, in which
TNF-a plays a critical role (Mohler et al., 1994a).

p38 and its downstream MAPK Activated Protein Kinase 2 (MK-2) have been
shown to be involved in the production of several cytokines and chemokines (ie. TNF-a,
macrophage inflammatory protein-2 [MIP-2], interleukin 6 [IL-6]) (Hitti et al.,

2006c;Neininger et al., 2002d;Numahata et al., 2003b) and in neutrophil activation (Nick

100

:1

th

int:

inj 1

 

et al., 1997;Mocsai et al., 2000). Thus, p38 activation is a potential upstream signal that
leads to production of cytokines/chemokines and subsequently to downstream cascades
that contribute to LPS/RAN-induced liver injury (Luyendyk et al., 2006f). Here we tested
the hypothesis that p38 is necessary for TNF-a production, neutrophil activation and
subsequent liver injury caused by LPS/RAN cotreatment. These studies elucidated

intracellular signaling events that are crucial to the initiation of LPS/RAN induced-liver

injury.

101

3.3 .\1

3.3.1 I

Louis.
L-ZSS
studie:

purcha

3.3.2 A

Labo ra

Chow ’1

They \i

 

 

 

 

3.3 Methods
3.3.1 Materials

Unless otherwise noted, all chemicals were purchased from Sigma-Aldrich (St.
Louis, MO). LPS derived from Escherichia Cali serotype 0552B5 (catalog number
L-2880) with activity of 13 x 106 EU/mg (lot number 43K4112) was used for these
studies. This activity was determined using a QCL Chromogenic LAL endpoint assay

purchased from Cambrex Bio Science, Inc. (Baltimore, MD).

3.3.2 Animals

Male, Sprague-Dawley rats (CrlzCD (SD)IGS BR; Charles River Breeding
Laboratories, Portage, MI) weighing 250 to 350 g were fed standard chow (rodent
chow/Tek 8640; Harlan Teklad, Madison, WI) and allowed access to water ad libitum.

They were allowed to acclimate for 1 week in a 12h light/dark cycle prior to use.

3.3.3 Experimental Protocol

Rats fasted for 24hr were given 2.5 x 106 EU/kg LPS or its saline vehicle (Veh) i.v.
at 5ml/kg, and food was then returned. Two hours later, 30 mg/kg RAN, 6mg/kg FAM or
their vehicle (sterile phosphate-buffered saline (PBS)) was administered (i.v.). RAN
solution was administered at 2 ml/kg at a rate of approximately 0.15 ml/min. The FAM
dose was selected as a pharmacologically equi-efiicacious dose based on relative

potencies of RAN and FAM in antagonizing H2-receptors (Luyendyk et al., 2006c). At

102

the ti

Plasn

coma

collec

and 31

0f 11’]:

forma

Was t1

3.3.4 '

(PAL

 

 

the time of sacrifice, rats were anesthetized with sodium pentobarbital (75 mg/kg, i.p.).
Plasma was collected by drawing 2 ml of blood from the vena cava into a syringe
containing sodium citrate (final concentration, 0.38%). Another portion of blood was
collected and allowed to clot at room temperature; serum was prepared from this fraction
and stored at -20°C until use. Representative slices (3—4-mm thick) of the ventral portion
of the left lateral liver lobe were collected and fixed in 10% (v/v) neutral buffered
formalin. For immunohistochemical analysis, a portion of the left medial lobe of the liver

was flash-frozen in isopentane cooled by liquid nitrogen.

3.3.4 Treatment with Inhibitors of p38, TACE or Plasminogen Activator Inhibitor-1
(PAI-l)

A p38 inhibitor, SB230963, or its vehicle (acidfied PBS, pH=6) was administered
(2.5mg/kg,iv) at the same time as RAN and again 2hr after RAN. Two administrations of
SB230963 were given because of its short half life (Barone et al., 2001). A TACE
inhibitor, EMS-561392 (60mg/kg, also named DPC-333, provided by Bristol-Myers
Squibb, Princeton, NJ) (Grootveld and McDermott, 2003;Qian et al., 2007), or its vehicle
(1% Tween 80 + 0.5% methylcellulose in PBS), was administered 15min before RAN
orally. This dose was shown in a preliminary study to reduce LPS (2.5X10"6
EU/kg,iv)-induced serum TNF—a increase in rats when given 30min before LPS. A PAI-l

inhibitor, WAY-140312 (10mg/kg, provided by Wyeth Research, Philadelphia, PA)

103

(Crane

admin

f.»
i»
0:

ALT ;
Them.
have s

HCCIOS

3.3.6 I

lysis b
iOUg'r
1mm

1101110,g

 

the SL‘

p38 a.

~—J

HCI‘CU

Phosp

 

 

 

(Crandall et al., 2004) or its vehicle (2.0% Tween 80+ 0.5% methylcellulose in PBS) was
administered orally 1hr before RAN and again 1hr and 3hr after RAN.
3.3.5 Hepatotoxicity Assessment

Hepatic parenchymal cell injury was estimated as an increase in serum ALT activity.
ALT activity was determined spectrophotometrically using Infinity-ALT reagent from
Thermo Electron Corporation (Louisville, CO). Previous studies in this LPS/RAN model
have shown that serum ALT activity reflects histopathologic evidence of hepatocellular

necrosis (Luyendyk et al., 2003h).

3.3.6 Evaluation of Hepatic Phospho-p38 and Phospho-MK-Z

A liver section of 5mm length from the right lateral lobe was homogenized in lml
lysis buffer (1mM EDTA, 0.5% Triton X-100, 5mM NaF, 6M urea, lOug/ml leupeptin,
lOug/ml pepstatin, 100uM PMSF, 3ug/ml aproptinin, 2.5 mM sodium pyrophosphate,
1mM activated sodium orthovanadate). After sitting on ice for 15min, the tissue
homogenates were sonicated for 10 sec. After centrifugation at 14,000 RPM for 10min,
the supematants were collected. Phospho-p38 and total p38 in the tissue homogenates
were evaluated using a commercial ELISA kit (R&D systems, Minneapolis, MN) and
p38 activation was expressed as ratio of phospho-p38 to total-p38.

The protein concentration was determined by a Bradford assay (Bio-rad laboratories,
Hercules, CA). lOug protein was loaded and separated on SDS-PAGE gel.

Phospho-MK-2 and total—MK-2 were detected by western analysis using rabbit

104

poht:
[Danxw

expre

3;&7.

coagu
innnui
(3VVX1

a con

(South

 

 

 

polyclonal anti phospho-MK-2 (Thr222) and anti total-MK-Z antibody (Cell Signalling,
Danvers, MA) (both at 1:1000 dilution, overnight at 4 OC). MK-2 activation was

expressed as the density ratio of phospho-MK-2 to total MK-2.

3.3.7 Evaluation of Coagulation System Activation and Plasma PAI-l

Plasma thrombin-antithrombin (TAT) concentration was used as a marker for
coagulation activation. Plasma TAT concentration was determined by an enzyme-linked
immunosorbent assay (ELISA) using a kit from Dade Behring Inc. (catalog number
OWMG15). The concentration of functionally active PAI-l in plasma was assessed using
a commercially available ELISA kit purchased from Molecular Innovations Inc.

(Southfield, MI).

3.3.8 Serum TNF-a and MIP—2 Evaluation
Serum TNF-a concentration was evaluated using a commercial ELISA kit (BD
Biosciences, San Diego, CA). Serum MIP-2 concentration was also evaluated by

commercial ELISA kit (Biosources, Camarillo, CA).

3.3.9 Fibrin Immunohistochemistry and Quantification
Fibrin immunohistochemistry and quantification were performed as described
previously (Copple et al., 2002b). This protocol solubilizes all fibrinogen and fibrin

except for cross-linked fibrin (Schnitt et al., 1993b). Therefore, only cross-linked fibrin

105

stains

positi

3.3.10

quanti
Stainir
(clone
epiIOp.
does n
were f;
1716 51
then b
(dilute
With g

I:lLlI'o .

“(lieg-

 

 

 

exami-

 

and p,

 

stains in sections of liver. The positive area fraction refers to the percentage of area with

positive staining in the total microscopic field.

3.3.10 Evaluation of Liver PMNs and PMN Activation

PMNs accumulated in liver were visualized by immunohistochemical staining and
quantified as described previously (Yee et al., 2003a). PMN activation was measured by
staining of hypochlorous acid (HOCl)-protein adducts in liver. The monoclonal antibody
(clone 2D10G9, subtype IgG2bk) is specific for hypochlorous acid (HOCl)—modified
epitopes generated in vivo (Malle et al., 1997b) and in vitro (Malle et al., 1995b) and
does not crossreact with other oxidative protein modifications. Frozen sections of liver
were fixed in 4% (v/v) formalin for 10 minutes at room temperature with gentle rocking.
The slides were washed 3 times, 5 minutes each, with PBS (phosphate-buffered saline),
then blocked for 1 hour at room temperature with 3% goat serum in PBS. Antibody
(diluted 1:1 in 3% goat serum) was added and incubated for 2 hr at room temperature
with gentle rocking. The slides were washed 3 times, 5 minutes each, with PBS. Alexa
Fluro 488-labeled goat anti-mouse secondary antibody (diluted 1:500 in 3% goat serum,
Molecular Probes, Carlsbad, Califomia) was applied, and the slides were incubated for 3
hr at room temperature. After washing 3 times, 5 minutes each with PBS they were
examined microscopically. Staining was quantified as for fibrin staining described above

and presented as positive area fraction.

106

3.3.11 Evaluation of Hepatic TNF-a mRNA and TACE activity

Total RNA was extracted from frozen liver tissues using the MELT Total Nucleic
Acid isolation system (Ambion, Austin, TX) according to the manufacturer’s instructions.
F irst-strand cDNA was synthesized from isolated RNA using oligo(dT)12—l8 primer and
Superscript II Reverse Transcriptase (Invitrogen, Carlsbad, CA). In the real time PCR
step, cDNA was amplified using the TaqMan universal PCR master mix and TaqMan
pre-developed gene expression assay reagents for rat TNF-or (Applied Biosystems, Foster
City,CA). B -actin was measured as an endogenous control, using TaqMan endogenous
control assay Rat ACTB (VIC® / MGB Probe, Primer Limited, Pismo Beach, CA).
Quantification was conducted on the Applied Biosystems StepOne Real-Time PCR
System, according to the manufacturer's protocols. A standard curve for each gene was
made of 4-fold serial dilutions of total RNA from one sample. The curve was then used to
calculate the relative amounts of target mRNA in the samples. The ratio between TNF-cc
mRNA and B -actin was used as an indicator for TNF-cc mRNA abundance. The TNF-cc
mRNA level in each liver sample was expressed as ratio vs one VehNeh-treated liver for
Fig 6 and vs one LPSNeh/Veh-treated liver for Fig 7B.

Protein extraction for TACE activity measurement was conducted as follows:
Protein was extracted from liver tissue by homogenization in RIPA buffer (50 mM
Tris pH 7.2, 150mM NaCl, 1% deoxycholic acid, 1% Triton X—100, and 0.1% SDS)
containing lOmM 4-nitrophenyl phosphate, 20 mM B-glycerophosphate, 500uM

Pefabloc, 2 ug/mL aprotinin, SOuM sodium orthovanadate and 0.5ug/mL leupeptin.

107

Homoi

centrif

COHCCI‘

commi

3.3.12

A

all dat;

l'ICSI \\

 

Homogenization was followed by a 30-minute tumbling period and a 10- minute
centrifugation at 14,000g (both at 4 ° C). Supernatant was collected and protein
concentration determined by Bradford assay. The TACE activity was measured using a

commercial TACE activity kit (Calbiochem, San Diego, CA).

3.3.12 Statistical Analysis
A two way analysis of variance (ANOVA) with Tukey’s post hoc test was used for
all data analyzed, except where only two groups were present, in which case student’s

t-test was applied. The criterion for statistical significance for all studies was p<0.05.

