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LSU Historical Dissertations and Theses
Graduate School
1995
Elucidation and Modulation of CEB Metabolism in
Fischer 344 Rats: Possible Mechanisms of Toxicity
and Anticarcinogenicity.
Judith Sara Prescott-mathews
Louisiana State University and Agricultural & Mechanical College
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Prescott-mathews, Judith Sara, "Elucidation and Modulation of CEB Metabolism in Fischer 344 Rats: Possible Mechanisms of
Toxicity and Anticarcinogenicity." (1995). LSU Historical Dissertations and Theses. 6045.
http://digitalcommons.lsu.edu/gradschool_disstheses/6045
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ELUCIDATION AN D MODULATION OF CEB METABOLISM
IN FISCHER 344 RATS:
POSSIBLE MECHANISMS OF TOXICITY AND ANTICARCINOGENICITY
A Dissertation
Submitted to the Graduate Faculty of the
Louisiana State University and
Agricultural and Mechanical College
in partial fulfillment of the
requirements for the degree of
Doctor of Philosophy
in
The Interdepartmental Program in Veterinary Medical Sciences
through the Department of Veterinary Pathology
by
Judith S. Prescott-Mathews
B.S., University of Tennessee, 1987
D.V.M., Louisiana State University, 1991
August, 1995
UMI Number: 9609119
OMI Microform 9609119
Copyright 1996, by OMI Company. All rights reserved.
This microform edition is protected against unauthorized
copying under Title 17, United States Code.
UMI
300 North Zeeb Road
Ann Arbor, MI 48103
Dedicated to my husband,
George
ACKNOWLEDGMENTS
I am greatly indebted to a number of individuals whose guidance,
expertise and assistance have been instrumental towards the completion of
my dissertation research and in nurturing my career in veterinary clinical
pathology. I would like to thank my major professor, Dr. Jan VanSteenhouse,
for her support and guidance. I would additionally like to thank Dr.
VanSteenhouse, as well as Dr. Stephen Gaunt, for their insight and experience
in clinical pathology which have proven invaluable in my residency training
program.
I am also appreciative of the other members of my advisory committee,
Dr. Wayne Taylor, Dr. David Swenson, Dr. Lamar Meek, and Dr. Steven
Barker, for the advice and guidance regarding these investigations and this
dissertation manuscript. I am additionally thankful to Dr. Steven Barker for
his expertise in analytical techniques and for his patience, guidance, and
encouragement. I would like to thank Connie David for graciously assisting
with the GC-MS analysis.
My appreciation is extended to Julie Millard for her devoted friendship,
support, and assistance. Her technical expertise proved invaluable to these
investigations. I am also thankful to Pam Stratton for her able secretarial
assistance throughout my graduate student career and to Mae Lopez and
Micheal Broussard for their assistance in preparing photomicrographs and
graphics for this manuscript, respectively.
I would like to acknowledge Dr. Fred Enright for providing access to
the instrumentation necessary for the histomorphometric measurements and I
am grateful to Dr. John Lynn for spending considerable time and effort
establishing the histomorphometric technique. Additionally, Dr. Tamara
Schaller and Dr. Mark McLaughlin are appreciated for synthesizing the CEB.
Finally I would like to thank my family for their support and
encouragement. I am grateful to my parents for teaching me the value of
education and for their guidance and reassurance. I am also thankful to my
in-laws for readily taking m e into their family and for believing in me much
more than is realistic I am eternally indebted to my husband, George, who
can find a positive aspect in the most negative situation. I am thankful to
George for his unconditional love, support, and friendship.
TABLE OF CONTENTS
DEDICATION.........................................................................................................
ii
ACKNOWLEDGMENTS......................................................................................
iii
ABSTRACT..................................................................
vii
CHAPTER 1 INTRODUCTION, LITERATURE REVIEW,
AND OBJECTIVES........................................................................
Introduction....................................................................................
Influence of Dietary Constituents on Carcinogenesis
Cruciferous Vegetables....................
Glucosinolates ..............................................
Functions of Glutathione............................................................
Glutathione Metabolism and Transport....................................
N itriles
............................................................................
Objectives.............................................................
1
1
2
26
31
40
48
73
83
CHAPTER 2 PROTECTIVE EFFECT OF AMINOOXYACETIC ACID ON
CEB-INDUCED NEPHROTOXICITY IN FISCHER 344
RATS..............................................................
87
Introduction.................................................................................... 87
Materials and M ethods...........................................
90
Animal Maintenance and Treatment..................
90
Materials.............................................................................. 91
93
Tissue Preparation..................................................
Biochemical A ssa y s
.......
93
Histological and Histomorphometric Examination.... 94
Statistical Analysis............................................................
95
R esults............................................................................................. 95
Renal and Hepatic G SH ................................................... 95
Renal and Hepatic GCS A ctivity.................................... 98
Histological Evaluation.................................................... 101
Karyometry......................................................................... 101
D iscussion
........................................................................... 116
CHAPTER 3 EFFECT OF CEB ON RENAL AND HEPATIC XENOBIOTIC
METABOLIZING ENZYMES IN FISCHER 344 RATS
125
Introduction ............................................................................... 125
Materials and M ethods................................................................ 131
Animal Maintenance and Treatment............................ 131
v
Materials..............................................................................
Tissue Preparation.............................................................
Biochemical A ssa y s...................
.................................................................................
R esults
Renal and Hepatic GCS....................................................
Renal and Hepatic GST....................................................
Renal and Hepatic Cytochrome P-450, EROD, PROD,
andECO D ...........................................................................
Renal and Hepatic Epoxide H ydrolase........................
D iscussion........................................ ..............................................
CHAPTER 4 IDENTIFICATION OF A CEB-DERIVED URINARY
MERCAPTURIC ACID IN FISCHER 344 RATS.....................
Introduction....................................................................................
Materials and M ethods.................................................................
Animal Maintenance and Treatment.............................
Materials.................................................
Biochemical A ssa y s.........................................
Gas Chromatography-Mass Spectrometry...................
Sample Preparation..............................................
Gas Chromatography-Mass Spectrometry
Statistical Analysis
....................................................
R esults.............................................................................................
Urinary Total Nonprotein Thiols....................................
Urinary Thioethers...................
Gas Chromatography-Mass Spectrometry...................
D iscussion.......................................................................................
132
134
135
138
138
138
138
143
143
152
152
157
157
158
160
161
161
161
162
162
162
164
164
180
CHAPTER 5 GENERAL CONCLUSIONS........................................................ 187
LITERATURE CITED...................
196
APPENDIX............................................................................................................... 262
VITA......................................................................................
266
ABSTRACT
The ability of acivicin (AT-125), probenecid, and aminooxyacetic acid
(AOAA), to ameliorate l-cyano-3,4-epithiobutane (CEB)-induced
nephrotoxicity and karyomegaly was evaluated by histopathology and
histomorphometry. Epithelial cell necrosis and karyomegaly were observed
in the renal proximal tubules at 24 and 48 hours in all of the treated groups
w ith the exception of AOAA pretreated rats at 24 hours. A predominant
urinary metabolite of CEB was identified as the mercapturate, N-acetyl-S- (4cyano-2-thio-l-butyl)-cysteine. Evidence of other urinary metabolites was also
demonstrated.
The influence of inhibitors on the GSH-enhancing effect of CEB was
determined through measurement of tissue GSH levels and y-glutamylcysteine synthetase (GCS) activity. Hepatic and renal GSH levels were
significantly elevated in rats treated with CEB alone or CEB in addition to any
of the inhibitors, with the exception of hepatic GSH at 24 hours in probenecidpretreated rats. Renal GCS activity was significantly decreased at 24 hours in
all of the treated groups with the exception of AOAA-pretreated rats.
Activities of xenobiotic metabolizing enzymes were evaluated. No
significant alterations were observed in renal or hepatic cytochrome F-450,
ethoxyresorufin O-deethylase (EROD), pentoxyresorufin O-depentylase
(PROD), and epoxide hydrolase (EH) activities. Renal and hepatic
ethoxycoumarin O-deethylase (ECOD) and glutathione S-transferase (GST)
activities, however, were significantly decreased in rats administered CEB.
These results indicate a protective effect of AOAA against CEB-induced
nephrotoxicity at 24 hours. AOAA inhibits the formation of a reactive thiol
vii
suggesting that such a metabolite may be responsible for CEB-induced
nephrotoxicity. The reactive thiol also appears to inhibit renal GCS activity.
The slight but statistically significant decrease in ECOD and GST are of
questionable biological significance. However, diminished ECOD activity
may be associated with the anticarcinogenic potential attributed to cruciferous
vegetables.
Identification of the urinary mercapturate represents detection of a
unique compound and provides further evidence of the importance of GSH
conjugation in CEB metabolism.
CHAPTER 1
INTRODUCTION, LITERATURE REVIEW,
AND OBJECTIVES
Introduction
This review focuses on various constituents of the human diet,
particularly compounds derived from cruciferous vegetables, that have
demonstrated anticarcinogenic potential in vitro and in vivo. Though
com pounds derived from cruciferous plants have demonstrated toxic
effects w hen fed in high concentrations, these compounds may act as
potential chernoprotective agents by inducing or inhibiting various
xenobiotic metabolizing enzymes and enhancing intracellular glutathione
(GSH) levels.
l-Cyano-3,4-epithiobutane (CEB) has been shown to be derived from
glucosinolates found in cruciferous vegetables. Previous investigations
have demonstrated nephrotoxicity as well sustained elevations in renal
GSH in Fischer 344 rats following administration of CEB (VanSteenhouse,
et al., 1989). This dissertation research was therefore designed to elucidate
the mechanism of nephrotoxicity and GSH-enhancing effect of CEB as w ell
as the effect of CEB on xenobiotic metabolizing enzymes.
The anticarcinogenic potential of various dietary constituents, the
influence of these compounds on xenobiotic metabolizing enzym es, and
the effect of these enzymes on anticarcinogenicity w ill be reviewed.
Additionally, the functions, metabolism, and transport of GSH and the
GSH-enhancing effect and toxicity of cyanoepithioalkanes similar to CEB
w ill be described in detail.
1
2
Influence o f Dietary Constituents
on Carcinogenesis
It has been estimated that approximately 35% of human cancer in the
Western world may relate to the diet (Doll, 1992). Statistics such as this
indicate a necessity for in depth evaluation of dietary constituents. Indeed,
a myriad of chemical constituents from the human diet have been
identified as natural carcinogens or anticarcinogens.
Since it has become apparent that complete avoidance of dietary
carcinogens is realistically impossible, there has recently been an emphasis
on the elucidation of dietary anticarcinogens. Deliberate consum ption of
anticarcinogens may directly or indirectly counteract the effects of
carcinogens in the human diet (Ferguson, 1994). More than 500 chemical
com pounds have been evaluated thus far as potential chemoprotective
agents (Boone et ah, 1992).
Natural cancer chemoprotective agents have been classified as dietary
inhibitors of mutagenesis or dietary inhibitors of. Mutagenesis is a welldefined phenomenon referring to the induction of a permanent
transmissible alteration in the genetic code (Cotran et a l, 1989).
Carcinogenesis, on the other hand, is a complex multistep process
consisting of at least two stages - initiation and promotion. The
anticarcinogenic mechanisms of dietary components are usually unclear
and the inherent complexity of carcinogenesis makes it difficult to pinpoint
the specific process that is inhibited.
Although initiation is probably synonym ous with mutation, an
antimutagenic agent should not be automatically considered
anticarcinogenic. Ellagic acid, a plant polyphenol, demonstrated
antimutagenic potential against benzo(a)pyrene diol epoxide in the Ames
3
Salmonella assay and Chinese hamster V79 cells (Wood et al., 1982). In
v i v o testing in mice with ellagic acid, however, demonstrated an absence of
inhibition towards benzo(a)pyrene-induced pulmonary tumors (Chang et
al., 1985) and a lack of inhibition towards 3-methylcholanthrene-induced
skin tumor formation (Smart et ah, 1986). Adsorption of ellagic acid to nongenetic substances such as proteins and oxidative breakdown of its
polyphenolic structure may interfere with its effects (Hayatsu et al., 1988).
Many anticarcinogens have demonstrated antimutagenic effects,
however, and may be generally classified as bioantimutagens or
desmutagens (Kuroda, 1990). Bioantimutagens or "true" antimutagens, as
defined by Clarke and Shankel (1975), act on the repair and replication
processes of damaged DNA. Desmutagens, on the other hand, are defined
as apparent antimutagens and pertain to any factor that directly inactivates
mutagens or their precursors.
Mechanisms of desmutagens include binding or adsorbtion of dietary
mutagens, scavenging of free radicals, and inactivation of chemically
reactive metabolites (Hartman and Shankel, 1990). Chlorophyllin binds
planar compounds through the formation of molecular complexes
(N egeshi et al., 1989) while some dietary fibers adsorb mutagens from
solution (Roberton et al., 1991). Desmutagens, such as carotenoids and
retinoids, scavenge free radicals and prevent oxidative damage (Ames,
1983). Finally, desmutagens may inactivate chemically reactive metabolites
or prevent the formation of reactive species through the induction or
inhibition of phase I and II biotransforming enzyme systems. The phase I
and II metabolizing enzyme systems are responsible for the biochemical
processes that convert lipophilic compounds to more hydrophilic
4
metabolites. Lipophilic compounds are more likely to diffuse into cellular
membranes and become w ell distributed throughout the body whereas
hydrophilic compounds are more easily excreted in the urine, bile, feces,
expired air and perspiration. (Sipes and Grandolfi, 1986).
Although phase I and II enzymes are localized mainly in the liver,
the literature indicates that every tissue analyzed thus far has demonstrated
activity towards some xenobiotic. The liver receives blood from the
splanchnic area which contains not only nutrients but various foreign
substances, or xenobiotics, as well. These compounds are metabolized by
the liver prior to release into the systemic circulation. Other tissues
involved in biotransformation reactions include lung, kidney, intestine,
skin, and gonads; however, these tissues are limited with respect to the
diversity of chemicals they can detoxify as compared to the liver (Sipes and
Gandolfi, 1986).
The need to metabolize various hydrophobic foreign com pounds
presents organisms with a unique challenge. Since it would be impractical
to produce one enzyme for each compound an organism comes into contact
with, families and subfamilies of enzymes with generally broad substrate
specificity are encoded to metabolize a diversity of xenobiotics.
Phase I enzymes are localized primarily in the endoplasmic
reticulum and inner mitochondrial membrane (Nebert et al., 1982; Black
and Coon, 1987). These membrane-bound enzyme complexes are readily
accessible to their lipophilic substrates. Phase I enzymes generally
participate in oxidation, reduction, or hydrolysis reactions and may add or
expose functional groups (eg. -OH, -SH, -NH 2 , -COOH) to form hydrophilic
metabolites (Sipes and Gandolfi, 1986).
5
One of the m ost important and, certainly, the most well-know n and
extensively studied phase I enzyme system is the cytochrome P450 (P450)
system. This system is less commonly referred to as the polysubstrate
monooxygenase system or the mixed-function oxygenase system. The
Nomenclature Committee of the International Union of Biochemistry
prefers the term "heme-thiolate" protein instead of "cytochrome" for P450
since these proteins are not "cytochromes" in the true meaning of its
terminology (Palmer and Reedijk, 1989). "Cytochrome P450" represents the
original term designated by Sato and Omura (1961).
Sato and Omura (1961) identified an unusual protein in liver
microsomal (smooth endoplasmic reticulum) fractions capable of carrying
out mixed-function oxidation reactions. Mixed-function oxidation
reactions involve the combination of one oxygen atom with two hydrogen
atoms to form water while the second oxygen atom is incorporated into the
substrate. This protein was shown to bear a heme group which, when
bound to a molecule of carbon monoxide (CO), produced an absorbance
spectrum with a peak at a wavelength of 450 nm. Sato and Omura (1961)
therefore referred to this hemoprotein as "cytochrome P450".
Coon and Lu (1968) later identified two proteins in rabbit liver
microsomes involved in the electron-transport chain of the m ixed function
oxidation reaction. These investigators demonstrated that the flow of
electrons began with NADPH and proceeded to a flavoprotein. The
electrons were subsequently transferred to the P450 molecule. They
additionally demonstrated the necessity of phospholipids for the functional
activity of these proteins. Lu and coworkers (1970) later identified these
proteins in rat liver.
6
The P450 system is actually a membrane-bound coupled enzym e
complex composed of an NADPH-cytochrome P450 reductase flavoprotein
and cytochrome P450 molecule. Cytochrome bs and cytochrome bs
reductase are also associated with this complex.
The substrate for this enzyme system binds to the oxidized form of
the P450 (Fe+3) molecule to form a substrate-P450 complex. This complex
initiates the first electron reduction and accepts an electron from NADPH
or cytochrome bs via NADPH-cytochrome P450 reductase or cytochrome bs
reductase, respectively. The iron in the P450 is hence reduced to the ferrous
(Fe+2) state. The ferrous form of the enzyme combines with a m olecule of
oxygen gas, and accepts a second electron from NADPH or cytochrome bs in
addition to a proton (H+). Heterolytic cleavage of the oxygen molecule
results in the release of water (H 2 O) leaving a (FeO) + 3 complex that directly
oxidizes the substrate (Guengrich, 1993). Although substrate binding
initiates the catalytic cycle, reduction and cleavage of molecular oxygen is
rate-limiting (White and Coon, 1980).
Each P450 enzyme appears to be encoded by a separate gene. The few
exceptions to this rule occur where functional alternative splicing might
exist (Lacroix et a l, 1990; Lephart et a l, 1990; Miles et al., 1990; Means et al.,
1991). To date, more than 40 different P450 genes have been identified in
the rat and at least 20 P450s have been identified in humans (Guengrich,
1993). Though the enzymes are encoded by different genes, many exhibit
overlapping substrate specificities.
Until recently the classification of P450 genes has been ambiguous
and inconsistent with investigators utilizing diverse systems of
nomenclature. Recently, Nebert et al. (1989 and 1991) classified P450 genes
7
on the basis of structural an d /or evolutionary relationships.
Complementary DNA sequences with at least 40% identity were grouped
into families and given an Arabic number designating the P450 family (1,2,3
etc). Sequences within families with at least 60% identity were grouped into
subfamilies designated with letters (A, B, C, etc.) while individual enzym es
were given specific Arabic numbers (1, 2, 3, etc.) unless only one gene is
present within the subfamily. Some confusion remains regarding P450
nomenclature, however, since the last number may refer in one instance to
a specific animal species but in other instances may apply to all species
(Guengrich, 1993). Additionally, cytochrome P450 genes may be given the
prefix "cyp" for the corresponding gene in mice or "CYP" for genes isolated
in other species (Nebert et ah, 1991).
As early as the 1950's investigators demonstrated induction
(upregulation) of cytochrome P450 enzyme activity (Brown et al., 1954). A
variety of compounds such as drugs, ethanol, substances in cigarette smoke,
and certain constituents of food have since been found to induce or
suppress the activity of cytochrome P450 genes (Diaz et a l, 1990; Murray,
1992; Guengrich, 1993). Each type of inducer increases the synthesis of a
characteristic range of P450s with each enzyme revealing a distinct, but not
necessarily unique, substrate and reaction specifity (Guengerich et al., 1982).
There does not, however, appear to be a close association between the ability
of a chemical to induce a cytochrome P450 enzyme and its identity as a
substrate for that particular enzyme.
In 1980, Bresnick demonstrated increased levels of mRNA specific for
P450 enzym es following administration of phenobarbitol (PB) or 3methylcholanthrene (3MC) to rats. These agents have since been shown to
8
differentially induce P450 enzymes with distinct but overlapping substrate
specificities in a variety of species (Yang and Lu, 1987). 3MC-pretreatment of
rats caused selective induction of P450 enzymes with spectral and catalytic
properties distinctly different from the PB-inducible P450s (Lu et al., 1973).
One of the 3MC-inducible enzymes exhibited high catalytic turnover for
benzo(a)pyrene and produced an absorbance spectrum with a peak at a
wavelength of 448 nm instead of 450 nm. This enzyme was thus referred to
as cytochrome P448. Other terms for this 3MC-inducible enzym e include
cytochrome Pi-450 and aryl hydrocarbon hydoxylase (AHH; Sipes and
Grandolfi, 1986).
The polycyclic aromatic hydrocarbons, including 3MC, have since
been shown to induce the 1A subfamily of P450s whereas PB induces the 2B
subfamily as well as other subfamilies (Okey, 1990). Interest in the
regulation of P450s originally stemmed from the observation that
administration of structurally diverse chemicals resulted in induction of
one or more distinct P450 enzymes. Various agents have been synthesized
to serve as model substrates for the assay of microsomal O-dealkylation as a
measurement of induced P450 enzymes. Liver microsomes from the rat
and hamster O-deethylated ethoxyresorufin to resorufin in an NADPH-,
0 2
dependent reaction involving NADPH-cytochrome reductase and
cytochrome P450 (Burke and Mayer, 1974). In both the rat and the hamster,
ethoxyresorufin O-deethylation was considerably induced by 3MC (50-70
fold) while its specific activity was induced less than 3-fold by PB.
Ethoxyresorufin O-deethylase (EROD) activity has, accordingly, been shown
to be highly selective for 3MC-induced P450 enzymes or the 1A subfamily
(Guengerich et al., 1982).
-
9
The pentyl homologue of ethoxyresorufin, pentoxyresorufin, appears
to also be highly selective. In contrast to ethoxyresorufin, pentoxyresorufin
is selective for PB-induced P450s (Burke and Mayer, 1983) since its activity is
enhanced follow ing administration of PB. Pentoxyresorufin O-depentylase
(PROD) activity therefore correlates with induction of the IB subfamily.
Alternatively, the O-dealkylation of 7-ethoxycoumarin was induced 3-fold
by 3MC and 7-fold by PB in hepatic microsomes from C57BL/6-J mice
(Greenlee and Poland, 1978). 7-Ethoxycoumarin O-deethylase (ECOD)
activity is therefore a measure of induction of both the 1A and 2B
subfamilies.
The selective induction of P450 enzymes is regulated by the
expression of P450 genes under the control of a diverse multiplicity of
regulatory mechanisms. P450 genes may be regulated at the level of
transcription, processing, mRNA stabilization, translation, or enzym e
stability in the presence of various inducing agents (Gonzalez et al., 1993).
The mechanism of induction of certain P450 enzymes by polycyclic
aromatic hydrocarbons has spurred tremendous scientific interest.
Cytochrome P450 1A1 monoxygenase activity, often measured as aryl
hydrocarbon hydroxylase (AHH) activity, was induced in certain inbred
strains of mice (e.g. C57BL/ 6 -J) following administration of SMC, while
other inbred strains (e.g. D B A /2) failed to respond (Nebert et al., 1972;
Thom as et al., 1972). The trait, referred to as aromatic hydrocarbon
responsiveness, was found to be inherited in a sim ple autosomal dominant
mode. The locus was designated the "Ah" locus (Thomas et al., 1972).
2,3,7,8-Tetrachlorodibenzo-p-dioxin (TCDD), a model halogenated aromatic
hydrocarbon, induced AHH or P450 1A1 activity in all strains of mice tested;
10
however, the 3MC-nonresponsive mice required a 10-fold greater dose of
TCDD as compared to the 3MC-responsive strain (Poland and Glover, 1975).
It was then discovered that TCDD, as w ell as other polycyclic aromatic
hydrocarbons, bind to an "Ah" receptor, a product of the "Ah" locus
(Poland et al., 1976).
Under normal conditions, the Ah receptor is complexed w ith a 90 kD
heat-shock protein (hsp 90) within the cytoplasm (Poland et al., 1976). Once
TCDD binds to the "Ah" receptor, the hsp 90 is displaced (Pongratz et al.,
1992). The TCDD-receptor complex then binds another protein, the Ah
receptor nuclear translocator (ARNT) molecule (Reyes et al., 1992), which
must be phosphorylated to an activated form (Carrier et al., 1992). The
activated complex then enters the nucleus and binds to a specific region on
the P450 1A1 gene referred to as the dioxin or xenobiotic regulatory
element. The activated protein-receptor complex has been show n to bind to
the regulatory sequence in gel shift and footprinting experiments (Denison
et al., 1985,1988). Binding of the complex to this enhancer sequence allows
other proteins to access the promoter region of the gene thereby enhancing
synthesis of mRNA and inducing the cytochrome P450 1A1 protein.
Although stimulation of transcription increases P450 mRNA
concentrations, enhanced mRNA stability has also been show n to elevate
P450 activity (Kimura et al., 1986).
The mechanism of induction of the P450 2B subfamily as w ell as
other subfamilies by PB also occurs through transcriptional activation. To
date, however, no receptor has been identified in mammals (Waxman and
Azaroff, 1992).
11
Other phase I metabolizing enzyme systems include the amine
oxidase system, epoxide hydrolase, esterases and aminidases, and alcohol,
aldehyde and ketone oxidation-reduction systems (Sipes and Gandolfi,
1986). Amine oxidase is an FAD-flavoprotein, localized in the endoplasmic
reticulum, capable of oxidizing nucleophilic nitrogen and sulfur atoms.
Nonspecific esterases and amidases hydrolyze ester or amide linkages,
respectively. Esterases liberate a carboxyl group and the corresponding
alcohol while amidases release a carboxyl group and an amine or ammonia.
Esterases and amidases are localized in both the endoplasmic reticulum and
the cytosol (Sipes and Gandolfi, 1986).
Alcohol and aldehyde dehydrogenases, aldehyde and ketone
reductases, and aldehyde oxidase are all involved in the oxidation or
reduction of aldehydes, ketones, and alcohols produced during the
oxidation of carbon or hydrolysis of ester linkages. These various enzymes
have been identified in the endoplasmic reticulum as well as the cytosol
(Sipes and Grandolfi, 1986).
Epoxide hydrolases have recently been reviewed by Guenther (1990)
and represent a group of catalytically-related but distinct enzym es that
catalyze the hydration of arene oxides and aliphatic epoxides to the
corresponding dihydrodiols. Epoxide hydrolases are believed to have
evolved as detoxifying enzymes which serve to protect cells from the
deleterious effects of epoxides or oxiranes formed by virtue of the
cytochrome P450 enzyme system or lipid peroxidation.
Discrepancies plague the nomenclature of this enzyme. Early
investigators referred to it as 'epoxide hydratase’ or 'epoxide hydrase*.
When the Nomenclature Committee of the International Union of
12
Biochemistry suggested the enzyme be renamed 'epoxide hydrolase' (EG
3.3.2.3) in 1978, it was not widely recognized that several forms of epoxide
hydrolase existed. Two distinct forms of epoxide hydrolase were later
isolated. One was found primarily within the microsomal fraction and the
other w ithin the cytosol and have since been referred to as "microsomal
EH" or "mEH" and "cytosolic EH" or "cEH", respectively (Guenther, 1990).
Inconsistencies in this nomenclature arose, however, w hen mEH,
also referred to in the literature as 'mEHb' (Oesch, et al., 1984), 'EHT
(Guenther and Oesch, 1983), and 'mCSO' (Finley and Hammock, 1988), was
isolated not only from the endoplasmic reticulum but also from the nuclear
membrane, plasma membrane and golgi (Stasiecki et al., 1980) as w ell as the
cytosolic fraction from human liver (Wang et al., 1982) and lung (Guenther
and Karnazis, 1986). mEH demonstrates a broad substrate specificity and
catalyzes the hydrolysis of a wide variety of arene and alkene oxides to
dihydrodiols (Guenther, 1990). The best substrates for mEH are oxiranes
with one or two hydrophobic substituents such as styrene oxide (Oesch et
al., 1971a). Czs-disubstituted oxiranes are generally better substrates for mEH
as compared to frans-disubstituted oxiranes (Oesch et al., 1971b).
mEH exhibits a wide tissue distribution and has been isolated from
numerous organs in the rat. The highest activity of mEH in the rat was
found in the liver, followed by the testis, kidney, ovary, and lung (Oesch et
al., 1977; Seidegard and DePierre, 1983). The order of activity in the mouse
w as testis>liver>lung>skin>kidney>intestine (Oesch et al., 1977).
Similarly, cEH, also referred to as 'EH2 ' (Guenther and Oesch, 1983)
and 'cTSO' (Finley and Hammock, 1988), has demonstrated the greatest
activity in the cytosol but has been identified in small amounts from the
13
microsomal fraction (Guenther and Oesch, 1983) and nuclear membrane
(Guenther and Karnezia, 1986). Like mEH, cEH demonstrates a broad
substrate specificity; however, fraws-disubstituted oxiranes are rapidly
hydrolyzed by cEH in contrast to cis-disubstituted oxiranes (Gill et ah, 1983a;
Hammock and Hasagawa, 1983). The complementary substrate preference
between cEH and mEH may aid in the detoxification of an extremely w ide
range of endogenous and exogenous compounds.
cEH activity is also observed in various tissues but differs markedly
in organ distribution and activity between species. Hepatic cEH activity is
highest in the mouse, intermediate in humans, rabbits, and guinea pigs,
and low in the rat (Gill and Hammock, 1980; Ota and Hammock, 1980; Gill
et ah, 1983a, Gill et ah, 1983b; Meijer et ah, 1987; Meijer and DePierre, 1988;
Mertes et al., 1985). In mice, the order of activity in various organs has been
show n to be liver > kidney > lung > testis > spleen (Gill and Hammock,
1980; Loury et ah, 1985) while in the rat the order is heart = kidney > liver >
brain > lung > testis > spleen (Gill et ah, 1983b).
Additionally, a unique microsomal EH, known as cholesterol 5,6oxide hydrolase, 'chEH' (Guenther, 1990), 'mEHch' (Oesch, et ah, 1984), or
'mCE' (Finley and Hammock, 1988), catalyzes the hydration of steroid 5,6oxides. cEH is located predominantly in the endoplasmic reticulum with
smaller amounts present in the mitochondrial fraction, but not the cytosol
(Aringer and Eneroth, 1974; Astrom et ah, 1986; Sevanian and McLeod,
1986). chEH exhibits a narrow substrate specificity. Steroid 5,6-oxides are
the only known substrates while analogs of cholesterol 5,6-oxides and the
3,5,6-triol product are the only known inhibitors of the enzym e (Guenther,
1990; Nashed, et ah, 1985; Sevanian and McLeod, 1986). chEH is structurally
14
and catalytically distinct from mEH as demonstrated by different substrate
and inhibitor specificities (Levin et al., 1983; Oesch et al., 1984; Kaur and
Gill, 1986; Palakodety et a l, 1987) and a lack of cross-reactivity between chEH
and mEH-specific antibodes (Levin et a l, 1983; Oesch et a l , 1984).
A unique cytosolic epoxide hydrolase involved in leukotriene
biosynthesis has additionally been characterized (Evans et a l, 1985; McGee
and Fitzpatrick, 1985; Ohishi et a l, 1987). Leukotriene A 4 (LTA4 ) hydrolase
or LTA4 H hydrolyzes the oxirane ring of LTA4 to form LTB4. LTA4 H is
located exclusively in the cytosol and differs catalytically from both cEH and
mEH in that the addition of water by LTA4 H occurs at a position distant
from the oxirane ring due to delocalization of the carbocation in the
conjugate triene structure of the substrate (Guenther, 1990). Like chEH,
LTA4 H demonstrates a narrow substrate specificity since only LTA4 and
closely related analogs of LTA4 act as substrates (Evans et a l, 1985; McGee
and Fitzpatrick, 1985; Ohishi et a l, 1987). LTA4 H differs from mEH and cEH
by virtue of its molecular weight, stability, amino acid composition,
substrate specificity, mechanism of action, and suicide inactivation via its
substrate, LTA4 (McGee and Fitzpatrick, 1985; Ohishi et a l, 1987; Guenther,
1990).
LTA4 H has been isolated from a variety of tissues from the guinea
pig with the highest activity found in the small intestine and lesser
amounts in the adrenal, liver, lung, kidney, leukocytes, stomach, heart,
brain, and spinal cord (Izumi et al., 1986). LTA4 H was also present in rodent
(Evans et al., 1985; Haeggstrom et a l, 1985; Pace-Asciak et a l, 1985; Medina et
a l , 1987) and human (Evans et al, 1985; Haeggstrom et al., 1985; Ohishi et
15
al., 1987) lung, liver, kidney, and neutrophil as well as human erythrocytes
(McGee and Fitzpatrick, 1985).
Phase II biotransforming enzyme systems are usually localized in the
cytosol and catalyze biosynthetic or conjugation reactions (Sipes and
Gandolfi, 1986). Various high-energy intermediates or cofactors are
activated to accomplish these transfer reactions. Covalent linkage of
endogenous m olecules to the parent compound or a phase I-derived
metabolite not only confers increased water solubility to facilitate excretion
but may also contribute a functional moiety enabling the metabolite to
participate in various transport systems (Aitio, 1978). Phase II enzym es
include glucuronosyltransferases, sulfotransferases, methyltransferases,
N-acetyl transferases, ATP-dependent acid:CoA ligases, rhodaneses and
glutathione S-transferases (Sipes and Gandolfi, 1986).
Glutathione S-transferases (GSTs; EC 2.5.1.18) are an extensive family
of multifunctional isoenzym es found in various species including bacteria,
yeast, trematodes and nematodes, insects, fish, rodents, and humans
(Ketterer and Mulder, 1990; Buetler and Eaton, 1992). Although the primary
function of GSTs involves catalyzing the conjugation reaction between
GSH and a variety of endogenous and exogenous electrophiles, these
enzym es may also catalyze organic nitrate and thiocyanate reduction,
thioester formation, (Chasseaud, 1979) and the isomerization of
prostaglandins (Christ-Hazelhof and Nutgeren, 1979). Alternatively, GSTs
may function as intracellular carrier proteins by binding non-substrate
hydrophobic molecules including heme, bile acids, bilirubin, thyroid
hormones, and various drugs such as steroids, sulfobromophthalein, and
16
iodinated contrast agents (Mannervik and Danielson, 1988; Ishigaki et al.,
1989; Deleve and Kaplowitz, 1990).