108

Figure 3.] p38 activation after LPS/RAN treatment. Rats were given either LPS or its
vehicle and cotreated with RAN, FAM (at a dose equiefficacious to RAN) or vehicle 2
hrs later. Hepatic p38 activation was evaluated 1 hr after drug treatment. * significantly
different from the respective treatment without LPS. # significantly different from the

respective treatment without the drug. n=5-9.

109

Phospho-P38 (ratio vs. total P38)

—m

 

 

.30 -

.25 -

 

 

 

. .20 ‘
j ther Us 01

.15 -

. - 1
or t'CfliCit‘ .

, “.1 .
signlllwlul

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Phospho-P38 (ratio vs. total P38)

Veh LPS

110

Figure 3. 2 Effects of a p38 inhibitor on MK-2 phosphorylation. Rats were given LPS
and RAN as in Fig. 3.1. A p38 inhibitor, SB 239063, was administered at the same time
as RAN and again 2hr later. Hepatic phospho-MK-Z protein and total MK-2 were
measured 2hr after RAN treatment as an indicator of p38 activation. # significantly

different from the respective treatment without SB 239063. n=3-4.

lll

LPSNeh/RAN LPS/SBIRAN

 

 

Phospho-
MK-2 —" - N... .___
Total __.
MK—2 , ~ “-— - - W uln- ‘.
0.50 -

0.40 4

Phospho MK-2
(ratio vs total MK-2

.c .o p .c
8 a s a

 

 

 

 

ll2

Figure 3.3 Effects of a p38 inhibitor on serum ALT activity. Rats were given LPS and
RAN as in Fig. 3.1. A p38 inhibitor. SB 239063, was administered at the same time as
RAN and again 2hr later. Serum ALT activity was evaluated 6hr after RAN. #

significantly different from the respective treatment without SB 239063. n=5-7.

113

 

SB 230963

Veh

 

 

 

00000000
5050505
332211

:5. 5<

114

Figure 3.4 Effects of a p38 inhibitor on biomarkers of hemostasis. Rats were treated
with LPS, and 2hrs later they were treated with RAN or its vehicle. SB 239063 or its
vehicle was administered as in Fig 3.3. TAT and active PAl-l were measured 2hrs and
6hrs after RAN or its vehicle. # significantly different from all the groups at the same

time. a significantly different from the respective treatment at 2hr. n=4-6.

115

350 -
300 -

NM
DUI
OO

TAT(ngImL)
;
O

100 ‘
50 .

600

U’l
O
O

h
C
O

Active PAI-1 (nglml)

 

 

300 -

200 -

100 -

 

 

 

 

2hr

 

 

 

_ LPSNeh
m LPS/SB
— LPS-RANNeh
[:3 LPS-RANISB

 

 

 

 

 

 

 

lgii'

 

— LPSNeh
E: LPS/SB
_ LPS-RANNeh
1:1 LPS-RANISB

 

 

 

#

 

116

 

6hr

3.4 Results
3.4.1 p38 activation after LPS/RAN treatment

Rats were given either LPS or its vehicle and cotreated 2 hrs later with RAN, F AM
(at a dose equiefficacious to RAN) or vehicle. Hepatic p38 activation was evaluated 1 hr
after drug treatment. Neither RAN nor FAM alone caused p38 activation as reflected by
the ratio of phospho-p38 to total p38 (Fig 3.1). LPS alone increased p38 activation.
Treatment of LPS-exposed rats with RAN caused greater p38 activation compared to LPS

alone, whereas F AM cotreatment had no additional effect (Fig 3.1).

3.4.2 Effects of p38 inhibitor on hepatotoxicity

Rats were given LPS and RAN as described above and a p38 inhibitor, SB 239063,
was administered at the same time as RAN and again 2hr after RAN. Hepatic
phospho-MK-2 protein was measured 2hrs after RAN treatment as an indicator of p38
activation, and serum ALT activity was evaluated at 6hrs as a marker of hepatotoxicity.
The p38 inhibitor reduced phospho-MK-2 protein (Fig 3.2), indicating attenuation of p38
activation. The inhibitor also significantly reduced the increase in serum ALT activity

(Fig 3.3).

3.4.3 Effects of p38 inhibitor on biomarkers of hemostasis and liver PMNs
Since previous findings showed that the hemostatic system and PMN activation are

crucial for liver injury caused by LPS/RAN (Luyendyk et al., 2004d;Deng et al., 2007f),

117

the effect of p38 inhibition on these events was evaluated. Two hours and six hours after
RAN or its vehicle, TAT and active PAI-l were measured as biomarkers of coagulation
and fibrinolysis, respectively (Levi et al., 2003b). At 2hr, LPS/RAN treatment caused a
greater increase in plasma TAT and active PAI-l compared to LPSN eh treatment 2hrs
after RAN (Fig 3.4), confirming previous results (Luyendyk et al., 2004c). SB 239063
reduced both plasma TAT and active PAI-l to the same levels as LPS/V eh treatment. At
6hrs, active PAI-l remained elevated after LPS/RAN treatment, and SB 239063
eliminated this increase. SB 239063 had no effect on plasma TAT or active PAI-l after
LPS/V eh treatment at either 2hrs or 6hrs (Fig 3.4).

In inflammatory liver injury, PMNs accumulate in sinusoids then transmigrate into
parenchyma in response to stimuli and become activated to injure hepatocytes (Springer,
1994c). LPS/RAN treatment did not change hepatic PMN accumulation compared to
LPSN eh treatment at either 2hrs or 6hrs, and SB 239063 had no effect on hepatic PMN
accumulation (Fig 3.5A). Serum MIP-2, a PMN chemokine, was elevated at 2hr by RAN
in LPS treated rats, and this increase was prevented by SB 239063 (Fig 3.53). Upon
activation, PMNs release myoperoxidase to form HOCl, which forms protein adducts in
liver. Thus, HOCl adduct staining was measured as a marker for PMN activation 6hrs
after RAN (Fig 3.5C), since PMNs are not activated until 3hrs after RAN (Deng et al.,
2007c). SB 239063 reduced HOCl adduct staining after LPS/RAN treatment, suggesting

attenuation of PMN activation.

118

3.4.4 Hepatic TNF-a mRNA after LPS/RAN treatment

Rats were treated with LPS or Veh and cotreated with RAN or Veh as in Fig 3.1.
Hepatic TNF-a mRNA was evaluated 1, 2 and 3hr after the drug treatment. RAN alone
did not affect TNF-0t mRNA at any of the times examined (Fig 3.6). By contrast, LPS
alone increased TNF-u mRNA at all the times examined. LPS/RAN treatment decreased
TNF-a mRNA compared to LPS/V eh treatment at 1hr after RAN but had no significant

effect at 2hr or 3hr (Fig 3.6).

3.4.5 Effects of p38 inhibitor on serum TNF-a protein and hepatic TNF-a mRNA
after LPS/RAN treatment

Rats were treated with LPS/RAN or LPS/Veh, and S3 239063 or its vehicle was
administered as in Fig 3.4. Serum TNF-u protein and hepatic TNF-u mRNA were
evaluated at 2hrs after RAN or Veh treatment. LPS/RAN treatment caused a significant
increase in serum TNF-0t concentration compared to LPS/Veh treatment (Fig 3.7A). SB
239063 reduced serum TNF-a concentration both after LPS/RAN and after LPSNeh
treatment. In contrast, LPS/RAN treatment did not affect hepatic TNF-a mRNA
compared to LPS treatment, and SB 239063 did not change TNF-u mRNA after either

treatment (Fig 3.73).

3.4.6 Hepatic TACE activity after LPS/RAN treatment

119

Figure 3.5 Effects of a p38 inhibitor on PMN biomarkers. Rats were treated with LPS,
and 2hrs later they were treated with RAN or its vehicle. SB 239063 or its vehicle was
administered as in Fig 3.3. A: PMNs in the liver were enumerated 2hrs and 6hrs after
RAN or its vehicle. 3: Serum MIP-2 was measured 2hrs after RAN or its vehicle. C:
HOCl adduct staining was measured 6hrs after RAN. * significantly different from the
respective treatment without RAN. # significantly different from the respective treatment

without SB 239063. n=4-6.

 

 

 

 

 

6hr
LPS/RAN

 

— LPS-RANNeh
J: [:1 LPS-RANISB

— LPSNeh

— LPS/SB

 

 

 

 

 

 

 

 

2hr
LPS/Veh

 

 

 

 

A

 

 

 

 

£583. .2. s 2?. 33.. =53... «.5:

C

 

83 239063

121

Veh

 

 

 

4

1

I1

3...: x 5:02“. $2 228...

9.2.3... 6:28 Go...

Figure 3.6 Hepatic TNF-a mRNA after LPS/RAN treatment. Rats were treated with
LPS or Veh and cotreated with RAN or Veh as in Fig 3.1. Hepatic TNF-a mRNA was
evaluated 1hr, 2hr and 3hr after the drug treatment. * significantly different from the

respective treatment without LPS. # significantly different from the respective treatment

without RAN. n=4-8.

 

 

 

* . «i(.‘fl\~.. . ...l.u.ml.10,w..wr
I .h.-.. but? 1....LP 9;... .h. t...

 

 

 

 

 

 

 

 

 

 

 

 

 

*
NN
wmwm
vu-v-u *I ..
awn» . ;.
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#
* TIJ
0 0 0 0 0 0
W 8 6 4 2

Eo>.:o> m> 039:
<sz u-....z._.

3hr

123

Figure 3.7 Effects of a p38 inhibitor on serum TNF-a protein and hepatic TNF-a
mRNA. Rats were treated with LPS/RAN or LPS/V eh, and SB 239063 or its vehicle was
also administered as in Fig 3.4. Serum TNF-u protein and hepatic TNF-0t mRNA were
evaluated at 2hrs after RAN or Veh treatment. * significantly different from the
respective treatment without RAN . # significantly different from the respective treatment

without SB 239063. n=4-7.

TNF-La mDM/l

TNF-a mRNA
(ratio vs LPSNehNeh)

O

 

 

 

 

 

 

bioIh-b'oo'cioih

_l

 

 

LPSNeh

 

LPSNeh

125

 

LPS/RAN

 

 

 

 

 

 

LPS/RAN

Figure 3.8 Hepatic TACE activity after LPS/RAN treatment. Rats were treated with
LPS/RAN or LPS/Veh, and SB 239063 or its vehicle was also administered as in Fig 3.4.
Hepatic TACE activity was measured 2hrs after RAN or Veh treatment. * significantly
different from the respective treatment without RAN . # significantly different from the

respective treatment without the S3 compound. n=4-7.

 

 

14000 -
12000 -
10000 -
8000 .
6000 -
4000 -
2000 ~

TACE actIVIty
( RFUI mg protein)

 

 

- Veh
SB

 

 

 

 

i

 

VehNeh LPSNeh LPS/RAN

127

He;
trea
trea
SB

FM

3.4.

caus
trea‘
not
inhi
leve

LPs

3.4,.

 

 

 

Hepatic TACE activity was measured 2hrs after RAN or Veh treatment. LPS/V eh
treatment caused a significant increase in hepatic TACE activity as compared to Veh/Veh
treatment (Fig 3.8). RAN treatment enhanced TACE activity in livers of LPS-treated rats.
SB 239063 had no effect on hepatic TACE activity after LPS/V eh treatment but reduced

TACE activity after LPS/RAN treatment to the same level as LPS/V eh treatment.

3.4.7 Effects of TACE inhibitor on serum TNFa and liver injury

A selective TACE inhibitor (BMS-561392; DPC-333) or its vehicle was administered
15 min before RAN or its vehicle. Serum TNFu concentration was measured 1hr after
RAN treatment, and serum ALT activity was measured at 6hr. LPS/RAN treatment
caused a significant increase in serum TNF-a concentration as compared to LPS/V eh
treatment (Fig 3.9A), confirming previous results (Tukov et al., 2007g). The inhibitor did
not affect serum TNF-a concentration after LPS/V eh treatment. By contrast, the TACE
inhibitor decreased serum TNF-a concentration after LPS/RAN treatment to the same
level as LPS/V eh treatment. The TACE inhibitor also reduced serum ALT activity after

LPS/RAN treatment, indicating attenuation of liver injury (Fig 3.93).

3.4.8 Effects of TACE inhibitor on plasma PAI-l
Rats were treated with LPS/RAN or LPS/V eh and with the TACE inhibitor as in Fig
3.9. Since previous results showed that TNF-a was important for PAI-l production

(Tukov et al., 2007f), plasma active PAI-l concentration was measured 2hrs after RAN

128

treatment. Confirming previous results (Luyendyk et al., 2006d), LPS/RAN treatment
caused a significant increase in plasma active PAI-l concentration as compared to
LPS/Veh treatment (Fig 3.10). The TACE inhibitor decreased plasma active PAI-l

concentration after LPS/RAN treatment to almost the same level as LPS/V eh treatment.

3.4.9 Effects of PAH inhibitor on hepatotoxicity and markers of hemostasis and
PMNs

A PAI-l inhibitor, WAY-140312, was administered 1hr before RAN and again 1hr
and 3hr later. Hepatotoxicity, plasma active PAI-l concentration, hepatic fibrin
deposition and liver HOCl adduct were measured 6hrs after RAN treatment. The PAI-l
inhibitor reduced plasma active PAI-l concentration and serum ALT activity (Fig 3.11A
and 113). It also decreased hepatic fibrin deposition and HOCl adduct staining (Fig
3.11C and 11D). The PAI-l inhibitor did not affect hepatic PMN accumulation or serum

CINC-l or MIP-2 concentrations after LPS/RAN treatment (data not shown).

129

Figure 3.9 Effects of a TACE inhibitor on serum TNF-a and liver injury. Rats were
treated with LPS/RAN or LPS/V eh as in Fig 3.4. A selective TACE inhibitor,
EMS-561392, or its vehicle was administered 15 min before RAN or its vehicle. (A)
Serum TNF-a was measured 1hr after RAN treatment and (3) serum ALT activity was
measured at 6hr. * significantly different from the respective treatment without RAN. #

significantly different from the respective treatment without BMS-56l392. n=6-10.

I‘

_-- 'F‘Il- _ l _ _

I. I

 

 

 

 

 

 

 

 

 

 

1400 «
— Veh *
E, 1000 ~
3
up 000~
u.
E 600*
E
= 400-
h
a:
w 200«
0 - w---
LPSNeh LPS/RAN

350 - B

300 J

250 J
2 i
a 200
'3 150 4
<

100 i

50 ~

 

 

O

 

 

Veh BMS

131

Figure 3.10 Effects of TACE inhibitor on plasma active PAI-l. Rats were treated with
LPS/RAN or LPS/V eh and with the TACE inhibitor as in Fig 3.9. Plasma active PAI-l
was evaluated 2hrs after RAN treatment. * significantly different from the respective

treatment without RAN . # significantly different from the respective treatment without

BMS-561392. n=6-10.

132

 

 

 

 

 

 

 

 

 

 

400 ~

0 0 0

0 0 0
3 2 1

:55: V 35. 0>=u<

 

LPS/RAN

LPSNeh

133

Figure 3.11 Effects of a PAI-l inhibitor on hepatotoxicity and markers of hemostasis
and PMN activation. Rats were treated with LPS and cotreated with RAN 2hrs later. A
PAI-l inhibitor, WAY-140312, was administered 1hr before RAN and again 1hr and 3hr
later. Hepatotoxicity, plasma active PAI-l, hepatic fibrin deposition and liver HOCl
adduct were measured 6hrs after RAN treatment. # significantly different from the

respective treatment without WAY-140312. n=6-9.

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Discussion

At the dose used in the present study, LPS rapidly induced serum TNFu production
but failed to cause liver injury (Tukov et al., 2007e). The serum TNFa concentration
peaked at about 1.5 hours and rapidly decreased after that, returning almost to normal by
8 hours. This indicates that an increase in TNFa of this magnitude and duration is not
sufficient to cause hepatocellular damage. RAN-cotreated rats had a longer lasting serum
TNF-0t increase compared to rats given LPS alone (Tukov et al., 2007d). Pretreatment
with agents that reduced serum TNF-u (ie. pentoxyfylline or etanercept) before LPS
challenge protected rats from LPS/RAN-induced liver injury, suggesting the necessity of
TNF-a in producing the hepatocellular injury. However, since these agents were given
before the LPS challenge, those results could not differentiate whether the LPS-induced
TNF elevation or the RAN-induced prolongation of the TNF-a response to LPS was
crucial for the pathogenesis. In addition, the mechanism of RAN-induced prolongation of
TNF-a production after LPS exposure remains unknown.

p38 and its downstream MAPK Activated Protein Kinase 2 (MK-2) are involved in
the production of several cytokines and chemokines including TNF-u (Hitti et al.,
2006b;Neininger et al., 2002e;Numahata et al., 2003a). Interestingly, the p38-dependent
cytokine/chemokines, such as TNF-a, MIP-2 and IL-6, are selectively upregulated in
LPS/RAN-treated rats compared to rats treated with LPS/FAM or only with LPS
(Luyendyk et al., 2006c;Tukov et al., 2007c). Furthermore, TNF-0t has been shown to be

critical for activation of the coagulation system, PMN chemokine production and

136

subsequent hepatotoxicity after LPS/RAN treatment (Tukov et al., 2007b). All of the
above results suggested the possibility that p38 MAPK activation might be an upstream
signal leading to the pathogenic cascade in LPS/RAN hepatotoxicity. Indeed, RAN, but
not FAM, selectively augmented p38 activation early after LPS treatment. A p38
inhibitor given at the same time as the drug reduced the hepatotoxicity. This suggests that
the p38 activation after RAN cotreatment in LPS-treated rats is critical for the liver
injury.

The p38 inhibitor also reduced activation of the hemostatic system and PMN
activation. The effect on the hemostatic sytem was reflected in the reduction of plasma
TAT and PAI-l concentration, whereas the effect on PMN activation was marked by the
reduction in serum MIP-2 concentration and hepatic HOCl-adduct staining. Similar
effects were observed after treatment with agents that reduced serum TNF-a, such as
pentoxyfylline or etanercept (Tukov et al., 2007a). These results are consistent with the
possibility that p38 activation was responsible for the prolonged TNF-a production
caused by RAN cotreatment and that TNF-0. precipitated the downstream effects on the
hemostatic system and PMNs.

RAN did not increase the hepatic TNF-0t mRNA level after LPS treatment nor it did
not increase the TNF-u mRNA stability at least during 1 hr-3 hr after RAN treatment,
suggesting a post-transcriptional mode of action. Moreover, the reduction in serum
TNF-0t protein concentration after inhibition of p38 MAPK was not accompanied by

diminished hepatic TNF-a mRN A, suggesting that p38 regulated TNF-a production after

137

RAN treatment in a post-transcriptional manner. p38 and its downstream activation of
MK-2 have been shown to regulate TNF-a production in macrophages mostly by
increasing mRNA translation (Neininger et al., 2002f;Hitti et al., 2006a). This was
mediated by the AU-rich 3'-untranslated region of TNF-a mRNA. This finding is
consistent with our finding that a p38 inhibitor reduces TNF-a production in a
post-transcriptional manner.

As mentioned in the introduction, increase in TNF-0t protein can arise from the
cleavage of pro-TNF-a by TACE (Aggarwal et al., l985a;Mullberg et al., 2000b).
Accordingly, another possibility is that p38 activates TACE, leading to increased TNF-a
protein release into the circulation from Kupffer cells or other nonparenchymal cells in
liver. Indeed, p38 is essential for ectodomain shedding of TNF-u in CHO cells (Fan and
Derynck, 1999b). In the present study, LPS/RAN treatment increased hepatic TACE
activity as compared to LPS/V eh treatment (Fig 8) the p38 inhibitor reduced hepatic
TACE activity after LPS/RAN treatment to the same level as LPSNeh treatment.
Furthermore, a TACE inhibitor similarly reduced serum TNF-a concentration and liver
injury. All of these results suggest that RAN prolonged TNF-a production after LPS
treatment mainly through augmented p38-dependent TACE activity. However, further
studies need to be conducted to check whether RAN could also prolong TNF-a
production in some part by p3 8-induced TNF-a translation.

As mentioned above, previous results could not differentiate whether the

LPS-induced TNF elevation or the RAN-induced prolongation of the TNF-0. response to

138

LPS was crucial for the pathogenesis. In the present study, the TACE inhibitor was given
immediately before RAN treatment so that the inhibitor did not affect serum TNF-0t level
until after RAN was administered. This treatment regimen was effective in reducing
hepatocellular injury, suggesting that the RAN-induced prolongation of the TNF-0t
response was critical for LPS/RAN-induced hepatotoxicity. In contrast to RAN, FAM did
not prolong TNF-u response after LPS treatment. Thus, the increase in LPS-stimulated
TNF-u production could distinguish a drug that causes human IADRs from one that does
not.

In human case reports of RAN-induced idiosyncratic hepatotoxicity, no direct
evidence for enhanced serum TNF-0t or other cytokines has been reported. However,
most clinical samples are taken after the hepatotoxicity develops when the peak of TNF-a
has likely passed. Furthermore, it is interesting that in 24 out of 34 human cases of RAN
idiosyncratic hepatotoxicity, prodromal signs consistent with endotoxemia (i.e., increased
LPS cone. in blood) or inflammation (e.g. diarrhea, fever, nausea/vomiting, and/or
abdominal pain) were present (Luyendyk et al., 2003i). These clinical signs are consistent
with increased production of TNF-0t and other inflammatory cytokines.

The TACE inhibitor also reduced plasma active PAI-l concentration after LPS/RAN
treatment to almost the same level as after LPS/V eh treatment, suggesting enhanced
PAI-I production as a possible downstream effect of RAN-augmented TNF-0t production.
Since PAI-l is an important negative regulator of the fibrinolytic system (Levi,

2OOSa;Levi et al., 2003a), this is consistent with the previous finding that fibrin

139

deposition resulting from the perturbed hemostatic system is crucial for the liver injury
caused by LPS/RAN (Luyendyk et al., 2004b). Indeed, in the present study a PAI-l
inhibitor reduced the hepatocellular injury caused by LPS/RAN cotreatment. The
inhibitor also reduced hepatic fibrin deposition and PMN activation. Fibrin deposition
can lead to hypoxia, which occurs early in this model (Deng et al., 2007d;Luyendyk et
al., 20050. Hypoxia could potentiate the killing of hepatocytes by proteases (e.g.
elastase) released from PMNs after their activation (Luyendyk et al., 2005u). Indeed,
proteases released from activated PMNs have been shown to be important in the
pathogenesis (Deng et al., 2007c).

Although the PAH inhibitor decreased PMN activation, it did not affect hepatic
PMN accumulation or serum PMN chemokine concentration, suggesting a direct effect of
PAH on PMN activation. A recent study showed that PAI-l directly potentiates
LPS-induced PMN activation through a Jun-N-terminal kinase (JNK)-dependent pathway
(Kwak et al., 2006b). In the LPS/RAN model, LPS causes PMN accumulation in the
liver, and RAN somehow activates the hepatic PMNs (Deng et al., 2007b). RAN itself
did not enhance PMN activation. In fact, it has been shown to reduce PMN activation in
vitro (Okajima et al., 2002c;Okajima et al., 2000c). All of these results suggest that RAN
might induce activation of PMNs accumulated in the liver after LPS exposure indirectly
by augmenting PAI-l production. Polymorphisms in PAI-l in the human population have
been identified (Lane and Grant, 2000), and thus PAI-l could represent a potential

interaction between genetic and environmental factors (i.e. inflammatory stress) in

140

RAN-induced IADRs. For instance, patients with a more active PAI-l-producing allele
might be more susceptible to RAN-induced IADRs caused by endotoxin exposure or
some other inflammatory stress.