In a similar manner, GSTs may act as a nucleophilic target for
electrophilic metabolites thereby removing toxic compounds through noncatalytic covalent binding (Jakoby, 1978; Listowsky et al., 1988). This has
been proposed as a 'suicidal' defense mechanism since the enzym e
m olecule is often inactivated in the process (van Bladeren and van
Qmmen, 1991; Pabst et al., 1974; Corrigall et al., 1989; Adam s and Sikakana,
1990).
Over 100 GST isoenzymes have been characterized from a variety of
species (Buetler and Eaton, 1992). Multiple forms of cytosolic GST arise
from various dimeric combinations of separately functioning subunits. To
date, 13 subunits have been identified in human tissues (Awasthi et al.,
1994) while 13 different subunits have been demonstrated in rats (Sternberg,
1991).
Various physical and biochemical criteria have been utilized to
catagorize GST isoenzymes including substrate specificity (Boyland and
Chasseaud, 1969), subunit molecular weight values (Taylor et al., 1987 Dao
et al., 1982), relative motility on sodium dodecyl sulfate-polyacrylamide gel
electrophoresis (McLellen and Hayes, 1987) or starch gels (Board, 1981),
elution from carboxymethyl cellulose ion-exchange columns (Jakoby, 1978),
and isoelectic point focusing of homodimers (Warholm et al., 1981,1983;
Singh et al., 1988).
The use of diverse criteria has resulted in a complex, convoluted
system of nomenclature. In 1984, Jakoby and coworkers devised a
nomenclature in rats using arabic numerals for subunits identified
17
chronologically. This naming system proved somewhat advantagous;
however, Mannervik, et al. (1985) later recommended classifying
mammalian, including rat, mouse and human, GSTs into three major
classes based on partial sequence homologies, imm unological similarities,
and kinetic properties. These three classes were designated a , p, and re.
Although the majority of GST proteins in human muscle belong to
the GST p and re classes, a small but significant portion of GSH conjugating
activity of human muscle is represented by isoenzym e GST £ which is
imm unologically distinct from the a, p, and re classes of GST (Singh et al.,
1988). This isoenzyme appears to be specific to skeletal muscle and
expresses interindividual variations.
Buetler and Eaton (1992) have established a fifth class of GST
isozym es, o, (in addition to a, p, and re) to include S-crystallins from
mollusk lens derived from squid and octopus. Their classification of GSTs
is based upon computational analysis of cDNA sequences. According to this
classification system, three recently purified GSTs, one from human and
two from rat (Meyer et ah, 1991; Hussey and Hayes, 1992), have been placed
in yet another GST class, 0. These GST isoenzymes are distantly related to
cytosolic a, p, and % class GSTs. This new class also includes many of the
non-mammalian GST isoenzym es from bacteria, yeast, insects, and plants
and has become the 'catch all' class of archaic GSTs. It is theorized that the
gene for the 0 class GST may be the parent gene from which the various
classes of GST have evolved (Buetler and Eaton, 1992).
It should be noted that the o and 0 class do not appear to be
universally accepted by all investigators as several authors do not include
them in the classification of GSTs. These should be considered minor
18
classes since greater than 90% of GST proteins from human tissue are
isoenzym es of the a, ji, and n classes (Awasthi et al., 1994).
Though GSTs are found primarily within the cytosol, two
membrane-bound forms have been identified in the endoplasm ic
reticulum and the outer mitochondrial membrane from both humans and
rats (Kraus, 1975; Morgenstem and DePierre, 1988; Wahllander et al., 1979;
M orgenstern et al., 1982). Membrane-bound GSTs have been found in all
mammalian species studied thus far and constitute 3% and 5% of the total
protein in the endoplasmic reticulum and the outer mitochondrial
membrane, respectively (Morgenstern et al., 1984).
Though rat liver exhibits several 100 times more enzym e activity
than extrahepatic tissues, membrane-bound GST proteins have been
demonstrated via Western blot analysis in the intestinal mucosa, adrenal,
testes, thymus, lung, and spleen; however, essentially none of this protein
has been identified in the heart, brain, or kidney (Morgenstern et al., 1984).
Conversely, Northern blot analysis demonstrated mRNAs expressed at 20%
of the hepatic level in the kidney and testes, with much lower levels in the
lung, spleen, and brain (Dejong et al., 1988). Though the reason for the
discrepancies between Western and Northern blot analysis are not known,
it is suggested that a different subcellular localization of membrane-bound
GST proteins may occur in different organs since only the microsomal
fraction was analyzed. Additionally, there may be different turnover rates
for the protein and the mRNA (Andersson et al., 1994).
Membrane-bound GSTs have been shown to be structurally distinct
and share minimal sequence homology with cytosolic GSTs (Buetler and
19
Eaton, 1992). Additionally, microsomal GSTs were not induced by common
inducers of drug-metabolizing enzym es (Morgenstern et al., 1980).
GST subunits within a given class form homo- and heterodimers or,
in the case of microsomal GSTs, trimers with each subunit maintaining
functional independence. Each subunit has a complete active site
(Danielson and Mannervik, 1985; Tahir and Mannervik, 1986) with two
adjacent binding sites. The glutathione-binding site, or G site, is highly
specific with the only other endogenous thiol substrate being yglutam ylcysteine (Sugimoto et a l, 1985). The GSH thiol moiety is thought
to be activated or deprotonated to a reactive thiolate ion by the hydroxyl
group on a highly conserved active-site tyrosine residue (Graminski et al.,
1989).
The electrophile-binding site, or H site, is located in the same general
region as the G site; however, there is considerable variance in the
associated amino acid residues (Gilliland, 1993). Variant amino acid
residues dictate the substrate specificity for each isozyme and an increased
degree of variance is obviously associated with a w ide range of substrate
specificity for the GSTs in general. It is proposed that the H site activates the
substrate in a similar manner to the activation of GSH at the G site
(Mannervik and Danielson, 1988). To increase electrophilicity, and activate
the substrate, an amino acid residue must protonate the substrate.
Although GSH reacts with electrophilic carbon, oxygen, nitrogen, and
sulfur atoms, the term GSH conjugation generally refers to products formed
by the attack of GSH on an electrophilic carbon (Ketterer and Mulder, 1990).
Although a diverse range of compounds act as substrates for GSTs
(Chasseaud, 1979; Vos and van Bladeren, 1990), the typical reactions of GST
20
conjugation include nucleophilic diplacement of good leaving groups,
typically halides, from saturated and aromatic carbons, Michael additions,
and nucleophilic attack on strained ring structures (Ketterer and Mulder,
1990). Nucleophilic displacement from a saturated carbon occurs when
GSH conjugates a-bromoisovalerylurea and displaces the bromide to yield
N-(aminocarbonyl-2-glutathion-S-yl)-3-methylbutanamide (Mulder and te
Koppele, 1988). Bromide is also displaced from ethylene dibromide
(Peterson and Guengrich, 1988) while chloride is displaced from
hexachlorobutadiene during GST-catalyzed conjugation with GSH (Wolfe
et al., 1984). Nucleophilic displacement reactions can also occur at aromatic
carbons. This type of reaction has been observed with l-chloro-2,4dinitrobenzene and l,2-dichloro-4-nitrobenzene (Habig et ah, 1974; Keen et
al., 1976). Alternatively, GSTs may catalyze Michael additions with
substrates such as diethyl maleate (Boyland and Chasseaud, 1967) and the
toxic metabolite of acetominophen, N-acetyl-p-benzoquinone imine
(Hinson et a l, 1982; Coles et a l, 1988).
Reactions of GSH w ith substrates containing strained ring structures
have typically been observed with oxirane rings or epoxides. Benzo(a)pyrene (BP) is metabolized to various epoxides including BP-7,8-diol-9,10oxide, which is selectively conjugated by GSTs (Cooper et al., 1980; Hesse et
al., 1982), and BP-4,5-oxide (Nemoto and Gelboin, 1975; Nemoto et a l, 1975;
Hernanandez et a l, 1980; Plummer et a l, 1980), the most abundant epoxide
metabolite of BP. The strained oxirane ring creates an electrophilic carbon
susceptible to attack by GSH.
It should be noted that GSH conjugation reactions can occur
spontaneously and need not always be catalyzed by GSTs. Soft electrophiles,
21
compounds with a low charge density or readily polarizable reactive
centers, are more likely to react spontaneously with GSH since GSH is a soft
nucleophile. Conversely, hard electrophiles have a high charge density or
polarized reactive center and more typically require catalysis by GSTs
(Ketterer et al., 1986; Deleve and Kaplowitz, 1990).
Spontaneous reactions with GSH have been demonstrated with
quinones such as 1,4-naphthoquinone (Takahashi et al., 1987) and Nacetylbenzoquinone imine (Ketterer et al., 1986). Spontaneous reactions
have additionally been demonstrated with dihaloethanes and acrylonitrile
and its metabolite, 2-cyanoethylene oxide (Peterson and Guengerich, 1988).
l-Chloro-2,4-dinitrobenzene (CDNB), a commonly utilized substrate for
measuring GST activity, undergoes noncatalytic conjugation with GSH;
however, the spontaneous reaction rate of CDNB with GSH is only 0.01% of
the total rate of conjugation (Ketterer, 1982; Ketterer et al., 1983).
Spontaneous GSH conjugation with soft electrophiles typically occurs
at a much slower rate relative to GST catalyzed conjugation. Catalysis by
GSTs not only increases the rate of conjugation, but affords independence
over the intracellular GSH concentration. This allows GSH conjugation to
occur rapidly in cells with low GSH concentrations (Ketterer et al., 1986).
Though all mammalian tissues possess some cytosolic GST activity
(Mannervik, 1985), there is usually a spectrum of isoenzym es which is
characteristic for the specific tissue. The isoenzyme pattern of distribution
as w ell as the isoenzym e concentration may be strikingly different from one
tissue to another. For example, rat liver contains the greatest amount of
total GST protein (3-5% of total soluble protein) as compared to other
organs and has significant amounts of all subunits except subunit 7 which is
22
only minimally expressed (Vos and van Bladeren, 1990). Similarly, human
liver exhibits high levels of a and n class GSTs but only trace amounts of n
which is analogous to rat subunit 7 (Awasthi et al., 1994). Substantial
amounts of GST n, however, have been demonstrated in humans in the
adrenal gland, cornea, gastric and small intestinal mucosa, testis, uterus,
pancreas, retina, and kidney (Awasthi et al., 1994) and subunit 7 is found in
moderate to large amounts in rat kidney, lung, and intestine (Vos and van
Bladeren, 1990).
Isoenzyme distribution patterns also vary depending on species, sex,
age, genetic composition, and exposure to inhibiting or inducing agents. In
human fetal liver, as opposed to adult liver, GST n, as well as a GSTs, are
expressed in substantial quantities (Awasthi et ah, 1994).
H Class GSTs exhibit extensive genetic polymorphism. Only 40-50%
of adult humans possess this class of GSTs (Vos and van Bladeren, 1990).
The interindividual variation of /j class GSTs has been associated with
variable susceptibility to various xenobiotics (Warholm et al., 1983).
When Seidegard and colleagues (1986) measured frans-stilbene oxide
activity, most specific for GST p, in lymphocytes of patients with lung
cancer and controls matched for smoking habits, a greater proportion of the
control smokers (without cancer) exhibited GST n activity as compared to
lung cancer patients. Additionally, fewer patients with adenocarcinomas
appeared to to be positive for GST ]i as compared to patients diagnosed with
squamous cell carcinomas or small or large cell carcinomas. A similar
population-based, case-control study measuring cytosolic activity towards
frans-stilbene oxide, l-chloro-2,4-dinitrobenzene, and benzo(a)pyrene-4,5oxide in patients with smoking-related cancer, demonstrated
23
proportionately fewer cancer patients with intermediate or high GST
activity towards frans-stilbene oxide as compared to controls. In this latter
study, however, the difference was plausibly due to chance (Heckbert et al.,
1992). These studies suggest that fi class GSTs may protect smokers from
poly cyclic aromatic hydrocarbons and that n class proteins may serve as
genetic markers for susceptibility to lung cancer. In fact, polymerase-chain
reaction-based methods for the detection of genetic polymorphisms in
human n class GST activity have been developed for rapid, accurate
detection of the GST n gene (Bell, 1991).
GST isoenzym e distribution patterns are also subject to hormonal
influences. Distinct sex differences have been noted in rats, mice, and
humans (Igarashi et al., 1987; McLellen and Hayes, 1987). Both the male rat
and m ouse exhibit higher enzyme activity in the liver than their female
counterparts and male rat liver contains higher levels of subunits 3 and 4
while female rat livers preferentially express subunits 1 and 2 (Igarashi et
al., 1987). In humans, there are gender related differences in the expression
of GSTs within the colon (Singhal et al., 1992) and skin (Singhal et a l, 1993).
GST isoenzym e expression is also mediated by various endogenous
and exogenous inhibitors. An extensive list of inhibitors of GST activity
has been published by Mannervik and Danielson (1988) and the various
types of inhibition have been reviewed recently by van Bladeren and van
Ommen (1991). GST activity may be inhibited in either an irreversible or
reversible manner, the latter of which occurs most commonly.
Reversible inhibitors of GST include GSH analogs as w ell as "second
substrate" analogs which bind to the G site or H site of GSTs, respectively.
Obviously, 'second substrate' analogs include a diverse range of inhibitors
24
consistent with the broad substrate specificity of GST isoenzymes.
Essentially, the only common feature of this group of inhibitors is their
lipophilic nature (van Bladeren and van Ommen, 1991). Many of the
substrates for GST also act as reversible inhibitors.
GST activity may additionally be reversibly inhibited by
'nonsubstrate ligands' such a heme, bilirubin, and bile acids. Interestingly,
this binding occurs at a site distinct from the G or H binding sites and may
cause conformation changes in the enzyme which alter its activity. Binding
of nonsubstrate molecules to GST is believed to be a transport or storage
function of the enzyme although no convincing evidence exists concerning
these functions (van Bladeren and van Ommen, 1991).
Alternatively, few compounds have demonstrated irreversible
inhibition of GST activity. Various quinones are among the strongest of
these types of inhibitors with both tetrachloro- 1,4-benzoquinone and its
GSH conjugate inhibiting rat and human GST (van Ommen et al., 1988;
Ploem en et al., 1991). Most GSH conjugates are inhibitors of GSTs and the
more hydrophobic the S-substituent, the greater the inhibitory effect
(Ketterer and Mulder, 1990).
Inhibition of GST may have multiple effects in vivo (van Bladeren
and van Ommen, 1991). While diminished GST activity may be of
therapeutic benefit in allergic reactions via the inhibition of leukotriene
formation, inhibition of prostaglandin synthesis may have toxic effects
since these com pounds are necessary for maintaining vasodilation within
tissues (Cotran et al., 1989). GST catalyzed conjugation of electrophilic
xenobiotics exhibits a similar dualism. Decreased GST activity may prove
detrimental due to decreased detoxification of alkylating agents. However,
25
increased GST concentrations have been documented in various tumor
cells and especially tumors that have become resistant to cancer
chemotherapeutic agents (Waxman, 1990). It is plausible that elevated
levels of GST isoenzym es are associated with the drug resistant state.
Ethacrynic acid, a known inhibitor of (i class GSTs, has been shown to
potentiate the cytotoxicity of several cancer chemotherapeutics in tumor
cell lines (Tew et al., 1988; Smith et al., 1989; Hansson et al., 1991) and
implanted tumor cells in mice (Clapper et al., 1990).
Conversely, a large and diverse group of compounds, both naturally
occurring and synthetic, induce GST isoenzymes. Induction of cytosolic
GSTs in rat liver has been demonstrated following treatment with
phenobarbital (Ding et al.,1986; Pickett et al., 1987; Hales and Neims, 1977;
Ding and Pickett, 1985; Pickett et al., 1984; Grasl-Kraupp et al., 1993), 3methylcholanthrene (Pickett et al., 1987; Hales and Neims, 1977; Ding and
Pickett, 1985; Pickett et al., 1984), frans-stilbene oxide, and
hexachlorobenzene (Di Simplicio et al., 1983; Vos et al., 1988). Phenobarbital
additionally induces GST activity in rat small intestine and testis (Clifton
and Kaplowitz, 1978; Sheenan et al., 1984) while 3-methylcholathrene
induces GSTs in rat lung, small intestine, and kidney (Stewart and Boston,
1987; Clifton and Kaplowitz, 1978). Microsomal GST activity in rat liver is
similarly induced by treatment with phenobarbital (Aniya et al., 1993).
Naturally occurring compounds may also induce GST activity.
N um erous investigators have demonstrated elevations in GST activity in
animals fed cruciferous diets or degradation products derived from these
plants (Sparnins et al., 1982a, b; Aspry and Bjeldanes, 1983; Chang and
Bjeldanes, 1985; Salbe and Bjeldanes 1985; Ansher e t a l , 1986; Kensler et al.,
26
1987; Vos et al., 1988; Bogaards et al., 1990; Wortelboer et al., 1992; Zhang et
al., 1992).
The induction of GSTs is believed to be an important mechanism of
protection against chemical carcinogenesis (Prochaska et al., 1985; Prochaska
and Talalay, 1988; Talalay et al., 1988) while inhibition of GST is theorized to
increase human cancer risk (Schneider, 1992). Rat and human GST a
inhibits covalent D N A binding of food-borne carcinogens (Lin et al., 1994).
Similarly, the dietary antioxidants, 2(3)-ferf-butyl-4-hydroxyanisole and 1,2dihydro-6-ethoxy-2,2,4-trimethylquinoline, enhance hepatic GST activities
in mice and decrease the level of mutagenic metabolites formed from
benzo(a)pyrene (Benson et al., 1978). It is similarly believed that induction
of GST may be at least partially responsible for the anticarcinogenic effect of
cruciferous vegetables.
Cruciferous Vegetables
The family Cruciferae, characterized by four petals standing opposite
one another in a square cross (Bailey, 1949), consists of well over 2,000 plant
species (Harberd, 1976). As a result, the classification of these plants poses
major difficulties to the taxonomist. Fortunately, the economically
important members are relatively few in number including those
cultivated for human an d /or animal consumption and grown as
ornamentals. The Brassica genus encompasses the majority of the
economically important members including koch and black mustard (B.
nigra varieties), cabbage, cauliflower, broccoli, kale, kohl-rabi, cole wart, and
Brussels sprouts (B. oleracea varieties), rape and rutabagas (B. napus
varieties), turnip rape and Chinese cabbage (B. campestris varieties), and
mustard (B. jancea) (Prakash and Hinata, 1980).
27
Though in ancient times crucifers were used primarily as medicinals
(Fenwick et al., 1983), cruciferous crops now account for a significant
portion of the diets of people throughout parts of the world. Brassica
species, including B. napus (rape) and B. campestris (turnip rape) are
cultivated for their high oil content. These oilseeds are utilized in the
production of margarine and soap, as a lubricating oil, and as a diesel fuel
substitute (Fenwick et al., 1983). Rapeseed and crambe or Abyssinian kale
(Crambe Abyssinian) are also cultivated as major oilseed crops for plastics
and processed foods (Lenman, 1992).
The seed meal that remains following oil extraction contains 40-45%
protein and is an excellent protein source for livestock and poultry
(Lenman, 1992). The use of this meal, however, is limited by its pungent
odor and taste as well as its toxic effects observed in the liver, kidneys, and
thyroid (Tookey et al., 1980; Fenwick et al., 1983).
Cruciferous plants are additionally utilized in condiments including
mustard (B. juncea, B. nigra, and Sinapis alba) and horseradish (Am oracia
rusticana) and the roots (radish, turnip), stems (kohl-rabi), flower buds
(cauliflower, broccoli), and leaves (cabbage, Brussels sprouts, Takana,
N ozawana, Hiroshimana) are commonly consumed as vegetables
(MacLeod, 1976; Tookey et al., 1980; Fenwick et al., 1983; Uda and Maeda,
1986).
Cruciferous vegetables including cabbage, broccoli, turnips,
cauliflower, and Brussels sprouts have become widely regarded as potential
cancer chemoprotectives. Based on extensive experimental testing of crude
vegetable extracts and epidemiologic data, both the National Research
Councils' Committee on Diet, Nutrition, and Cancer (1982) and the
28
American Cancer Society (1984) have recommended increased
consumption of cruciferous vegetables to decrease human cancer incidence.
While occasional studies do not corroborate an inverse relationship
between cancer and cruciferous vegetables (MacClure and Willett, 1990;
McKeown-Eyssen, 1984), an abundance of epidemiologic studies indicate
that the consumption of cruciferous vegetables reduces the risk for
developm ent of cancer at various sites including the colon, rectum, bladder,
lung, thyroid, pancreas, skin, stomach, and m esothelium (Graham et al.,
1978; Graham et a l, 1983; Kune et al., 1987; Hoff et al., 1988; Schiffman et al.,
1988; Young and Wolf, 1988; Lee et al., 1989; LeMarchand et al., 1989; Olsen
et al., 1989; Chyou et al., 1990; Kolonel et al., 1990; Wohlleb et al., 1990; Olsen
et al., 1991; Kune et al., 1992; Swanson et al., 1992).
In experimental animal models, investigators have reported a
reduced incidence of mammary cancer in dimethylbenz(a)anthracenetreated rats fed diets containing cabbage and cauliflower (Wattenberg, 1983).
There was also a significant decrease in the number of pulmonary
metastases in mice fed cabbage and collards after being injected
intraveneously with BALB/ c mammary carcinoma cells (Scholar et al.,
1989). Animals fed cruciferous diets have similarly demonstrated
anticarcinogenic effects against aflatoxin Bi-induced neoplasia (Boyd et al.,
1982).
Similarly, supernatant from crude homogenates of cruciferous
vegetables have demonstrated antimutagenic activity against 2 -amino- 3 m ethylimidazo[4,5-f]quinoline, 2-amino-3,4-dimethylimidazo[4,5-f]quinoline, 2-amino-3,8-dimethylimidazo[4,5-f]quinoxaline in the Am es test
(Edenharder et al., 1994) while boiled extracts of crucifers demonstrated
29
significant "anti-tumor-promoter and radical-scavenging" activities in the
phorbol myristate acetate/Epstein-Barr virus/B lymphocyte system (Maeda
et a l, 1992).
Despite the well-established inverse relationship between cruciferous
vegetables and the incidence of various types of human cancer, the specific
constituents responsible for this protective effect have not been well
defined. Although vegetable fiber has been advocated as a risk-lowering
factor for some types of cancer such as colon cancer, there has been a failure
to establish a consistent inverse relationship between cancer and dietary
fiber (Doll, 1992). Similarly, vitamin A and (3-carotene may contribute to
the protective effects of crucifers against lung cancer in humans (National
Research Council, 1982); however, experimental data regarding the indices
of vitamin A intake were derived from food that may contain other
anticarcinogenic agents. It is therefore plausible that dietary constituents
other than vitamin A or (3-carotene may act as risk-reducing factors
(National Research Council, 1982). In fact, a population-based study of diet
and lung cancer incidence suggested that other components of crucifers
appear to have a stronger inverse relationship with risk than (3-carotene
(LeMarchand et al., 1989).
Chemoprotective effects are often attributed to induction of phase II
xenobiotic metabolizing enzymes. Induction of GST activity in liver and
small intestinal mucosa has been demonstrated in rats and female mice fed
semi-synthetic diets supplemented with raw or cooked Brussel sprouts
(Sparnins et ah, 1982b; Salbe and Bjeldanes 1985; Bogaards et ah, 1990;
W ortelboer et ah, 1992). Similarly, diets supplemented with dried
powdered preparations of cabbage increased GST activity in the liver and
30
small intestine of female mice (Sparnins et al., 1982b) while broccoli
supplemented diets elevated GST levels in rat liver (Aspry and Bjeldanes,
1983). Occasional investigators observed no induction in GST activity in
animals fed cruciferous diets (Hendrich and Bjeldanes, 1986; Nijhoff et al.,
1993).
Various constituents of cruciferous vegetables have been show n to
induce GST activity. Isothiocyanates (Bogaards et al., 1990; Sparnins et al.,
1982a; Vos et al., 1988; Zhang et al., 1992), indole-3-carbinol (Sparnins et al.,
1982a), goitrin (Bogaards et al., 1990; Chang and Bjeldanes, 1985) and
dithiolthiones, such as oltipraz (Kensler et al., 1987; Ansher et al., 1986) are
derived from cruciferous vegetables and have been shown to induce GST
activity in rat and mouse liver and small intestine. Oltipraz induces
hepatic GST activity via changes in steady state levels of GST mRNA and
rates of GST gene transcription (Davidson et al., 1990).
Several natural substances derived from cruciferous vegetables
including isothiocyanates, indoles, and dithiolthiones have been show n to
possess anticarcinogenic properties (National Reseach Council, 1982).
Indole-3-carbinol, a naturally occurring indole found in cruciferous plants,
inhibits carcinogen-induced neoplasms of the forestomach, mammary
gland, and tongue in mice and rats (Wattenberg and Loub, 1978; Tanaka et
a l, 1992).
Similarly, a variety of organic isothiocyanates have demonstrated
chemoprotective effects against various types of cancer. A recent review by
Zhang and Talalay (1994) outlines the ability of isothiocyanates to protect
against carcinogen-induced tumors in rats and mice at various locations
including liver, mammary gland, esophagus, forestomach, and lung.
31
Occasional studies have examined the chemoprotective effects of
glucosinolates. Administration of glucobrassicin (indolylmethylglucosinolate) or glucotropaelin (benzylglucosinolate) prior to DMBA
exposure decreased both the incidence and multiplicity of mammary
tumors in rats (Wattenberg et al., 1986). The interpretation of such studies
is complex. It is not known to what extent the glucosinolates are degraded
in vivo. Additionally, if the glucosinolates are hydrolyzed, it is unclear
whether endogenous or microbial enzymes are responsible and what
products are formed. Finally, it is not known whether it is the intact
glucosinolate, a hydrolysis product, or a metabolite of either of these that is
responsible for the anticarcinogenic effects.
Glucosinolates
Glucosinolates are thioglucosides found predominantly, but not
exclusively, in plants of the Cruciferae family. These compounds occur
ubiquitously throughout the order Capparales including not only the
Cruciferae but the Capparaceae, Moringaceae, Resedaceae, and Tovariaceae
Families as well (Fenwick et al, 1983). Glucosinolates also occur
sporadically in other families including the Caricaceae, Euphorbiaceae,
Gyrostemonaceae, Limnanthaceae, Salvadoraceae, and Tropalolaceae
(Fenwick et al., 1983).
Glucosinolates, or their degradation products, are responsible for the
pungent flavor of many of the cruciferous vegetables (Tookey et al., 1980).
Though the biological function of these compounds is still unkown, it is
theorized that glucosinolates or gucosinolate-derived compounds act as
insect attractants or stimulants. Glucosinolates have been shown to affect
the egg-laying behavior and feeding patterns of various insects
32
(Thorsteinson, 1953; Coaker, 1969). It is proposed that these compounds
may act as protectants against predators and parasites since glucosinolates
and associated hydrolysis products have been reported to be fungistatic,
bacteriostatic, and toxic to insects as well as mammals (Erickson and Feeney,
1974; Fenwick et al., 1983; Chew, 1988a) Alternatively, glucosinolates may
serve as a sulphur storage pool since the glucosinolate concentration in
vegetative tissue increases with addition of sulphur fertilizer (Lenman,
1992).
The evolution of the nomenclature of glucosinolates has been
reviewed by Fenwick et al. (1983). In the early 1800's, the first glucosinolates
isolated from cruciferous vegetables were given classical names such as
sinalbin and sinigrin, derived from white mustard seed (Sinapis alba L.)
and seeds of black mustard (Brassica nigra Koch), respectively. In 1960, a
new system of nomenclature was adopted in which the glucosinolate was
given the prefix "gluco" followed by part of the Latin species or botanical
name of the plant from which it was originally isolated. Finally, a system
was suggested with regards to the chemical structure of the glucosinolate
(Ettlinger and Dateo, 1961) in which a chemically-descriptive prefix proceeds
"glucosinolate".
The basic structure of most glucosinolates (Figure 1.1) consists of a pD-thioglucose sugar and a sulfonated oxime moiety both bound to the
terminal carbon of a variable organic side chain "R" (Ettlinger and
Lundeen, 1956). The variable side chains of glucosinolates most commonly
consist of alkenyls, alkyls, aryls, and indolyls as well as hydroxylated or
sulfated alkyls and alkenyls (Muuse and Van der Kemp, 1987; Fenwick et
al., 1983). Though most glucosinolates possess the same basic structure,
H
R1 \
Figure
1 .1
NOSO3
Basic structure of glucosinolates.
34
occasional glucosinolates, such as p-hydroxybenzyl-glucosinolate isolated
from radish seed (Rapltanus sativus), contain a 6 -sinapoylthioglucose
moiety instead of the typical thioglucose group (Linscheid et al., 1980). The
ramifications of this alternative group are still unknown. It is possible that
such glucosinolates could produce novel breakdown products that cause
distinct physiological effects.
Glucosinolates are biosynthesized within the plant from amino acids
including tryptophan, tyrosine, leucine, valine, alanine, and methionine
(Kjaer, 1976; Moller 1981). Amino acids are oxidatively decarboxylated to an
aldoxime which is sulfurated to a thiohydroximate. The latter intermediate
is subsequently glycosylated to a desulfoglucosinolate which undergoes
sulfation to the glucosinolate. 3'-Phosphoadenosine 5'-phosphosulfate
(PAPS) has been shown to act as the sulfur donor in the final step of
glucosinolate biosynthesis (Glendening and Poulton, 1988).
Glucosinolates are found within all tissues of Cruciferae including
the root, stem, leaf, and seed (Fieldsend and Milford, 1994a). The type and
amount of glucosinolate as well as the location of the glucosinolate within
the plant are highly variable (Josefsson, 1967) and are dependent on various
environmental and agronomic conditions. Glucosinolate levels and the
types of gluconsinolates present are dependent on the developmental stage
of the plant (Cole, 1978; Sukhija et al, 1985; Fieldsend and Milford, 1994b),
the part of the plant examined (Joseffesson, 1967; Lenman, 1992), the
processing conditions (Kondo et al., 1985; de Vos and Blijleven, 1988),
storage conditions (Fales et al., 1987), cultivation conditions (Heaney and
Fenwick, 1985; Fieldsend and Milford, 1994a), and the genetic composition
(Paik et al., 1980; Bell, 1984; Martland and Butler, 1984; Fieldsend and
35
Milford, 1994a). Considerable variation in glucosinolate levels may be seen
not only among species, but among varieties or cultivars within a species
(Josefsson, 1967; Fieldsend and Milford, 1994a; Fieldsend and Milford,
1994b).
Glucosinolates normally remain intact within the plant but undergo
enzymatic hydrolysis when plant cells are disrupted by crushing, cutting, or
chewing (Tookey et al., 1980; de Vos and Blijleven, 1988). Glucosinolates
have also been shown to be degraded chemically (Lanzani et al., 1976;
Gronowitz et al., 1978) and thermally upon cooking (MacLeod et al., 1981).
Thioglucosidase glucohydrolase (EC 3.2.3.1), normally stored
separately from glucosinolates in plant tissues (Lenman, 1992), represents a
group of isoenzymes responsible for the hydrolysis of glucosinolates. The
hydrolytic products include an organic aglucone moiety representative of
the variable side chain and equimolar quantities of glucose and sulfate ions
(de Vos and Blijleven, 1988; Tookey et al., 1980). As with many enzymes,
especially those constituting large families of isoenzymes, there is some
confusion within the literature regarding their nomenclature. This
enzyme is referred to as 'glucosinolase' (Ettinger et al., 1961) in the older
literature and is commonly termed 'myrosinase' throughout recent articles
although the International Union of Biochemistry recommended
thioglucosidase glucohydrolase as the preferred name in 1965 (Florkin and
Stotz, 1965). Frequently, the name of the enzyme is truncated to
thioglucosidase in many articles although thioglucosidases are not
exclusive to plants and have been identified in fungi (Reese et al., 1958),
bacteria (Oginsky et al., 1965), and mammals (Goodman et al., 1959).
36
Thioglucosidase glucohydrolase produces a variety of hydrolytic
products from the variable side chain of their glucosinolate substrates. The
spectrum of products formed depends upon the chemical structure of the
original glucosinolate, the conditions of hydrolysis, the characteristics of the
thioglucosidase glucohydrolase isoenzyme, and the presence of various
compounds which modify enzyme activity (Tookey et al., 1980; Fenwick et
al., 1983; de Vos and Blijleven, 1988; Lenman, 1992).
Depending on these factors, glucosinolate hydrolysis may produce
isothiocyanates, thiocyanates, oxazolidine- 2 -thiones, nitriles or
hydroxynitriles, amines, or epithionitriles (Ettlinger and Lundeen, 1957;
Fenwick et al., 1983). In some cases, isothiocyanates appear to simply
rearrange to give rise to epithionitriles by sulfur migration (Cole, 1976).
Under certain conditions of processing, however, relatively greater
amounts of epithionitriles, rather than the expected isothiocyanates, may be
formed from cruciferous plants. The autolysis of crambe or rapeseed meal
formed a predominance of nitriles and epithionitriles including l-cyano- 2 hydroxy-3,4-epithiobutane and 1-cyano-2-hydroxy-3-butene (Daxenbichler et
al., 1967; Daxenbichler et al., 1968). Similarly, the hydrolysis of seeds or
meal from Brassica campetris (turnip rape) under selected conditions
produced l-cyano-3,4-epithiobutane as a major product (Cole, 1975; Kirk and
MacDonald, 1974).
It has also been demonstrated that the formation of epithionitriles,
including cyanoepithioalkanes, is dependent on the presence of ferrous ion
(Fe+2) and a 30-40 kilodalton protein referred to as epithio specifer protein
(ESP) which directs the inclusion of sulfur into a terminal unsaturation
(Tookey and Wolff, 1970; Tookey, 1973; Tookey et al., 1980).
37
Cyanoepithioalkanes are formed from the rearrangement of alkenyl
isothiocyanates derived from alkenyl glucosinolates (Cole, 1976) such as 2propenyl glucosinolate (sinigrin), 3-butenyl glucosinolate (gluconapin), and
4-pentenyl glucosinolate (glucobrassica nipin) (deVos and Blijleven, 1988).
According to this observation, l-cyano-3,4-epithiobutane (CEB) would be
derived from 3-butenyl glucosinolate which is found in relatively high
concentrations in Brussels sprouts, swede, turnip rape, Yellow Sarson, and
turnips (Josefsson et al., 1967; Josefsson and Appelqvist, 1968; Downey et al.,
1969; Kondra and Downey, 1969; Heaney and Fenwick, 1985). In fact, CEB
has been demonstrated as a major hydrolysis product from B. campestris
(turnip rape and Chinese cabbage), Alyssum perenne (madwort), B. rapa
(turnip), B. napus (rapeseed), Hirchfieldia incana (hoary mustard),
Lobularia maritima (sweet alyssum), Cakile maritima Scop, (sea rocket), and
Turritis glabra (tower-mustard) (Kirk and MacDonald, 1974; Cole, 1975; Cole,
1976; Paik et al., 1980) as well as the leaves of B.campestris L. var. rap if era
(Nozawan) and B.campestris L. var. pekinensis (Hiroshimana) used for
pickling in Japan (Uda and Maeda, 1986).
While the administration of intact glucosinolates has resulted in
various toxic effects in vivo, it is generally believed that the hydrolysis
products derived during processing or by the intestinal microflora are
responsible for glucosinolate-related toxicoses. Diets rich in glucosinolates
cause marked decreases in feed intake, weight gain, and food conversion in
poultry (Turner, 1946; Clandinin et al., 1966; Campbell and Smith, 1979;
Mawson et al., 1944a), swine (Nordfeldt et al., 1954; Aherne and Lewis, 1978;
Bell et al., 1991; Mawson et al., 1944a), rodents (Kennedy and Purves, 1941;
Bell, 1957; Belzile et al., 1963; Vermorel et al., 1987; Nugon-Baudon et al.,
38
1990; de Groot et al., 1991; Smith and Bray, 1992; Mawson et al., 1944a), and
ruminants (DePeters and Bath, 1986; Mawson et al., 1944a) and have
demonstrated goitrogenic effects in pigs (Bell and Belzile, 1965; Ochetim et
al., 1980; Rundgren, 1983; Mawson et al., 1944b), rats (Kennedy and Purves,
1941; Vermorel et al., 1988; Mawson et al., 1944b), poultry ( Bell and Belzile,
1965; Elwinger, 1986; Mawson et al., 1944b), ruminants (Iwasson et al., 1973;
Seim iya et al., 1991; Mawson et al., 1944b), and rabbits (Chesney et al., 1928).
Dietary gluconsinolates additionally cause hepatotoxicity in poultry
(Jackson, 1969; Umenura et al., 1977, Martland and Butler, 1984) and rats
(Mawson et al., 1944b) and hemolytic anemia in ruminants (Greenhalgh et
al., 1969) They also cause enlargement of the liver and kidneys in swine
(Nordfeldt et al., 1954; Bowland et al., 1963) and rodents (Miller and
Stoewsand, 1983; Vermorel et al., 1986; de Groot et al., 1991).
Due to the adverse effects of glucosinolates, the Food and Drug
Administrations' Center for Veterinary Medicine has provided strict
guidelines for the use of cruciferous meals as a feedstuff for livestock and
poultry (Price et al., 1993). Plant breeders have developed cultivars
containing low levels of glucosinolates (0.5% as compared to 5-6% for other
cultivars) which are referred to as 'single low' varieties if they are low in
glucosinolates or 'double low' if they are genetically low in both
glucosinolates and erucic acid (Lenman, 1992). 'Single low' and 'double
low' cultivars can be fed at higher concentrations and are excellent protein
sources for livestock and poultry.
Despite the toxic effects of glucosinolates, numerous studies have
revealed a protective effect against naturally occurring and synthetic
carcinogens in animals administered products of glucosinolate degradation
39
(Wattenberg, 1977; Wattenberg and Loub, 1978; McDanell et a l, 1988;
Ramsdell and Eaton, 1988; Morse et al., 1989; Ishizaki et al., 1990;
Wattenberg, 1990; Bailey et al., 1991; Bradfield and Bjeldanes, 1991; Rao et al.,
1991; Steinmetz and Potter, 1991; Stoner et al., 1991; Chung et al., 1992;
Tanaka et al., 1992; Benson, 1993; Chung et al., 1993; Preobrazhenskaya et al.,
1993; Smith et a l, 1993; Sugie et al., 1993; Jiaro et al., 1994; Kojima et al., 1994;
Stresser et a l, 1994; Zhang and Talalay, 1994). Many investigators have
theorized, and some have actually demonstrated, that the anticarcinogenic
effect of these compounds is the result of induction and /or inhibition of
phase I and phase II metabolizing enzyme systems (McDanell et a l, 1988;
N ugon-Baudon et a l, 1990; Smith et al., 1990; Rao et al, 1991; Steinmetz and
Potter, 1991; Baldwin and LeBlanc, 1992; Loft et a l, 1992; Ferguson, 1994;
Stresser et a l, 1994; Zhang and Talalay, 1994). Although elevations in
cytochrome P450 enzymes may accelerate the metabolism and excretion of
toxic xenobiotics, induction can either activate or detoxify carcinogenic
compounds depending on the carcinogen and the specific P450 enzyme.
Recent studies, however, support the conclusion that inactivation of phase
I enzym es may diminish the formation of toxic metabolites thereby
reducing DNA-adduct formation and carcinogenesis (Ishizaki et a l, 1990;
Smith et a l, 1990; Zhang and Talalay, 1994).
In tandem with the inhibition/ activation of phase I enzym es,
products of glucosinolate hydrolysis may block carcinogenesis through the
induction of phase II enzymes, particularly GST. It is assumed that
glucosinolates must be hydrolyzed during processing or by the intestinal
microflora to attain their inducing effect since intact glucosinolates do not
themselves induce xenobiotic metabolizing enzymes in germ-free rats fed
40
glucosinolate-rich diets (Rabot et al., 1993). In addition, several
glucosinolate degradation products induce GSTs in the lung, liver, and
kidney (Sparnins et al., 1982a, 1982b; Rao et al., 1991; Smith and Bray, 1992;
Benson, 1993; Kore et al, 1993).
Functions of Glutathione
Glutathione ( y-glutamylcysteinylglycine; Figure 1.2) is a ubiquitous
tripeptide widely distributed in animal tissues, plants, and many
microorganisms (Larsson et al., 1983). Glutathione occurs primarily
intracellularly at concentrations ranging from 0.1- 10 mM (Meister, 1988a)
with the highest concentrations existing in ocular lens epithelium and
cortex (Hockwin and Korte, 1990). As such, it is the most prevalent
intracellular thiol in almost all biological species studied and accounts for
>90% of the total non-protein thiol concentration within most cells
(Meister, 1983).
Most of the intracellular glutathione exists in the reduced, or thiol,
form (GSH), although mixed disulfides (G-SS-protein), thioethers, and, to a
lesser extent, the glutathione disulfide (GSSG) contribute to the total
cellular pool of glutathione. The ratio of reduced glutathione (GSH) to
oxidized glutathione (GSSG) within the cell is typically 300 ± 60 (Qian and
Murphy, 1993). GSSG is accumulated within the cell to a limited extent due
to the action of GSSG reductase (EC 1.6.4.2). This reductase is a flavoprotein
which utilizes NADPH as an electron donor for the reduction of GSSG to
GSH (Stryer, 1988).
The majority of intracellular GSH is found within the cytosol despite
the existence of small but significant mitochondrial and nuclear pools
(Reed and Olaffsdottir, 1989; Romero and Galaris, 1990; Jevtovic-Todorovic
Figure
1 .2
Structure of glutathione (y-glutamylcystemylglycine).
42
and Guenther, 1992). Additionally, GSH and GSSG are also present in
various body fluids including plasma, bile, and urine, although at much
lower 0/M) concentrations (Anderson and Meister, 1980; Moldeus and
Quanguan, 1987). In rodents about 90% of the total GSH within the plasma
is in the reduced form (Anderson et al., 1980).
In its predominant state, GSH acts as a critical reducing agent in a
variety of biological functions including protein and DNA synthesis,
maintenance of protein conformation and enzyme activity, leukotriene and
prostaglandin synthesis, amino acid transport, and protection of cells from
free radical damage and carcinogenesis. GSH is also involved in
maintenance of membrane integrity and cytoskeletal organization (Larsson
et al., 1983; Meister and Anderson, 1983; Meister, 1988b; Fettman, 1991).
One of the best known functions of GSH resides in its ability to
protect cells from free radical and organic peroxide-mediated protein and
D NA damage or lipid peroxidation. Superoxide radicals and hydrogen
peroxide formed in biological systems produce reactive species that lead to
organic peroxide formation (Chance et al., 1979). Some reduction of
hydroxy radicals and hydroperoxides occurs nonenzymatically (Krinsky,
1992; de Groot, 1994); however, glutathione peroxidase (EC 1.11.1.9), first
identified in erythrocytes (Mills, 1957 and 1959), is a GSH-dependent
cytosolic enzyme involved in the reduction of hydrogen peroxide and
organic peroxides (including ethyl hydroperoxide, cumene hydroperoxide,
and linoleic acid hydroperoxide) to water and their corresponding alcohols,
respectively (Sies et al., 1980).
Both selenium -dependent (Se-GSH-Px) and selenium -independent
(Se-I-GSH-Px) glutathione peroxidases have been described (Flohe' et al..,
43
1973; Lawerence and Burke, 1976; Burke et al., 1978). Se-GSH-Px is thought
to require phospholipase A 2 (PLase A 2 ) which converts potentially harmful
phospholipid hydroperoxides (PLOOH) to free fatty acid hydroperoxides
(LOOH). Se-GSH-Px subsequently converts LOOHs to harmless fatty acid
alcohols (van Kuijk et al., 1987). Conversely, Se-I-GSH-Px acts directly on
phospholipid hydroperoxides, without the necessity of hydrolyzing the free
fatty acid hydroperoxide from the phospholipid. Accordingly, it is referred
to as phopholipid hydroperoxide glutathione peroxidase (Ursini et al., 1982).
Se-I-GSH-Px can also reduce membrane-associated cholesterol
hydroperoxides (Thomas, 1990).
GSH also provides reducing capacity for cellular macromolecules and
intracellular thiol-containing proteins. Proteins involved in membrane
stability, DN A and protein synthesis, and other enzymatic reactions are
maintained in the reduced state by thiol transfer reactions catalyzed by thiol
transferases (Racker, 1955; Chang and Wilken, 1966; Nagai and Black, 1968).
Glutaredoxin, a thiol transferase homologous to thioredoxin, transfers
electrons from NADPH to ribonucleotide reductase via a glutathione
electron donor. Ribonucleotide reductase catalyzes the reduction of
ribonucleoside diphosphate to deoxyribonucleoside diphosphate and is
therefore essential for DNA synthesis (Stryer, 1988).
Glutaredoxin also provides protection against oxidative damage
through the reduction of dehydroascorbate to the powerful antioxidant,
ascorbic acid (Meister, 1992). GSPI may also keep other antioxidants such as
a-tocopherol (Niki et al., 1982; Reddy et al., 1982; Leedle and Aust, 1990) and
p-carotene (Jialal and Grundy, 1991) in the reduced form, either by a direct
reaction or by thiol transferases.
44
GSH also reduces coenzyme A, the universal donor of acyl groups.
Coenzyme A possesses a terminal sulfhydryl moiety essential to the citric
acid and glyoxalate cycles (Stryer, 1988). Once the acyl group is donated, the
terminal sulfur must be reduced by a GSH-dependent thiol transferase in
order to bind another acyl group (Chang and Wilken, 1966). Similarly,
cystine and homocystine are reduced to cysteine and homocysteine,
respectively, by thiol transferases (Racker, 1955; Meister, 1988b).
GSH-insulin transhydrogenase (GIT; EC 5.3.4.1) is yet another protein
dependent on GSH for its reducing power. GIT catalyzes the first step in the
degradation of insulin by cleaving three disulfide bonds (Tomizawa and
Halsey, 1959; Varandani, 1972).
GSH is additionally involved in the metabolism of prostaglandins
and leukotrienes (Huber and Keppler, 1988; Ketterer et a l, 1988).
Arachadonic acid, found within cellular membranes, is converted to 5hydroperoxy-eicosatetraenoic acid (5-HPETE) by 5-lipoxygenase which, in
turn, is converted to leukotriene A 4 (LTA4 ). LTA4 is metabolized by
glutathione S-transferase to its GSH conjugate, LTC4 , which is cleaved at its
y-glutamyl linkage to yield LTD4 . The cysteinyl-glycine linkage of LTD4 is
then cleaved to produce LTE4 . Leukotrienes are important in vascular
permeability and chemotaxis (Cotran et al, 1989).
GSH contributes to the regulation of prostaglandin and thromboxane
synthesis and metabolism. The effect of GSH on prostaglandin and
thromboxane levels and patterns of synthesis are dependent on the tissue
or organ as well as the level of GSH utilized in the experiment (Mimata et
a l , 1988; Hidaka et al, 1990; Buckley et al, 1991; Nejad et a l, 1991; Mimata et
a l , 1993).
45
Decreases in GSH by administration of diethyl maleate tend to
suppress prostaglandin synthesis as a whole but may dramatically alter the
prostaglandin pattern or profile within the tissue (Mimata et a l, 1988;
Hidaka et al., 1990; Nejad et al., 1991). Increases in GSH appear to stimulate
the synthesis of PGE2 relative to other prostaglandins in the urinary bladder
epithelium (Mimata et ah, 1988) and renal medullary homogenates (Nejad
et a l, 1991).
The effect of GSH on prostaglandin biosynthesis may be mediated
through GSH conjugation. Glutathione S-transferases have been shown to
reduce PGH2 to PGF20 U presumably due to GSH peroxidase activity (Burgess
et a l, 1987) and may be involved in isomerization of PGH 2 to PGE2 and
PGD2 (Meyer and Ketterer, 1987).
GSH has been shown to function as a coenzyme in various biological
reactions. First discovered was the glyoxylase reaction responsible for the
conversion of methylglyoxal to lactate. GSH is essential for the formation
of a hemimercaptal from methylglyoxal (Meister, 1988b). Glyoxylase then
converts the hemimercaptal to lactate. Methylglyoxal theoretically retards
cell growth (Meister, 1988a) so that increases in GSH may actually stimulate
cell growth.
Formaldehyde dehydrogenase also utilizes GSH as a coenzyme in the
conversion of formaldehyde to formic acid (Meister, 1988b). The actual
substrate is believed to be the hemimercaptal formed from the interaction
of formaldehyde with GSH.
The reducing power of GSH is not only utilized intracellularly but
has a variety of similar functions extracellularly. Extracellular GSH has
been shown to protect retinal pigment epithelial (RPE) cells, unable to take
46
up intact GSH, from oxidative damage. Isolated RPE cells were exposed to
diethyl maleate, buthionine sulfoximine, and AT-125 to deplete
intracellular GSH, prevent de novo synthesis of GSH, and impede
degradation and uptake of plasma GSH, respectively. The presence of small
concentrations of extracellular GSH protected these cells from subsequent
exposure to t-butyl hydroxyquinolone (Sternberg et ah, 1993).
Extracellular GSH may also maintain antioxidants, such as ascorbate
and vitamin E, and membrane-bound enzymes in the reduced state
(Meister, 1992; Sternberg et al., 1993). For example, ATPase possesses a
sulfhydryl group on both the cytoplasmic and external aspect which must be
kept in the reduced form to be functional (Qian and Murphy, 1993).
GSH has also been advocated as a storage form of cysteine since the
intracellular concentration of GSH far exceeds that of cysteine with a ratio of
>100:1 (States and Segal, 1990). GSH provides a "bound" form of cysteine
residues since high concentrations of intracellular cysteine are considered
toxic (Bannai and Tateishi, 1986). GSH is an appropriate alternative since
the sulfhydryl group of GSH is more stable than that of cysteine (Bannai
and Tateishi, 1986). Additionally, the tripeptide contains a large number of
hydrophilic functional groups which increase its water solubility (Kosower,
1976). Cystine, the disulfide form of cysteine, has a low water solubility and
may be biologically unsuitable for a thiol-disulfide system in which the two
oxidation states are interconverted.
GSH also functions in the transport of amino acids by moving readily
across cell membranes possessing particular enzyme systems (Meister,
1988b). Translocation of extracellular amino acids into the cell appears to be
linked to efflux of GSH (Griffith et al., 1979).
47
GSH is intimately associated with phase II conjugation reactions
which render nonpolar xenobiotics to more hydrophilic compounds for
elimination in the urine and the bile. GSH conjugates and detoxifies a
variety of classes of electrophilic, hydrophobic compounds. The ability to
detoxify a diversity of xenobiotics is probably the most obvious
physiological benefit of GSH.
N-methyl-4-am inoazobenzene (MAB) and N-acetyl-2-aminofluorene
(AAF) are potent hepatic carcinogens detoxified by GSH conjugation
(Ketterer et al., 1983). N-Acetylbenzoquinone imine (NAPQI) is a reactive,
soft electrophile produced by cytochrome P450 oxidation of paracetamol
(Hinson, 1983). NAPQI may react with soft nucleophilic sites in cellular
macromolecules or may be metabolized to a toxic phenoxy radical (Fisher
and Mason, 1984). GSH can compete with these reactions either by
conjugating NAPQI or by reducing it back to paracetamol (Hinson, 1983).
Biotransformation of benzo(a)pyrene (BP) by cytochrome P450s forms
reactive metabolites which may be detoxified by conjugation with GSH.
The toxic effects of BP appear to be mediated by BP quinones and epoxides
(Huberman et al., 1976; Wislocki et al., 1976). GSH conjugation of
electrophilic BP epoxides and quinones detoxifies these compounds and
enhances their excretion in the bile (Plummer et al., 1980; Chipman et al.,
1981; Elmhirst et al., 1985).
Similarly, the addition of GSH to S9 liver
fractions from the rat results in the reduction of BP and BP 7,8-diol-induced
cytotoxicity while the presence of GSH in addition to glutathione Stransferase eliminates BP-induced cytotoxicity and reduces the mutagenicity
of BP (Recio and Hsie, 1987).
48
Similar to BP, the powerful hepatotoxin aflatoxin Bi exerts its
carcinogenic effect through microsomal activation to a reactive epoxide
(Sw enson et al., 1977). This epoxide reacts with DNA primarily at the N-7
of guanine bases in DNA (Lin et al., 1977). Detoxification of the epoxide by
GSH conjugation appears to depend on the presence of cytosolic GSH Stransferases (Degen and Neumann, 1978; Emerole et al., 1979).
Glutathione Metabolism and Transport
GSH possesses two structurally characteristic features: 1) the yglutamyl linkage between the y-carboxyl group of glutamate and the amino
group of cysteine that protects the cysteinyl-glycine linkage from attack by
intracellular peptidases and 2 ) a reactive thiol that provides the reducing
power of the tripeptide. These components not only promote the
intracellular stability of glutathione but are intimately associated with its
functions (Meister, 1988a).
The y-glutamyl cycle (Figure 1.3), present in various types of cells
including renal and intestinal epithelial cells, hepatocytes, and erythrocytes
(Meister, 1994), is responsible for the degradation and resynthesis of cellular
GSH. GSH utilized by this cycle is derived from the plasma or is directly
transported out of the cell (Meister, 1988b). y-Glutamyltranspeptidase (GGT;
EC 2.3.2.2), present on the external cell membrane, catalyzes the initial step
of GSH degradation and cleaves the y-glutamyl linkage releasing
cysteinylglycine and a y-glutamyl moiety. Since GGT is the only known
enzym e able to cleave the y-glutamyl linkage (Meister, et al., 1981), GSH is
protected from intracellular degradation. The cleaved y-glutamyl moiety
binds to an extracellular amino acid and the resultant y-glutamyl amino
acid is transported into the cell. Though other neutral amino acids such as
49
GSH CONJUGATES
(GSSG)
NADPH
(GSH REDUCTASE)
NADP+
<fH2SH
RCHCOOH
fO-NHCHCO-NHCH2COOH
(CH2)2
CHNH2 (GSH)
I
NH2
amino acids
ADP+P;
I
(GSH
SYNTHETASE)
COOH
(y -GLUTAMYL
TRANS PEPTIDASE)
ATP
CYSTEINYL-GLYCINE
c h 2s h
i 2
f H 2SH
H2NCHCO-NHCH2COOH
CO-NHCHCOOH
(pO-NHCHCOOH
( f H 2)2
(DffEFTIDASE)
CHNH2
GLYCINE
H 2NCH2COOH
COOH
y - GLUTAMYL AA
(fHz)2
chnh2
I
2
COOH
y - GLUTAMYL
CYSTEINE
A
(Y-GLUTAMYL
CYCLOTRANSFERASE
A D P+P:
CH2SH
I 2
H2NCHCOOH
CYSTEINE
RCHCOOH
<pOOH
NH2
(Y-GLUTAMYL
CYSTEINE
SYNTHETASE)
(CH2)2
AMINO ACID
<pHNH2
N
COOH
ATP
COOH
GLUTAM ATE
5-OXOPROLINE
(5-OXOPROLINASE)
ADP+P
Figure 1.3
Illustration of the y-glutamyl cycle depicting the degradation
and resynthesis of glutathione.
50
methionine and glutamine may bind to the y-glutamyl moeity, cystine and
cysteine are the most active amino acid acceptors for y-glutamyl
transpeptidase (Meister, 1988b). Once within the cell, y-glutamyl
cyclotransferase converts the y-glutamyl amino acid to 5-oxoproline and
releases the amino acid. 5-oxoproline is then decyclized by 5-oxoprolinase to
glutamate in an ATP-dependent step.
Coincidental to the transport of the y-glutamyl amino acid, cysteinylglycine may be transported into the cell where it is further degraded by
cysteinylglycine dipeptidase (EC 3.4.13.6) or aminopeptidase M (EC 3.4.11.2)
to cysteine and glycine. Alternatively, membrane-bound dipeptidase on the
exterior of the cell may also cleave cysteinyl-glycine (Ketterer and Mulder,
1990).
GSH resynthesis is then accomplished in two ATP-dependent steps
from its glutamate, cysteine, and glycine constituents. The initial step is
catalyzed by y-glutamylcysteine synthetase (GCS, EC 6.3.2.2). This forms the
y-glutamyl linkage between the y-carboxyl group of glutamate and the
amino group of the cysteine residue. y-Glutamylcysteine synthetase is
feedback inhibited by glutathione (Richman and Meister, 1975) and is the
rate-limiting enzyme in GSH synthesis (Meister, 1974). This suggests a
physiologically significant feedback control on GSH synthesis. The synthesis
of GSH is then completed by addition of a glycine moiety in an ATPdependent reaction catalyzed by glutathione synthetase (EC 6.3.2.3).
Those organs expressing high enzyme activity for the y-glutamyl
cycle are actively involved in GSH metabolism and transport as either "net
exporters" or "net importers" of GSH. The liver actively synthesizes large
quantities of GSH and is a" net exporter" of the tripeptide. Hepatic
51
turnover (t 1 / 2 = 2 hours) is determined by the rate of GSH efflux (Bartoli et
al., 1978; Lauterberg et al., 1984). Efflux of GSH from cells was first observed
in perfused liver preparations (Bartoli and Sies, 1978) and has subsequently
been shown to be translocated from hepatocytes into the plasma and the
bile. The rate of hepatic efflux of GSH into the plasma is approximately
four times greater than that translocated into the bile (Lauterburg et al.,
1984).
Though GSH efflux has been observed in a variety of cells including
lymphoid cells (Griffith et al, 1979b), fibroblasts (Bannai and Tsukeda, 1979)
and macrophages (Rouzer et al., 1982), efflux of GSH from the liver
accounts for greater than 90% of the total GSH within the circulation
(Bannai and Tateishi, 1986). Studies on the plasma levels of GSH in
various blood vessels in the rat provide evidence supporting GSH efflux
across the sinusoidal membrane (Anderson et al., 1980). Relative to the
hepatic artery or inferior vena cava, the hepatic vein had a much higher
level of plasma GSH. This reflects translocation of hepatic GSH across the
sinusoidal membrane into the plasma. Under physiological conditions,
GSH sinusoidal transport was observed as zero order with a rate near
maximum (80% of Vmax). In the rat, sinusoidal export of GSH has been
show n to be a saturable, carrier-mediated process that can be inhibited by
organic anions such as bilirubin, BSP, and the sulfobromophthalein-GSH
conjugate and can be stimulated by adenosine 3',5'-cyclic monophosphatedependent hormones (Lu et al., 1990). Using hepatic sinusoidal membrane
vesicles, investigators have described a multispecific, electrogenic transport
system for GSH with low affinity kinetics (Ookhtens et al., 1985; Aw et al.,
1986; Ookhtens et a l, 1988; Garcia-Ruiz et al., 1992) although Inoue et al.
52
(1984) characterized two kinetically distinct, N a+-independent transport
systems, one with a high affinity and the other with a low affinity for GSH.
GSSG and GSH S-conjugates inhibited the latter transport system suggesting
that GSH, GSSG, and some GSH conjugates are transported by the same
system.
Though it has been w idely accepted for some time that translocation
of intact GSH across the hepatic sinusoidal membrane is a undirectional
process and plasma GSH cannot be taken up by hepatocytes, recent evidence
supports the theory that bidirectional transport of intact GSH actually occurs
in rat and human hepatocytes. Bidirectional GSH transport across the
sinusoidal membrane has been demonstrated in freshly isolated rat
hepatocytes, membrane vesicles, and in Xenopus laevis oocytes expressing
rat liver mRNA encoding the sinusoidal GSH transporter (Garcia-Ruiz et
al., 1992; Fernandez-Checa et al., 1993) as well as in human hepatoblastomaderived Hep G2 cells (Sze et al., 1993). Since both the efflux and uptake
systems exhibit low affinity for GSH and the intracellular GSH
concentration is several orders of magnitude greater than the extracellular
GSH concentration, a net efflux of GSH occurs under normal physiological
conditions. Additionally, low extracellular concentrations of GSH and
GSSG have been shown to fra/zs-stimulate GSH efflux (Garcia-Ruiz et al.,
1992) while GSH uptake in Hep G2 cells, as well as rat hepatocytes, exhibit
no N a+-dependence suggesting there is no driving force for net uptake
against the intracellular concentration (Garcia-Ruiz et al., 1992; Sze et al.,
1993).
The activity of the sinusoidal GSH transporter in rats has also been
shown to be modulated by the thiol-disulfide status of the cell and its
53
environment (Lu et al., 1993). In cultured rat hepatocytes and perfused
livers, GSH efflux was stimulated by the presence of dithiols and inhibited
by disulfides. Similarly, GSH uptake was inhibited by the presence of
dithiols. Although it is not certain whether thiols exert their stimulatory
effect on the outside of the cell, within the cell membrane, or inside the cell,
it seems clear that thiol reduction converts a bidirectional GSH transporter
to a preferentially unidirectional exporter of GSH by inhibiting uptake and
stimulating efflux. These data are also consistent with the observation that
the GSH transporter operates as a net efflux pump under normal
physiological conditions since the majority of plasma and intracellular GSH
is in the reduced state (Anderson et al., 1980).
Approximately 15-20% of hepatic GSH efflux is directed into the
biliary system (Kretzschmar and Klinger, 1990). Though Inoue et al. (1983,
1984) characterized a high affinity, N a+-independent, saturable, carriermediated system using canalicular membrane vesicles, no saturation of
GSH efflux into bile has been observed in vivo or in rat perfused liver
(Kaplowitz et al, 1983; Lauterbergef al, 1984; Ookhtens and Maddatu, 1991).
Studies utilizing mutant Eisai hyperbilirubinemic rats have demonstrated a
distinct low affinity electrogenic carrier (Fernandez-Checa et al., 1992) for
canalicular GSH transport; however, recent evidence suggests the presence
of high- and low-affinity transport systems for GSH across the canalicular
membrane (Ballatori et al., 1994). The high affinity system is c/s-inhibited
by GSH conjugates, other y-glutamyl compounds, and BSP and is thought to
be a multispecific organic anion transporter.
Though intact GSH is unidirectionally secreted from hepatocytes into
bile, moderate amounts of GGT and dipeptidases are present along the
54
canalicular membrane, bile duct epithelium and, to a lesser extent, in bile
secretions (Hirata et al., 1984). This permits the degradation of biliary GSH
and subsequent reabsorbtion of amino acid precursors. The amino acids
may then be utilized for GSH resynthesis or in the synthesis of other
peptides or proteins.
GSSG and GSH S-conjugates are also translocated across hepatic
sinusoidal and canalicular membranes. At least two organic anion
transport systems have been demonstrated for GSH conjugate transport
across the sinusoidal membrane based on kinetics, substrate specificity, and
chloride dependence (Stremmel and Diede, 1990; Min et al., 1991). A
chloride-independent GSH conjugate efflux system appears to be shared
with GSH (Sze et al., 1993). The fact that the efflux of GSH across the
sinusoidal membrane is inhibited by GSH S-conjugates (Lu et al., 1990) also
suggests a shared transport system.
Although small but significant quantities of GSH S-conjugates are
released from the liver into the plasma (Sies et al., 1978; Wahllander and
Sies, 1979), GSH S-conjugates are preferentially released across the
canalicular membrane into the bile (Sies et al., 1978; Sies et al., 1980;
Akerboom etal., 1982a; Akerboom etal., 1982b). Sinusoidal transport is
electrogenic (Kobayashi et al., 1990) and may augment hepatic excretion of
GSH conjugates when their concentrations exceed the capacity of the
transport processes on the canalicular membrane. Many GSH conjugates
exhibit physiochemical properties favoring their excretion into the bile.
Compounds with a molecular weight 325 ± 50 or greater and possessing a
strong polar group are preferentially excreted in the bile (Hirom et al., 1972;
Sipes and Gandolfi, 1986).
55
A high affinity ATP-dependent transport system across the
canalicular membrane has been demonstrated for BSP-SG (Kitamura et al.,
1990), S-(2,4-dinitrophenyl) glutathione (DNP-SG; Kobayashi et al., 1990;
Akerboom et al., 1991), and cysteinyl leukotrienes (Ishikawa et al., 1990).
This system actually mediates the efflux of xenobiotics conjugated with
GSH, glucuronate, and possibly sulfate (LeBlanc, 1994) and is important not
only in the elimination of xenobiotics and GSSG, but also in the regulation
of biologically active GSH conjugates such as leukotriene C4 . This
transporter is also present in erythrocytes and cardiac muscle and is
believed to be mediated by a 90 kDa protein with ATPase activity (Pikula et
a l , 1994).
ATP-dependent uptake of DNP-SG was czs-inhibited in a competitive
manner by other GSH conjugates and GSSG but not by GSH (Kobayashi et
a l , 1990; Akerboom et al, 1991) and the inhibitory effect of GSH conjugates
increased with increasing chain length (Kobayashi et al , 1990). In addition,
secretion of GSH was fjans-stimulated by GSH conjugates such that GSH
efflux may drive the uptake of organic anions into the hepatocyte with
significant toxicological consequences.
In contrast, two ATP-independent GSH conjugate transport
mechanisms have recently been identified (Ballatori et al , 1994) at the
canalicular membrane. While one ATP-independent transport mechanism
appears to be fairly specific for GSH conjugates, the other ATP-independent
carrier is analogous to the high affinity system that transports GSH as well
as other y-glutamyl compounds. This latter system apparently transports
GSH conjugates of low molecular weight since ethyl-GSH was a substrate
for this carrier and not the ATP-dependent mechanism. Conversely,
56
DNP-GSH is not only a substrate for the high affinity, ATP-dependent
transporter, but acts as a substrate for at least two other ATP-independent
mechanisms as w ell (Ballatori et al., 1994). It appears that the
physiochemical properties of the S-moiety may determine whether the
compound is a substrate for a particular transport mechanism. Data
throughout the literature suggest that multiple transport mechanisms exist
for GSH and GSH conjugates in both the sinusoidal and canalicular
membranes.
GSSG also competes with GSH conjugates for transport across the
canalicular and sinusoidal membranes. As previously mentioned, Inoue et
al. (1984) demonstrated a low affinity transporter in the hepatic sinusoidal
membrane involved in the transport of GSH, GSSG, and some GSH
conjugates.
Lindwall and Boyer (1987) reportedly demonstrated a specific
transport system for GSH S-conjugates which was not inhibited by GSSG;
however, ATP-dependent carrier systems have been demonstrated for the
transport of both GSSG (Akerboom etal., 1991) and GSH S-conjugates
(Kobayashi et al., 1990) across the canalicular membrane. It has been
demonstrated that the ATP-dependent GSH conjugate export pump
transports GSSG as well as various conjugates of GSH out of the cell
(Ishikawa, 1992). Additionally, a common biliary secretory mechanism is
shared by GSSG and bilirubin in rats (Kuromma et al., 1993) but whether
this is a multispecific organic anion transporter shared by GSH conjugates is
not yet clear.