As summarized in Fig 3.12, RAN augmented TNF-a production after LPS treatment
in a post-transcriptional manner by enhancing p38 activation. The increase in TNF-a
protein appears to occur through the p38-dependent activation of TACE. The
prolongation of LPS-induced TNF-0t production by RAN appears to be crucial for the
liver injury caused by LPS/RAN cotreatment. The injury appears to involve PAI-l, which
is important for both the hepatic fibrin deposition and PMN activation. The hypoxia
resulting from hepatic fibrin deposition could act synergistically with toxic proteases

released from activated PMNs to kill the hepatocytes.

141

Figure 3.12 Diagram of pathogenic mechanism contributing to hepatocellular injury
in LPS/RAN model. RAN augments TNF-0t production after LPS treatment in a
post-transcriptional manner by enhancing p38 activation. The increase in TNF-a protein
appears to occur through the p38-dependent activation of TACE. The prolongation of
LPS-induced TNF-u production by RAN causes more PAl-l production, which is
important for both the hepatic fibrin deposition and PMN activation. The hypoxia
resulting from hepatic fibrin deposition could act synergistically with toxic proteases

released from activated PMNs to kill the hepatocytes.

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CHAPTER FOUR
Xiaomin Deng, Robert F. Stachlewitz, Michael J. Liguori, Eric A.G. Blomme, Jeffrey F.
Waring, James P. Luyendyk, Jane F. Maddox, Patricia E. Ganey and Robert A. Roth.
(2006) Modest inflammation enhances diclofenac hepatotoxicity in rats: role of

neutrophils and bacterial translocation. J Pharmacol Exp Ther. 319(3):1191-9.

144

4.1.

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4.1 Abstract

Idiosyncratic adverse drug reactions (lADRs) represent an important human health
problem, yet animal models for preclinical prediction of these reactions are lacking.
Recent evidence in animals suggests that some IADRs arise from drug interaction with an
inflammatory episode that renders the liver sensitive to injury. Diclofenac (DCLF) is one
of those drugs for which the clinical use is limited by idiosyncratic liver injury. We tested.
the hypothesis that modest inflammation triggered in rats by a small dose of
lipopolysaccharide (LPS) renders a nonhepatotoxic dose of DCLF injurious to liver.
Cotreatment of rats with nonhepatotoxic doses of LPS and DCLF resulted in elevated
serum alanine aminotransferase (ALT) activity and liver histopathologic change 6 hours
after DCLF administration. Neither LPS nor DCLF alone had such an effect. Gene array
analysis of livers revealed a unique gene expression pattern in the LPS/DCLF-cotreated
group compared to groups given either agent alone. Antiserum-induced. neutrophil
(PMN) depletion in LPS/DCLF-cotreated rats protected against liver injury,
demonstrating a crucial role for PMNs in the pathogenesis of this LPS/DCLF interaction.
Gut sterilization of LPS/DCLF-treated rats did not protect against liver injury. In contrast,
gut sterilization did attenuate liver injury caused by a large, hepatotoxic dose of DCLF,
suggesting that hepatotoxicity induced by large dose of DCLF is caused in part by its
ability to increase intestinal permeability to endotoxin or other bacterial products and to

cause their translocation. These results demonstrate that inflammation-DCLF interaction

145

 

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precipitates hepatotoxicity in rats and raise the possibility of creating animal models that

predict human IADRs.

146

4.2 Introduction

Idiosyncratic adverse drug reactions (IADRs) remain a challenging human health
problem (Boelsterli, 2003e;Kaplowitz, 2005a). They are one of the leading causes for
drug development failures and removal of medicines from the market. A well known
example is troglitazone, which was removed from the market by the Food And Drug
Administration in 2000, because of its potential to cause idiosyncratic liver injury
(Chojkier, 2005a). IADRs appear to be independent of dose, and the onset of injury
varies relative to the onset of drug treatment. Unfortunately, the mechanisms of
idiosyncratic reactions are poorly understood despite the large number of drugs
associated with these reactions. One of the common targets of idiosyncratic toxicity is
liver. Animal models to predict these adverse reactions are lacking. It is therefore of
interest to establish models that mimic human IADRs. In addition, mechanistic studies of
such models could provide biomarkers for IADRs or strategies to prevent them.

There are two conventional hypotheses to explain IADRs. One is that the reactions
occur as a consequence of drug metabolism polymorphisms, which result in different
levels of toxic drug metabolites among patients (Williams and Park, 2003a). The other
one argues that they arise from a specific immune response to a hapten formed by a drug
or its metabolites (Pirmohamed et al., 2002a). However, convincing evidence for these
hypotheses is lacking for the majority of drugs associated with idiosyncratic toxicity. It is
equally plausible that other, unrecognized events render tissues susceptible to

toxicity during drug therapy.

147

Results of several studies in experimental animals indicate that modest
inflammation, triggered by a small dose of lipopolysaccharide (LPS), augments
hepatotoxicity induced by several classes of xenobiotic agents. For example,
coadministration of nonhepatotoxic doses of LPS and trovafloxacin (TVX), a quinolone
antibiotic associated with hepatic IADRs, results in liver damage that resembles human
TVX idiosyncrasy. By contrast, cotreatment with LPS and levofloxacin, a quinolone
antibiotic without idiosyncratic liability, did not produce liver injury at a dose equipotent
to that of TVX (Waring et al., 2006a). Episodes of modest, subclinical inflammation are
commonplace in people. Since they occur irregularly and may go unnoticed during drug
therapy, their interaction with drugs could explain the erratic temporal and dose
relationships that characterize IADRs (Roth et al., 2003a).

Non steroidal anti-inflammatory drugs (N SAIDs) comprise another class of drugs,
some of which cause hepatic IADRs. For example, diclofenac(DCLF) has caused rare but
sometimes serious hepatotoxicity in humans (Boelsterli, 2003d). Although the apparent
incidence of severe DCLF-induced hepatic adverse reactions is quite low (from 1 to 2
cases per million prescriptions or 6 to 18 cases/ 100,000 person-year.), the large number
of patients treated with DCLF makes the absolute number of cases impressive (Walker,
1997a). In addition, cases of severe injury leading to liver transplantation comprise a
large proportion of the reported cases of DCLF-induced hepatotoxicity (Lewis et al.,
2003). The scarceness of liver biopsies from patients and the diverse histopathlogical

presentations in available samples make it difficult to draw clues about mechanisms from

148

human liver pathology alone (Zimmerman et al., 1999). Thus, the pathogenesis of this
low-incidence/ high-severity DCLF hepatotoxicity is largely unknown. Several
mechanisms have been proposed, including formations of reactive drug metabolites,
oxidative stress (Cantoni et al., 2003b), mitochondrial injury (Masubuchi et al., 2002b)
and immune-modulated hypersensitivity (Greaves et al., 2001b). Experimental data
supporting such mechanisms are incomplete, especially at the in vivo level (Cantoni et al.,
2003c;Boelsterli, 2003c).

Based on the observation that a number of drugs that have the ability to cause
IADRs in human patients cause liver injury in LPS-treated rats and are not hepatotoxic in
naive rats, we tested the hypothesis that a modest inflammatory episode in rats increases
the sensitivity of the liver to hepatotoxic effects of DCLF. We found that pretreatment
of rats with a small, nonhepatotoxic dose of LPS renders a nontoxic dose of DCLF
injurious to liver. Furthermore, we determined whether gene expression profiles could
distinguish a LPS/DCLF interaction that causes hepatotoxicity from the nontoxic

treatments with LPS or DCLF alone and provide clues about the mode of action.

149

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4.3 Methods
4.3.1 Experimental Design

Rats received humane care according to the criteria outlined in the “Guide for the
Care and Use of Laboratory Animals” prepared by the National Academy of Sciences,
and procedures were approved by the Michigan State University Committee on Animal
Use and Care. Male, Sprague-Dawley rats (CrlzCD (SD)IGS BR; Charles River, Portage,
MI) weighing 250 to 350 grams were used for these studies. Animals were fed standard
chow (Rodent chow/Tek 8640, Harlan Teklad, Madison, WI) and allowed access to water
ad libitum. They were allowed to acclimate for 1 week in a 12-hr light/dark cycle before
use.

Rats were fasted for 16 hrs then given a nonhepatotoxic dose of LPS (29 X 106
endotoxin units (EU)/kg; Sigma-Aldrich, Inc., St. Louis, MO, Cat. No. L-2880; lot
072K4095) or sterile saline (i.v.). This activity was determined using a QCL
Chromogenic LAL Endpoint Assay from Cambrex (East Rutherford, NJ). Two hours
later, they were given 20 mg/kg DCLF (Sigma-Aldrich, Inc., St. Louis, MO) or sterile
saline, ip. For the high dose DCLF study, 100mg/kg DCLF was given without LPS
administration. Rats remained fasted and were sacrificed 6 hrs after DCLF treatment for
evaluation of liver injury, histopathology and gene expression. They were anesthetized
with sodium pentobarbital (50 mg/kg, i.p.). Blood was collected from the dorsal aorta and
a portion of the blood was placed into a tube containing sodium citrate (final

concentration, 0.38%) for collection of plasma. Another portion was allowed to clot at

150

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room temperature, and serum was collected and stored at -20°C until use. Representative
(3-4-mm) slices of the ventral portion of the left lateral liver lobe were collected and
fixed in 10% neutral-buffered formalin. A portion of the right medial lobe of the liver

was flash-frozen in liquid nitrogen for subsequent gene expression analysis.

4.3.2 PMN Depletion

PMNs were depleted by intravenous administration of 0.25m] rabbit anti-rat PMN
serum (anti-PMN serum, Inter-cell Technologies, Jupiter, FL), diluted 1:1 with sterile
saline, sixteen hours before administration of LPS. Normal rabbit serum (normal serum)
was administered to some animals as a control. Previous studies in which anti-PMN

serum was administered. to rats demonstrated a selective depletion of PMNs .

4.3.3. Gut Sterilization

Gut sterilization was achieved by treating rats with polymyxin B (150 mg/kg) and
neomycin (450 mg/kg) orally for 4 days before LPS/DCLF treatment or high dose DCLF
alone. This treatment has been shown to completely abolish the gram-negative bacterial
growth in rat fecal culture and to reduce plasma endotoxin concentration after chronic

ethanol treatment (Adachi et al., 1995).

4.3.4 Bacterial Culture in Fecal and Liver Homogenates

151

Fecal and liver bacterial cultures were performed 6 hrs after DCLF or saline
treatment. Rat fecal pellets (400mg) were collected directly from the anus and
homogenized in 4 mL trypticase soy broth containing 15% glycerol. A lOul fecal slurry
was plated on MacConkey plates (Becton, Dickinson Co., Sparks, MD). For the liver
bacterial culture, 100mg of the liver from the left lateral lobe was collected in 0.5 ml
trypticase soy broth containing 15% glycerol and homogenized. Liver homogenates
were diluted serially to 1:106, and 100 pl of each dilution was plated on MacConkey
plates. The culture plates were incubated for 24 hrs at 37°C in an atmosphere of 95% air
and 5% C02. The presence or absence of bacterial colonies was examined for the fecal
homogenates. The colonies of bacteria were counted for each dilution of liver

homogenate and averaged for each animal.

4.3.5 ALT Activity and Histopathology Assessment

Hepatic parenchymal cell injury was estimated by quantifying serum alanine
aminotransferase (ALT) activity. ALT activity was determined spectrophotometrically
using Infinity-ALT from Thermo Electron Corp. (Louisville, CO). Formalin-fixed liver
samples were routinely processed and stained with hematoxylin and eosin (H&E).