A second, distinct transport mechanism has been demonstrated at
the sinusoidal membrane which is thought to secrete excess intracellular
57
GSSG when canalicular transport becomes saturated (Akerboom etal.,
1982b; Ballatori and Troung, 1989; Jaeschke, 1990). Sinusoidal GSSG efflux is
driven by a K+ concentration gradient which requires ATP only indirectly to
maintain the intracellular K+ concentration via a N a+-K+ pump (Ozaki et
a l , 1994).
The biliary concentration of GSSG is approximately 20-fold higher
than the intrahepatic concentration (Akerboom etal., 1982a) and
intrahepatic concentrations of GSSG are typically very low under normal
physiological conditions. During periods of oxidative stress GSSG efflux
increases dramatically and measurement of plasma GSSG is considered a
reliable index of cellular oxidative stress (Kretzschmar and Klinger, 1990).
Increased efflux of GSSG during oxidative stress has been reported in the
ocular lens (Srivastra and Beutler, 1968), erythrocytes (Srivastra and Beutler,
1969), perfused heart (Ishikawa and Sies, 1984; Ishikawa et al., 1986) and
liver (Akerboom etal., 1982b; Ballatori and Troung, 1989; Jaeschke, 1990;
Younes and Strubelt; 1990), and isolated hepatocytes (Oude Elferink et al.,
1990).
Release of GSSG is thought to be an "emergency" response to high
intracellular levels of GSSG and loss of reducing capacity. Sinusoidal GSSG
release during oxidative stress, however, results in a net loss of cellular
GSH which may play a significant role in the subsequent development of
toxicity.
The accumulation of intracellular GSH protects cells from oxidative
damage (Hagen et al., 1988). The kidney is the primary organ for clearance
of plasma GSH (McIntyre and Curthoys, 1980; Orrenius et al., 1983) and is
capable of extracting 80-90% of circulating GSH in a single pass (Haberle et
58
al., 1979). This results in extremely low levels of plasma GSH (/iM) and a
11 / 2
of only 2 minutes (Wendel and Cikryt, 1980).
Cells exhibiting high levels of GGT activity cleave the y-glutamyl
linkage and accumulate GSH intracellularly. Renal proximal tubular
epithelial cells display the highest levels of GGT activity (Goldberg et al.,
1960); however, an appreciable amount is also evident in pancreatic acinar
and ductile epithelium as well as epithelial cells of the jejunum, biliary
canaliculi, epididymus, seminal vesicles, bronchioles, thyroid follicles, and
the retinal pigment epithelium, choroid plexus and ciliary body (Meister et
al., 1976).
Although it was previously believed that renal GGT was only present
on the luminal aspect of the tubular epithelial cell, electron micrograph
studies and immunochemical staining (Spater et al., 1982) identified the
transpeptidase on the basolateral membrane as well. Additionally,
Anderson and coworkers (1980) observed that the GSH levels in the renal
vein contained approximately
20%
of the levels found in the renal arterial
plasma. This suggests that the kidney removes GSH from the plasma not
only by glomerular filtration, which would account for only 20-30% of the
GSH removed, but also by a mechanism at the basolateral membrane.
Furthermore, AT-125, an inhibitor of GGT, decreased the amount of plasma
GSH cleared by the kidney to levels approximating that removed through
glomerular filtration. This supports a role for GGT in the renal clearance
and uptake of GSH. GSH in the plasma or filtered by the glomerulus is
degraded to its constituent amino acids by membrane-bound GGT and
aminopeptidase M or cysteinylglycine dipeptidase (Ketterer and Mulder,
1990).
59
GGT is not only capable of catalyzing transpeptidation reactions, but
may additionally catalyze autotranspeptidation of y-glutamyl peptides or
amino acids and hydrolytic reactions resulting in the formation of
glutamate (Laperche et al., 1990).
Despite the fact that the kidney exhibits high levels of GGT and is a
"net importer" of GSH, 80-90% of renal GSH is actually derived from de
novo synthesis within the tubular epithelial cell (Meister and Anderson,
1983). Renal GSH is rapidly synthesized from its amino acid constituents by
GCS and GSH synthetase and the rate of turnover of cytosolic GSH within
the kidney is approximately 20 minutes (Sekura and Meister, 1974).
Large amounts of GSH are actively secreted from renal proximal
tubular epithelial cells into the tubular lumen (Hsu et al., 1984). Under
normal physiological conditions the normal level of GSH within the
filtrate is approximately 0.6 //M in the rat. Following administration of AT125, the urinary GSH concentration increased to 3mM indicating that > 99%
of the tubular GSH is normally reabsorbed (Curthoys, 1990).
Plasma GSH is not only accumulated in the renal proximal tubular
epithelial cell via degradation and resynthesis, but may be transported intact
across the basolateral membrane (Lash and Jones, 1983). Intact GSH is taken
up by a Na+-dependent, probenecid-sensitive electrogenic transporter
thought to be a general transport mechanism for y-glutamyl com pounds
including GSSG and GSH conjugates (Lash and Jones, 1984; Rankin et al.,
1985; Hagen et al., 1988). GSH transport exhibits a Na+:GSH stoichiometry
of 2:1 (Lash and Jones, 1984b) and exhibits a rate of GSH accumulation
approximately 3-fold greater than the GSH synthetic rate from amino acid
precursors (Hagen et al., 1988). The system appears to have a sufficient
60
driving force to allow uptake of GSH into cells against a GSH concentration
gradient.
The significance of renal influx of intact GSH is highly controversial.
Though Abbott and coworkers (1984) demonstrated barely detectable
transport of intact GSH in the presence of extensive degradation of the
tripeptide at the basolateral membrane, studies using isolated proximal
tubular epithelial cells revealed protection by the uptake of intact GSH
against the potent antioxidant, t-butylhydroperoxide (Hagen et al., 1988).
Several other epithelial cells have also demonstrated uptake of intact GSH
by either N a+-dependent (alveolar type II epithelial cells and enterocytes) or
N a +-independent (buccal epithelial cells, colonocytes, and enterocytes)
transport systems (Hagen et ah, 1986; Lash et al., 1986b; Vincenzini et al.,
1992). Conversely, erythrocytes, macrophages, retinal pigment epithelial
cells, and alveolar type I cells did not exhibit uptake of intact GSH (Foreman
and Skelton, 1990; Sternberg et al., 1993).
Intact GSH is transported across the basolateral membrane from the
vasculature into enterocytes by a common transport system shared with
GSH conjugates (Lash et al., 1986b; Vincenzini et al., 1992). The transport of
GSH into intestinal epithelial cells is thought to protect these cells from
oxidative or xenobiotic injury and may act as the first line of defense against
dietary electrophiles. Transport of GSH conjugates from the vasculature
into enterocytes may also serve as a potential route of excretion.
Translocation of intact GSH and GSH conjugates across intestinal brush
border membranes into the luminal space has been demonstrated in rats
using small intestine brush-border membrane vesicles (BBMV; Linder et
al., 1984; Vincenzini et al., 1992) and vascularly perfused jejunum (Hagen
61
and Jones, 1987). Although Linder and coworkers (1984) described transport
across the brush-border membrane as Na+-dependent and carrier-mediated,
Vincenzini et al. (1992) characterized a Na+-independent carrier system.
Basolateral transport of GSH is competitively inhibited by probenecid while
the brush-border system is not, suggesting two distinct transport
mechanisms (Hagen et al., 1987).
Intestinal transport of GSH across the brush-border membrane
exhibits cis-inhibition and fraus-stimulation in the presence of GSH
conjugates such that intracellular GSH conjugates inhibit GSH efflux and
stimulate GSH uptake while extracellular GSH conjugates inhibit GSH
uptake (Vincenzini et al., 1992). Alternatively, as GSH conjugates cross the
epithelial cell from the vasculature, GSH is transported from the lumen
into the cells and finally to the plasma where it is available to other organs.
Trans-stimulation of intestinal transport of GSH across BBMV was
also observed with amino acids (Corcelli et al., 1982; Lash and Jones, 1984a).
Trans-stimulation by amino acids would allow increased efflux of GSH into
the intestinal lumen following a meal. Once within the lumen, GGT may
release the y-glutamyl moiety which accepts an amino acid prior to
transport into the cell. By this mechanism amino acid transport could be
enhanced in the presence of high concentrations of dietary amino acids. It
also appears that the formation of GSH conjugates within intestinal
epithelial cells stimulates the uptake of GSH into cells where it is needed
due to possible depletion from GSH conjugation. GSH may be utilized
intracellularly or transported to the luminal surface.
Within the intestinal lumen, extracellular GSH, from the bile or
intestinal secretions, may conjugate electrophiles resulting from microbial
62
metabolism or the diet. In vivo studies by Hagen and coworkers (1990)
revealed that GSH was present within the intestinal lumen under all
conditions even without supplementation of GSH in the diet. The
combined activities of y-glutamyl transpeptidase and the GSH uptake
system were not sufficient to remove all the luminal GSH. In fact, a
mechanism for reduction of GSSG to GSH was observed in the intestinal
lumen and it was theorized that GSH within the lumen may detoxify
xenobiotics in the food or bile to protect the intestinal epithelium from
injury.
Numerous xenobiotics, or their corresponding metabolites, are
detoxified by GSH conjugation. GSH conjugation of vinylidene chloride,
bromobenzene, aflatoxin By styrene, 1-nitropyrene, paracetemol, and
monocrotaline (Jollow et al., 1974; D'Souza and Andersen, 1988; Wilson et
al., 1992; Kataoka et al., 1991; James et al., 1993; Morgan et al., 1993; GopalanKriczky et al., 1994) prevents the toxicity and carcinogenic effects of these
xenobiotic metabolites.
The interorgan translocation of GSH conjugates is complex. The
liver, receiving the portal circulation from the gastrointestinal tract, is
responsible for the formation of the majority of GSH conjugates. GSH
conjugates may be translocated from the hepatocyte into the bile or
circulation (Dekant et al., 1988). Once present in the circulation, GSH
conjugates may be taken up intact by the kidney (Lash and Jones, 1984b) or
the intestinal epithelial cell (Lash et al., 1986b) or may be degraded at the
basolateral membrane of the kidney and transported into the renal
epithelial cell as a y-glutamyl amino acid and the cysteine conjugate
(Rankin et al., 1983; Abbott, et al., 1984). Intact intracellular GSH conjugates
63
may be secreted into the urine from the renal epithelial cell (Griffith and
Meister, 1979; Griffith, 1981; Scott and Curthoys, 1986).
Plasma GSH conjugates may also be filtered by the glomerulus
(Dekant et al., 1988). Following degradation of the GSH conjugate by yglutamyltranspeptidase and cysteinyl-glycine dipeptidase or aminopeptidase
M, the cysteine conjugate within the filtrate may be transported into the
renal epithelial cell from the urinary filtrate or it may be excreted in the
urine. In the renal epithelial cell, the cysteine conjugate may be excreted
back into the urine (Dekant et al., 1988) or N-acetylated to the mercapturic
acid via cysteine conjugate N-acetyltransferase (Duffel and Jakoby, 1982).
Alternatively, the cysteine conjugate can be converted to a reactive thiol in
addition to ammonia and pyruvate by renal cysteine conjugate p-lyase
(Elfarra and Anders, 1984). The potentially reactive thiol is then capable of
being converted to the methyl thiol derivative by thiol S-methyl
transferase; the methyl thiol derivative may be excreted in the urine. The
cysteine conjugate may also be released into the plasma and transported to
the liver where it undergoes N-acetylation to the mercapturic acid (Dekant
et al., 1988). It should be noted that at any point mecapturic acids may be
deacetylated back to the cysteine conjugate (Suzuki and Tateishi, 1981).
GSH conjugates, mercapturates, and cysteine conjugates in the
plasma may alternatively be taken up by intestinal epithelial cells
(Vincenzini et al., 1991). Cysteine conjugates and mercapturates excreted
into the intestinal lumen may be excreted in the feces or they may be
further metabolized. Cysteine conjugates can be converted to the
corresponding mercapturic acid or reactive thiol via N-acetyltransferase
(Meister, 1988b) and bacterial p-lyase (Stevens and jakoby, 1985),
64
respectively. The thiol can be methylated to the methyl thiol derivative
which is excreted in the feces. At any point in the metabolism of these
compounds a metabolite may undergo enterohepatic recirculation to the
liver.
GSH conjugates in the plasma that undergo transcellular
translocation into the intestinal lumen are subject to the same fate as GSH
conjugates secreted in the bile from the liver. GSH conjugates in the bile
and intestinal secretions are either transported intact into the intestinal
epithelium (Dekant et al., 1988) or are degraded to the cysteine conjugate by
y-glutamyltranspeptidase and dipeptidases present along the biliary
epithelium and canalicular membrane of hepatocytes (Inoue et al., 1983;
Meier et al., 1984; Ballatori et al., 1986). Once formed, the cysteine conjugate
may undergo enterohepatic recirculation or it may be transported to the
kidney (Dekant et al., 1988). Alternatively, the cysteine conjugate may be
further metabolized within the intestinal lumen as previously described.
GSH conjugates may also be directly formed in the intestinal
epithelial cell which may serve as the first line of defense against dietary
mutagens and carcinogens (Vincenzini et al., 1991). GSH conjugates
translocated into the plasma from the intestinal epithelial cell are subject to
uptake and metabolism by renal epithelial cells (Lash and Jones, 1984b).
Although GSH-dependent reactions that terminate in the formation
and excretion of mercapturic acids play a major role in the detoxification of
xenobiotics, it has become increasingly obvious that GSH conjugation also
constitutes an important mechanism of bioactivation for several classes of
compounds (Monks and Lau, 1987; Monks et al., 1990; Vamvakas and
Anders, 1990; Koob and Dekant, 1991; Dekant and Vamvakas, 1993). GSH
65
conjugates may be either direct acting toxic agents which are noxious
without further metabolism or they may require further metabolism to
express toxicity.
Ethylene dibromide is a direct-acting GSH conjugate
w idely used as a lead scavenger for gasoline and as a soil fumigant. This
compound forms the half-sulfur mustard, S-(2-bromethyl)GSH, also
referred to as a thiiranium or episulfonium (Ozawa and Guengerich, 1983;
Koga et al., 1986). The half-sulfur mustard reacts with the N 7 of guanine in
D N A to form the S-[2-(N 7 -guanyl)ethyl]-GSH adduct and is responsible for
the carcinogenic effects of this vicinal dihaloalkane (Inskeep et al., 1986).
Similarly, the genotoxicity of ethylene dichloride is mediated through GSH
conjugation and episulfonium ion formation (Rannug et al., 1978;
Guengerich et al., 1980; Storer and Conolly, 1985; Inskeep et al., 1986).
Although l,2-dibromo-3-chloropropane is metabolized by
cytochrome P450 to a mutagenic metabolite (2-bromoacrolein) and the
presence of GSH and GST decrease the mutagenicity of this compound
(Omichinski et al., 1988), the GSH conjugate, 2-(3-chloro-2-bromopropyl)GSH, is an intermediate that spontaneously cyclizes to an
episulfonium ion. This later metabolite has been shown to be responsible
for the acute toxicity observed following administration of l,2-dibromo~3chloropropane (Pearson et al., 1990).
Unlike some of the halogenated hydrocarbons, the majority of
xenobiotics bioactivated by GSH conjugation require further metabolism to
express toxicity . Many of these compounds appear to target the kidney with
resultant nephrotoxicity. Because of their structure and physiological
function, renal epithelial cells are exposed to more chemicals than most
other organs and tend to concentrate xenobiotics due to the normal
66
functions of filtration, reabsorbtion, and secretion. The kidneys comprise
only 0.4% of the body weight of most mammals, but receive 25% of the
cardiac output (Monks and Lau, 1987). In particular, the renal cortex (where
the majority of the proximal tubules are located) receives greater than 90%
of the renal blood flow (Maher, 1976). Additionally, the specialized
functions of renal proximal tubular epithelial cells may enhance toxicity
through the concentration of xenobiotics. Renal proximal tubular
epithelial cells possess the highest level of GGT activity (Goldberg et al.,
1960) and are active in the uptake and concentration of metabolites of GSH
conjugates.
Nephrotoxicity through episulfonium formation has been
demonstrated for direct-acting compounds including ethylene dichloride
(Elfarra et al., 1985), l,2-dibromo-3-chloropropane (Pearson et al., 1990), and
tris(2,3-dibromopropyl) phosphate (Dekant and Vamvakas, 1993).
Nephrotoxicity following GSH conjugation, however, is more typically
mediated through the formation of cysteine conjugates, mercapturic acids,
or reactive thiols. Due to the expression of extremely high concentrations
of GGT along the basolateral and luminal membranes, the kidney actively
concentrates cysteine conjugates within proximal tubular epithelial cells.
Once within the cell, the cysteine conjugate may be directly nephrotoxic or
may require further metabolism to the corresponding mercapturic acid or
reactive thiol by N-acetyltransferase or cysteine conjugate (3-lyase,
respectively (Nelson, 1992).
The cysteine conjugate of bromohydroquinone, a toxic metabolite of
bromobenzene (Lau et al., 1984), appears to be responsible for
bromobenzene-induced nephrotoxicity. Substitution of
67
bromohydroquinones with cysteine lowers the redox potential and renders
bromohydroquinone more susceptible to oxidation and increases the
formation of toxic quinones (Monks and Lau, 1990; Lau et al., 1988a, b).
Additionally, the reduction of bromohydroquinone-induced nephrotoxicity
by simultaneous administration of ascorbic acid suggests that processes
involving oxidation to a reactive quinone and, possibly, peroxidative
mechanisms are involved in the nephrotoxic effects (Lau et al., 1988b).
The GSH conjugate [S-(l,2-dichlorovinyl)GSH or DCVG] and the
cysteine conjugate [S-(l,2-dichlorovinyl)-L-cysteine or DCVG] of
trichloroethylene as w ell as the cysteine conjugate (S-pentachlorobuta-1,3dienylcysteine) and mercapturic acid [N-acetyl-(S-l,l,2,3,4- pentachlorobutadienyl)-L-cysteine] of hexachlorobutadiene have demonstrated
nephrotoxic effects in vivo and in vitro (Terracini and Parker, 1965; Jaffe et
al., 1983; Elfarra and Anders, 1984; Reichert and Schutz, 1986). The
nephrotoxicity of these compounds, however, has been shown to be
mediated through activation via renal p-lyase (Elfarra and Anders, 1984;
Lash and Anders, 1986; Dekant and Vamvakas, 1993).
Similarly, haloalkene-induced nephrotoxicity involves bioactivation
by renal 0-lyase. The cysteine conjugate of chlorotrifluoroethene, S-(2chloro-l,l,2-trifluoroethyl)-L-cysteine, is metabolized by 0 -lyase to 2-chloro1,1,2-trifluoroethane thiol which forms a thionoacetyl fluoride (Dekant et
al., 1987). This latter metabolite reacts with cellular macromolecules with
resultant nephrotoxicity.
Inhibition of mitochondrial function appears to be the mechanism of
toxicity for many of these cysteine conjugates (Lash and Anders, 1986; Lash
and Anders, 1987; Banki and Anders, 1989). Mitochondria are readily
68
targeted by reactive thiols since these organelles contain significant
quantities of renal p-lyase (Lashef al., 1986a). p-lyase-mediated toxicity
inhibits cellular oxygen consumption and decreases ATP concentrations.
Renal cysteine conjugate p-lyase is a pyridoxal phosphate (PLP)dependent enzyme present in both cytosolic and mitochondrial cellular
fractions (Dohn and Anders, 1982; Stevens, 1985a). Renal cytosolic p-lyase
is identical to glutamine transaminase K (EC 2.6.1.64; Stevens et al., 1986).
Glutamine transaminase K , found in the cytosol and mitochondria,
catalyzes transamination reactions between a-keto acids and amino acids
(Ricci et al., 1986).
Mitochondrial P-lyase has been localized to the outer mitochondrial
membrane (Lash et al., 1986a) and mitochondrial matrix (Stevens et al.,
1988a). Mitochondrial p-lyase found within the matrix is inhibited by
antibodies raised to the renal cytosolic enzyme while the enzyme located in
the outer membrane was not (Stevens et al., 1988a). Thus, there appear to
be at least two mitochondrial p-lyases in the rat kidney.
The catalytic mechanism of renal p-lyase involves, after Schiff base
or aldimine formation with PLP, proton abstraction from the a-carbon, a pelimination reaction, and subsequent hydrolysis to eventually yield
ammonia, pyruvate, and a potentially reactive thiol thought to be
responsible for the toxicity (Davis and Metzler, 1972). Pyruvate and the
reactive thiol may also be formed via a C-S cleavage reaction which is not
catalyzed by renal p-lyase (Tomisawa et al., 1986). Alternatively, p-lyase
has been demonstrated to catalyze a half-transamination reaction rather
than a p-elimination reaction and produce an a-keto acid analog of the
cysteine conjugate. Additionally, L-amino acid oxidase (EC 1.4.3.2),
69
identified in the cytosol and mitochondria, may catalyze the oxidative
deamination of the cysteine conjugate (Stevens et al., 1986).
P-lyase activity is also found in the liver. Hepatic p-lyase is
different from renal p-lyase activity (Tateishi and Shimizu, 1980; Stevens
and Jackoby, 1983; Stevens, 1985b), however, and is actually identical to
kynureninase (Stevens, 1985b).
Renal cysteine conjugate p-lyase activity has been demonstrated to be
responsible for the formation of mutagenic metabolites from cysteine
conjugates of various halogenated alkenes (Green and Odum, 1985; Dekant
et al., 1988; Commandeur et al., 1991). The mutagenicity of these
com pounds was diminished in the presence of aminooxyacetic acid
(AOAA). AOAA (Figure 1.4) is a competitive inhibitor of pyridoxal
phosphate-dependent enzymes (Wallach, 1960; Beeler and Churchich, 1976)
including renal p-lyase (Figure 1.5; Elfarra et al., 1986).
Other inhibitors, such as p-di-/z-propylsulfamoyl benzoic acid
(probenecid) and (aS, 5S)-a-amino-3-chloro-4,5-dihydro-5-isoxazoleacetic
acid (acivicin or AT-125) have been extensively utilized to elucidate the
ultimate toxic metabolite for various classes of compounds. Probenecid
(Figure 1.6) has been used therapeutically to prolong the half-life of
penicillin (Beyer, 1950; Beyer et al., 1951) and as an uricosuric agent in the
treatment of gout (Gutman and Yu, 1957).
Probenecid is secreted into the tubular lumen as the glucuronide
conjugate by the organic anion transport system (OATS; Israeli et al., 1972;
Dayton et al., 1973; Cunningham et al., 1981). Probenecid inhibits
glucuronidation of various compounds (Vree et al., 1993; Vree et al., 1994)
70
h 2n —
— c h 2— c
2
/ / °
\
OH
Figure 1.4
Structure of aminooxyacetic acid (AOAA).
71
GLUTATHIONE
X
(GSH - S - TRANSFERASE)
X
y- g l u
- c y s - gly
(Y - GLUTAMYL TRANSPEPTIDASE)
glutamate-
<r
CYS - GLY
I
X
(DIPEPTIDASE)
glycini
AOAA
CY!
(B - LYASE)
PYRUVATE
NH,
A REACTIVE THIOL
(N - ACETYL - TRANSFERASE)
N - ACETYL - CYS
I
X
(MERCAPTURIC ACID)
Figure 1.5
Biochemical pathway illustrating the metabolism of
glutathione conjugates and the inhibitory effect of aminooxyacetic acid
(AOAA) on renal cysteine conjugate p-lyase.
72
CH3CH2CH;
// \\'> —COOH
' n SO —( '
CH3CH2CH2^
Figure
1 .6
Structure of probenecid (p-di-n -propylsulfamoyl benzoic acid)
73
and competitively inhibits renal elimination of weak organic acids such as
p-aminohippurate and salicylic acid (Figure 1.7; Gutman et al., 1955).
Evidence suggests that there are at least three distinct basolateral
OATS in rat proximal tubular epithelial cells (Ullrich and Rumrich, 1988;
Burckhardt and Ullrich, 1989). Once inside the cell, organic anions may be
transported into the tubular lumen in exchange for another anion. In the
process of tubular secretion, renal proximal tubular cells can concentrate
organic anions to levels several hundred times greater than that of the
interstitial fluid (Goldstein, 1993). Renal secretion via the OATS is confined
to the proximal convoluted tubule and appears most intense in the
52-
segment for most organic anions (Shimomura et ah, 1981).
Conversely, AT-125 is a glutamine analog (Figure 1.8) that
irreversibly inactivates y-glutamyl transpeptidase (GGT; Gardell and
Tate,1980; Reed et al., 1980) as well as various glutamine amidotransferases
(Tso et al., 1980) and glutaminases (Jayaram et al., 1975). AT-125 inhibits
GGT by covalently attaching to the y-glutamyl region of the active site
(Figure 1.9).
AOAA, probenecid, and AT-125 have been utilized individually as
w ell as in tandem to elucidate toxic metabolites responsible for renal
toxicity for various compounds.
Nitriles
Nitriles are organic compounds with a terminal cyano (CN) moiety
w idely distributed in the environment both as natural and industrially
produced chemicals (Egekeze and Oehme, 1980). Synthetic aliphatic and
aromatic nitriles are used extensively in the manufacture of plastics,
solvents, synthetic fibers, resins, pharmaceuticals, vitamins, and
74
LIVER
GS-R
PROBENECID
NAC-CSY-R
BILE
GSH + R
PLASMA
CYS-R
ENTEROHEPATIC CIRCULATION
NAC-CSY-R
NAC-CSY-R
GS-R
NAC-CSY-R
PROBENICID
CYS-R
CYS-R
EXCRETION
EPITHELIUM
INTESTINE
BASOLATERAL
KIDNEY
LUMENAL
Figure 1.7
Diagram depicting the inhibitory effect of probenecid (p-di-npropylsulfamoyl benzoic acid) on the organic anion transport system
(OATS) at the sinusoidal membrane of the hepatocyte and the basolateral
membrane of the renal proximal tubular epithelial cell.
75
COOH
I
H2N - CH
COOH
*
H2N— CH
^2
HC— 0 N
i
Hr
I
Cl
AT-125
4
CH2
o
I
nh2
GLUTAMINE
Figure 1.8
Structure of the glutamine analog, AT-125 [acivicin or (aS, 5S)a-amino-3-chloro-4,5-dihydro-5-isoxazoleacetic acid], in comparison with
the structure of glutamine.
76
RENAL PROXIMAL TUBULE
f
yGLU-CYS-GLY
R-SH MA
AT-125
\
CY^-GLY-
GSH
/
CYS-R
GLY
AA(CYS;
yGLU
YGLU-AA
BASOLATERAL
MEMBRANE
BRUSH-BORDER
MEMBRANE
Figure 1.9
Diagram illustrating the inhibitory effect of AT-125 [acivicin
or (aS, 5S)-a-amino-3-chloro-4,5-dihydro-5-isoxazoleacetic acid] on yglutamyltranspeptidase at the basolateral membrane of the renal proximal
tubular epithelial cell.
77
agricultural insecticides (Whitehill and Smith, 1981). Organonitriles are
produced from the catabolism of plant cyanogenic glucosides or are derived
from glucosinolates in plants. There are approximately 2000 species of
plants that have been shown to be cyanogenic, releasing hydrogen cyanide
when macerated (Conn, 1980; Vennesland et al, 1982).
Exposure of humans and experimental animals to organic nitriles
has been documented to cause neurotoxicity ( Farooqui and Mumtaz, 1991)
in addition to various lesions within the thyroid, liver, adrenal glands, and
gastrointestinal tract (Silver et al., 1982; Sipes and Gandolfi, 1986; Villarreal
et al., 1988). Organic nitriles have also been shown to cause teratogenic
effects in rats (Whitehill et al, 1981a, b).
The acute toxicity of nitriles has been correlated with the in vivo
release of cyanide (Willhite and Smith, 1981; Farooqui and Mumtaz, 1991)
with the extent of liberation dependent on the nitrile, animal species, route
of exposure, and chemical structure of the parent compound (Ahmed and
Farooqui, 1982; Silver et al., 1982; Tanii and Flashimoto, 1984a, b).
Urinary
excretion of high levels of thiocyanate (SCM~) were observed following
administration of saturated aliphatic nitriles consistent with detoxification
of the cyanide ion by rhodanese activity (Sipes and Grandolfi, 1986; Wallig
et al., 1988a). Additionally, many of these compounds cause classical signs
of cyanide toxicosis which were ameliorated by standard therapy for cyanide
intoxication (Pozzani et al., 1968).
Unsaturated aliphatic nitriles such as acrylonitrile, methacrylonitrile,
crotonitrile, and cinnamonitrile are partially metabolized to the cyanide
ion, but have also been shown to undergo GSH conjugation in vitro and in
v i v o (van Bladeren, et al., 1981a; Cote et al., 1984; Day et al., 1988).
78
Administration of unsaturated aliphatic nitriles to experimental animals
resulted in depletion of tissue GSH levels (van Bladeren, et al., 1981a; Cote
etal., 1984; Day et al., 1988) and the formation of mercapturic acids which
were excreted in the urine (Langvardt et al., 1980; Muller et al., 1987;Tardif
et al, 1987).
Nitriles, including l-cyano-3,4-epithiobutane (CEB), l-cyano-2hydroxy-3,4-epithiobutane (CHEB), and l-cyano-2-hydroxy-3-butene (CHB),
derived from cruciferous plants cause only mild increases in urinary
thiocyanate (SCN~) excretion and are extensively conjugated with GSH in
vivo (Wallig et al., 1988 a, b; VanSteenhouse et al., 1989). Conjugation with
GSH bioactivates these organonitriles with resultant toxicity to specific
target organs.
(S)-l-cyano-2-hydroxy-3-butene (CHB) was selectively pancreatotoxic
at low dosages (Wallig et al., 1988b; Maher et al., 1991) but acutely
pancreatotoxic and hepatotoxic at higher concentrations (Nishie and
Daxenbichler, 1980). Administration of CHB caused exocrine pancreatic
atrophy with acinar cell apoptosis and hepatocellular necrosis with biliary
hyperplasia. CHB markedly elevated tissue GSH levels in the liver and
pancreas while renal GSH was unaffected (Wallig and Gould, 1986; Wallig
et al., 1992).
Conversely, 2S-l-cyano-2-hydroxy-3,4-epithiobutane (CHEB) was
acutely nephrotoxic causing dose- and time-dependent renal proximal
tubular epithelial cell necrosis (Gould et al., 1985). When administered
chronically, however, CHEB demonstrated hepatotoxicity and
pancreatotoxicity as well as nephrotoxicity (Gould et al., 1980). Hepatic
lesions were characterized by bile duct hyperplasia, fibrosis, megalocytosis,
79
and hepatocellular necrosis while pancreatic alterations consisted of
karyomegaly of acinar cells. Lesions within the kidneys were characterized
by cytomegaly consisting of cytoplasmic enlargement as well as striking
karyomegaly. CHEB caused marked elevations in renal, hepatic, and
pancreatic GSH (Wallig and Gould, 1986).
l-Cyano-3,4-epithiobutane (CEB; Figure 1.10) has demonstrated
teratogenic effects in pregnant Holtzman rats (Nishie and Daxenbichler,
1980) and nephrotoxic effects in male Fischer 344 rats (VanSteenhouse et al.,
1989). CEB was shown to be selectively nephrotoxic with no lesions evident
in the liver or pancreas. Administration of a single high dose (125 m g /k g
body weight) of CEB resulted in epithelial cell necrosis and karyomegaly
within the pars recta of the renal proximal tubules; administration of a
single low dose (50 m g /k g body weight) of CEB did not result in epithelial
cell death but caused karyomegaly within the renal proximal tubule.
In rats administered the high dose of CEB, an initial phase of
depletion in hepatic GSH was observed, followed by increased GSH by 12
hours post-CEB administration. Conversely, increased hepatic GSH was
observed by 12 hours post-CEB administration with no phase of depletion
in rats treated with low doses of CEB. With both doses, renal GSH was
elevated at all time periods with no phase of depletion recognized. Urinary
excretion of nonprotein thiols was increased with both doses. There were
no significant alterations in pancreatic GSH.
Further investigations by VanSteenhouse et al. (1991) revealed that
pretreatment with buthionine sulfoximine (BSO; Figures 1.11 and 1.12), an
inhibitor of y-glutamylcysteine synthetase and a GSH-depleting agent,
greatly diminished tubular necrosis and karyomegaly induced by CEB.
/
s \
N = C- CH2- CH2- CH — CH2
Figure 1.10
Structure of l-cyano-3,4-epithiobutane (CEB).
Figure 1.11 Structure of buthionine sulfoximine (BSO) in its
phosphorylated form.
82
'(GSSG)
MERCAPTURIC ACIDS
(GSH REDUCTASE)
(GST)
GLUTATHIONE
(GSH)
amino acu
(Y - GLUTAMYL
TRANS PEPTIDASE)
CYSTEINYL-GLYCINE
ADP+Pj
(GSH
SYNTHETASE)
ATP
(DIPEPTIDASE)
Y - GLUTAMYL AA
GLYCINE
(Y-GLUTAMYL
CYCLO»
TRANSFERASE)
y - GLUTAMYL
CYSTEINE
,/ADP+Pi
CYSTEINE
I (y-GLUTAMYL
I CYSTEINE
\ SYNTHETASE)
amino acids
BSO
■ATP
GLUTAMATE
5-OXOPROLINE
(5-QXOPROLINASE)
ATP
ADP+Pj
Figure 1.12 Diagram of the y-glutamyl cylce depicting the inhibitory effect
of buthionine sufoximine (BSO) on y-glutamylcysteine synthetase.
83
Depletion of hepatic GSH, increased urinary excretion of nonprotein thiols,
and decreased nephrotoxicity following pretreatment with BSO suggest that
GSH conjugation is an important pathway of bioactivation in CEB
metabolism with the conjugate, or a metabolite of the conjugate, being
involved in the nephrotoxicity. Since CHEB is also selectively nephrotoxic
while CHB is not, it is possible that the epithiol moiety of CEB and CHEB
may be involved in the nephrotoxicity associated with these compounds.
Objectives
Objectives for subsequent research have focused on elucidation of the
nephrotoxic metabolite(s) of CEB. To evaluate the role of the GSH
conjugate, or metabolite of the GSH conjugate, in CEB-induced
nephrotoxicity and karyomegaly, various inhibitors of GSH transport and
metabolism were administered to male Fischer 344 rats. The protective
effect of these inhibitors against the toxicity was evaluated via
histopathology and karyometry.
Fischer 344 rats were administered a single dose of CEB following
pretreatment with one of three selective inhibitors of GSH transport and
metabolism including acivicin (AT-125 or a-amino-3-chloro-4,5-dihydro-5isoxazoleacetic acid), probenecid (p-di-n-propylsulfamoyl benzoic acid), and
aminooxyacetic acid (AOAA). Additionally, treatment of rats with one of
the selective inhibitors alone was essential to elucidate the effect, if any, on
the experimental parameters in the absence of CEB.
AT-125 is a glutamine analog and irreversible inhibitor of y-glutamyl
transpeptidase (GGT; Allen et al., 1980; Griffith and Meister, 1980). GGT is
the only known enzyme able to cleave the y-glutamyl moiety of GSH and
GSH-conjugates (Goldberg et al, 1960). Inhibition of GGT will prevent
84
subsequent cleavage of the cysteinyl-glycine conjugate and uptake of the
cysteine conjugate into the renal tubular epithelial cell. Protection against
CEB-induced nephrotoxicity afforded by AT-125 would incriminate the
cysteine conjugate, or a metabolite derived from the cysteine conjugate, in
the nephrotoxic effect.
Conversely, probenecid competitively inhibits the organic anion
transport system (OATS; Gutman et al., 1955). Probenecid prevents
secretion of the N-acetylated cysteine conjugate or mercapturic acid from
hepatocytes and inhibits subsequent renal uptake at the basolateral
membrane. Mercapturate biosynthesis is accomplished by the transfer of an
acetyl group from acetyl coenzyme A to the cysteine S-conjugate by cysteine
conjugate N-acetyltransferase (Duffel and Jakoby, 1982). If the mercapturic
acid is responsible for CEB-induced nephrotoxicity in Fischer 344 rats,
probenecid may diminish the histopathological lesions.
Finally, AOAA inhibits pyridoxal phosphate-dependent enzymes
including renal cysteine conjugate (3-lyase (Jones et al., 1986). (3-lyase has
been shown to catalyze the conversion of cysteine conjugates to potentially
reactive thiols in addition to ammonia and pyruvate (Tateishi and
Shimizu, 1980). If the reactive thiol is responsible for CEB-induced
nephrotoxicity, administration of AOAA prior to CEB treatment will
ameliorate the nephrotoxic effects.
Renal and hepatic GSH levels were also determined following
pretreatment with these inhibitors. GSH levels were evaluated to assess the
possibility of the involvement of a CEB-derived metabolite in the GSHenhancing effect. Enhanced renal GSH may prove anticarcinogenic by
conjugating and detoxifying other electrophiles which may be potential
85
mutagens an d /or carcinogens. Additionally, elevated renal GSH levels
may scavenge free radicals associated with oxidative stress and free radicalinduced mutagenesis and tumorigenesis. The free radical-scavenging
ability of GSH may also prove beneficial for renal transplant recipients
during periods of extreme oxidative stress following ischemic injury and
during the reperfusion phase of renal transplantation (Paller et al, 1984;
Ouriel etal., 1985).
y-Glutamylcysteine synthetase (GCS) activity was measured to assess
the effect of CEB on the rate-limiting enzyme in GSH synthesis. An
induction of GCS activity in the presence of CEB suggests that elevated GSH
levels may be due to increased de novo synthesis of GSH as opposed to
increased transport of GSH from the liver to the kidney.
Measurement of urinary mercapturates both following
administration of CEB alone and following pretreatment with the various
inhibitors was also of interest. Mercapturic acids are metabolic endproducts of GSH conjugation. Increased excretion of mercapturates into the
urine would further demonstrate the importance of GSH conjugation in
CEB metabolism.
Finally, investigations evaluating the effect of CEB on phase I
xenobiotic metabolizing enzyme systems were pertinent to the elucidation
of CEB metabolism in its entirety. Although CEB is metabolized through
GSH conjugation, this does not preclude involvement of phase I
m etabolizing enzymes such as cytochrome P450 and epoxide hydrolase.
Measurement of total cytochrome P450 levels as well as the activity of more
specific cytochrome P450 enzymes such as ethoxyresorufin O-deethylase,
pentoxyresorufin O-depentylase, and 7-ethoxycoumarin O-deethylase, were
86
evaluated in the liver and kidney. Renal and hepatic epoxide hydrolase
activities were also measured following administration of a single dose (125
mg / kg body weight) of CEB.
The activity of glutathione S-transferases (GSTs) toward l-chloro-2,4dinitrobenzene (CDNB) was also evaluated following administration of
CEB. CDNB reacts with a broad spectrum of GSTs and is considered a
general substrate for evaluation of GST activity. Since CEB is conjugated
with GSH in vivo, it is of interest to assess the effect of CEB on enzymes
involved in the catalytic reaction.
CHAPTER 2
PROTECTIVE EFFECT OF AMINOOXYACETIC ACID ON CEB-INDUCED
NEPHROTOXICITY IN FISCHER 344 RATS
Introduction
Cruciferous plants are cultivated extensively for use as livestock feed, in
the manufacturing of plastics, and for human consumption as condiments,
vegetables, and oilseed. Glucosinolates, derived from cruciferous vegetables,
undergo autolytic hydrolysis to yield various cyanoepithioalkanes including 1 cyano-3,4-epithiobutane (CEB; Luthy and Benn, 1980).
CEB, the structure of which was initially proposed by Kirk and
MacDonald (1974), has demonstrated teratogenic effects in pregnant Holtzman
rats (Nishie and Daxenbichler, 1980) and nephrotoxic effects in male Fischer
344 rats (VanSteenhouse, et al., 1989). Administration of CEB to male Fischer
344 rats resulted in epithelial cell necrosis and karyomegaly within the pars
recta of the proximal convoluted tubules. CEB treatment also caused an early
phase of depletion of hepatic glutathione (GSH) followed by a rebound
elevation. Renal GSH levels, however, were elevated at all time periods
evaluated with no identifiable phase of depletion. Additionally, CEB-induced
nephrotoxic and karyomegalic changes were diminished following
pretreatment with buthionine sulfoximine (BSO), an inhibitor of yglutamylcysteine synthetase (GCS) and GSH-depleting agent (VanSteenhouse,
e ta l, 1991).
GSH, a ubiquitous tripeptide (y-glutamylcysteinyl glycine), conjugates
and detoxifies a variety of electrophilic, hydrophobic compounds (Mitchell et
al., 1973; Sinsheimer et al, 1987; Mannervik et al, 1989). GSH conjugation may,
however, lead to the bioactivation of xenobiotics with resultant cytotoxic or
genotoxic effects (Elfarra and Anders, 1984; Inskeep etal, 1986; Koga, etal.,
87
88
1986; Guengerich et al., 1987; Anders, 1990; Koob and Dekant, 1991). The rapid
depletion of hepatic GSH and reduction of CEB-induced nephrotoxicity
following BSO pretreatment strongly suggest that GSH conjugation and
bioactivation is a significant pathway in CEB metabolism and toxicity.
The present study was devised to further elucidate the role of the GSH
conjugate, or a metabolite thereof, in CEB-induced nephrotoxicity and altered
renal GSH. Elevations of renal GSH may partially explain the chemoprotective
effects associated with cruciferous vegetables.
Elevations in renal GSH may prevent carcinogenesis in a twofold
manner. GSH may conjugate and detoxify electrophilic mutagens and
carcinogens and may scavenge free radicals associated with oxidative stress
and free radical-induced mu tagenesis and tumorigenesis (Feig et al., 1994;
Satoh and Lindahl, 1994). The free radical-scavanging ability of GSH may also
prove beneficial for renal transplant recipients during periods of extreme
oxidative stress following ischemic injury and during the reperfusion phase of
renal transplantation (Paller et al., 1984; Ouriel et al., 1985).
Fischer 344 rats were administered a single dose of CEB following
pretreatment with one of three selective inhibitors of GSH transport and
metabolism including acivicin (AT-125 or a-amino-3-chloro-4,5-dihydro-5isoxazoleacetic acid), probenecid (p-di-»-propylsulfamoyl benzoic acid), and
aminooxyacetic acid (AOAA). AT-125 is a glutamine analog and irreversible
inhibitor of y-glutamyl transpeptidase (GGT; Allen etal., 1980; Griffith and
Meister, 1980). GGT is the only known enzyme able to cleave the y-glutamyl
moiety of GSH and GSH-conjugates (Goldberg et al., 1960). Inhibition of GGT
will prevent subsequent cleavage of the cysteinyl-glycine conjugate and uptake
of the cysteine conjugate into the renal tubular epithelial cell. Protection
89
against CEB-induced nephrotoxicity afforded by AT-125 would incriminate the
cysteine conjugate, or a metabolite thereof, in the nephrotoxic effect.
Conversely, probenecid competitively inhibits the organic anion
transport system (OATS; Gutman etal., 1955) preventing secretion of the N acetylated cysteine conjugate or mercapturic acid from hepatocytes and
subsequent renal uptake of the mercapturate at the basolateral membrane.
Mercapturate biosynthesis is accomplished by the transfer of an acetyl group
from acetyl coenzyme A to the cysteine S-conjugate catalyzed by cysteinethioester N-acetyltransferase (Duffel and Jakoby, 1982). If the mercapturate is
responsible for CEB-induced nephrotoxicity in Fischer 344 rats, probenecid
may diminish the histopathological lesions.
Finally, AOAA inhibits pyridoxal phosphate-dependent enzymes
including renal cysteine conjugate p-lyase (Jones et al., 1986). p-lyase has been
shown to catalyze the conversion of cysteine conjugates to potentially reactive
thiols in addition to ammonia and pyruvate (Tateishi and Shimizu, 1980). If
the reactive thiol was responsible for CEB-induced nephrotoxicity,
administration of AOAA prior to CEB treatment will ameliorate the
nephrotoxic effects.
Renal and hepatic GSH levels were also determined following
pretreatment with these inhibitors. GSH levels were evaluated to assess the
possibility of the involvement of a CEB-derived metabolite in the GSHenhancing effect.
GCS activity in the liver and kidney was similarly measured to assess
the effect of CEB on the rate-limiting enzyme in GSH synthesis. If renal GCS
activity is induced in the presence of CEB, elevated GSH levels may be due to
90
increased de novo synthesis of GSH as opposed to increased transport of GSH
from the liver to the kidney.
Finally, rats were administered one of the three selective inhibitors alone
to evaluate the effect of these compounds on the experimental parameters in
the absence of CEB.
Materials and Methods
Animal Maintenance and Treatment
One hundred and thirty eight male, 5-7 week old Fischer 344 rats were
obtained from Harlan Sprague Dawley, Inc. (Indianapolis, IN, USA). Rats
were housed in an AAALAC (American Association for Accreditation of
Laboratory Animal Care)-accredited facility. Rats were initially housed in
groups of 3-5 under controlled conditions of 12-hour light/ dark cycle at 21 °C
and 50-70 % relative humidity. For one week prior to treatment rats were fed a
modified, semi-purified diet, AIN-76A, with cornstarch substituted for sucrose
to prevent hepatic lipidosis (Hamm etal, 1982). Throughout experimental
trials rats were housed individually in metabolic cages to facilitate estimation
of feed and water intake. Water was allowed ad lib. Pair-feeding was
instituted to abrogate the effects of dietary intake on hepatic GSH levels.
At 8-9 weeks of age rats were divided into one of eight treatment groups
in a random fashion. The eight groups consisted of a negative, saline control
group; a positive, CEB-treated control group; an AT-125 pretreated group, a
probenecid pretreated group, and an AOAA pretreated group; and three
treatment groups administered one of the three inhibitors alone. Each group of
rats was further divided into
2
groups (minimum of 6 rats / group) for
evaluation at 24 or 48 hours post-CEB administration.
91
An intraperitoneal (IP) injection of 10 mg AT-125/ kg body weight was
administered in sterile saline, pH 6.5, two hours prior to CEB treatment while
200 mg probenecid/kg body weight was administered IP in 0.1 M sterile
potassium phosphate buffer, pH 7.4,30 minutes prior to and 5.5 hours
following CEB administration. One hour prior to CEB administration, the
AOAA pretreated group received an IP injection of 32.8 mg A O A A /kg body
weight in 0.1 M sterile phosphate buffer, pH 7.0.
All rats were then orally gavaged with 125 mg CEB/kg body weight or
the same relative volume of deionized, distilled water with respect to body
weight. All gavaging was performed between 8 and 9 AM.
Twenty-four or 48 hours following CEB administration, rats were
anaesthetized under isoflurane inhalation and euthanized via cardiac
exsanguination. Liver and kidney samples were prepared for histological and
histomorphometric examination as well as GSH and y- glutamylcysteine
synthetase (GCS) determination.
In accordance with the Animal Welfare Act, the experimental protocol
was submitted to and approved by the Louisiana State University Institutional
Animal Care and Use Committee.
Materials
CEB was graciously synthesized by Dr. Mark McLaughlin and Tamara
Schaller from the Department of Chemistry at Louisiana State University. A
modified method of Luthy and Benn (1980) was utilized to synthesize the CEB.
The bromo group of 4-bromo-l-butene (Janssen Chimica, N ew Brunswick, NJ,
USA) was replaced with a cyano moiety from sodium cyanide (Aldrich,
Milwaukee, WI, USA) to yield l-cyano-3,4-butene. l-Cyano-3,4-butene was
then oxidized at the unsaturated double bond with magnesium
92
monoperoxyphthalate hexahydrate (Aldrich, Milwaukee, WI, USA) yielding 1 cyano-3,4-epoxybutane which was subsequently converted to a thiouronium
salt in the presence of thiourea (Aldrich, Milwaukee, WI, USA). Final
conversion to l-cyano-3,4-epithiobutane was achieved in an alkaline
environment, pH 9.0. The identity of the product was confirmed by nuclear
magnetic resonance and the purity of l-cyano-3,4-epithiobutane was
demonstrated to be 98% or greater by gas chromatography (see Appendix).
L-glutamate, L-a-aminobutyrate, adenosine triphosphate, disodium
salt, bovine serum albumin, reduced glutathione, glutathione reductase
[NADPH: oxidized-glutathione oxidoreductase (GSSG reductase); EC 1.6.4.2],
reduced (3-nicotinamide adenine dinucleotide phosphate, 5,5'-dithiobis-(2nitrobenzoic acid) (DTNB), aminooxyacetic acid, AT-125 (acivicin), probenecid,
and a diagnostic kit for determination of inorganic phosphorus (Procedure no.
670) were obtained from Sigma Chemical Co. ( St. Louis, MO, USA). Procedure
23225X-BCA Protein Assay Reagent method for the determination of total
protein was obtained from Pierce (Rockford, IL, USA). Ethylene diamineacetic
acid, disodium salt and magnesium chloride were obtained from Ameresco
(Solon, OH, USA) and Mallinckrodt, Inc. (Paris, KY, USA), respectively. Saline
was from Baxter Healthcare Corp. (Deerfield, IL, USA). The dietary
components were obtained from ICN Nutritional Biochemical (Costa Mesa,
CA, USA).
Samples were prepared using a TissumizerR rotor-sator generator
(Tekmar, Cinncinnati, OH, USA) and a Beckman Model TJ- 6 centrifuge
(Beckman Instruments, Inc., Fullerton, CA, USA). A Hitachi U-2000 doublebeam U V /V is spectrophotometer (Hitachi Instruments, Inc., San Jose, CA,
USA) was utilized to analyze GSH levels and GCS activity.
93
Tissue Preparation
Samples of liver and kidney for GSH and GCS determination were
collected from the median liver lobe and cortex of the left kidney. The right
kidney and the remainder of the median liver lobe were utilized for
preparation of tissue sections for histological and histomorphometric
evaluation.
Liver and kidney samples taken for histological and histomorphometric
examination were immediately fixed in neutral buffered 10% formalin. Fixed
tissues were trimmed, processed, embedded in paraffin, and cut in 3-4 //m
sections. Sections were stained with haematoxylin and eosin.
A 150-200 mg sample of liver or kidney was collected for GSH analysis
and added to 4 mis 0.02M EDTA in a stoppered tube on ice. The specimens
were homogenized for 20-30 seconds using a TissumizerR rotor-sator generator
(Tekmar, Cinncinnati, OH, USA). Four mis 10% trichloroacetic acid were
added to the homogenate. The homogenate was allowed to stand for 15
minutes prior to centrifugation at 4°C for 15 minutes at 3000g. The
supernatant was saved and frozen at -20°C until analysis.
To determine GCS activity, a 150-200 mg sample of liver or kidney was
placed in 2 mis of 0.1 M Tris-HCl buffer (pH 8.2). Tissues were homogenized
for 20-30 seconds. Samples were centrifuged at 1800 rpm for 20 minutes at 4°C
and the fresh supernatant assayed for GCS activity.
Biochemical Assays
Total GSH levels were determined on the supernatants utilizing the
colorimetric recycling method of Tietze (1969). A 200 //I supernatant sample
was added to 700 //I of 0.3 mM NADPH and 100 /d of 6.0 rnM DTNB. Samples
were incubated for 5 minutes at 30°C and 20 //Is of GSSG reductase
94
(25 U nits/m l buffer) were added to initiate the reaction. The change in
absorbance was recorded at 412 nm. GSH activity was expressed in mmol
GSH /100 g tissue.
GCS activities were evaluated on fresh supernatant using a modified
method of Sekura and Meister (1977). Kidney and liver samples were diluted
1:20 and 1:5, respectively. A 200 //I supernatant sample was added to 200 //I of
10 mM glutamate, 100 //I of 10 mM L-a-aminobutyrate, 200 //I of 20 mM MgCl^
100 //I of 2 mM N a 2 EDTA, 100 //I of 5 mM Na 2 ATP, and 20 //g BSA. The
samples were incubated at 37°C for 30 minutes and the reaction stopped with
1.0 ml of 20% TCA. Samples were allowed to stand at room temperature for 510 minutes and centrifuged at 1800 rpm for 10 minutes at 4°C. The liberation
of inorganic phosphorous was determined by the use of Sigma Diagnostic
Procedure No. 670. The absorbance of the samples was read at 660 nm. An
appropriate sample blank was assayed to compensate for ATPase activity.
GCS activity was expressed in //moles G CS/hr/m g protein.
Total protein levels were determined by the Pierce Procedure 23225XBCA Protein Assay Reagent method. The absorbance of the samples was
recorded at 562 nm. Total protein levels were expressed in mg protein/ml.
GSH, GCS, and protein determinations were performed on a Hitachi U-2000
double-beam U V /V is spectrophotometer.
Histological and Histomorphometric Examination
Hematoxylin and eosin stained tissue sections were evaluated for
hepatic and renal lesions on a Leitz Labolux 12 transmitting-light microscope
(Leica, Inc., Rockleigh, NJ, USA) at magnifications x 100 and 400. Areas
containing the tips of the medullary rays within the renal cortex were
delineated with ink using a Wild M3 typ 376788 stereomicroscope (Leica, Inc.,
95
Rockleigh, NJ, USA). The cross-sectional area of one-hundred and twenty
proximal tubular epithelial cell nuclei within these regions, excluding affected
cells with pyknotic nuclei, were measured with a Zeiss axioplan microscope
(Carl Zeiss, Inc., Thornwood, USA), lOOOx magnification, equipped with a
semi-automated IM-Series System, Version 3.0 data program (Analytical
Imaging Concepts, Irvine, CA, USA).
Statistical Analysis
GSH levels and GCS activity were evaluated by analysis of variance
with respect to treatment group. The means of each treatment group were
compared to one another by Fisher's least significant difference test for
multiple comparisons at a 95% confidence level.
Mean nuclear size and frequency distributions of nuclear area were
determined for each animal and averaged for each respective treatment group
at 24 and 48 hours. The mean values for the treatment groups were then
evaluated by analysis of variance using Fisher's least significant difference test
for multiple comparisons at a 95% confidence level. Comparisons between 24
and 48 hour CEB treated groups were analyzed by an unpaired t-test with
respect to time. P-values <0.02 were considered significant.
Results
Renal and Hepatic GSH
Hepatic and renal GSH levels were significantly elevated at 24 and 48
hours in rats treated with CEB alone or CEB in addition to any of the inhibitors
with the exception of hepatic GSH at 24 hours in rats pretreated with
probenecid (Figures 2.1 and 2.2). Additionally, rats pretreated with AT-125
revealed significantly greater levels of GSH in the kidney at 24 hours as
compared to AOAA-pretreated rats and rats administered with CEB alone. At
96
♦
□ 24 h o u r s
S 4 8 h o u rs
C o n tr o l
AOAA
P r o b e n e c id
A T -1 2 5
C EB
Treatment
Figure 2.1
Effect of inhibitor pretreatment or treatment with CEB alone on
renal GSH levels at 24 and 48 hours post-CEB administration. Value bars
represent mean GSH levels. Error bars represent the standard errror of the
mean; n = minimum of 6 rats per group. ♦ represents P = 0.0001 as compared
to controls; t represents P = 0.0001 as compared to CEB treated group; *
represents P = 0.0001 as compared to AOAA-pretreated rats.
97
□ 24 H o u rs
□ 48 H ours
C o n tr o l
AOAA
P r o b e n e c id
A T -1 2 5
CEB
Treatment
Figure 2.2
Effect of inhibitor pretreatment or treatment with CEB alone on
hepatic GSH levels at 24 and 48 hours post-CEB administration. Value bars
represent mean GSH levels. Error bars represent the standard errror of the
mean; n = minimum of 6 rats per group. ♦ represents P <0.001 as compared to
controls; + represents P < 0.001 as compared to CEB treated group; * represents
P <0.001 as compared to AOAA-pretreated rats.
98
48 hours, AT-125 pretreated rats exhibited elevated renal GSH levels as
compared to CEB controls while rats pretreated with probenecid had increased
renal GSH levels as compared to CEB controls and AOAA-pretreated rats.
Hepatic GSH was significantly greater in rats treated with CEB alone or rats
pretreated with AOAA or AT-125 as compared to probenecid-pretreated rats at
24 hours. At 48 hours, rats pretreated with any of the inhibitors exhibited
significantly elevated hepatic GSH levels as compared to CEB treated rats.
Additionally, AT-125-pretreated rats revealed elevated GSH levels in the liver
at 48 hours as compared to rats pretreated with AOAA. None of the rats
pretreated with an inhibitor revealed significantly decreased renal or hepatic
GSH as compared to CEB treated rats.
Significant elevations in renal GSH were also observed at 24 hours in
rats treated with AOAA alone (0.244 ± 0.02; p = 0.0001) and at 24 (0.333 ± 0.02;
p = 0.0001) and 48 hours (0.290 ± 0.03; p = 0.001) in rats treated with AT-125
alone; however, the magnitude of the elevation of renal GSH in rats receiving
CEB in addition to AOAA or AT-125 was statistically greater (p = 0.0001) than
those rats receiving only AOAA or AT-125.
Renal and Hepatic GCS Activity
Rats administered CEB alone, or pretreated with AT-125 or probenecid
in addition to CEB, exhibited significantly decreased renal GCS activity as
compared to controls at 24 hours post-CEB administration; however, renal GCS
activity at 24 hours in rats treated with AOAA in addition to CEB was not
significantly different from saline controls. No significant differences were
seen in renal GCS activity at 48 hours or in hepatic GCS activities at 24 or 48
hours (Figures 2.3 and 2.4). Additionally, there was no significant difference in
99
12 i
□ 24 h o u rs
10
-
6
-
H 4 8 h o u rs
Q,
CD
E
>_
JC
w
o
o
o
E
5.
4 -2
-
C o n tro l
AOAA
P r o b e n e c id
A T -125
CEB
Treatment
Figure 2.3
Effect of inhibitor pretreatment or treatment with CEB alone on
renal GCS activities at 24 and 48 hours post-CEB administration. Value bars
represent mean GCS activites. Error bars represent the standard errror of the
mean; n = minimum of 6 rats per group. ♦ represents P <0.02 as compared to
controls. No significant differences were observed between CEB treated and
inhibitor pretreated groups.
100
□ 24 h o u rs
H 48 h o u rs
C o n tr o l
AOAA
P r o b e n e c id
A T -1 2 5
CEB
Treatment
Figure 2.4
Effect of inhibitor pretreatment or treatment with CEB alone on
hepatic GCS activities at 24 and 48 hours post-CEB administration. Value bars
represent mean GCS activities. Error bars represent the standard errror of the
mean; n = minimum of 6 rats per group. N o significant differences were
observed at 24 or 48 hours.
101
renal or hepatic GCS activity in rats treated with any of the inhibitors alone at
24 or 48 hours.
Histological Evaluation
Histologic lesions were confined to the kidney and characterized by
intensely eosinophilic cells with dense, pyknotic nuclei occurring individually
or in clusters within the pars recta of the proximal tubular epithelium at the
tips of the medullary rays. Vesiculated, karyomegalic nuclei were observed in
many of the neighboring epithelial cells. These cells exhibited normal
cytoplasmic eosinophilia but possessed a large vesiculated nucleus with a thin
rim of condensed chromatin and a prominent nucleolus.
Epithelial cell necrosis and karyomegaly were observed sporadically at
24 hours and consistently at 48 hours in rats treated with CEB alone (Figures
2.5-2.7). Rats pretreated with AOAA prior to CEB administration revealed no
histologic changes at 24 hours and were similar to saline treated controls
(Figure 2.8). By 48 hours, however, significant histological lesions and
karyomegaly were observed in rats pretreated with AOAA (Figure 2.9). Those
rats pretreated with AT-125 or probenecid, however, exhibited more severe
renal lesions as compared to AOAA-pretreated rats at 48 hours.
Rats pretreated with AT-125 or probenecid revealed characteristic
histopathological lesions at both 24 and 48 hours (Figures 2.10-2.13). Animals
treated with AT-125, AOAA, or probenecid alone exhibited no significant
histopathological lesions. No significant histopathological lesions were
observed in the liver in any treatment group.
Karyometry
Renal karyometric evaluation revealed a marked increase in the nuclear
size of the cells of the proximal tubules at both 24 and 48 hours in CEB-treated
Figure 2.5
Photomicrograph of normal proximal tubular epithelial cells from a control rat kidney collected at 24
hours. Tissue sections from kidneys collected at 48 hours from control rats were essentially identical to this
photomicrograph. Hematoxylin and Eosin stain. Bar = 10 nm.
Figure 2.6
Photomicrograph depicting renal lesions at 24 hours in the pars recta of the proximal tubule from a rat
treated with CEB alone. Renal lesions are characterized by dense eosinophilic epithelial cells (thin arrow) and
karyomegalic nuclei (broad arrow). Hematoxylin and Eosin stain. Bar = 10 fim.
Figure 2.7
Photomicrograph depicting renal lesions at 48 hours in the pars recta of the proximal tubule from a rat
treated with CEB alone. Lesions reveal the same morphologic changes as illustrated in Figure 2.6, but are of greater
severity. Hematoxylin and Eosin stain. Bar = 10 /zm.
Figure 2.8
Photomicrograph of the pars recta of the proximal tubules at 24 hours from a rat treated with AOAA
prior to CEB administration. Note the absence of significant morphological changes. Hematoxylin and Eosin stain.
Bar = 10 (im.
Figure 2.9
Photomicrograph of the pars recta of the proximal tubules at 48 hours from a rat treated with AOAA
prior to CEB administration. Dense eosinophilic cells (thin arrow) and karyomegalic nuclei (broad arrow) are evident.
Hematoxylin and Eosin stain. Bar = 10 jzm.
Figure 2.10 Photomicrograph of the pars recta of the proximal tubules at 24 hours from a rat treated with AT-125 in
addition to CEB. Note the presence of significant renal lesions. Hematoxylin and Eosin stain. Bar = 10 fjm.
Figure 2.11 Photomicrograph of the pars recta of the proximal tubules at 48 hours from a rat treated with AT-125 in
addition to CEB. Note the presence of significant renal lesions. Hematoxylin and Eosin stain. Bar = 10 f«n.
Figure 2.12 Photomicrograph of the pars recta of the proximal tubules at 24 hours from a rat treated with probenecid
in addition to CEB. Note the presence of significant renal lesions. Hematoxylin and Eosin stain. Bar = 10 fiin.
Figure 2.13 Photomicrograph of the pars recta of the proximal tubules at 48 hours from a rat treated with probenecid
in addition to CEB. Note the presence of significant renal lesions. Hematoxylin and Eosin stain. Bar = 10 pm.
o
I ll
rats as compared to controls (Figure 2.14). Mean nuclear size as well as the
number of nuclei > 55 //m 2 were significantly increased at 24 and 48 hours in
rats treated only with CEB as compared to controls. Additionally, the
frequency distributions of nuclear area for the control rats were significantly
greater than CEB-treated rats for the 35-55 /<m2 range at 24 hours and 25-55
;/m 2 range at 48 hours. These findings reveal a general shift toward larger
nuclear area in CEB-treated rats (Figures 2.15 and 2.16).
At 24 hours, rats pretreated with AOAA prior to CEB administration
revealed no significant difference in mean nuclear size as compared to controls.
AOAA-pretreated rats also exhibited significantly decreased mean nuclear area
as compared to rats treated with CEB alone. Similarly, no significant
differences between control rats and AOAA-pretreated rats were observed in
the frequency distributions of nuclear area with the exception of those nuclei
ranging from 45-55 jum2 which were significantly increased in controls as
compared to AOAA-pretreated rats.
By 48 hours AOAA pretreatment resulted in significantly increased
mean nuclear area. The mean nuclear area of AOAA-pretreated rats was,
however, significantly less than that of rats given CEB alone. The number of
nuclei falling in the 55-85 j/m 2 range at 48 hours were significantly greater for
AOAA-pretreated rats as compared to controls; yet, the number of nuclei >85
//m 2 in AOAA rats were significantly lower than CEB-treated rats. Animals
pretreated with AT-125 or probenecid demonstrated significantly greater mean
nuclear areas as compared to negative controls and were equivalent to CEB
control rats. At 24 and 48 hours the number of nuclei within the 35-55 ;«m2
range was greater in control rats as compared to rats pretreated with AT-125 or
probenecid. The number of nuclei within the 55-85 |im 2 and > 85 /»m 2 range
□ 24 h o u rs
8 48 h o u rs
C o n tro l
AOAA
P r o b e n e c id
A T -1 2 5
C EB
Treatment
Figure 2.14 Effect of inhibitor pretreatment or administration of CEB alone
mean nuclear size at 24 and 48 hours as measured by histomorphometiy.
Value bars represent mean nuclear area. Error bars represent the standard
error of the mean; n = minimum of 6 rats per group; 120 nuclei per rat. ♦
represents P < 0.001 as compared to controls; t represents P < 0.001 as
compared to CEB treated group.
113
♦ - ■ • C ontrol
•
— AOAA + CEB
* — -C E B
Nuclear Area (jim2)
Figure 2.15 Frequency distribution of proximal tubule epithelial cell nuclear
size w ith respect to treatment group at 24 hours. AT-125 and probenecid
pretreated groups were deleted for clarity as these groups were essentially the
same as animals treated with CEB alone. Data points represent the relative
number of nuclei within each range of nuclear area. Error bars represent the
standard error of the mean; n = minimum of 6 rats per treatment group; 120
nuclei per rat. .
114
0.6
>
-r
0.5 -
■■
-
0.4 -or
■ - C ontrol
* — AOAA + CEB
A
CEB
0.3 -0.2
- -
0.1
-
<25
25-35 35-45 45-55 55-65 65-75 75-85
>85
Nuclear Area (nm2)
Figure 2.16 Frequency distribution of proximal tubule epithelial cell nuclear
size with respect to treatment group at 48 hours. AT-125 and probenecid
pretreated groups were deleted for clarity as these groups were essentially the
same as animals treated with CEB alone. Data points represent the relative
number of nuclei within each range of nuclear area. Error bars represent the
standard error of the mean; n = minimum of 6 rats per treatment group; 120
nuclei per rat.
115
was greater in rats pretreated with AT-125 and probenecid at 24 and 48 hours
as compared to controls with the exception of the 75-85 /im 2 range in rats
pretreated with AT-125 at 24 hours. Rats pretreated with AT-125 or
probenecid revealed karyomegalic changes equivalent to CEB controls.