Slides were read by a pathologist (EB) blinded to the treatment.

4.3.6 RNA Preparation

152

Frozen liver samples (approximately 100 mg of tissue per sample) were immediately
added to 2 mL of TRIzol reagent per sample (Invitrogen Life Technologies, Carlsbad,
California) and homogenized using a Polytron 300D tissue grinder (Brinkman
Instruments, Westbury, NY). One mL of the tissue homogenate was transferred to a
microfuge tube, and total RNA was extracted with chloroform followed by nucleic acid
precipitation with isopropanol. The pellet was washed with 80% ethanol and
resuspended in molecular biology grade water. Nucleic acid concentration was
determined spectrophotometrically at 260 nm (Smart-Spec, Bio-Rad Laboratories,
Hercules, CA), and RNA integrity was evaluated using an Agilent bioanalyzer (Agilent

Technologies, Model 2100, Foster City, CA).

4.3.7 Gene Array Analysis

Microarray analysis was performed using the standard protocol provided by
Affymetrix, Inc. (Santa Clara, CA). Briefly, approximately 15 pg of total RNA was
reversed transcribed into cDNA using a Superscript II Double-Strand cDNA synthesis kit
(Invitrogen Life Technologies, Carlsbad, California) according to the manufacturer’s
instructions with the exception that the primer used for the reverse transcription reaction
was a modified T7 primer with 24 thymidines at the 5’ end (Affymetrix). The sequence
was 5’GGCCAGTGAATTGTAATA- CGACTCACTATAGGGAGGCGG-(dT)24-3’.
cDNA was purified via phenol/chloroform/isoamylalcohol (Invitrogen Life Technologies,

Carlsbad, California) extraction and ethanol precipitation. The purified cDNA was

153

resuspended in molecular biology grade water. Following this procedure, biotin-labeled
cRNA was synthesized according to the manufacturer’s instructions from the cDNA
using the Enzo RNA Transcript Labeling Kit (Affymetrix). The labeled cRNA was
then purified using RNeasy kits (Qiagen, Valencia, CA). Subsequently, cRNA
concentration and integrity were evaluated. Approximately 20 ug of cRNA was then
fragmented in a solution of 40 mM Tris-acetate, pH 8.1, 100 mM potassium acetate, and
30 mM magnesium acetate at 94°C for 35 minutes. Fragmented, labeled cRNA was
hybridized to an Affymetrix rat genome RAE230A array, which contains sequences
corresponding to roughly 15,900 transcripts, at 45°C overnight using an Affymetrix
Hybridization Oven 640. The array was subsequently washed and stained twice with
strepavidin-phycoerythrin (Molecular Probes, Eugen, OR) using a Gene-Chip Fluidics
Workstation 400 (Affymetrix). The array was then scanned using the Affymetrix
GeneChip® Scanner 3000.

The microarray scanned image and intensity files (.cel files) were imported into
Rosetta Resolver gene expression analysis software version 4.0 (Rosetta Inpharmatics,
Seattle, WA). Error models were applied and ratios were built for each treatment array
versus its respective vehicle control (pooled in silico). Using Rosetta Resolver, a
p-value is calculated for every fold change, using the Rosetta Resolver error model

(Rajagopalan, 2003).

4.3.8 Statistical Filtering of Genes Changed After LPS/DCLF Treatment

154

Genes changed after treatment with LPS/DCLF, LPS/saline or saline/DCLF were
identified with saline/saline as baseline using the following criteria: P<0.01 in at least 3
out of 4 animals (groups LPS/saline and LPS/DCLF); P<0.01 in at least 4 out of 5
animals (group saline/DCLF). To identify the genes that changed after LPS/DCLF
treatment relative to LPS/saline group or saline/DCLF group, a second filter was applied
with the LPS/DCLF group as baseline using the same criteria above.

The genes expressed to a greater degree after LPS/DCLF treatment than after LPS/saline
and saline/DCLF treatment were classified based on the relationship of their gene
products to one or more functions. The classification was based on the gene fimction
described in Entrez Gene site (http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=gene).
Furthermore, this list of genes was imported into Genomatix Bibliosphere 5.13 software
(Miinchen, Germany). This yielded a list of biological process terms ranked by their
probability of over-representation or under-representation in a particular gene list.
Functional annotations were based on the Genomatix knowledge base. “Expected” refers
to the numbers of gene expression changes expected by chance in the gene list. The “Z
Score” ranks the deviation of the observed number of genes in one biological process
category of the Genomatix database found in the imported list of genes from the numbers
expected to occur by chance. That is, it ranks the probability that the genes in this

category are over-represented or under- represented in the gene list.

4.3.9 Serum MIP-2 Concentration and Blood/Liver Leukocyte Evaluation

155

Serum MIP-2 concentrations were determined using a commercially available
ELISA kit (Biosource International, Camarillo, CA). Total blood leukocytes were
quantified using a Unopette white blood cell (W3C) determination kit
(Becton-Dickinson, Franklin Lakes, NJ) and a hemacytometer. Slides were prepared from
whole blood and stained using the Hema 3® Staining System (Fisher Scientific,
Middletown, VA), and differential counting was performed. Immunohistochemical
staining for PMNs was performed on formalin-fixed liver sections as described
previously (Kim et al., 2005d). Hepatic PMN accumulation was evaluated by counting

PMNs in 20 randomly selected, high-power fields (HPF, 400X).

4.3.10 Statistical Analysis

All data are expressed as mean 1 SEM. ALT activity, ELISA and circulating and
liver leukocyte data were analyzed by two-way ANOVA with Tukey's posthoc test. For
Figures 1 and 5 and Table 5, one-way ANOVA was applied. Data were transformed if
they did not pass the normality and equal variance tests for ANOVA. For microarray
analysis, error models were applied and ratios were built for each treatment array versus
its averaged respective vehicle control using the Rosetta Resolver system. Gene
expression was considered significantly changed if the p—value was less than or equal to
0.01. Agglomerative cluster analysis was performed using the average link heuristic

criteria and the Euclidean distance metric for similarity measure.

156

4.4 Results
4.4.1 Hepatotoxicity from DCLF/LPS Interaction

In a preliminary study, a large dose of DCLF caused liver injury within 6 hrs.
Accordingly, in an initial dose response study, rats were given DCLF (i.p) at doses from
0-100mg/kg, and serum ALT activity was evaluated 6 hrs after DCLF treatment. Doses
up to 40mg/kg did not cause elevation in serum ALT activity, indicating no liver injury.
Administration of DCLF at doses of 50mg/kg and 100mg/kg caused a significant increase
in ALT activity (Fig 4.1), indicating hepatocellular injury.

To determine the influence of a nonhepatotoxic dose of LPS on DCLF
hepatotoxicity, rats were pretreated with LPS (29X106 EU/kg, iv) or sterile saline, Two
hrs later they were treated with DCLF (20mg/kg, ip) or saline. Serum ALT activity was
evaluated 6 hrs later. Neither LPS nor DCLF when given alone altered ALT activity.
However, cotreatment with LPS and DCLF caused a significant increase in serum ALT
activity (Fig 4.2).

Livers from both saline/saline-treated and saline/' DCLF-treated rats had no or
minimal histopathological changes (Table 4.1). LPS/saline treatment caused moderate
leukocyte infiltration, occasional hepatocellular apoptosis, and modest parenchymal
edema and hemorrhage. LPS/DCLF cotreatment enhanced hepatocellular apoptosis,

parenchymal edema and hemorrhage relative to LPS/saline treatment.

4.4.2 Microarray Analysis of Livers from Rats Treated with LPS/DCLF

157

 

Figure 4.1 Dose-responsive DCLF hepatotoxcity. DCLF was given at doses ranging
from 0-100mg/kg, i.p. and ALT was measured 6 hrs later. n=3 -4; * significantly different

from saline-treated group. p<0.05.

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Figure 4.2 LPS potentiates DCLF hepatotoxicity. LPS (29X106 EU/kg, i.v) or vehicle
was administered 2 hrs before DCLF (20mg/kg. i.p) or vehicle. ALT was measured 6 hrs

after DCLF or its vehicle. n=4-5; * significantly different from all other groups. p<0.05.

160

 

 

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LPS/Saline 1.6 i 0.25 1.2 i 0.2 1.25 i 0.22 1.8: 0.38
Saline/DCLF 0.8 i 0.2 0.0 i 0 0.6 1; 0.24 0.0 i 0
LPS/DCLF 3.0 ; 0 2.25i_0.25 2.25 $0.25 2: 0

 

 

 

Table 4.1 Histopathological Evaluation of Livers from LPS/DCLF-treated Rats.LPS
(29X106 EU/kg, i.v) or saline was administered 2 hrs before DCLF (20mg/kg, i.p) or
saline. Livers were taken 6 hr after DCLF and stained with H&E. Liver histopathology
was evaluated on a scale of 0-4 based on severity (0: normal; 1: minimal; 2: mild; 3:

moderate; 4: severe). n=4-5.

162

Rat livers were subjected to RNA extraction and subsequent microarray analysis for
global gene expression. Hierachical clustering analysis revealed mostly treatment-related
clusters (Fig 4.3). Saline/DCLF treatment caused few gene expression changes. All rats
treated with LPS irrespective of cotreatment clustered together. Within this major cluster,
LPS/DCLF-treated rats formed a distinct subcluster. One LPS/saline-treated rat clustered
with the LPS/DCLF-treated group. Interestingly, this animal had the largest serum ALT
value in the LPS/saline group.

The number of gene expression changes after LPS/saline, saline/DCLF and
LPS/DCLF treatments were identified with saline/saline-treated rats as the baseline. The
numbers of genes changed in expression are depicted as a Venn diagram (Fig 4.4). A
large number of gene expression changes occurred after either LPS/DCLF or LPS/saline
treatment, but many more expression alterations occurred in the former group. A large
number of gene expression changes occurred after both of these treatments (L and LD
intersection in Fig 4.4). Numerous genes were differentially expressed specifically in the
LPS/DCLF group (LD in Fig 4.4). DCLF alone caused few changes in gene expression
based on the criteria used.

The genes altered in expression after LPS/DCLF treatment were further filtered by
their expression pattern. Normalization to LPS/DCLF as the baseline was used to identify
the genes changed to a greater degree after LPS/DCLF treatment compared with LPS

alone and DCLF alone. This group of genes was interesting because liver injury occurred

163

 

 

Figure 4.3 Hierachical cluster analysis of hepatic gene expression after LPS/DCLF
treatment. LPS (29X106 EU/kg, i.v) or vehicle was administered 2 hrs before DCLF
(20mg/kg, i.p) or vehicle. Livers were taken at 6hr after DCLF, and mRNA was extracted
and subjected to Affymetrix RAE230A gene array analysis. Gene expression changes
were determined using saline/saline group as baseline and hierachical cluster analysis

was performed by Rosetta Resolver using Euclidean average link distance.

164

 

 

 

 

165

LPS+Veh
LPS+Veh
LPS+Veh
LPS+Veh
LPS+DCLF
LPS+DCLF
LPS+DCLF
LPS+DCLF
V%h+DCLF
Vbh+DCLF
Veh+DCLF
Vbh+DCLF
Vbh+DCLF

 

Figure 4.4 Venn Diagram of gene expression changes caused by LPS/DCLF. Gene
array analysis was performed as described in Fig 3. Active genes were identified as
described in Methods, using saline/saline as the baseline. L indicates LPS/saline, LD
indicates LPS/DCLF and D indicates saline/DCLF. The intersection of treatment groupS
indicates the gene expression changed after both or all three of the indicated treatments-

Up arrow indicates numbers of genes with enhanced expression and down arrow

indicates reduced expression.

166

 

 

167

 

 

 

 

 

 

 

 

 

 

, Cell death .

Expressnon Inflammation and Stress Hemostasis .Hypoxla Total

Pattern , Inducible

survnval

LD&L,
LD>L and 15 8 6 2 4 104

LD>D

LD

only,LD>L 10 5 3 3 l 111
and LD>D

 

Table 4.2 Functional Classification of Genes Expressed to Greater Degree after

LPS/DCLF Treatment than after LPS or DCLF Given Alone. Genes with two

expression patterns were identified from the sets of genes in the Venn Diagram, either in

the LD only group and LD intersect L group (refer to Fig 4.). Those two groups

comprised genes expressed to a greater degree after LPS/DCLF treatment than after

treatment with either agent alone. Genes were classified based on the relationship of their

gene products to one or more functions described in the Entrez Gene web site. A gene

can be assigned to one or more than one function.

168

 

Table 4.3 Gene Ontology (GO) Biological Process Categories for Which Gene
Expression Was Affected to a Greater Degree after LPS/DCLF Treatment than
after Treatment with LPS or DCLF Alone Genes with the expression pattern [LD>L
and LD>D] were identified as in Table 2. This gene list was imported into Genomatix
Bibliosphere 5.13. Functional annotations were based on the Genomatix knowledge
base. “Expected” refers to the numbers of gene expression changes expected by chance
in the gene list. The “Z Score” ranks the deviation of the observed number of genes in
one biological process category of the Genomatix database found in the imported list of
genes from the numbers expected to occur by chance. It indicates the ranking of whether
the genes in this biological process category are over-represented or under-represented in

the imported gene list.

169

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Term GO ID Observed Expected ZScore

neutrophil chemotaxis GO:0030593 2 0 35.25
immune cell chemotaxis GO:0030595 2 0 32.63
taxis GO:0042330 3 0.01 28.94
chemotaxis GO:0006935 3 0.01 28.94
immune cell migration GO:0050900 2 0 28.76
inflammatory response GO:0006954 3 0.02 19.25
innate immune response GO:0045087 3 0.02 19.04
response to chemical substance GO:0042221 3 0.03 17.72
response to wounding GO:0009611 3 0.04 14.67
response to pest/pathogen/parasite GO:0009613 3 0.04 14.13
response to abiotic stimulus GO:0009628 3 0.05 13.79
cell migration GO:0016477 2 0.02 13.4
immune response GO:0006955 3 0.08 10.74
defense response GO:0006952 3 0.08 10.23
cell motility GO:0006928 2 0.04 10.05
response to biotic stimulus GO:0009607 3 0.1 9.19
response to stress GO:0006950 3 0.12 8.61
response to external stimulus GO:0009605 3 0.17 7.03
response to stimulus GO:0050896 3 0.2 6.43
organismal physiological process GO:0050874 3 0.22 6.15
signal transduction GO:0007165 2 0.33 3.08
cellular process GO:0009987 3 0.92 2.61
cell communication GO:0007 154 2 0.45 2.5
cellular physiological process GO:0050875 2 0.58 2.09
physiological process GO:0007582 3 1.23 2.07
biological _process GO:0008150 3 1.49 1.74

 

170

 

in the LPS/DCLF group but not in the LPS alone or the DCLF alone group. Accordingly,
genes changed to a greater degree in the LPS/DCLF group than in the LPS/saline and
saline/DCLF groups are associated with liver injury. These genes were classified based
on the relationship of their gene products to one or more biological functions (Table 4.2).
Many genes related to inflammation, cell death/survival, stress response, hemostasis or
hypoxia were selectively upregulated by LPS/DCLF treatment.

The genes changed to a greater degree after LPS/DCLF treatment compared to LPS
alone or DCLF alone were imported into Genomatix Bibliosphere 5.13. This yielded a
list of gene ontology biological process terms ranked by their probability of

over-representation or under-representation on the gene list (Table 4.3).

4.4.3 Neutrophil-related Gene Expression Changes after LPS/DCLF Treatment and
Effects of Neutrophil Depletion on LPS/DCLF-induced Hepatotoxicity

Transcripts for the genes encoding the neutrophil chemokines such as MIP-2 and
MIP-la and the adhesion molecule ICAM-l were changed to a greater degree in the
LPS/DCLF cotreated group compared to LPS or DCLF given alone (Fig 4.5). Serum
MIP-2 protein levels in both saline/saline and saline/DCLF groups were below the
detection limit of the assay (Fig 4.6). LPS/saline treatment increased serum MIP-2
protein concentration, whereas LPS/DCLF treatment caused a much greater increase (Fig
4.6).

Both LPS/saline and LPS/DCLF treatment caused hepatic PMN accumulation (Table

171

 

Figure 4.5 Neutrophil-related gene expression changes after LPS/DCLF treatment.
Gene array analysis was performed as described in Fig 3. The fold changes in MIP-2,
MIP-la and ICAM-1 transcripts were calculated with the saline/saline group as baseline.
n=4-5; * significantly different from saline/DCLF group; # significantly different from

LPS/saline group. p<0.05.

172

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173

 

I: 1i. 11. .

 

Figure 4.6 Serum MIP—2 protein concentration after LPS/DCLF treatment. LPS
(29X106 EU/kg, i.v) or saline was administered to rats 2 hr before DCLF (20mg/kg, i.p)
or saline. Serum MIP-2 was measured 6 hr after DC LF treatment. n= 4-5; * significantly
different from group without LPS; # significantly different from group without DCLF.

p<0.05.

174

MIP-2 (pg/ml)

2500 -

 

 

 

 

 

 

* #
2000 ~ i
1500 - i
1000 -
500 - *
. , I.
LPS -- -- + +
DCLF -- + -- +

175

Figure 4.7 Effects of PMN depletion on LPS/DCLF-induced liver injury. Rats were
pretreated with normal rabbit serum or rabbit anti-PMN serum 16 hrs before LPS
(29X106 EU/kg, i.v) or its vehicle. DC LF (20mg/kg. i.p) or its vehicle was administered 2
hr after LPS. Serum ALT activity was evaluated 6hr after DCLF. n= 3-7; * significantly
different from respective group without LPS/DCLF; # significantly different from

respective group without antiserum. p<0.05.

176

 

— Normal Serum
200 - c:::i Anti-PMN Serum *

 

 

 

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3.160
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VehNeh LPS/DCLF

177

4.1 and 4.4). To investigate the requirement for neutrophils in LPS/DCLF hepatotoxicity,
neutrophil numbers in liver were reduced by treating rats with rabbit anti-rat PMN serum
16 hrs before LPS treatment. This treatment selectively reduced circulating and hepatic
PMNs (Table 4.4). The anti-PMN serum alone did not alter ALT activity but did
attenuate LPS/DCLF hepatotoxicity, as reflected by reduction in serum ALT activity (Fig
4.7). LPS/DCLF cotreatment reduced circulating lymphocytes (Table 4.4), similar to

effects of LPS alone (data not shown).

4.4.4 Effects of Gut Sterilization on LPS/DCLF Hepatotoxicity

DCLF is known to cause gastrointestinal injury and bacterial translocation from
intestine to liver (Kim et al., 2005c). The observation in the gene array study that the rat
with the largest serum ALT activity after LPS/saline treatment clustered with the
LPS/DCLF-treated rats suggested that the LPS/DCLF interaction might be due to
bacteria/endotoxin exposure that resulted from a DCLF-induced intestinal permeability
increase, which added to the effects of exogenous LPS given before DCLF exposure. To
test this hypothesis, rats were treated with the nonabsorbable antibiotics, polymyxin B
and neomycin, orally for 4 days before LPS/DCLF treatment. This antibiotic treatment
completely abolished the colony growth from homogenized rat fecal samples on
MacConkey agar plates, which are selective for gram-negative bacteria (data not

shown).Gut sterilization did not significantly affect LPS/DCLF hepatotoxicity (Fig 4.8).

178

This result suggested that the LPS/DCLF interaction is not due to DCLF enhancing

bacteria/endotoxin translocation from intestine to liver.

4.4.5 Effects of Gut Sterilization on Liver Injury in Rats Treated with a Hepatotoxic
Dose of DCLF.

Large doses of DCLF have been shown to cause bacterial translocation from the
intestine to liver (Kim et al., 2005b;Seitz and Boelsterli, 1998b). Consistent with this,
DCLF (100mg/kg) did increase the level of gram-negative bacteria in the liver (Table
4.5). Although gut sterilization had no significant effect on LPS/DCLF hepatotoxicity,
the possibility remained that LPS or bacterial translocation from intestine to liver plays
an important role in the hepatotoxicity from a large, toxic dose of DCLF (100mg/kg). To
test this hypothesis, rats were treated with nonabsorbable antibiotics as above before they
were given a toxic dose of DCLF (100mg/kg). This antibiotic treatment attenuated the
gram-negative bacterial colonies detected in liver (Table 4.5), and it reduced the serum
ALT activity after administration of a hepatotoxic dose of DCLF, indicating attenuation

of liver injury (Fig 4.9).

179

 

 

Tot" Blood PMNs Blow Liver PMN
Treatment Leukocytes

Lymphocytes
(#lul) (#lul) (#lul) (# per field)
NS/Veh/Veh 4350 1; 153 1105 _-I_-_ 50 2684 i 527 1.5 i 0.3

AS/Veh/Veh 2817 i 394 # 45 i 23 # 2356 i 503 1.7 i 0.5

 

 

 

NS/LPS/DCLF 2479 j; 283 * 1228 i 234 1063 i 95 * 45.8 i 6.0 *

 

 

 

 

 

 

 

AS/LPS/DCLF 1183 : 223 # 122 254 # 1001 i 263 * 26.2 + 1.8 *# ,

 

Table 4.4 Effects of Anti-PMN Serum on Circulating Leukocytes and Liver PMNs.
Rats were pretreated with normal rabbit serum (NS) or rabbit anti-PMN serum (AS) 16
hrs before treatment with LPS (29X106EU/kg, i.v) or its vehicle. DCLF (20mg/kg, i.p) or
its vehicle was administered 2 hr after LPS. Total leukocytes, PMNs and lymphocytes in
blood and liver PMNs were counted 6hr after DCLF. n= 3-7; * significantly different
from respective group without LPS/DCLF; # significantly different from respective group

without antiserum. p<0.05.

180

 

 

Figure 4.8 Effects of gut sterilization on LPS/DCLF-induced liver injury. Rats were
pretreated with polymyxin B (150mg/kg/day) and neomycin (450mg/kg/day) orally for 4
days before LPS (29X106 EU/kg) or its vehicle. DCLF (20mg/kg, i.p) or its vehicle was
administered 2 hr after LPS or vehicle. Serum ALT activity was evaluated 6hr after
DCLF or its vehicle. n=3-8; * significantly different from all other groups with the same

antibiotic treatment. p<0.05.

181

 

400 i — Saline
12:1 Antibiotics *

 

 

 

09
O
O

a.

200 - 1

Serum ALT activity (UIL)

 

 

 

 

 

 

 

 

 

t 11

VehNeh LPSNeh Veh/DCLF LPS/DCLF

182

 

l-h vu—

Figure 4.9. Effects of gut sterilization on hepatotoxicity induced by a large dose of
DCLF. Rats were pretreated with polymyxin 3 (lSOmg/kg/day) and neomycin
(450mg/kg/day) orally for 4 days before administration of DCLF (100mg/kg, i.p) or
saline. Serum ALT activity was evaluated 6hr after DCLF or vehicle. n=3-6; *
significantly different from respective group without DCLF; # significantly different

from respective group without antibiotics. p<0.05.