Additionally, rats pretreated with probenecid revealed significantly
elevated mean nuclear areas as compared to AOAA-pretreated rats at 24 and
48 hours. AT-125-pretreated rats, however, exhibited significantly increased
mean nuclear areas as compared to AOAA-pretreated rats only at 24 hours. At
24 hours the number of nuclei >85 /jm 2 was significantly greater in rats
pretreated with probenecid as compared to AOAA-pretreated rats. At 48
hours the number of nuclei within the 65-75 /«m2 range and > 85 /im 2 were also
significantly greater in probenecid-pretreated rats while the number of nuclei
within the 45-55 /<m2 range were significantly decreased as compared to
AOAA-pretreated rats. Similarly, rats pretreated with AT-125 exhibited
significantly decreased numbers of nuclei within the 45-55 //m 2 range and
significantly increased numbers of nuclei within the 55-75 //m 2 range and >85
//m 2 as compared to AOAA-pretreated rats.
There was also a time-dependent increase in the mean nuclear area and
frequency distributions for CEB-treated rats. Mean nuclear area and the
number of nuclei falling in the 75-85 //m 2 and >85 /mi2 ranges were
significantly greater at 48 hours as compared to 24 hours in rats given only
CEB.
No significant differences were detected for mean nuclear area or
frequency distibutions for rats treated with any of the inhibitors given alone as
compared to controls with the exception of rats pretreated with probenecid
116
which revealed an increased number of nuclei within the 35-45 fitn 2 range as
compared to controls.
Discussion
The present study indicates that the reactive thiol may be at least
partially responsible for CEB-induced nephrotoxicity. A single dose of 125 mg
CEB/ kg body weight results in mild nephrotoxicity at 24 hours and more
severe lesions at 48 hours post-CEB administration. The lesions were
characterized by epithelial cell necrosis and karyomegaly as previously
described by VanSteenhouse et al. (1989). The fact that hepatic lesions were not
observed suggests that the parent compound (CEB) is not directly toxic but
requires metabolic activation to exert its deleterious effect.
Renal proximal tubular epithelial cells are frequent targets for toxic
xenobiotics. The kidneys receive 25% of cardiac output (Monks and Lau, 1987)
and > 90% of the renal blood flow is directed to the renal cortex where the
majority of the proximal tubules are located (Maher, 1976). Additionally,
proximal tubular epithelial cells possess specialized functions of transport and
metabolism which may enhance toxicity through the concentration and
bioactivation of xenobiotics. Proximal tubular epithelial cells possess the
highest concentrations of y-glutamyl transpeptidase (Goldberg et al., 1960), the
only known enzyme able to cleave the y-glutamyl linkage, and are active in the
uptake and concentration of metabolites of GSH conjugates.
Though CEB targets the renal proximal tubular epithelium,
pretreatment with AOAA prior to CEB administration exerts a protective effect
against nephrotoxicity at 24 hours. AOAA competitively inhibits pyridoxal
phosphate-dependent enzymes including renal cysteine conjugate (3-lyase
(Elfarra et al., 1986). Renal cysteine conjugate (3-lyase catalyzes P-elimination
117
reactions to yield a potentially reactive thiol in addition to pyruvate and
ammonia. Reactive thiols may be excreted as free thiols, detoxified by
methylation or glucuronidation, or converted to electrophilic thiol derivatives
that bind to essential protein macromolecules (Dekant, 1993). Bioactivation by
cysteine conjugate (3-lyase has been documented for various nephrotoxic and
nephrocarcinogenic compounds including several haloalkenes (Dekant et al.,
1990) and bromobenzene (Monks etal, 1985).
Numerous cells possess relatively high y-glutamyl transpeptidase
activity including pancreatic acinar and ductile epithelial cells, epithelial cells
of the renal proximal tubule, jejunum, bile duct, epididymus, seminal vesicles,
bronchioles, thyroid follicles, as well as the chroid plexus, ciliary body, and
retinal pigment epithelium (Meister etal., 1976). These cells may be active in
the uptake of metabolites of GSH conjugates; however, other factors may also
play a role in susceptibility of different tissues to toxicity. The relative
activities of cysteine conjugate N-acetyltransferase and N-deacetylase as well
as cysteine conjugate |3-lyase may contribute to tissue selectivity. The fact that
CEB is selectively nephrotoxic and renal cysteine conjugate |3-lyase is unique
to the kidney further supports the finding that the reactive thiol is the
nephrotoxic metabolite.
The protective effect of AOAA is diminished by 48 hours. Renal lesions
and karyomegaly are present by 48 hours in rats pretreated with AOAA and
are of equivalent severity to rats treated with CEB alone. The decreased
protective effect of AOAA seen at 48 hours is presumably due to the normal
metabolism and degradation of AOAA or due to the inability of AOAA to
competitively inhibit (3-lyase in the presence of increased molecular
equivalents of the cysteine conjugate. Delayed increases in the cysteine
118
conjugate may result from slow or protracted metabolism of CEB and /or
transport of mercapturates or conjugates of GSH or cysteine from the liver to
the kidney. Mercapturates may also be deacetylated in the kidney to form the
respective cysteine conjugate. Such a delay in the formation of the nephrotoxic
metabolite is supported by the fact that those rats treated with CEB alone
exhibited more severe renal lesions at 48 hours as compared to 24 hours.
Inhibition of y-glutamyl transpeptidase by AT-125 should also prevent
nephrotoxicity if the reactive thiol is responsible for the CEB-induced toxicity.
Renal lesions observed in animals pretreated with AT-125 may be due to a lack
of complete inhibition of y-glutamyl transpeptidase. Though AT-125 inhibits
renal y-glutamyl transpeptidase by >95% (Shapiro and Curthoys, 1981),
sufficient activity may have remained to allow uptake of an adequate amount
of the cysteine conjugate to cause toxicity.
Additionally, transport of intact GSH conjugates across the basolateral
and luminal aspects of renal tubular epithelial cells has been documented
(Ballatori e ta l, 1994). If the GSH conjugate were directly toxic, however,
AOAA could not have ameliorated the nephrotoxicity in rats pretreated with
this inhibitor.
Failure of probenecid to protect against CEB-induced nephrotoxicity
suggests that transport of the mercapturate from the liver to the kidney is not
responsible for renal lesions. This does not, however, exclude the
mercapturate as a potential nephrotoxin since mercapturic acids may be
formed within the proximal tubular epithelial cell via N-acetyltransferase. Yet,
if the mercapturic acid is directly nephrotoxic, AOAA should not have
diminished the histopathological lesions since |3-lyase and N-acetyltransferase
mutually compete for the cysteine conjugate as a substrate. Inhibition of
119
P-Iyase in the presence of AOAA would provide adequate substrate for
mercapturate formation.
The reactive thiol also appears to be involved in diminished GCS
activity as suggested by the fact that AOAA-pretreated rats did not
demonstrate decreased renal GCS activity at 24 hours. The reactive thiol may
reduce amino acid residues within the active site or disulfide bridges essential
for GCS activity. Conversely, a metabolite of the reactive thiol, such as a
methylthio or S-glucuronide derivative, could inhibit GCS activity. The
possibility that AOAA pretreatment actually induced GCS activity is
contradicted by the fact that AOAA given alone did not induce enzyme
activity.
Decreased levels of renal GCS activity at 24 hours do not appear to be
the result of feedback inhibition by elevated GSH since renal GSH levels were
elevated at 48 hours as well as 24 hours. Renal GCS activity recovered by 48
hours post-CEB administration despite significant elevations in renal GSH
levels.
At both 24 and 48 hours a marked shift towards larger nuclear area was
observed in rats treated with CEB alone. Renal kaiyomegaly has been
documented following administration of auranofin (Markiewicz, 1988),
hexachlorobutadiene (Kociba e ta l, 1977), N-(DL-2-amino-2-carboxyethyl)-Llysine (Reyniers etal., 1974), lead acetate (Choie and Richter, 1972), N-(4'fluoro-4-biphenylyl)acetimide (Dees etal., 1980a, b), 2S-l-cyano-2-hydroxy-3,4epithiobutane (Gould etal., 1980), aflatoxin Bi (Butler, 1964), trichloroethylene
(Maltoni etal., 1988), cisplatin (Goudie etal., 1994), l-(2-chloroethyl-4methylcyclohexyl)-l-nitrosourea (MeCCNU) and chlorozotocin (Kramer etal.,
120
1986) and nephrotoxic mycotoxins such as ochratoxin A (Huff, 1991; Mantle et
a l, 1991; Boorman etal., 1992; Mantle, 1994).
Karyomegaly may represent a preneoplastic change and is recognized
as one of the earliest discernable morphological features in model systems of
renal carcinogenesis (Dees et al., 1980a, b). There is evidence that nuclear size
increases during progression from hyperplasia to preneoplastic and neoplastic
lesions (Young etal., 1984). Karyomegaly preceeds tumorigenesis following
administration of N-(4'-fluoro-4-biphenylyl)acetimide (Dees et al, 1980a, b),
aflatoxin Bi (Butler, 1964), ochratoxin A (Boorman et a l, 1992), auranofin
(Markiewicz, 1988), trichloroethylene (Maltoni et al, 1988), and
hexachlorobutadiene (Kociba etal., 1977) the latter two of which are selective
nephrotoxins bioactivated by GSH conjugation and metabolism (Terracine and
Parker, 1965; Jaffe et al, 1983; Elfarra and Anders, 1984; Reichert and Schutz,
1986). Renal carcinogenesis subsequent to CEB-induced karyomegaly has not
been demonstrated and further investigations regarding chronic exposure to
CEB and DNA adduct formation are needed.
Several nephrotoxic agents cause renal karyomegaly unassociated with
any neoplastic change (Choie and Richter, 1972; Reyniers et al., 1974).
Karyomegaly may represent compensatory hyperplasia in response to renal
tubular epithelial cell necrosis. However, karyomegaly has been observed with
low doses of CEB in the absence of cell death (VanSteenhouse et al, 1989).
It is also unclear whether increased nuclear size is a direct effect of CEB
or the GSH conjugate of CEB or is mediated through elevated levels of renal
GSH. Increased cellular levels of GSH in cultured human bronchial epithelial
cells and K562 leukemia cells preceeded the major growth phase (Frischer et al,
1993; Atzori et al, 1994) while depletion of GSH decreased the proliferative
121
ability of these cells (Atzori et al., 1994). Additionally, elevations of GSH in
endothelial cells and fibroblasts prevented inhibition of cellular proliferation
by transforming growth factor-|3i (Das et a l, 1992; White et al., 1992) and
hyperbaric oxygen (Honda and Matsuo, 1989), respectively.
The mechanism of increased cellular proliferation in response to
elevated GSH levels is still unclear. GSH is involved as an electron donor at
the active site of ribonucleotide reductase via glutaredoxin (Stiyer, 1988).
Elevated levels of GSH may therefore increase DNA synthesis with resultant
karyomegaly. GSH also acts as a coenzyme for glyoxalase which catalyzes the
conversion of methylglyoxal to lactate (Ohmori et al, 1989). Accumulation of
methylglyoxal retards cell growth (Szent-Gyorgyi, 1965). Increased GSH may
increase the rate of methylglyoxal degradation and release the cell from its
inhibitory effects.
GSH may indirectly influence cell proliferation through the modulation
of peptide growth factors. GSH has been shown to be involved in the release
of fibroblast growth factor (FGF) from NIH 3T3 cells (Jackson et al, 1995). FGF
has been demonstrated to cause karyomegaly of glomerular podocytes when
administered to rats (Mazue etal., 1993).
Though diminished intracellular GSH stimulated cystine transport in
endothelial cells (Deneke et al, 1992), increased GSH may increase transport of
cystine, an essential amino acid for cell growth in culture. The efflux of GSH
from renal proximal tubular epithelial cells is coupled to transport and uptake
of y-glutamyl amino acids including y-gluta my Icystine (Griffith etal., 1979a;
Anderson and Meister, 1983). y-Glutamylcystine may then be reduced by
transhydrogenation with GSH to form cysteine and y-glutamylcysteine.
122
Cysteine may then be utilized for GSH synthesis or in the synthesis of proteins
or other peptides essential for cell growth.
Alternatively, the GSH conjugate, or a metabolite of the GSH conjugate,
may also have an effect on cellular proliferation. S-(l,2-dicarboxyethyl)
glutathione enhances DNA synthesis in the presence of epidermal growth
factor (Miyazaki e ta l, 1990). Additionally, cystine transport into human
erythrocytes was increased in the presence of extracellular l-chloro-2,4dinitrobenzene (CDNB). Cystine was incorporated into fractions of the GSH
conjugate of CDNB (Otsuka, 1989). Similarly, the GSH conjugate of CEB may
induce the transport of cystine and stimulate cell growth.
Elevations in renal and hepatic GSH do not appear to be the result of
induction of GCS activity. Alternative mechanisms for elevations in GSH
include enhanced transcriptional activation or stabilization of GCS mRNA and
increased cellular uptake of cysteine or cystine. Cysteine is the rate-limiting
substrate for GSH synthesis (Deneke and Fanburg, 1989).
Cyanohydroxybutene (CHB), a naturally occurring nitrile also derived
from cruciferous vegetables, elevates hepatic and pancreatic GSH but does not
cause significant elevations in hepatic or pancreatic GCS activity. Pancreatic
and hepatic cysteine equivalents and hepatic GCS mRNA concentrations,
however, were elevated in CHB-treated rats (Davis etal., 1993).
Increased transport of cystine and cysteine may be mediated by GGT.
GGT cleaves extracellular GSH and increases transport of y-glutamyl amino
acids. Increased transport of y-glutamylcystine may elevate intracellular GSH
levels through reduction of y-glutamylcystine to cysteine and y-glutamylcysteine. In this manner, increased cysteine transport may bypass y-glutamyl cysteine synthetase and increase intracellular GSH levels. AT-125 did not,
123
however, prevent the GSH-enhancing effect of CEB despite its ability to inhibit
GGT. Alternatively, other transport mechanisms for cysteine or cystine may be
involved.
Increases in GCS mRNA, as seen with CHB, do not necessarily correlate
with increased GCS activity. Mercury has been shown to increase GCS mRNA
without increasing GCS activity (Wood et al, 1992). CEB may actually inhibit
de novo protein synthesis or post-translational modifications of GCS thereby
preventing an increase in the active form of the enzyme. The mechanism of
GSH elevation, therefore, may prove to be multifaceted including increases in
substrate availability and induction at the genetic level, but does not appear
dependent on actual induction of GCS activity.
Pretreatment with probenecid appeared to prevent the GSH-enhancing
effect of CEB in the liver at 24 hours. According to Christensen (1984), amino
acid transport involves the transport of ions. Systems A, ACS, and L typically
transport amino acids in the zwitterionic, or dipolar, form (Bannai and
Tateishi, 1986). Cystine, however, is transported as a tripolar, or anionic, ion
in human fibroblasts (Bannai and Kitamura, 1980,1981) and hepatocytes
(Makowske and Christensen, 1982). Since probenecid is also an anion at
physiological pH and inhibits the organic anionic transport system, it is
possible that probenecid may also inhibit cystine uptake and subsequent GSH
synthesis. This suggests that a lack of substrate availability in probenecid
pretreated rats limits the rate of GSH synthesis in the liver at 24 hours. This
transport system is independent of GGT-mediated uptake of cystine and
would therefore not be affected by AT-125. This further supports a role of
increased cystine transport in elevated GSH levels. Additionally, renal GSH
was not affected by pretreatment with probenecid presumably since this
124
transport mechanism for cystine uptake has been identified only in fibroblasts
and hepatocytes but not in renal proximal tubular epithelial cells.
AOAA and AT-125 administered alone also increase hepatic and renal
GSH, though to a lesser extent than CEB in combination with these inhibitors.
AOAA and AT-125 may theoretically have an effect on amino acid transport or
influence GSH synthesis at the genetic level; however, the mechanism for the
GSH-enhancing effect of these compounds is undefined at this time.
The results of these investigations indicate that the reactive thiol, or a
metabolite of the thiol, is at least partially responsible for CEB-induced
nephrotoxicity. Despite the fact that the reactive thiol appears to be involved
in diminished GCS activity, this metabolite does not appear to influence renal
or hepatic GSH levels since pretreatment with AOAA did not prevent the
GSH-enhancing effect of CEB in the liver or the kidney.
CHAPTER 3
EFFECT OF CEB ON RENAL AND HEPATIC XENOBIOTIC
METABOLIZING ENZYMES IN FISCHER 344 RATS
Introduction
Humans and other animal species are constantly exposed to a myriad of
potentially toxic, lipophilic foreign compounds, or xenobiotics, readily
absorbed through the skin, mucus membranes, gastrointesinal tract, and across
pulmonary tissues. Though some xenobiotics may be excreted unchanged,
most compounds are converted to more polar metabolites for excretion in the
urine, bile, saliva, and expired air (Mulder, 1990). This process of
biotransformation is mediated by various cytosolic and membrane-bound
enzyme systems (Sipes and Gandolfi, 1986).
Biotransforming enzymes are grouped into two general categories
depending on their catalytic specificity (Sipes and Gandolfi, 1986). Phase I
enzymes typically catalyze oxidation, reduction, and hydrolysis reactions and
include the cytochrome P450 and epoxide hydrolase enzyme systems. Phase II
enzymes, on the other hand, involve conjugation or biosynthetic reactions
mediated by enzyme families such as the glutathione S-transferases.
The phase I and II xenobiotic metabolizing enzyme systems are
responsible for the activation of parent compounds and/or detoxication of
reactive metabolites (Aitio, 1978). Both synthetic and naturally occurring
carcinogens are typically inert and require metabolic activation to exert their
deleterious effects (Gonzalez and Gelboin, 1994). The cytochrome P450s
(P450s) are the principal enzymes involved in the metabolic activation
(Gonzalez and Gelboin, 1994) of xenobiotics while glutathione S-transferases
(GST; Ketterer and Mulder, 1990) and epoxide hydrolases (EH; Guenther,
1990) are more frequently involved in the detoxication of reactive P450-derived
125
126
metabolites or parent compounds. EHs (EC 3.3.2.3) represent a group of
catalytically-related but distinct enzymes which catalyze the hydration of arene
oxides and aliphatic epoxides to the corresponding dihydrodiols (Sipes and
Gandolfi, 1986). EHs are believed to have evolved as detoxifying enzymes
which serve to protect cells from the deleterious effects of epoxides or oxiranes
formed by cytochrome P450s or lipid peroxidation (Guenther, 1990). Similar to
EHs, GSTs (EC 2.5.1.18) are an extensive family of multifunctional isozymes
and usually detoxify xenobiotics or reactive metabolites through glutathione
conjugation and subsequent metabolism to more polar compounds (Ketterer
and Mulder, 1990).
The balance between detoxication and bioactivation of a compound
within a particular species or organ depends on the relative amounts and
activities of the different forms of xenobiotic metabolizing enzymes (Halpert et
al., 1994). Inducers of metabolizing enzymes such as GST and EH have been
shown to inhibit a wide range of potential carcinogenic compounds. Rat and
human GSTs inhibit covalent DNA binding of food-borne carcinogens (Lin et
al., 1994) while the dietary antioxidants, 2(3)-ferf-buty 1-4-hydroxyanisole and
l,2-dihydro-6-ethoxy-2,2,4-trimethylquinoline, enhance hepatic GST activities
in mice and decrease the level of mutagenic metabolites formed from
benzo(a)pyrene (Benson etal., 1978). Similarly, the genotoxicity of
chloromethyloxirane in vitro is reduced in the presence of increased mEH
activity (Rossi et al., 1983) while the addition of purified mEH reduces the
mutagenicity (Glatt et al., 1983) and DNA binding of K-region epoxides of
polycyclic aromatic hydrocarbons (Guenther and Oesch, 1981; Oesch and
Guenther, 1983) in vitro.
127
Cruciferous vegetables including cabbage, cauliflower, broccoli, and
turnips have become widely regarded as potential cancer chemoprotectives.
Based on extensive experimental testing of crude vegetable extracts and
epidemiologic data (Olsen et al., 1991; Kune et al., 1992; LeMarchand et al., 1989;
Swanson et al., 1992; Schiffman et al., 1988; Kunes et al., 1987; Hoff et al, 1988;
Chyou et al., 1990; Young et al., 1988), both the National Research Councils'
Committee on Diet, Nutrition, and Cancer (1982) and the American Cancer
Society (1984) have recommended increased consumption of cruciferous
vegetables to decrease human cancer incidence. Despite the well-established
inverse relationship between cruciferous vegetables and the incidence of
various types of cancer, the specific dietary constituents responsible for this
protective effect have not been thoroughly defined.
Various constituents of cruciferous vegetables have demonstrated
anticarcinogenic potential (Ishizaki et al., 1990; Michnovicz and Bradlow, 1990;
Smith etal., 1990; Smith etal., 1993;Kojima etal., 1994; Zhang and Talalay,
1994). Though the mechanisms of action of these chemoprotective agents are
poorly understood, most of these compounds appear to exert their
anticarcinogenic effect by inhibiting and/or inducing xenobiotic metabolizing
enzyme systems.
Animals fed cruciferous plants or their degradation products have
demonstrated elevations in GST activity in various tissues (Sparnins el al.,
1982a, b; Aspry and Bjeldanes, 1983; Chang and Bjeldanes, 1985; Salbe and
Bjeldanes 1985; Ansher et al., 1986; Kensler et al., 1987; Vos et al., 1988; Bogaards
et ah, 1990; Wortelboer et al., 1992; Zhang et al., 1992). Additionally, breakdown
products of crucifers such as indole-3-carbinol (I3C; Ferguson, 1994) and
isothiocyanates (Ishizaki et al., 1990; Smith et al., 1990; Guo et al., 1992; Smith et
128
al., 1993) have been shown to inhibit the activity of different forms of P450s
and, therefore, the activation of various carcinogenic agents. It is believed that
GST induction and the inhibition of P450s may at least partially explain the
anticarcinogenic effect of cruciferous vegetables.
Depending on the plant, the enzymatic activity within the plant, and the
conditions of hydrolysis, cruciferous vegetables may undergo autolytic
hydrolysis to yield various cyanoepithioalkanes including l-cyano-3,4epithiobutane (CEB; Luthy and Benn, 1980). CEB, the structure of which was
initially proposed by Kirk and MacDonald (1974), has demonstrated
teratogenic effects in pregnant Holtzman rats (Nishie and Daxembichler, 1980)
and nephrotoxic effects in male Fischer 344 (F344) rats (VanSteenhouse et al.,
1989). Administration of CEB to male F344 rats resulted in epithelial cell
necrosis and karyomegaly within the pars recta of the proximal convoluted
tubules. CEB treatment also caused an early phase of depletion of hepatic
glutathione (GSH) followed by a rebound elevation. Renal GSH levels,
however, were elevated at all time periods evaluated with no identifiable
phase of depletion.
GSH, a ubiquitous tripeptide (glu-cys-gly), conjugates and detoxifies a
variety of electrophilic, hydrophobic compounds (Sinsheimer, etal., 1987;
Mitchell et al., 1973). Elevations in renal GSH may prevent carcinogenesis in a
twofold manner. GSH may conjugate and detoxify electrophilic mutagens
an d /or carcinogens and scavenge free radicals associated with oxidative stress
and free radical-induced mutagenesis and tumorigenesis (Feigetal., 1994;
Satoh and Lindahl, 1994). The free radical-scavanging ability of GSH may also
prove beneficial for renal transplant recipients following ischemic injury and
129
during the reperfusion phase of renal transplantation (Paller et al., 1984; Ouriel
et al, 1985).
The present study was devised to further elucidate the in vivo
metabolism of CEB in Fischer 344 rats and demonstrate the role, if any, of
phase I and phase II metabolizing enzymes in the activation or
anticarcinogenic potential of CEB. Since exposure to crucifers occurs via the
oral route of administration, thereby exposing the liver to high doses of the
compound, and the kidney appears to be the target organ for CEB, hepatic and
renal enzymes were of primary interest for this investigation.
Although 25-30 distinct forms of P450s have been characterized in
humans and > 40 forms have been described in rats (Nelson et al., 1993), only
three gene families (CYP1, CYP2, and CYP3) are currently thought to be
responsible for the majority of xenobiotic metabolism (Wrighton and Stevens,
1992). As such, various substrates have been developed to measure the specific
activity of P450 enzymes belonging to these three gene families. In rats,
ethoxyresorufin O-deethylation was shown (Burke etal., 1985) to be cataylzed
by P450s induced by polycyclic aromatic hydrocarbons including 3methylcholanthrene (3MC). Antibody inhibition experiments indicated that
ethoxyresorufin O-deethylase (EROD) was catalyzed mainly by P450 1A l and,
to a lesser extent, 1A2 (Doostdar et al., 1993). In control rats EROD activity was
shown to be highest in the liver (de Waziers et al., 1990) while lower activity
(25% of liver) was evident in the duodenum. EROD activity in the rat was
barely detectable in the kidney and undetectable in the jejunum, ileum, and
colon. Burke and coworkers (1985) also demonstrated pentoxyresorufin Odepentylase (PROD)-specific activity for the phenobarbitol-inducible P450
enzymes, 2B1 and 2B2. PROD activity was greatest in the duodenum (125% of
130
liver) and lung (120% of liver) in untreated rats (de Waziers et al, 1990) and
appeared to be the main isoenzyme in the small intestine. Weak PROD activity
was also present in the kidney, although P450 2B1/2B2 was not detectable by
Western blot.
7-Ethoxycoumarin has also been utilized as a measure of P450dependent O-dealkylation in rats. 7-Ethoxycoumarin O-deethylase (ECOD)
activity is induced by both phenobarbitol and 3MC and, as such, is a measure
of both P450 1A l /1 A2 and 2B1 / 2B2.
Renal and hepatic EROD, PROD, and ECOD activities were determined
to assess the effect of CEB on the induction, or inhibition, of these P450
enzymes. Although CEB is conjugated with GSH in vivo, that does not
preclude the involvement of phase I enzymes in CEB metabolism.
Microsomal EH activity was additionally measured in the liver and
kidney to determine involvement of this enzyme system in CEB metabolism in
vivo. CEB possesses an electrophilic epithiol moiety similar to an epoxide ring
and may theoretically act as a substrate for mEH. Cytosolic EH was not
measured since cEH activity is extremely low in the rat as compared to other
species (Gill and Hammock, 1980; Ota and Hammock, 1980).
y-Glutamylcysteine synthetase (GCS), the rate-limiting enzyme in GSH
synthesis, was also measured since previous investigations revealed no
induction of activity at 24 or 48 hours despite enhanced GSH levels. Samples
for this study were collected at earlier time points (4 and 12 hours) as
compared to the previous investigation, so that an early induction in GCS
activity, if present, could be detected. GST activity was also measured to assess
the possibility of induction of this enzyme in the presence of high levels of
GSH.
131
Materials and Methods
Animal Maintenance and Treatment
A total of 6 8 male,
6 -8
week old, Fischer 344 rats were obtained from
Harlan Sprague Dawley, Inc. (Indianapolis, IN, USA). Rats were housed in an
AALAC (American Association for the Accreditation for Laboratory Animal
Care)-accredited facility. Rats were initially housed in groups of 3 rats per cage
under controlled conditions of 12-hour light/dark cycle at 21 °C and 50-70%
relative humidity. For one week prior to treatment rats were fed a modified,
semi-purified diet, AIN-76A, with cornstarch substituted for sucrose to prevent
hepatic lipidosis (Hamm, etal, 1982). Throughout experimental trials rats were
housed individually in metabolic cages to facilitate estimation of feed and
water intake. Water was allowed ad lib. Pair-feeding was instituted to
abrogate the effects of dietary intake on xenobiotic metabolizing enzyme
systems.
At 8-9 weeks of age rats were divided into two treatment groups in a
random fashion. One group was gavaged with 125 mg CEB/ kg body weight
while the control group was gavaged with deionized, distilled water at the
same relative volume with respect to body weight. All gavaging was
performed between
8
and 9 AM. Each treatment group was then subdivided
into one of five groups (minimum of 6 rats/group) for evaluation at 4,12, 24,
36, and 48 hours.
At 4,12, 24, 36, or 48 hours following CEB administration, rats were
anaesthetized under isoflurane inhalation and euthanized via cardiac
exsanguination. Liver and kidney samples were collected and prepared for
evaluation of P-450 levels as well as determination of glutathione S-transferase,
y-glutamylcysteine synthetase, ethoxyresorufin O-deethylase, pentoxyresorufin
132
O-depentylase, ethoxycoumarin O-deethylase, and epoxide hydrolase
activities.
In accordance with the Animal Welfare Act, the experimental protocol
was submitted to and approved by the Louisiana State University Institutional
Animal Care and Use Committee.
Materials
CEB was graciously synthesized by Dr. Mark McLaughlin and Tamara
Schaller from the Department of Chemistry at Louisiana State University. A
modified method of Luthy and Benn (1980) was utilized to synthesize the CEB.
The bromo group of 4-bromo-l-butene (Janssen Chimica, N ew Brunswick, NJ,
USA) was replaced with a cyano moeity from sodium cyanide (Aldrich,
Milwaukee, WI, USA) to yield l-cyano-3,4-butene. l-Cyano-3,4-butene was
then oxidized at the unsaturated double bond with magnesium monoperoxyphthalate hexahydrate (Aldrich, Milwaukee, WI, USA) yielding l-cyano-3,4epoxybutane which was subsequently converted to a thiouronium salt in the
presence of thiourea (Aldrich, Milwaukee, WI, USA). Final conversion to 1cyano-3,4-epithiobutane was achieved in an alkaline environment, pH 9.0. The
identity of the product was confirmed by nuclear magnetic resonance (NMR)
and the purity of l-cyano-3,4-epithiobutane was demonstrated to be 98% or
greater by gas chromatography (see Appendix).
p-Nitrostyrene oxide and p-nitrostyrene diol were kindly synthesized by
Dr. David Swenson of the Department of Physiology, Pharmacology, and
Toxicology, School of Veterinary Medicine, Loiusiana State University by the
method of Westkaemper and Hanzlik (1980).
L-glutamate, L-a-aminobutyrate, adenosine triphosphate, disodium salt
(Na 2 ATP), bovine serum albumin (BSA), reduced glutathione (GSH), reduced
133
P-nicotinamide adenine dinucleotide phosphate (NADPH),
tris(hydroxymethyl) aminomethane hydrochloride (Tris-HCl), glycerol,
glutathione S-transferase (GST; EC 2.5.1.18), l-chloro-2,4-dinitrobenzene
(CDNB), potassium phosphate dibasic and potassium phosphate monobasic,
ethoxyresorufin, pentoxyresorufin, resorufin, 7-ethoxycoumarin, 7hydroxycoumarin, dimethyl sulfoxide (DMSO), p-nitrobenzyl alcohol, and a
diagnostic kit for determination of inorganic phosphorus (Procedure no. 670)
were obtained from Sigma Chemical Co. (St. Louis, MO, USA). Procedure
23225X-BCA Protein Assay Reagent method for the determination of total
protein was obtained from Pierce (Rockford,IL, USA). Ethylene diamineacetic
acid, disodium salt (EDTA) was obtained from Amresco (Solon, OH, USA)
while methanol and magnesium chloride (MgCl2 )were obtained from
Mallinckrodt, Inc. (Paris, KY, USA). Trichloroacetic acid (TCA) was acquired
from Curtin Matheson Scientific, Inc. (Houston, TX, USA.) Sucrose was
obtained from Fisher Scientific, Inc. (Fair Lawn, NJ, USA) while potassium
chloride (KC1), dithionite, and carbon monoxide (CO) were obtained from
Aldrich Chemical Company, Inc. (Milwaukee, WI, USA). The dietary
components were obtained from ICN Nutritional Biochemical (Costa Mesa,
CA, USA).
Samples were prepared using a Tissumizer® rotor-sator generator from
Tekmar (Cinncinnati, OH, USA) for homogenizing, a Beckman Model TJ- 6
centrifuge from Beckman Instruments, Inc. (Fullerton, CA, USA), a Sorvall RC58 Refrigerated Superspeed and a Sorvall OTD-65 Ultra Centrifuge - Oil
Turbine Drive centrifuge obtained from DuPont, Clinical and Instrument
Systems Division (Wilmington, DE, USA). Enzyme activities and P450 levels
were analyzed with a Hitachi F-2000 Fluorescence Spectrophotometer and a
134
Hitachi U-2000 double-beam UV/Vis spectrophotometer from Hitachi
Instruments, Inc. (San Jose, CA, USA).
High performance liquid chromatography analysis was performed
using a Waters 6000 pump, U 6 K injector, and a Cs reverse phase, 4 j/m,
8
mm
cartridge column from Waters Chromatography (Milford, MA, USA) equipped
with a Perkin-Elmer LC75 Spectrophotometric Detector (Thurnhill, Ontario,
Canada).
Tissue Preparation
Liver and kidney samples were collected from the median liver lobe and
cortex of the left kidney for determination of GST and GCS activities. The
remainder of the liver and kidney was utilized for microsomal preparations.
A 150-200 mg sample of liver or kidney was placed in 2 mis of 0.1 M
Tris-HCl buffer (pH 8.2). Tissues were homogenized for 20-30 seconds.
Samples were centrifuged at 1800 rpm for 20 minutes at 4°C and the fresh
supernatant assayed for GCS activity.
For evaluation of GST activity, a 150-200 mg sample of liver or kidney
was placed in 3 mis of 0.25 M sucrose/ 0.02 M EDTA buffer and homogenized
for 20-30 seconds. Samples were then centrifuged at 1800 rpm for 15 minutes
and the resultant supernatant frozen at -20°C.
Renal and hepatic microsomes were prepared on ice. Tissues were
rinsed in 125 M KCl, blotted, weighed and diced. Diced tissues were rinsed
three times in 0.25 M sucrose/1.0 M EDTA/10.0 M Tris-HCl buffer. Liver and
kidney samples were transferred to 30 mis and 20 mis of 0.25 M sucrose/1.0 M
E D T A /10.0 M Tris-HCl buffer, respectively. Six individual kidneys from the
same group were batched to enhance detection of enzyme activities. The
135
tissues were homogenized for 20-30 seconds and placed on ice until
centrifugation.