183

 

 

 

 

 

 

 

 

 

 

*
S
.m
a
mm
rum
SA
0 0 0 0 0
0 0 0 0
4 3 2 1

3.3 33:8 hi anm

DCLF

Veh

184

 

Treatment CFU/g (liver)

 

 

 

Saline/Vehicle 209 j; 68
Saline/DCLF 5,845,596 i 1,580,295 *
Antibiotics/DCLF 264,943 : 167,660 * #

 

 

 

 

Table 4.5. Effects of Gut Sterilization on liver Bacterial Colony Growth after High
Dose DCLF Treatment. Rats were treated with polymyxin B and neomycin for 4 days
before DCLF (100mg/kg) or vehicle treatment. Liver bacterial growth was quantified as
described in Methods 6 hrs after DCLF or vehicle treatment. n= 6-8; * significantly
different from respective group without DCLF; # significantly different from respective

group without antibiotics. p<0.05.

185

4.5 Discussion

DCLF is well known to cause idiosyncratic hepatotoxicity (Boelsterli, 2003b). In
this study, we showed that pretreating rats with a small dose of LPS converts a dose of
DCLF that is normally noninjurious to one that is hepatotoxic. This result suggests the
possibility that human patients might become susceptible to DCLF hepatotoxicity during
a modest inflammatory episode. This appears even more intriguing, since most patients
take DCLF for inflammation-related diseases such as rheumatoid arthritis or
osteoarthritis. In fact, osteoarthritis has been associated with increased risk for
DCLF-induced liver injury (Banks et al., l995b).

Mechanisms of DCLF-induced liver injury remain incompletely understood.
Mitochondrial injury leading to cell death has been proposed as a mode of hepatocellular
injury, but by itself this does not explain the occurrence of injury only in a minority of
patients (Masubuchi et al., 2002c;Gomez-Lechon et al., 2003). In addition, to our
knowledge no direct evidence of mitochondrial injury in vivo has been reported. Given
our results, it would be interesting to see DCLF induces mitochondrial injury in vivo or in
vitro in the presence of inflammatory mediators.

Whether inflammation in human patients acts as a risk factor for DCLF-induced
idiosyncratic liver injury requires additional study. Interestingly, polymorphisms in genes
encoding cytokines IL-4 and lL-10 have been identified in patients developing
DCLF-induced liver injury (Aithal et al., 2004c). These polymorphisms cause less IL-10

and more IL-4 production. Since IL-10 is an anti-inflammatory cytokine and IL-4

186

participates in B cell activation, these changes could enhance hepatotoxic interactions
between DCLF and LPS or other inflammagens to which patients are exposed during
drug therapy.

The potentiation of DCLF induced hepatotoxicity by LPS might appear paradoxical,
since DCLF acts as a NSAID to dampen inflammatory responses. The primary
mechanism of action of DCLF in this respect is inhibition of the enzyme cyclooxygenase
(COX). COX catalyzes production of prostaglandins (PGs) and thromboxanes (Tx ) from
arachidonic acid. PGs and Tx have diverse biological actions, either proinflammatory or
anti-inflammatory. However, as a nonselective COX inhibitor (i.e., inhibits both isoforms
of the enzymes COX-1 and COX-2), it inhibits the synthesis of cytoprotective and
anti-inflammatory PGs produced by constitutive COX activity as well as products of
inducible COX-2, which are mostly proinflammatory. Moreover, the toxic interaction of
DCLF with LPS might arise from inflammatory mediators (eg., cytokines) that are not
products of the COX pathway and are not inhibited by NSAIDs.

Analysis of hepatic gene expression revealed a unique pattern of gene expression for
animals treated with LPS/DCLF. The genes expressed to a greater degree in the
LPS/DCLF group than in LPS/saline and saline/DCLF groups were selected for fiirther
functional analysis because liver injury followed this pattern. Interestingly, many genes
involved in inflammation, cell death/survival, and response to stress were expressed to a
greater degree in the LPS/DCLF group, consistent with the hypothesis that DCLF

exaggerates the inflammatory stress caused by LPS.

187

Neutrophil/immune cell chemotaxis was at the top of the biological process rankings
of the GO functional analysis for genes expressed to a greater degree in the LPS/DCLF
group, and transcripts representing several neutrophil chemokines and adhesion
molecules were upregulated to a great degree in this group, suggesting that PMNs might
be involved in the pathogenesis. Serum MIP-2 concentration reflected the increase in the
transcripts for this chemokine. The mechanism by which DCLF enhances MIP-2 protein
expression remains unknown. There are reports that COX-2 inhibition enhances
MCP-l/MIP-2 production in models of inflammation caused by radiation (Kyrkanides et
al., 2002b). In addition, it has been proposed that in the face of inhibition of COX,
arachidonic acid is metabolized more extensively by lipoxygenase. In this regard, it is
interesting that the lipoxygenase products derived from arachidonic acid, such as
l5-hydroxyperoxyeicosatetraenoic acid (IS-HPETE), can increase adhesion molecules
and transmigration of neutrophils across endothelial cell barriers (Sultana et al., l996a).
Thus, shifting of the generation of eicosanoids from the COX pathway to the
lipoxygenase pathway by DCLF during inflammation (eg. LPS exposure) could
contribute to enhancing the expression of MIP-2 and subsequent PMN activation.
Depletion of PMNs in LPS/DCLF-treated rats attenuated liver injury, indicating the
involvement of PMNs in the pathogenesis. PMNs cause cytotoxicity through the release
of reactive oxygen species and other toxic factors. It is interesting in this regard that the
addition of noncytotoxic concentrations of peroxidase/11202 to hepatocyte cultures

markedly increased DCLF cytotoxicity (Tafazoli et al., 2005). In fact, several novel

188

reactive metabolites of DCLF formed by incubation of the drug with PMN-derived
myeloperoxidase (Miyamoto et al., 1997c;Zuurbier et al., 1990a).

DCLF, as other NSAIDS, is known to cause gastrointestinal injury and bacteria
translocation from the intestine to liver (Kim et al., 2005a;3anks et al., 1995a;Seitz and
Boelsterli, 1998a). The resultant excessive bacteria/endotoxin exposure of liver could
combine with the small dose of administered LPS to cause liver injury. This is not likely,
because gut sterilization with nonabsorbable antibiotics did not dampen the hepatic
damage caused by coadministration of LPS and DCLF. In contrast, gut sterilization did
reduce liver injury caused by a larger, hepatotoxic dose of DCLF. It is possible that
DCLF becomes hepatotoxic only in the presence of bacteria/endotoxin or other
inflammagens and that the larger dose of DCLF is hepatotoxic because it causes
translocation of LPS or bacteria from the intestine to liver.

LPS can increase intestinal permeability by inducing epithelial barrier dysfunction.
This is reflected, for example, as an increase in dextran absorption (Moriez et al.,
2005;Han et al., 2004). In addition, LPS administration to rats can change drug
bioavailability through alterations in expression and activity of transporters and drug
metabolizing enzymes in the intestinal epithelium (Roth et al., 2003b). Accordingly, it
seems possible that LPS exposure could alter the toxicity of orally administered
diclofenac by increasing its absorption from the gastrointestinal tract. This potential

mode of action likely did not pertain to our results, since we administered the drug

189

intraperitoneally. However, in human patients who typically take diclofenac orally, this
could provide an additional mode by which LPS exposure could influence drug toxicity.

Diclofenac causes IADRs in people in a time frame varying from a few weeks to a
year after onset of therapy with the drug (Boelsterli, 2003). If drug-inflammation
interaction underlies some human IADRs, then a reaction would be expected to occur
only in patients with an inflammatory episode occurring during drug therapy and having
sufficient magnitude to precipitate a hepatotoxic interaction with the drug. Modest
inflammatory episodes likely vary in both timing of onset and magnitude among and
within people (Roth et al., 2003c). Accordingly, this hypothesis of drug-inflammation
interaction could explain the variable onset of diclofenac IADRs in human patients.

In summary, we established a potential model of DCLF-induced idiosyncratic liver
injury by pretreating rats with a small dose of LPS before an otherwise nonhepatotoxic
dose of DCLF. The molecular mechanisms of this inflammation-DCLF interaction
remain unknown, but gene array analysis and PMN depletion studies indicated a role for
PMNs. Gut sterilization of LPS/DCLF-treated rats failed to protect against liver injury,
suggesting that this interaction is not due to excessive bacteria/endotoxin translocation
from intestine to liver caused by DCLF. However, gut sterilization attenuated the liver
injury caused by a large, toxic dose DCLF, suggesting that DCLF hepatotoxicity is

partially due to bacteria/endotoxin translocation caused by injuring the intestinal barrier.

190

 

CHAPTER FIVE

Summary and Conclusions

191

5.1 Summary and Conclusions

The overall goal of the thesis project was to investigate how PMNs contribute to
hepatocellular injury in LPS/RAN-cotreated rats and the mechanism by which PMNs
become activated in the livers of LPS/RAN-cotreated rats. These studies will contribute
to the understanding of mechanisms by which inflammation acts as a susceptibility factor
for toxicity due to drugs and other xenobiotic agents. In addition, the completed study
could lead to improved strategies for preventing or treating conditions involving
PMN-dependent tissue injury.

In the thesis research conducted, a mechanism was proposed that PMNs contribute
to liver injury by enhancing hemostasis. Both PMN antiserum and CD18 antiserum
reduces the hepatic fibrin deposition at 6hrs after RAN treatment, confirming the role of
PMNs in the hemostatic system. Previous in vitro studies showed that PMNs could cause
thrombin activation and fibrin deposition by platelet-dependent and -independent
mechanisms and PMNs even express functional tissue factor upon stimulation
(MAUGERI et al., 2006;Goel and Diamond, 2003b;Goel and Diamond, 2003a); however,
there have been no reports on the contribution of PMNs to fibrin formation in vivo to our
knowledge. Data in this thesis showed that PMNs are involved in hepatic fibrin
deposition in an animal model of liver injury in vivo. The PMN antiserum did not reduce
fibrin deposition that occurs early after LPS/RAN treatment, suggesting that PMNs only
contribute to the late phase of sinusoidal fibrin deposition. Furthermore, the PMN

antiserum did not affect plasma TAT concentration, indicating this effect of PMNs on

192

 

fibrin formation is not due to thrombin activation. These findings contrast with the in
vitro findings mentioned above. Furthermore, neutrophil proteases inhibitor (eglin C)
studies showed that eglin C did reduced the fibrin deposition whereas had not effect on
thrombin activation. Thus in the LPS/RAN model, PMNs contribute to sinusoidal fibrin
deposition in a thrombin-independent manner.

An alternative mechanism by which PMNs could contribute to deposition of fibrin is
through inhibiting fibrinolysis by increasing active PAI-l. PMN lysosomal proteases
cathepsin B, D and G could increase PAI-l activity in the medium of human umbilical
vein endothelial cells by cleaving PAl-l from extracellular matrix (Kimura and
Yokoi-Hayashi, l996d;Pintucci et al., 1993b;Pintucci et al., 1992d). Consistent with this,
PMN depletion reduced active PAI-l at 6hr after RAN (Fig. 2.5). A PMN proteases
inhibitor also reduced active PAI-l (Fig. 2.12). This suggests that PMNs contribute to
fibrin deposition after 2hr by releasing proteases to activate PAH and thereby inhibit
fibrinolysis. The lack of earlier effects of PMNs on active PAI—l (i.e., 2hr) might be due
to PMNs are not being activated to release proteases during that time but begin to
degranulate thereafter. In this regard, the shedding of PAH from endothelial matrix by
PMNs proteases might play a more dominate role at later times (i.e., 6hr). This is
consistent with the results showing that formation of HOCl-modified epitopes/proteins, a
marker of PMN degranulation/activation, did not increase until 3hr after LPS/RAN

treatment.