Liver and kidney samples were centrifuged at 1800 rpm at 4°C for 20
minutes. The supernatant was collected and consecutively centrifuged at 9,200
rpm and 11,000 rpm at 4°C for 20 minutes. The collected supernatant was then
ultracentrifuged at 30,000 rpm at 4°C for 60 minutes. The pellet was
resuspended in cold 125 M KCl in 20% glycerol, aliquoted, and frozen at -70°C.
Biochemical Assays
GST activity was determined on supernatants utilizing the method of
Habig et al. (1974). 50 //I of 0.04 M l-chloro-2,4-dinitrobenzene (CDNB) and 0.2
ml liver or kidney sample were added to 1.65 ml of 0.121 M potassium
phosphate buffer (pH 6.5). The reaction was initiated with 0.1 ml of 0.02 M
GSH and the change in absorbance recorded for three minutes at a wavelength
of 340 nm. GST activity was expressed in EU GST/ mg protein.
GCS activity was evaluated on fresh supernatant using a modified
method of Sekura and Meister (1977). Kidney and liver samples were diluted
1:20 and 1:5, respectively. A 200 //I sample of the supernatant was added to 200
//I of 10 mM glutamate, 100 //I of 10 mM L-a-aminobutyrate, 200 //I of 20 mM
MgCli, 100 fil of 2 mM N a 2 EDTA, 100 //I of 5 mM Na 2 ATP, and 20 fig BSA.
Samples were incubated at 37°C for 30 minutes and the reaction stopped with
1.0 ml of 20% TCA, Samples were allowed to stand at room temperature for 510 minutes prior to centrifugation at 1800 rpm for 10 minutes at 4°C. The
liberation of inorganic phosphorous was determined by the use of Sigma
Diagnostic Procedure No. 670. The absorbance of the samples was read at 660
nm. An appropriate sample blank was assayed to compensate for intrinsic
ATPase activity. GCS activity was expressed in //moles GCS/ hr/ mg protein.
136
All GST and GCS determinations were preformed on a Hitachi U-2000 double­
beam U V /V is spectrophotometer.
Renal and hepatic microsomes were evaluated for cytochrome P-450
levels as well as EROD, PROD, and ECOD activities using a modified method
of Lake (1987). A 2.0 mg sample of microsomal protein was brought to a total
volume of 2.0 mis with 0.1 M potassium phosphate/ 1.0m M EDTA buffer (pH
7.4). Dithionite (20 mg) was added to the samples which were subsequently
divided into two microcuvettes (1 ml per cuvette). A baseline was recorded
from 400 to 500 nm and the sample cuvette was bubbled with carbon monoxide
for 30 seconds. A wavelength scan from 400 to 500 nm was then recorded
against the reference cuvette. Cytochrome P-450 levels were expressed in nmol
P-450/ mg protein.
For evaluation of EROD, PROD, and ECOD activities, 2.0 nmoles
ethoxyresorufin, 10 nmoles pentoxyresorufin, and 0.5 pmoles 7-hydroxy coumarin were added, respectively, to aliquots of microsomal protein (1 . 0 mg
microsomal protein from the kidney and 0.5 mg microsomal protein from the
liver were assayed for EROD and ECOD activity while 1.5 mg microsomal
protein from the liver or kidney was analyzed for PROD activity). Samples
were brought to 980 fd total volume with 50 mM Tris-HCl buffer (pH 8.4 for
EROD and PROD and pH 7.8 for ECOD). Samples were incubated for 5
minutes at 37°C and the reaction initiated with 20 fil of 50 mM NADPH. The
change in intensity per minute was recorded at an excitation of 530 nm (EX5 3 0 )
and an emission of 580 nm (EM5 8 0 ) for EROD and PROD activities and at
EX5 3 0 and EM4 5 2 for ECOD activity. The activities of these enzymes were
expressed in p m ole/m in /m g protein.
137
All EROD, PROD, and ECOD determinations were performed on a
Hitachi F-2000 Fluorescence Spectrophotomete. Cytochrome P-450 levels were
evaluated on a Hitachi U-2000 double-beam U V /V is spectrophotometer.
Total protein levels were determined by the Pierce Procedure 23225XBCA Protein Assay Reagent method. The absorbance of the samples was
recorded at 562 nm. Total protein levels were expressed in mg protein/m l.
Microsomal samples were analyzed for epoxide hydrolase activity using
a modified method of WestKaemper and Hanzlik (1980). A 50 //I aliquot of
renal or hepatic microsomal protein was added to 550 //I of 0.1 M potassium
phosphate buffer (pH 8.0) and the mixture preincubated at 37°C for 2 minutes.
The reaction was initiated with the addition of 30 pi of 10 mM p-nitrostyrene
oxide and samples were incubated at 37 °C for 5 minutes. The reaction was
stopped with 1 ml of 3.45 x 10-5 M p-nitrostyrene alcohol (internal standard),
tubes were placed on ice, and samples centrifuged for 5 minutes. A 100 pi
aliquot of the clear supernatant was injected for analysis on a Waters 6000 high
performance liquid chromatograph equipped with a Waters U6 K injector and a
Perkin Elmer LC75 spectrophotometric detector operating at a 280 nm
wavelength and a sensitivity of 0.1 AUFs. Samples were analyzed on a Cg
reverse-phase, 4 pm column with a methanokwater (30:70) mobile phase
eluting at 2.0 m l/ min. p-Nitrostyrene diol was utilized as a standard. Peak
height was used as a measure of the quantity of product formed.
Statistical Analysis
Activities of GST, GCS, EROD, PROD, ECOD, and epoxide hydrolase
and cytochrome P-450 levels were evaluated at each time point by analysis of
variance with respect to treatment group. The means of each treatment group
138
were compared to one another by Fisher's least significant difference test at a
95% confidence level. P-values <0.05 were considered significant.
CEB-treated groups were compared to one another with respect to time
to assess the effect of time on xenobiotic metabolizing enzyme activities.
However, when control groups were compared to one another with respect to
time, there was a significant amount of interassay variation between different
time groups. Any significant differences between CEB-treated animals from
different time points was, therefore, not considered a valid estimate of the
effects of CEB.
Results
Renal and Hepatic GCS
Animals treated with a single dose of CEB exhibited significantly
decreased (p<0.01) renal GCS activity at 12 and 24 hours post-CEB
administration. Renal GCS activity at 4, 36 and 48 hours and hepatic GCS
activity at all time periods were not significantly different from control rats
(Figures 3.1 and 3.2).
Renal and Hepatic GST
Renal GST activity towards CDNB was significantly lower (p<0.01) in
rats treated with CEB as compared to controls at 4 hours post-CEB
administration. Similarly, hepatic GST activity was decreased (p<0.01) in CEBtreated rats at 12 and 36 hours. No significant differences in hepatic or renal
GST were observed in any of the other time periods evaluated (Figures 3.3 and
3.4).
Renal and Hepatic Cytochrome P-450, EROD, PROD, and ECOD
Animals treated with a single high dose of CEB exhibited no significant
difference in cytochrome P-450 levels in the liver or kidney at any of the time
139
□ C o n tro l
□ CEB
Time (Hours)
Figure 3.1
Comparison of renal GCS activity at 4,12,24, 36 and 48 hours in
CEB-treated and control rats. Value bars represent mean GCS activity. Error
bars represent the standard error of the mean; n = minimum of 6 rats per
group, t represents P < 0.01 as compared to controls.
140
1.2
T
□ C ontrol
E3CEB
4
12
24
36
48
Time (Hours)
Figure 3.2
Comparison of hepatic GCS activity at 4 ,1 2 ,2 4 ,3 6 and 48 hours
in CEB-treated and control rats. Value bars represent mean GCS activity.
Error bars represent the standard error of the mean; n = minimum of 6 rats per
group. N o significant differences were observed between control and CEB
treated groups at any time period evaluated.
141
0.30 -r
□ C ontrol
c
0.25
‘3
2
0.20
a.
SC E B
03
£
0.15--
h
§
0.10
“
0.05 4
0.00
12
24
36
48
Time (Hours)
Figure 3.3
Comparison of renal GST activity at 4 ,1 2 ,2 4 ,3 6 and 48 hours in
CEB-treated and control rats. Value bars represent mean GST activity. Error
bars represent the standard error of the mean; n = minimum of 6 rats per
group, t represents P < 0.01 as compared to controls.
142
□ C ontrol
HCEB
Time (Hours)
Figure 3.4
Comparison of hepatic GSTactivity at 4,12, 24,36 and 48 hours in
CEB-treated and control rats. Value bars represent mean GST activity. Error
bars represent the standard error of the mean; n =minimum of 6 rats per group,
t represents P < 0.01 as compared to controls.
143
periods studied. Similarly, renal and hepatic EROD and PROD activities
exhibited no significant differences as compared to controls.
Renal and hepatic ECOD activities, however, were significantly
decreased in rats administered CEB. Renal ECOD activity in CEB-treated
animals was decreased (p<0.01) at 24 hours while hepatic ECOD activity was
diminished (p<0.05) at 12 and 24 hours post-CEB administration (Figures 3.5
and 3.6).
Renal and Hepatic Epoxide Hydrolase
N o significant differences were observed at any time point in renal or
hepatic microsomal epoxide hydrolase activity.
Discussion
Decreased renal GCS activity at 12 and 24 hours is consistent with a
previous study in which various inhibitors of GSH transport and metabolism
were utilized to assess the nephrotoxic effects of CEB (see chapter 2).
Decreased levels of renal GCS at 12 and 24 hours in rats treated with CEB do
not appear to be the result of feedback inhibition by elevated GSH. In the
previous study, renal GSH levels were elevated at 48 hours as well as 24 hours.
Renal GCS levels recovered by 48 hours post-CEB administration despite
significant elevations in renal GSH.
Decreased levels of GCS may be a direct effect of the reactive thiol as
suggested by previous investigations. Diminished renal GCS activity was
observed in all treated groups with the exception of those rats pretreated with
aminooxyacetic acid (AOAA). AOAA competitively inhibits pyridoxal
phosphate-dependent enzymes including renal cysteine conjugate |3-lyase
(Elfarra et al., 1986). Renal cysteine conjugate (5-lyase catalyzes |5-elimination
144
□ C ontrol
Q CEB
4
12
24
36
48
Time (Hours)
Figure 3.5
Comparison of renal ECOD activity at 4,12,24,3 6 and 48 hours
in CEB-treated and control rats. Value bars represent mean ECOD activity.
Error bars represent the standard error of the mean; n = minimum of 6 rats per
group, t represents P < 0.01 as compared to controls.
145
450 T
□ C ontrol
HCEB
4
12
24
36
48
Time (Hours)
Figure 3.6
Comparison of hepatic ECOD activity at 4 ,1 2 ,2 4 ,3 6 and 48 hours
in CEB-treated and control rats. Value bars represent mean ECOD activity.
Error bars represent the standard error of the mean; n = minimum of 6 rats per
group, t represents P < 0.05 as compared to controls.
146
reactions to yield a potentially reactive thiol in addition to pyruvate and
ammonia.
Conversely, a metabolite of the reactive thiol, such as a methylthio or Sglucuronide derivative could inhibit GCS activity. The possibility that AOAA
pretreatment actually induced GCS activity is contradicted by the fact that
AOAA given alone did not induce enzyme activity.
Although it is of questionable biological significance, the slight but
statistically significant decrease in ECOD activity may represent an
anticarcinogenic effect associated with CEB. Inhibition of P450 enzymes has
been associated with decreased xenobiotic bioactivation. Since decreased
ECOD activity represents inhibition of P450 1A and/or 2B isoenzymes, either
EROD or PROD activity should also be affected. However, the constitutive
levels of PROD activity in untreated rats in this study as compared to ECOD
activity were extremely low. Although 7-ethoxycoumarin and
pentoxyresorufin are both substrates for P450 2B1/2B2, the difference in the
constitutive activity for the two substrates indicates a much higher activity of
P450 2B1 / 2B2 towards 7-ethoxycoumarin. Due to the low constitutive activity
of PROD, it is possible that inhibition of PROD activity could not be detected
as it was beyond the lower limits of sensitivity of the assay.
Hepatic and renal ECOD activity in the control rats were approximately
350 p m o le/m in /m g protein and 25 p m ole/m in/m g protein, respectively,
while hepatic and renal PROD activity were 5 pm ole/ m in/ mg protein and 1
p m o le/m in /m g protein, respectively. EROD activity was 30 p m ole/m in /m g
protein and 2 p m ole/m in /m g protein in the liver and kidney, respectively.
These levels were consistent with values found in the literature although
constitutive levels of hepatic ECOD and PROD in the current study were
147
slightly lower than other investigators' findings (Kleinow et al., 1990; de
Waziers et al., 1990). However, these investigators fed commercial diets which
have been shown to induce hepatic P450s (Deloria and Mannering, 1986) and
used a different strain of rat (Sprague-Dawley).
Inhibition of ECOD activity in the absence of inhibition of EROD
activity and no conclusive PROD data suggest that the P450s 2B1 or 2B2 may
be inhibited by CEB, or a metabolite thereof. Various anticarcinogenic agents
have demonstrated selective inhibition of the P450 1A and 2B subfamilies.
(-)-Epigallocatechin-3-gallate inhibited the catalytic activity of several P450
enzymes, but most potently inhibited P450 1A and 2B enzymes. Diminished
P450 1A and 2B activity thereby inhibited 4-(methylnitrosamino)-l-(3-pyridyl)1-butanone oxidation and DNA methylation (Shi etal., 1994). Similarly,
furafyllin has been shown to be a potent mechanism-based inhibitor of P450
1A2 activity in human liver microsomes (Kunze and Traner, 1993) while 1aminobenzotriazole inhibited hepatic PROD activitity to a greater extent than
EROD activity in untreated and phenobarbital-treated guinea pigs (Knickle
and Bend, 1992). EROD and PROD activities have additionally been shown to
be inhibited by a series of natural coumarins in vitro (Cai et al, 1993).
Alternative mechanisms for the inhibition of ECOD activity must also be
considered. According to a process referred to as the "steal phenomenon", it is
possible that ECOD activity is diminished through substrate-dependent
competition of different P450 enzymes for NADPH-cytochrome P450
reductase. Depending on the substrate present, one P450 enzyme can affect the
function of another P450 enzyme when combined in a reconstituted system.
This has been demonstrated in the presence of benzphetamine and
pentoxyresorufin, both preferred substrates for P450 2B4 as compared to 1A2,
148
in which P450 1A2 was shown to inhibit the reaction by competing with P450
2B4 for NADPH-cytochrome P450 reductase molecules (Cawley et al., 1995). It
is therefore possible that, in the presence of CEB, another P450 enzyme could
have interferred with P450 2B function and inhibited ECOD activity.
Alternatively, CEB, or a metabolite of CEB, could be directly inhibiting
ECOD activity through interaction at the active site. Such inhibition would not
necessarily affect PROD activity since the active site for ECOD and PROD are
obviously slightly different as they utilize distinct substrates. Theoretically,
ECOD, being catalyzed by both P4501A and 2B, is less selective than PROD
and could more readily be affected by CEB. It is also possible that ECOD is
more susceptible to nonspecific inhibition with contaminants although several
different aliquots of 7-hydroxycoumarin were obtained from the manufacturer.
Although renal and hepatic ECOD activity was diminished, there was
no significant decrease in total P450 levels at any time period evaluated. Total
P450 levels may not be as readily affected as ECOD activity since total P450 is a
less sensitive indicator of induction or inhibition of specific P450 enzymes.
Additionally, P450 levels are indicative of the amount of enzyme present but
do not necessarily correlate with actual enzyme activity.
The slight but statistically significant decrease in GST activity in the
kidney and liver was also of questionable biological significance especially in
light of the fact that previous investigations by VanSteenhouse et al. (1989)
demonstrated no effect on GST activity in the liver or kidney at similar time
points.
Inhibition of GST activity has not been well documented in vivo,
however, extensive lists of effective in vitro GST inhibitors have recently been
compiled (Mannervik and Danielson, 1988; Flatguard etal., 1993). This diverse
149
group of GST inhibitors includes glutathione analogs, drugs from a variety of
classes, metal compounds, endogenous substances, and naturally occurring
compounds such as plant phenols. Although remarkable specificities towards
certain isoenzymes have been observed, the distinction between substrates and
inhibitors has not been well defined. In fact, any substrate, by virtue of its
binding capacity, may act as a competitive inhibitor for other substrates.
Ethacrynic acid (Ahokas et ah, 1985), bromosulphophthalein (Andersson et al.,
1988), and even CDNB (Adams and Sikakana, 1990) are substrates for GSTcatalyzed GSH conjugation which inhibit enzyme activity at high doses.
High doses of CEB may conceivably diminish GST activity towards
CDNB by simply inhibiting CDNB binding at the H site (electrophilic substrate
binding site) in a competitive manner. Alternatively, the GSH conjugate of
CEB may directly inhibit GST as suggested by the fact that the GSH conjugate
of ethycrynic acid is a strong inhibitor of GST isoenzymes (Ploemen et al.,
1990). The CEB-GSH conjugate may act at the G site as a GSH analog or at the
H site as a "second substrate" analog.
Inhibitors of GST may also act in a noncompetitive, irreversible manner.
This type of inhibition has been demonstrated with ethylene dibromide
(Ivanetich etal., 1984), a compound known to be activated by GSH conjugation
(van Bladeren et al., 1980). It is believed that the highly reactive half-sulfur
mustard initially formed upon GSH conjugation binds to GST and inactivates
the enzyme. A similar scenerio following CEB conjugation with GSH might be
expected to cause immediate decreases in GST activity. Upon GSH
conjugation, the CEB-GSH conjugate may undergo cyclization to an
iminothiolane-Jike compound which may exert a cytotoxic effect through
150
interaction with lysine residues on cellular proteins. In this manner, GST may
be inactivated through conjugate-mediated thiolation.
Finally, lipid peroxidation may mediate GST inhibition in vivo. In rats,
both cytosolic and microsomal hepatic GST activities were inhibited during
episodes of lipid peroxidation (Beckman and Greene, 1988; Harris and Stone,
1988; Tampo and Yonaha, 1990; Kawashima et al., 1994). Additionally, Rao and
colleagues (1982) have found that, in rats, liver tumors induced by peroxisomeproliferating agents do not demonstrate increased levels of GST as seen with
genotoxic agents.
Though the diminished activities of renal and hepatic ECOD and GST
are of debatable biological significance, it is plausible that decreased ECOD
activity may be associated with the anticarcinogenic effect of cruciferous
vegetables. The significance of CEB-induced inhibition of GST is obscure.
Unfortunately, none of these mechanisms of inhibition explain the relatively
sporadic inhibition of renal and hepatic GST. GSH conjugation catalyzed by
GST enzymes generally results in detoxication of xenobiotics although
numerous compounds have been shown to actually be bioactivated by GSH
conjugation (Dekant and Vamvakas, 1993). As more compounds are shown to
be bioactivated by GSH conjugation, particularly those which target the
kidney, decreased GST activity may actually prove antinephrotoxic and
possibly anticarcinogenic.
The absence of induction of GST activity in this study does not preclude
the possibility of GSH conjugation with CEB. Several electrophilic xenobiotics
have been shown to undergo nonenzymatic conjugation with GSH (Ketterer et
al., 1986; Takahashi et al., 1987; Peterson and Geungerich, 1988). Alternatively,
it is plausible that a microsomal GST is responsible for catalyzing the
151
conjugation reaction since cytosolic GST was measured in this experiment.
Finally, the GST isozyme responsible for GSH conjugation with CEB may not
be specific for the substrate (CDNB) utilized in the in vitro assay although
CDNB is a nonspecific substrate with broad reactivity for many of the GSTs.
Obviously, future investigations regarding the effect of CEB on xenobiotic
metabolizing enzymes are necessary to elucidate the biological significance of
these alterations in enzyme activity.
CHAPTER 4
IDENTIFICATION OF A CEB-DERIVED URINARY
MERCAPTURIC ACID IN FISCHER 344 RATS
Introduction
l-Cyano-3,4-epithiobutane (CEB; Figure 4.1) is a naturally occurring
nitrile derived from glucosinolates found in cruciferous vegetables (Fenwick
and McDonald, 1974; Cole, 1975). Cruciferous plants are cultivated
extensively for use as livestock feed, in the manufacturing of plastics, and for
human consumption as condiments, vegetables, and oilseed (Fenwicketal.,
1983; Lenman, 1992).
Previous investigations by VanSteenhouse etal. (1989, 1991) have
demonstrated bioactivation of CEB by glutathione (GSH; Figure 4.2)
conjugation. Administration of CEB to male Fischer 344 rats demonstrated
epithelial cell necrosis and karyomegaly within the pars recta of the proximal
convoluted tubules. Treatment with CEB also caused an early depletion of
hepatic GSH followed by a rebound elevation.
Additionally, CEB-induced nephrotoxicity and karyomegaly were
greatly diminished following pretreatment with buthionine sulfoximine (BSO;
Figure 4.3), an inhibitor of y-glutamylcysteine synthetase (GCS) activity and
GSH-depleting agent (VanSteenhouse etal., 1991). The rapid depletion of
hepatic GSH and reduction of CEB-induced nephrotoxicity following BSO
pretreatment strongly suggest that glutathione (GSH) conjugation is a
significant pathway in CEB bioactivation and toxicity.
The kidneys, particularly proximal tubular epithelial cells, are frequent
targets for toxic xenobiotics. The kidneys are exposed to numerous
xenobiotics and xenobiotic-derived metabolites since they receive 25% of
cardiac output (Monks and Lau, 1987). Additionally, the proximal tubular
152
Figure 4.1
Structure of l-cyano-3,4-epithiobutane (CEB).
SH
O
*
C
-
nh3
o
I
II
CH— CH2- CH2- C
-O
ch2
I
N H - C H - C - N H - CH2
II
O
Figure 4.2
Structure of glutathione (GSH).
154
ch3
I
ch2
o
o
1
II
II
CH2- C H ,- S = N - P — O '
I
ch2
o-
ch2
I
HC— NH,+
I
'O—c=o
Figure 4.3
Structure of buthionine sulfoximine (BSO). BSO acts as an
irreversible inhibitor by binding to the active site of GCS in the
phosphorylated form.
155
epithelium possesses specialized functions of transport and metabolism which
may enhance toxicity through the concentration and bioactivation of
xenobiotics. Proximal tubular epithelial cells possess the highest
concentration of y-glutamyl transpeptidase (Goldberg e ta l, 1960), the only
known enzyme able to cleave the y-glutamyl linkage (Meister etal., 1981), and
are active in the uptake and concentration of metabolites of GSH conjugation.
GSH is a ubiquitious tripeptide present in high concentrations (mM) in
most living cells and participates in a variety of biological functions (Larsson
etal., 1983; Meister and Anderson, 1983). GSH conjugation is usually
associated with the detoxication and excretion of reactive electrophiles;
however, conjugation occasionally results in bioactivation of xenobiotics with
resultant toxic consequences (Monks and Lau, 1987; Monks etal., 1990;
Vamvakas and Anders, 1990; Koob and Dekant, 1991; Dekant and Vamvakas,
1993).
Compounds conjugated with GSH may be excreted in the urine as their
corresponding mercapturic acids which are N-acetylated derivatives of the
cysteine S-conjugates (Tate, 1980). N-acetylation is catalyzed by N acetyltransferase which utilizes acetyl Coenzyme A as the acyl donor. Several
compounds bioactivated by GSH conjugation are selective nephrotoxins and
excrete mercapturic acids as the predominant urinary metabolite (van
Bladeren etal., 1980; Jones and Well, 1981; van Bladeren et al., 1981a; Monks et
al., 1985,1988; Lau etal., 1988b; Kim and Guengerich, 1989).
Since CEB is bioactivated by GSH conjugation and selectively
targets the renal proximal tubular epithelium, it is of interest to establish
whether a CEB-derived mercapturate is excreted as the primary urinary
metabolite. Elucidation of the structure of the mercapturic acid by gas
156
chromotography-mass spectrometry (GC-MS) represents identification of a
unique compound and provides further evidence for the importance of GSH
conjugation in CEB metabolism (Nelson, 1992). Detection of other CEBassociated urinaiy metabolites is also important to the elucidation of in vivo
CEB metabolism in its entirety.
Measurement of urinary thioethers by thiol reagents such as 5,5’dithiobis-2 -nitrobenzoic acid represents a less specific and sensitive assay than
GC-MS but serves as a useful screening technique and as a method of
demonstrating GSH conjugation of various compounds (Chasseaud, 1988).
Administration of various inhibitors of GSH transport and metabolism
followed by determination of urinary thiols and thioethers (mercapturates)
would elucidate the effect of these inhibitors on CEB metabolism and possible
mechanisms of toxicity.
In the present study, Fischer 344 rats were administered a single dose of
CEB (125 m g /k g body weight) alone or following pretreatment with one of
three selective inhibitors of GSH transport and metabolism: acivicin (AT-125
or a-amino-3-chloro-4,5-dihydro-5-isoxazoleacetic acid), probenecid (p-di-npropylsulfamoyl benzoic acid), and aminooxyacetic acid (AOAA). AT-125 is a
glutamine analog and irreversible inhibitor of y- glutamyl transpeptidase
(GGT; Allen et al., 1980; Griffith and Meister, 1980).
Probenecid competitively inhibits the organic anion transport system
(OATS; Gutman etal., 1955) preventing secretion of the N-acetylated cysteine
conjugate or mercapturic acid from hepatocytes and subsequent renal uptake
at the basolateral membrane. Conversely, AOAA inhibits pyridoxal
phosphate-dependent enzymes including renal cysteine conjugate
157
P-lyase (Jones et al., 1986) and therefore the formation of a potentially reactive
thiol (Tateishi and Shimizu, 1980).
Materials and Methods
Animal Maintenance and Treatment
One hundred and thirty eight, 5-7 week old Fischer 344 rats were
obtained from Harlan Sprague Dawley, Inc. (Indianapolis, IN, USA). Rats
were housed in an AAALAC (American Association for Accreditation of
Laboratory Animal Care)-accredited facility. Rats were initially housed in
groups of 3-5 under controlled conditions of 12-hour light/dark cycle at 21 °C
and 50-70 % relative humidity. For one week prior to treatment rats were fed
a modified, semi-purified diet (AIN-76A) with cornstarch substituted for
sucrose to prevent hepatic lipidosis (Hamm etal., 1982). Throughout
experimental trials rats were housed individually in metabolic cages to
facilitate urine collection and estimation of feed and water intake. Water was
allowed ad lib. Pair-feeding was instituted to abrogate the effects of dietary
intake on the metabolism of CEB.
At 8-9 weeks of age rats were divided into one of eight treatment
groups in a random fashion. The eight groups consisted of a negative, saline
control group; a positive, CEB-treated control group; an AT-125 pretreated
group, a probenecid pretreated group, and an AOAA pretreated group; and
three treatment groups administered one of the three inhibitors alone. Each
group of rats was further divided into 2 groups (minimum of 6 rats / group)
for evaluation at 24 or 48 hours post-CEB administration.
An intraperitoneal (IP) injection of 10 mg AT-125/kg body weight was
administered two hours prior to CEB treatment in sterile saline, pH 6.5 while
200 mg probenecid/kg body weight was administered IP in 0.1 M sterile
158
potassium phosphate buffer (pH 7.4) 30 minutes prior to and 5.5 hours
following CEB administration. One hour prior to CEB administration, the
AOAA pretreated group received an IP injection of 32.8 mg A O A A /kg body
weight in 0.1 M sterile phosphate buffer, pH 7.0.
Treated rats were gavaged with 125 mg CEB/ kg body weight while the
negative control group received an equivalent volume of deionized, distilled
water. All gavaging was performed between
8
and 9 AM.
In accordance with the Animal Welfare Act, the experimental protocol
was submitted to and approved by the Louisiana State University Institutional
Animal Care and Use Committee.
Materials
CEB was graciously synthesized by Dr. Mark McLaughlin and Tamara
Schaller from the Department of Chemistry at Louisiana State University. A
modified method of Luthy and Benn (1980) was utilized to synthesize the
CEB. The bromo group of 4-bromo-l-butene (Janssen Chimica, N ew
Brunswick, NJ, USA) was replaced with a cyano moiety from sodium cyanide
(Aldrich, Milwaukee, WI, USA) to yield l-cyano-3,4-butene. l-Cyano-3,4butene was then oxidized at the unsaturated double bond with magnesium
monoperoxyphthalate hexahydrate (Aldrich, Milwaukee, WI, USA) yielding
l-cyano-3,4-epoxybutane which was subsequently converted to a thiouronium
salt in the presence of thiourea (Aldrich, Milwaukee, WI, USA). Final
conversion to l-cyano-3,4-epithiobutane was achieved in an alkaline
environment (pH 9.0). The identity of the product was confirmed by nuclear
magnetic resonance (NMR) and the purity of l-cyano-3,4-epithiobutane was
demonstrated to be 98% or greater by gas chromatography (see Appendix).
159
5,5'-dithiobis-(2-nitrobenzoic acid) (DTNB), potassium hydroxide, Nnitroso-N-methylurea, L-cysteine, aminooxyacetic acid, AT-125 (acivicin),
probenecid, tris(hydroxymethyl) aminomethane hydrochloride (Tris-HCl),
and a diagnostic kit for determination of creatinine (Procedure no. 555) were
obtained from Sigma Chemical Co. (St. Louis, MO, USA). Trichloroacetic acid
(TCA) was acquired from Curtin Matheson Scientific, Inc. (Houston, TX, USA)
while hydrogen chloride (HC1) was obtained from Aldrich Chemical
Company, Inc. (Milwaukee, WI, USA). Nitrogen gas for the evaporation of
samples was obtained from Lincoln Big Three, Inc. (Baton Rouge, LA, USA).
Methanol, sodium hydroxide (NaOH), ethyl acetate, and ethyl ether were
acquired from Mallinckrodt, Inc. (Paris, KY, USA). Ethylene diamineacetic
acid, disodium salt (EDTA) was obtained from Amresco (Solon, OH, USA).
Saline was from Baxter Healthcare Corp. (Deerfield, IL, USA). The dietary
components were obtained from ICN Nutritional Biochemical (Costa Mesa,
CA, USA).
Urine samples were prepared using a Beckman Model TJ- 6 centrifuge
from Beckman Instruments, Inc. (Fullerton, CA, USA) and two ml ChemElut
columns from Analytichem International (Harbor City, CA, USA). Urinary
metabolites were analyzed by electron impact ionization and chemial
ionization (methane) using a Hewlett-Packard 5890A Gas Chromatograph
equipped with a 5970 Series Mass Selective Detector (Palo Alto, CA, USA) and
a TSQ G C /M S/M S/D S from Finnigan MAT (San Jose, CA, USA),
respectively. A Hitachi U-2000 double-beam U V /V is spectrophotometer from
Hitachi Instruments, Inc. (San Jose, CA, USA) was utilized to analyze urinary
thiols and mercapturate levels.
160
Biochemical Assays
Total nonprotein (NP) urinary thiols and urinary thioethers (including
mercapturates) were analyzed according to a modification of the combined
methods of Seutter-Berlage et al. (1977) and Ellman (1959). Aliquots were
analyzed from samples obtained at 12, 24, and 48 hours. One ml of rat urine
was deproteinated with an equivalent volume of 10% TCA. The mixture was
allowed to sit at room temperature for 15 minutes and samples were
centrifuged at 1800 rpm for 15 minutes at 4°C. A 0.8 ml aliquot of the clear
supernatant was added to 0.2 ml of 5N NaOH or 2.0 mis of 0.4 M Tris-HCl
buffer (pH 8.9) for the determination of thioethers or total thiols, respectively.
50 nl of 0.01 M DTNB was added to samples for NP thiol evaluation. The
mixture was inverted and the absorbance promptly recorded at 412 nm.
Samples determined for thioether levels were bubbled with nitrogen
gas, sealed, and placed in a boiling water bath for 50 minutes. Once removed
from the water bath, tubes were placed on ice and 0.1 ml of 10 N HC1 was
added to stop the reaction. Two mis of 0.4 M Tris-HCl buffer (pH 8.9) and 50
//I of 0.01 M DTNB were sequentially added to each sample tube. The
absorbance was recorded at 412 nm within 10 minutes of adding the DTNB.
Absorbances for NP thiol and thioether levels were measured on a
Hitachi U-2000 double-beam U V/Vis spectrophotometer. Cysteine in 0.02M
EDTA was utilized as a standard.
The thiol concentration from the nonhydrolyzed sample was subtracted
from the thiol concentration measured in the hydrolyzed sample to calculate
the level of thioethers present. Urinary NP thiols and thioethers were
expressed in //M per /(M creatinine. Urine creatinine was measured using a
161
quantitative colorimetric alkaline picric acid method developed by Sigma
Diagnostics (Procedure no. 555).
Gas Chromatography-Mass Spectrometry
Sample Preparation: To identify urinary metabolites in samples obtained at 4,
12,24,36, and 48 hours, a 0.5 ml sample of rat urine was prepared according
to a modified method of Brocker et al. (1984). The samples were thawed and
adjusted to pH 2.0 with 1 N HC1 and 5 N NaOH. Adjusted samples were
adsorbed onto a 2.0 ml ChemElut (diatomaceous earth) column. Metabolites
were extracted from the column with 6 . 0 mis ethyl acetate which was
subsequently evaporated under nitrogen gas.
The remaining residue was derivatized with diazomethane.