193

In LPS/RAN-treated rats, PMN depletion reduced. PIM staining at 2hr afier
LPS/RAN treatment, suggesting that PMNs contribute to early development of liver
hypoxia. This early contribution of PMNs to hypoxia did not depend on sinusoidal fibrin
deposition, since PMN deletion in LPS/RAN-treated rats did not affect liver fibrin at 2hr
after RAN (Fig. 2.2). Furthermore, PMNs accumulated in livers were not activated at 2hr
(Fig. 2.8), suggesting that enhancement of hypoxia by PMNs at this time does not require
their activation and might be mediated directly by plugging sinusoids. LPS-RAN
cotreatment caused a greater degree of tissue hypoxia than LPS given alone, although
these two treatments had a similar effect on PMN accumulation (Luyendyk et al.,
2004aiLuyendyk et al., 2005v). It is possible that RAN cotreatment causes PMNs to
adhere more firmly to sinusoidal endothelium and/or to undergo a shape change that
results in reduced sinusoidal blood flow and consequent hypoxia. In this regard, it would
be interesting to examine the rolling and adhesion of PMNs on vessel walls in rats treated
with LPS/RAN with intravascular microscopy. Also, flow cytometry could be applied to
examine the ciruculating PMN size change after LPS/RAN treatment.

In the LPS/RAN model, PMN accumulation in liver was caused largely by LPS
(Luyendyk et al., 2005w). However, LPS exposure alone did not lead to activation of the
accumulated PMNs. RAN did cause activation of the PMNs that accumulated in liver
after LPS treatment, as reflected by an increase in HOCl-protein adduct formation. RAN
can reduce PMN activation in vitro (Okajima et al., 2000b;Okajima et al., 2002b); thus,

the effects of RAN on hepatic PMN activation must be an indirect effect of RAN. This

194

might occur through MlP-2 or PAI-l, both of which are increased in LPS/RAN-treated
rats and are known to enhance PMN activation. Anti-CD18 serum did not alter PMN
accumulation but reduced PMN activation in liver (Fig. 2.9), suggesting that the early,
LPS-induced sinusoidal accumulation of PMNs does not require CD18 whereas their
activation is CD18-dependent. In addition, anti-CD18 serum markedly reduced
hepatocellular injury (Fig. 2.9), suggesting that PMN transmigration/activation is
required for LPS/RAN-induced liver injury.

There exists controversy regarding the mechanisms of activated PMN-induced cell
injury in the liver. Reactive oxygen species and PMN protease are both proposed to be
important for PMN-induced hepatocyte death (Jaeschke, 2006). PMN proteases can cause
hepatocelluar death directly in vitro (Luyendyk et al., 2005x), but the time frame for the
development of cell death is longer than that by which the liver injury develops in vivo
after LPS/RAN treatment. This suggests that PMN proteases either did not participate in
the hepatocyte killing in vivo or they need a second stress to manifest the process. The
observation that PMN proteases inhibitors protected from LPS/RAN-induced liver injury
ruled out the first possibility. Interestingly, hypoxia renders hepatocytes more sensitive to
killing by PMN protease (i.e. elastases), and the time frame reflects the one by which
liver injury develops in LPS/RAN-treated rats (Luyendyk et al., 2005y). Since hypoxia
occurs early in livers of LPS/RAN treated rats (Luyendyk et al., 20052;Deng et al.,
2007a), these results suggest that PMNs contribute to the pathogenesis by releasing

proteases that kill hepatocytes in an environment made hypoxic by fibrin deposition.

195

 

In the next section of the thesis, the role of RAN in TNFu production, PMN
activation was explored. The results suggested that RAN augmented TNFa production
after LPS treatment, and the prolongation of LPS-induced TNFu production by RAN
appears to be crucial for the liver injury caused by LPS/RAN cotreatment. This was
supported by the observation that a TACE inhibitor, given immediately before RAN,
reduced both serum TNFa protein and the hepatocellular injury. The augmentation of
TNF-01 by RAN occured in a post-transcriptional manner through enhanced
p38-dependent TACE activation. In addition, since RAN augmented hepatic p38
activation induced by LPS, p38 and downstream MK-2 may increase TNFu protein
production by accelerating translation as reported by Neinigers and the colleagues
(Neininger et al., 2002g).

The TACE inhibitor also reduced plasma active PAI-l concentration after LPS/RAN
treatment to almost the same level as after LPS treatment. Moreover, a PAI-l inhibitor
reduced the hepatocellular injury caused by LPS/RAN cotreatment. This suggested that
enhanced PAI-l production was a downstream effect of RAN-augmented TNF-a
production. Further studies showed that PAI-l was important for both hepatic fibrin
deposition and PMN activation. As mentioned above, hypoxia resulting from fibrin
deposition could potentiate the killing of hepatocytes by proteases (e. g. elastase) released
from PMNs after their activation (Luyendyk et al., 2005).

Although the PAH inhibitor decreased PMN activation, it did not affect hepatic

PMN accumulation or serum PMN chemokine concentration, suggesting a direct effect of

196

PAI-l on PMN activation. A recent study showed that PAI—l directly potentiates
LPS-induced PMN activation through a Jun-N-terminal kinase (JNK)-dependent pathway
(Kwak et al., 2006a). As mentioned above, RAN by itself did not enhance PMN
activation. In fact, it has been shown to reduce PMN activation in vitro (Okajima et al.,
2002a;Okajima et al., 2000a). All of these results suggest that RAN might induce
activation of PMNs indirectly by augmenting PAI-l production. It would be interesting to
see whether incubating primary rat PMNs with PAI-l can increase fMLP- or
PMA-induced degranulation of PMNs.

Although RAN augmented TNFu production and downstream cascades though
activating TACE, the mechanism behind the TACE activation by RAN is largely
unknown. The enhanced TACE activation after RAN treatment was p38 dependent; this
was consistent with the finding that p38 is essential for ectodomain shedding of TNFa in
CHO cells (Fan and Derynck, 1999a). Although ERK, another MAPK kinase, is able to
phosphorylate TACE (az-Rodriguez et al., 2002), there are no reports to our knowledge
that p38 directly phosphorylates and activates TACE. Thus, it is likely that p38
downstream kinases such as MK-2 participate in the activation of TACE. Further studies
need to be conducted to pursue this possibility.

In the LPS/RAN model, it seems plausible that RAN augmented the LPS-induced
inflammatory response to precipitate the hepatocellular injury. However, the initial
molecular target of RAN remains largely unknown in this model. The lack of liver injury

after cotreatment of LPS with FAM at a dose pharmacologically comparable to RAN

197

rules out the involvement of H2 receptor antagonism. Since RAN, not FAM, augmented
hepatic p38 activation induced by LPS and since p38 is crucial for the downstream
cascade of TNFa production, coagulation activation and PMN activation, it is possible
that the p38 activation pathway might be where RAN exerts its initial effect. RAN itself
did not activate p38, whereas it enhanced the p38 activation after LPS exposure. Thus,
RAN might perturb one or more of the signaling molecules in the pathway of
LPS-induced p38 activation. This pathway is well studied (Guha and Mackman, 2001).
Toll-like receptor 4 (TLR4) on macrophages serves as a pattern recognition receptor for
LPS. After recognizing LPS, TLR4 causes the recruitment of a set of intracellular
Toll/IL-l receptor (TIR) domain-containing adaptors, including myeloid differentiation
factor 88 (MyD88). The association of TLR4 and MyD88 activates p38 through a
signaling cascade involving IRAK4, TRAF6 and TAKl. It would be interesting to
examine the phosphorylation states of these signaling molecules after LPS/RAN
cotreatment to identify the possible molecular target of RAN.

The proposed pathogenic mechanism of LPS/RAN-induced liver injury is
summarized in Figure 5.1. RAN augments TNF-a production after LPS treatment in a
post-transcriptional manner by enhancing p38 activation. The increase in TNF-0t protein
appears to occur through the p38-dependent activation of TACE. The prolongation of
LPS-induced TNF-01 production by RAN appears to be crucial for the liver injury caused
by LPS/RAN cotreatment. TNF-01 leads to coagulation system activation and PAI-l

production, both of which increase hepatic fibrin deposition. PAI-l might also be directly

198

Figure 5.1 Hypothetical pathogenesis of LPS/RAN-induced liver injury. RAN
augments TNF-a production after LPS treatment in a post-transcriptional manner by
enhancing p38 activation. The increase in TNF-a protein appears to occur through the
p38-dependent activation of TACE. and the resultant prolongation of LPS-induced
TNF-u production by RAN appears to be crucial for the liver injury caused by LPS/RAN
cotreatment. TNF-a leads to coagulation system activation and PAI-l production, both of
which cause hepatic fibrin deposition. PAl-l might also be directly responsible for the
activation of hepatic PMNs accumulated after LPS exposure. The hypoxia resulting from
hepatic fibrin deposition could act synergistically with toxic proteases released from
activated PMNs to kill hepatocytes. PMN proteases are also involved in enhancing PAI-l

production and fibrin deposition.

199

 
  
  
 
   

TF \
Thrombin F'b _
1 rm
***

1 1*
/?ti

proteases

  

200

responsible for the activation of hepatic PMNs accumulated after LPS exposure. The
hypoxia resulting from hepatic fibrin deposition could act synergistically with toxic
proteases released from activated PMNs to kill hepatocytes. PMN proteases are also
involved in enhancing PAI-l production and fibrin deposition.

In addition to the LPS/RAN model, PMNs are involved in the liver injury caused by
LPS/DCLF cotreatment. This raises the possibility that PMNs are one of the universal
participants in liver injury in LPS-drug interaction models. Since PMNs cause
cytotoxicity through the release of reactive oxygen species and other toxic factors, it is
interesting in this regard that the addition of noncytotoxic concentrations of
peroxidase/11202 to hepatocyte cultures markedly increased DCLF cytotoxicity
(Miyamoto et al., 1997b). In fact, several novel reactive metabolites of DCLF formed
when the drug was incubated with PMN-derived MPO (Miyamoto et al., 1997a;Zuurbier
et al., 1990b). Thus, PMNs accumulated and activated in liver after LPS exposure are
likely to render DCLF more cytotoxic by MPO-derived metabolism. It would be
interesting to examine these MPO-derived metabolites in vivo after LPS/DCLF
cotreatment.

The mechanism of PMN activation in the LPS/DCLF model is largely known. In a
gene array study, transcripts representing several neutrophil chemokines (MIP-2 and
MIP-la) and adhesion molecules (ICAM-l) were upregulated to a greater degree in livers
of LPS/DCLF-treated rats than in rats treated with LPS alone. Serum MlP-2

concentration reflected the increase in the transcripts for this chemokine. There are

201

 

reports that COX-2 inhibition enhances MCP-l/MIP-2 production in models of
inflammation caused by radiation (Kyrkanides et al., 2002a). In addition, it has been
proposed that in the face of inhibition of COX, arachidonic acid is metabolized more
extensively by lipoxygenase. In this regard, it is interesting that lipoxygenase products
derived from arachidonic acid, such as 15- hydroxyperoxyeicosatetraenoic acid
(lS-HPETE), can increase adhesion molecules and transmigration of neutrophils across
endothelial cell barriers (Sultana et al., 1996b). Thus, shifting of the generation of
eicosanoids from the COX pathway to the lipoxygenase pathway by DCLF during
inflammation (eg. LPS exposure) might contribute to enhancing the expression of MIP~2
and subsequent PMN activation.

In summary, the research conducted in this thesis revealed the crucial roles of
PMNs in models of idiosyncratic hepatotoxicity resulting from LPS-drug interactions. In
the LPS/RAN model, LPS induced the accumulation of PMNs in the liver, and RAN
caused the activation of these PMNs. This might occur through TNFa-dependent
production of PAH. Activated PMNs can release toxic proteases to kill hepatocytes or to
enhance fibrin deposition, both of which could lead to hepatocellular injury. In addition,
PMNs are also important in the liver injury caused by LPS/DCLF cotreatment. These
results contribute to the understanding of mechanisms by which inflammation could act
as a susceptibility factor for IADRs and to the possible identification of biomarkers for

IADRs resulting from inflammation-drug interaction. In addition, the conducted study

202

could lead to improved strategies (i.e., proteases inhibitors) for preventing or treating

conditions involving PMN-dependent tissue injury.

203

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