Diazomethane was prepared by adding 20 mis ethyl ether to 20 mis 45%
potassium hydroxide. N-nitroso-N-methylurea (0.5 gm) was added and the
layers stirred until the ether (top) layer formed a bright yellow color. A 1-2 ml
aliquot of the ether layer was added to each sample residue until the solution
no longer cleared with standing. Samples were allowed to stand at room
temperature for 10-15 minutes. The ether was then evaporated under nitrogen
gas.
Gas Chromatography-Mass Spectrometry: GC-MS analysis (electron impact
ionization) was performed on a Hewlett-Packard 5890A Gas Chromatograph
equipped with a 5970 Series Mass Selective Detector and a 30-m DB5 capillary
column (i.D. 0.25 mm) dynamically coated with 5% phenylmethyl silicone
(0.25 //m film thickness) stationary phase. A 3 /d aliquot of sample residue
dissolved in 150 /d of ethyl acetate was injected via a Hewlett-Packard 7673A
automatic sampler into a split/splitless injection port set in the splitless mode.
Helium was utilized as the carrier gas at a flow rate of 1 m l/ min and a head
162
pressure of 15 psi. The temperature of the column was programmed from
50°C, holding for one minute, rising to 300°C at a rate of 30°C/min.
Temperatures of the injection port and detector were 250°C and 300 °C,
respectively. The source temperature was 150°C; pressure 5 x 10-5 torr;
electron energy 70 eV.
GC-MS analysis (chemical and electron impact ionization) was also
performed on a Finnigan TSQ G C/M S/M S/DS 4500. The same column and
detector conditions were applied as previously described. Methane was
utilized as the soft ionization gas at a pressure of
1
torr.
Statistical Analysis
Urine total thiols and thioethers were evaluated by analysis of variance
with respect to treatment group. The means of each treatment group were
compared to one another by Fisher's least significant difference test for
multiple comparsions at a 95% confidence level. P-values <0.02 were
considered significant.
Results
Urinary Total Nonprotein Thiols
Rats administered CEB in addition to any of the inhibitors exhibited
significant elevations in the excretion of urinaiy NP thiols as compared to
controls at 12 hours post-CEB administration (Figure 4.4). At 24 hours urinary
NP thiol levels remained sgnificantly elevated in rats pretreated with AT-125
but were not significantly increased in rats pretreated with AOAA or
probenecid or given CEB alone. By 48 hours urinary NP thiols were 10 times
greater in animals administered AT-125 in addition to CEB as compared to
controls; however, the difference between the two groups was not statistically
significant. Rats treated with CEB alone did not exhibit significantly elevated
163
1.2
Q)
C
'E 1.0 +
B 12 h o u rs
□ 24 h o u rs
148 h o u rs
CO
® 0.8 +
o
2 0.6 +
O
0 .4 +
jE
Jjj 0.2
^ 0.0
4C o n tro l
AOAA
P r o b e n e c id
A T -125
CEB
Treatment
Figure 4.4
Urinary nonprotein thiol excretion at 12,24, and 48 hours postCEB administration in rats pretreated with one of three inhibitors in addition
to CEB or CEB alone as compared to controls. Values represent means ±SEM;
n=minimum of 6 per group; + represents P = 0.0001 as compared to controls; ♦
represents P = 0.0001 as compared to CEB treated group.
164
urinary NP thiol levels although the mean was 10 times greater for the CEB
treated group as compared to controls. Rats treated with AT-125 in addition
to CEB also exhibited increased thiol levels at 12 and 24 hours as compared to
rats treated with CEB alone.
Rats given AT-125 alone demonstrated significantly elevated levels of
urinary NP thiols at 12 and 24 hours as compared to saline controls (Figure
4.5). No significant differences were observed in rats treated only with AOAA
or probenecid alone or at 48 hours in animals given only AT-125.
Urinary Thioethers
Rats administered a single dose of CEB alone or CEB in addition to any
of the inhibitors exhibited significant elevations in the excretion of urinary
thioethers as compared to controls at 12 and 24 hours (Figure 4.6). This
elevation, however, extended out to 48 hours only in AT-125 pretreated
animals. Additionally, rats pretreated with AT-125 exhibited significantly
elevated urinary thioethers as compared to CEB treated rats at 12 and 48
hours.
Animals given AT-125 alone demonstrated significantly elevated levels
of urinary thioethers at 12 and 24 hours (Figure 4.7). By 48 hours this group of
animals did not exhibit significantly different levels of thioethers as compared
to controls. N o significant differences were observed between controls and
animals treated with AOAA or probenecid alone.
Gas Chromatography-Mass Spectrometry
GC-MS analysis of urine samples revealed a single predominant
metabolite with a retention time of 10.61 ± 0.16 minutes which was unique to
the CEB-treated rats and absent in the urine of control animals (Figures 4.84.13). The metabolite was also present in the urine of rats pretreated with
165
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0 .3 0 $
S 12 hours
□ 24 hours
■ 48 hours
0 .2 5 +
0.20
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3L 0 .1 5 -■
0.10
-
0 .0 5 -
C o n tro l
AOAA
P r o b e n e c id
A T -125
Treatment
Figure 4.5
Urinary nonprotein thiols excreted at 12,24, and 48 hours posttreatment in rats administered one of the three inhibitors alone. Values
represent means ± SEM; n=minmum of 6 per group; t represents P = 0.0001 as
compared to controls.
166
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CEB
Treatment
Figure 4.6
Excretion of urinary thioethers at 12,24, and 48 hours post-CEB
administration in rats pretreated with one of three inhibitors in addition to
CEB or CEB alone. Values represent means ± SEM; n=minimum of 6 per
group; t represents P < 0.05 as compared to controls; ♦ represents P < 0.05 as
compared to CEB treated group.
167
0.35 -r
H 12 h o u rs
0.3 -0.25 --
□ 24 h o u rs
48 h o u rs
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- -
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P r o b e n e c id
A T -125
Treatment
Figure 4.7
Excretion of urinary thioethers at 12, 24, and 48 hours post-CEB
administration in rats treated with one of the three inhibitors alone. Values
represent means ± SEM; n= minimum of 6 per group, t represents P < 0.005 as
compared to controls.
168
File >JM405
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TIC
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Figure 4.8
Total ion chromatogram from a CEB control rat demonstrating a
unique peak at 10.64 minutes (*). Electron impact positive ionization using
Hewlett Packard instrumentation.
File >JM4Q1
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Figure 4.9
Total ion chromatogram from a saline control rat demonstrating
the absence of the unique peak at 10.64 minutes (*). Peaks eluting around
10.64 minutes in the control urine had dramatically different mass spectra as
compared to the mercapturate. Electron impact positive ionization using
Hewlett Packard instrumentation.
169
CEB CONTROL # 4 4
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240
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TIME
Figure 4.10 Electron impact ionization (Hewlett Packard) mass spectrum of
the unique peak at 10.64 minutes. This spectrum was consistently observed in
CEB treated and AOAA and AT-125 pretreated urine samples but was absent
from the negative control urines as well as the probenecid pretreated samples.
170
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266
268
270
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TIME
Figure 4.11 Selected ion tracing (extracted ion currents) of the electron
impact ionization (Hewlett Packard) chromatograph in a CEB control rat.
These ions are characteristic of the mercapturate which has a retention time of
10.64 minutes (*).
177 .0 0
C0Ntl?0L # 2 7 F 3 4 4 UR JSPtl
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Figure 4.12 A similar ion tracing of the electron impact ionization (Hewlett
Packard) chromatogram in a saline control rat demonstrates the absence of the
metabolite at 10.64 minutes (*).
127
100-1
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M/Z
301
300
314
328
343
356
350
475
405
400
450
Figure 4.13 Electron ionization mass spectrum from a CEB control sample
analyzed via Finnigan instrumentation. A similar ionization pattern is
observed for a unique peak at the same relative retention time as observed
with the Hewlett Packard GC-MS.
172
AT-125 and AOAA but not probenecid pretreated animals. The positive
electron impact ionization pattern of this metabolite was consistent with the
N-acetylated cysteine conjugate (mercapturic acid) of CEB or N-acetyl-S-(4cyano-2-thio-l-butyl)-cysteine (Figure 4.14).
Chemical ionization (Cl) with methane revealed a unique peak at the
same relative retention time as compared to the El chromatograms (4.15-4.16).
Additionally, a search of the Cl chromatogram for the 305 ion revealed a
single large peak at scan # 367 (Figure 4.17). The 305 ion represents the
calculated molecular weight of the mercapturate plus a positive charge (MW +
H +; 304+1). The mass spectrum of scan # 367 revealed M + 1 (305.2), M + 29
(333.2), and M + 41 (345.2) peaks consistent with a molecular weight of 304.2
and the masses for the methane conjugates of this mass, respectively (Figures
4.18-4.19).
In CEB-treated rats, the majority of the mercapturate was excreted by
12 hours post-CEB administration as demonstrated by relative quantification
(peak area) of the mercapturate at different time points (Figure 4.20).
Minimal levels of the mercapturate were excreted by 24 hours post-CEB
administration and essentially none was observed at 36 and 48 hours. Small
quantities were inconsistently observed in the urine at 36 and 48 hours. Four
of 6 rats and 2 of 6 rats revealed detectable levels of the mercapturate at 36 and
48 hours, respectively.
Pretreatment with both AT-125 and AOAA appeared to decrease the
relative amount of mercapturate excreted in the urine at 12 hours. A valid
interpretation of the effects of different treatments on mercapturic acid levels
could not be attained at 24, 36, or 48 hours since the relative quantities did not
exhibit large enough variations between experimental groups.
Figure 4.14 Proposed structure for the methylated CEB-associated urinary
metabolite. The fragment weights of various structural components consistent
with the electron impact ionization pattern are denoted by numbers in
parentheses. This structure is consistent with the methylated mercapturic acid
derived from CEB [N-acetyl-S-(4-cyano-2-thio-l-butyl)-cysteine].
174
366
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300
1 0 :0 0
350
1 1 :4 0
400
1 3 :2 0
445
468
450
1 5 :0 0
5 0 0 SCA N
1 6 :4 0 TIM E
Figure 4.15 Total ion chromatogram (methane chemical ionization) from a
CEB treated rat obtained using Finnigan instrumentation The unique peak at
scan # 368 occurs at the same relative retention time as the mercapturic acid
on the electron ionzation chromatogram.
175
1 0 0 .0
-,
270
r 2842620
234
222
255
374
302
162
195
332
119
351
361
RIC
305
414
100
3 :2 0
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250
8 :2 0
300
1 0 :0 0
350
1 1 :4 0
400
1 3 :2 0
461
450
1 5 :0 0
490
5 0 0 SCA N
1 6 :4 0 TIME
Figure 4.16 Total ion chromatogram (methane chemical ionization) from a
saline control rat obtained using Finnigan instrumentation. The peak
observed at scan # 368 in treated rats (figure 4.12) is absent from control
samples.
176
100
367
r 16 5 3 7 6
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164
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250
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6 :4 0
8:20
10:00
350
1 1 :4 0
400
1 3 :2 0
445 486
450
1 5 :0 0
5 0 0 SCA N
1 6 :4 0 TIME
Figure 4.17 A scan of the chemical ionization chromatogram for the 305 ion
(consistent with the molecular weight of the mercapturate + 1 ) reveals a
single, predominant metabolite at scan # 367. This peak occurs at the same
relative retention time as the peak for the mercapturate in the electron
ionization chromatogram.
ABU NDA NCE
305.091
± 0.5
5 0 .0 -
177
176
100
r 196352
144
305
263
114
128
190
160
5 0 .0 -
245
257
M/z 100
100
200
150
250
300
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lull11
M/Z
355
350
378 387 397
400
410 419
432
449 458
450
475
493 504
■
- K
500
516 527
■
i
540
■
550
Figure 4.18 Mass spectrum of scan # 367 from the chemical ionization
chromatogram.
A BU NDA NCE
% IN TEN SITY
273
178
305
r 146432
ABUNDA NCE
% IN TEN SITY
1 0 0 .0-1
5 0 .0 333
345
301
M /Z
300
311
310
314
319
320
325
328
331
330
339
340
343
351
350
Figure 4.19 A closer view of the mass spectrum of scan # 367 from the
chemical ionization chromatogram reveals the M + 1 (305), M + 29 (333), and
M + 41 (345) ions consistent with a molecular weight of 304.
179
RELATIVE QUANTIFICATION OF
URINARY MERCAPTURATE
2500000
— ■—
CEB
— • - “ AT-125
2000000
“
A - AOAA
1500000
1000000
500000
0
0
4
12
24
36
48
T im e (H o u r s)
Figure 4.20
Relative quantification of the mercapturic acid in urine
samples at 4,12,24,36, and 48 hours post-CEB administration. The
mercapturic acid was not observed in the saline control rats and animals
administered probenecid. The quantity of the metabolite is estimated by the
peak area from the electron ionization chromatogram.
180
An extensive search of the electron impact ionization chromatogram for
the 127/128 ion (associated with the CEB moiety of the mercapturic acid)
revealed the existence of other urinary mercapturates or similar compounds
related to CEB metabolism (Figure 4.21). After thorough and extensive
investigation of these peaks using pooled samples of urine to increase the
response and sensitivity of the analysis, however, no consistent, unique
metabolites could be demonstrated. These peaks were not consistently
evident in CEB control animals, were not unique to the CEB-treated groups, or
were present beyond the lower level of sensitivity of the analytical technique.
The electron impact ionization and methane chemical ionization
chromatograms were additionally traced for unconjugated CEB and
"anticipated" metabolites derived from CEB. No detectable levels of any of
these compounds were observed.
Discussion
Determination of urinary nonprotein (NP) thiols represents
measurement of GSH, cysteine, and cysteinylglycine, as well as GSH and
cysteine S-conjugates of CEB and the corresponding mercapturic acid.
Normally, GSH conjugates and their metabolites would not be considered
thiols; however, CEB, due to its epithiol moiety, forms a thiol upon GSH
conjugation. An elevation in urinary NP thiols at 12 hours in groups treated
with CEB in addition to an inhibitor and an absence of such an increase in rats
treated with CEB alone suggests that the inhibitors have a pronounced effect
on excretion of urinary thiols.
A profound increase in thiol excretion was observed in AT-125
pretreated rats or rats given AT-125 alone at 12 and 24 hours. AT-125 causes a
mild glutathionemia and pronounced glutathionuria in mice due to inhibition
181
1 2 8 .0 0
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.
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TIME
b)
128 .00
20000
0 - . . i . .A . ..
1 2 7 .0 0
1000001
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150
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UR JS PM
300
350
400
4000000
2000000
5 .0
6 .0
7 .0
8 .0
9 .0
TIME
Figure 4.21 a and b
Comparison of selected ion tracings of the total ion
chromatogram (electron impact ionization) from a) CEB-treated rat urine and
b) control rat urine for ions associated with the CEB moiety (127 and 128)
indicating possible CEB-derived metabolites.
182
of renal y-glutam yl transpeptidase (GGT) at the basolateral and luminal
membranes (Griffith and Meister, 1979), GSH and GSH conjugates are freely
filtered at the glomerulus with greater than 80% of circulating GSH being
cleared on a single pass through the kidney (Rankin etal., 1985). The majority
of GSH and GSH conjugates in the filtrate are normally absorbed as cysteine
or cysteine conjugates, respectively, following degradation by y-glutamyl
transpeptidase (GGT) and cysteinylglycine dipeptidase on the luminal
epithelium. Administration of AT-125 and inhibition of GGT prevents
degradation of GSH and GSH conjugates thereby increasing urinary loss of
these compounds.
AOAA and probenecid pretreatment also increase the excretion of
urinary NP thiols though to a lesser extent as compared to AT-125. AOAA
inhibits the conversion of cysteine conjugates to reactive thiols in addition to
pyruvate and ammonia (Jones et ah, 1986). The cysteine conjugate of CEB may
therefore be excreted into the urine or serve as a substrate for Nacetyltransferase and increase the formation and urinary excretion of the
corresponding mercapturic acid.
Probenecid inhibits the renal uptake of mercapturates at the basolateral
membrane and, though controversial, evidence suggests it also inhibits
transport of mercapturic acids from the renal epithelial cell into the filtrate
(Moller and Sheikh, 1983). If this latter point proves valid, inhibition of
secretion of mercapturates formed within the tubular epithelial cell may
increase deacetylation of the compound to the corresponding cysteine
conjugate. Thus, elevations in urinary thiols in probenecid pretreated rats
may be the result of increased excretion of the cysteine conjugate. Inhibition
of renal uptake of the mercapturate at the basolateral border may increase the
183
filtration of this compound through the glomerulus; however, probenecid also
inhibits secretion of mercapturates from the hepatic sinusoidal membrane
thereby decreasing transport from the liver to the kidney.
The fact that AOAA and probenecid when administered alone did not
increase excretion of urinary NP thiols supports the idea that the cysteine
conjugate of CEB and / or the corresponding mercapturate are responsible for
elevations observed in animals pretreated with these inhibitors. Formation of
large quantities of the cysteine conjugate and mercapturic acid following
administration of CEB and inhibition of GSH metabolism or transport by
AOAA and probenecid results in increased excretion of urinary thiols.
It also seems plausible that rats treated with CEB alone should also
exhibit elevated thiol levels simply due to the formation of cysteine
conjugates, GSH conjugates, and mercapturic acids. Though there was no
statistically significant difference between the CEB treated group and the
control group, there was a distinct trend towards elevated levels of urinary
thiols in CEB treated rats at 12 and 24 hours. The detection limit of this assay
may be limited by the naturally occurring background of thiols. A larger
sample size may actually be needed to detect statistically significant
differences between these groups.
Although the thioether assay is useful for assessing exposure to
electrophiles in biological monitoring studies and for demonstrating GSH
conjugation of experimental compounds, it should not be considered a true
quantification of urinary mercapturates as it was originally proposed (SeutterBerlage etaL, 1977). Urinary thioethers (R-S-R') include GSH conjugates,
cysteine conjugates and mercapturates as well as products of further
metabolism (S-methylation, glucuronidation, etc). Spectrophotometric
184
evaluation of urine samples for thioethers is therefore a nonspecific assay and
the chemical structures of the thioethers determined are generally unknown.
Relative levels of thioethers generally paralleled those of NP thiols.
Rats administered CEB alone exhibited elevated thioether levels at 12 and 24
hours presumably due to excretion of mercapturates as well as GSH and
cysteine conjugates. Administration of any of the inhibitors in combination
with CEB also elevated urinary thioethers at 12 and 24 hours due to excretion
of cysteine conjugates and mercapturates as previously described. Once
again, AT-125 caused a profound elevation in urinary metabolites at all time
points indicating inhibition of GGT and filtration of the GSH conjugate at the
glomerulus. Typically, compounds with a molecular weight < 68,000 may be
freely filtered at the glomerulus (Osborne and Stevens, 1981). The GSH
conjugate of CEB has a calculated molecular weight of 420.5 and therefore
would be expected to be readily filtered from the plasma at the glomerulus.
Elevations in urinary NP thiols and thioethers at 12 and 24 hours were
most pronounced in animals pretreated with the inhibitors indicating effective
inhibition at these time points. The inhibitory effects of AOAA and
probenecid appear to have diminished by 48 hours while the effect of AT-125
extended out to 48 hours.
Increased levels of thioethers in rats treated with CEB alone also
provides further evidence of the role of GSH conjugation in the metabolism of
CEB. The presence of urinary thioethers suggests excessive excretion of GSH
conjugates, cysteine conjugates, and the corresponding mercapturic acids.
GC-MS analysis of urine samples can provide a selective and sensitive
method for identifying urinary metabolites. A single, predominant
mercapturate was identified in the urine in the present study although
185
evidence for other CEB-derived metabolites was also demonstrated. Absolute
identification will require synthesis of the urinary mercapturate and
reexamination under the conditions descibed here.
Mercapturic acids are metabolic end-products of GSH conjugation.
Many mercapturates derived from GSH conjugation are predominantly
excreted in the urine as a consequence of the high activity of GGT on proximal
tubular epithelial cell membranes. High levels of GGT expression allow these
cells to cleave the y-glutamyl moiety of GSH conjugates and transport the
cysteine conjugate into the cell providing substrates for mercapturic acid
formation.
Excretion of urinary mercapturates has been demonstrated in rats,
rabbits, and primates (including humans) following exposure to a wide range
of electrophilic compounds (te Koppele et al., 1986; Zoetemelk etal., 1986;
Winter etal., 1987; Chasseaud, 1988; Polhuijs etal., 1989; Monks and Lau, 1990;
Polhuijs et al., 1991; Nelson, 1992; van Welie et al., 1992; Jensen et al., 1993).
Though conjugation with GSH generally detoxifies electrophiles, several
xenobiotics are actually bioactivated by GSH conjugation. 1,2-Dibromoethane
(DBE) is an extensively utilized industrial chemical and nematocide
bioactivated by GSH conjugation either directly or following oxidation. In
rats, the major urinary metabolite of DBE is N-acetyl-S-(2-hydroxyethyl)-Lcysteine (van Bladeren et al., 1980; van Bladeren et al., 1981b). GSH conjugates
of DBE also form DNA adducts in the liver and kidney which have been
identified as (2-[N-7-guanyl]ethyl)-glutathione (van Welie etal., 1992).
Interestingly, this adduct is also excreted in the urine as N-acetyl-S-2-[N-7guanyl]ethy!)-L-cysteine (Kim and Guengerich, 1989). Ethylene oxide, a
186
potential human carcinogen, similarly forms N-acetyl-S-(2-hydroxyethyl)-Lcysteine following GSH conjugation (Jones and Well, 1981).
Conjugation of benzoquinones with GSH results in the formation of
selective nephrotoxicants which target proximal tubular epithelial cells
(Monks et a l, 1985,1988; Lau et al., 1988b). 2-bromo-3-(N-acetylcystein-S-yl)hydroquinone was excreted as a major urinary metabolite and the tissue
selectivity is a consequence of high GGT activity on renal proximal tubular
epithelial cells (Monks and Lau, 1990).
The selective nephrotoxicity of CEB also appears to be the result of
high renal epithelial cell GGT activity. Though it is not clear whether the
mercapturate is involved in the toxicity, identification of this metabolite in the
urine provides evidence for the importance of GSH conjugation in CEB
metabolism and represents preliminary identification of a unique compound.
Based on mass spectral data obtained from this study, the
mercapturic acid derived from CEB may be chemically sythesized and
analyzed for a more definitive identification of the metabolite. GC-MS
analysis of the synthetic compound should reveal an identical mass spectrum
to the mercapturate identified in CEB-treated rat urine.
CHAPTER 5
GENERAL CONCLUSIONS
l-Cyano-3,4-epithiobutane (CEB) is a naturally occurring nitrile derived
from cruciferous plants that is bioactivated by glutathione (GSH) conjugation
in Fischer 344 rats. A single oral exposure to CEB results in epithelial cell
necrosis and karyomegaly in the pars recta of the renal proximal tubules in
Fischer 344 rats (VanSteenhouse, et al, 1989). Administration of CEB to rats
also caused a sustained elevation in renal GSH which may be associated with
the anticarcinogenic effect characteristic of cruciferous vegetables.
The results of these investigations indicate that the reactive thiol, or a
metabolite of the thiol, is at least partially responsible for CEB-induced
nephrotoxicity. Aminooxyacetic acid (AOAA) inhibits renal cysteine conjugate
p-lyase and exerts a protective effect, though short-lived, against CEB-induced
epithelial cell necrosis and karyomegaly within the renal proximal tubules.
The reactive thiol also appears to be involved in diminished v-glutamylcysteine synthetase (GCS) activity. Rats treated with CEB alone or pretreated
with AT-125 or probenecid exhibited significantly decreased renal GCS activity
at 24 hours as compared to controls. AOAA-pretreated rats, however, did not
demonstrate diminished renal GCS activity at 24 hours. Since AOAA inhibits
the formation of the reactive thiol, these findings suggest that the thiol may be
involved in decreased GCS activity.
The reactive thiol may inhibit GCS activity through reduction of
disulfide bridges or amino acid residues within the active site which are
essential for GCS activity. Conversely, a metabolite of the reactive thiol, such
as a methylthio or S-glucuronide derivative, could inhibit GCS activity. The
187
188
possibility that AOAA pretreatment actually induced GCS activity is
contradicted by the fact that AOAA given alone did not induce enzyme
activity.
Decreased levels of renal GCS at 12 and 24 hours do not appear to be the
result of feedback inhibition by elevated GSH since renal GSH levels were
elevated at 48 hours as well as 24 hours. Renal GCS levels recovered by 48
hours post-CEB administration despite significant elevations in renal GSH.
The mechanism of karyomegaly remains elusive at this time.
Karyomegaly may represent a preneoplastic change or a compensatory
regenerative response to renal tubular epithelial cell necrosis. It should be
noted, however, that karyomegaly was observed with low doses of CEB in the
absence of cell death (VanSteenhouse et al., 1989). Increased nuclear size may
be the result of elevated GSH levels or may be a direct effect of CEB or the GSH
conjugate of CEB.
Renal and hepatic GSH levels were significantly elevated at 24 and 48
hours in rats administered CEB alone or following administration of CEB in
addition to any of the inhibitors with the exception of hepatic GSH at 24 hours
following pretreatment with probenecid. Possible mechanisms for elevations
in renal and hepatic GSH include increased cellular uptake of cysteine or
cystine since cysteine is the limiting substrate for GSH synthesis (Deneke and
Fanburg, 1989) and enhanced transcriptional activation or stabilization of GCS
mRNA.
Cyanohydroxybutene (CHB), a naturally occurring nitrile also derived
from cruciferous vegetables, elevates hepatic and pancreatic GSH but does not
cause significant elevations in hepatic or pancreatic GCS activity. Pancreatic
189
and hepatic cysteine equivalents and the hepatic GCS mRNA concentration,
however, were elevated in CHB-treated rats (Davis et al., 1993).
It appears that elevated GSH levels in the liver and the kidney may be
under the control of multiple regulatory mechanisms. The lack of enhanced
hepatic GSH in rats pretreated with probenecid actually supports the theory
that increased cystine (or cysteine) uptake is involved in elevated intracellular
levels of GSH. Since cystine is transported as a tripolar, of anionic, ion in
human fibroblasts (Bannai and Kitamura, 1980, 1981) and hepatocytes
(Makowske and Christensen, 1982), probenecid may inhibit the organic anionic
transport system and the uptake of cystine into hepatocytes. This transport
mechanism has not been described in renal proximal tubular epithelial cells
and probenecid did not prevent the CEB-induced increase in renal GSH.
Results from these studies also suggest that CEB exerts an inhibitory
effect on some xenobiotic metabolizing enzymes including ethoxycoumarin Odeethylase (ECOD) and glutathione S-transferase (GST). The slight but
statistically significant decrease in ECOD and GST activity may be of
questionable biological significance especially considering the fact that
previous investigations by VanSteenhouse etal. (1989) demonstrated no effect
on GST activity in the liver or kidney following administration of CEB at a
similar dose and time.
Inhibition of cytochrome P450 enzymes, such as ECOD, may be
associated with the anticarcinogenic effect characteristic of cruciferous
vegetables. Decreased ECOD activity represents inhibition of P450 1A and/ or
2B enzymes suggesting that either EROD or PROD activity should also be
affected. The constitutive levels of PROD activity in untreated rats as
compared to ECOD activity, however, were extremely low. It is therefore
190
possible that inhibition of PROD activity could not be detected as it was
beyond the lower limits of sensitivity of the assay.
Alternatively, inhibition of ECOD activity may be due to a process
referred to as the "steal phenomenon". Depending on the substrate present,
one cytochrome P450 enzyme can interfere with the function of another
cytochrome P450 enzyme by competing for NADPPI-cytochrome P450
reductase molecules (Cawley et al., 1995). It is therefore possible that another
P450 enzyme could have interfered with P450 2B function and inhibited ECOD
activity in the presence of CEB.
Finally, it is also possible that CEB, or a metabolite of CEB, could
directly inhibit ECOD activity through interaction at the active site. Such
inhibition would not necessarily affect PROD activity since the active site for
ECOD and PROD are obviously different since they utilize distinctly different
substrates.
The slight but statistically significant decrease in renal and hepatic GST
activity was observed at 4 hours and at 12 and 36 hours, respectively. Any
substrate, by virtue of its binding capacity, can inhibit GST activity towards
another substrate. Ethacrynic acid (Ahokas et al, 1985), bromosulphophthalein
(Andersson et al., 1988), and CDNB (Adams and Sikakana, 1990) are substrates
for GST-catalyzed GSH conjugation which inhibit enzyme activity at high
doses. High doses of CEB may diminish GST activity towards CDNB by
inhibiting CDNB binding at the H site (electrophilic substrate binding site) in a
competitive manner. It is plausible that lower doses of CEB could actually
enhance GST activity; however, previous investigations regarding CEB
metabolism in F344 rats by VanSteenhouse etal. (1989) revealed no alteration in
GST activity at lower doses (50 m g/k g body w eigh t) of CEB.
191
Alternative mechanisms for inhibition of GST activity include direct
inhibition of GST activity by CEB or the CEB-GSH conjugate, nonsubstrate
ligand binding resulting in a conformational change in the enzyme, and
inhibition of GST in the presence of lipid peroxidation. None of these
mechanisms of inhibition, however, readily explain the relatively sporadic
nature of the inhibition of GST in the liver and the kidney.
It should be noted that a lack of induction and decreased activity of GST
towards CDNB does not preclude the possibilty of GSH conjugation. GSH
conjugation may occur nonezymatically. Spontaneous reactions with GSH
have been demonstrated for various quinones, dihaloethanes, and acrylonitrile
(Ketterer etal., 1986;Takahashi etal., 1987; Peterson and Guengerich, 1988;
Buffinton et al., 1989). Alternatively, the GST isoenzyme responsible for
catalyzing GSH conjugation with CEB may be a microsomal GST. Induction of
a microsomal GST isoenzyme would not have been detected in these studies
since GST activity was measured only in the cytosolic fraction of the liver and
kidney. Finally, it is plausible that the GST isoenzyme involved in GSH
conjugation with CEB may not exhibit activity towards the substrate, CDNB,
utilized in the in vitro assay.
Analysis of urine samples for thiol and thioether levels demonstrated
elevated urinary nonprotein thiols and thioethers in rats treated with CEB; yet,
a more profound increase in metabolite excretion was observed in rats
pretreated with the inhibitors. Urinary thiols include GSH, cysteine, and
cysteinylglycine, as well as GSH conjugates, cysteine S-conjugates, and the
corresponding mercapturic acids. Urinary thioethers (R-S-R') include GSH
conjugates, cysteine conjugates and mercapturates as well as products of
further metabolism (S-methylation, glucuronidation, etc).
192
AT-125 increases urinary excretion of GSH and GSH conjugates by
virtue of its inhibitory effect on y-glutamyl transpeptidase (GGT) at the
basolateral and luminal membranes (Griffith and Meister, 1979). Inhibition of
GGT prevents degradation of GSH and GSH conjugates since the y-glutamyl
linkage can not be cleaved.
Aminooxyacetic acid (AOAA) inhibits the conversion of cysteine
conjugates to reactive thiols thereby allowing excretion of the cysteine
conjugate into the urine. The cysteine conjugate may alternatively be
converted to the corresponding mercapturic acid which is subsequently
excreted in the urine.
Probenecid inhibits the renal uptake of mercapturates at the basolateral
membrane and, though controversial, evidence suggests it also inhibits
transport of mercapturic acids from the renal epithelial cell into the filtrate
(Moller and Sheikh, 1983). Inhibition of secretion of mercapturates formed
within the tubular epithelial cell may increase deacetylation of the compound
to the corresponding cysteine conjugate. The cysteine conjugate may then be
secreted into the urine.
The fact that AOAA and probenecid when administered alone did not
increase excretion of urinary NP thiols and thioethers supports the concept that
the cysteine conjugate of CEB and/or the corresponding mercapturate are
responsible for elevations observed in animals pretreated with these inhibitors.
Increased levels of thioethers in rats treated with CEB alone provides
further evidence of the role of GSH conjugation in the metabolism of CEB. The
presence of urinary thioethers suggests excessive excretion of GSH conjugates,
cysteine conjugates, and the corresponding mercapturic acids.
193
Analysis of urine samples by gas chromatography-mass
spectrometry demonstrated the presence of a single predominant metabolite
although evidence for trace levels of other metabolites was also demonstrated.
The metabolite was identified as N-acetyl-S-(4-cyano-2-thio-l-butyl)-cysteine
according to electron impact and chemical ionization mass spectral data.
Identification of the mercapturate in the urine provides further evidence for the
importance of GSH conjugation in CEB metabolism and represents
identification of a unique compound.
Future research endeavors which may be considered include studies
evaluating chronic administration of CEB to Fischer 344 rats to assess potential
carcinogenic effects of the compound. These investigations could include
evaluation of karyomegalic changes induced by CEB. The possibility of
increased DNA synthesis may be assessed from Feulgen-stained renal tissues.
Alternatively, immunohistochemical staining for proliferating cell nuclear
antigen (PCNA) or bromodeoxyuridine (BrDU) would reveal the relative
number of cells in the S phase of DNA synthesis. The amount of staining in the
renal proximal tubules may actually be quantitated through
histomorphometric techniques.
The mechanism of elevated GSH levels in the liver and kidney could be
further investigated by measuring cellular uptake of cystine and cysteine.
Cysteine is the rate-limiting substrate for GSH synthesis. It would also be of
interest to measure GCS mRNA levels in the liver and kidney to assess
transcriptional activity following administration of CEB. CEB may induce the
production of the GCS protein and/or mRNA without inducing actual enzyme
activity.
194
The mechanism of GSH conjugation with CEB could be further
elucidated through in vitro investigations. Nonenzymatic conjugation may
occur between GSH and CEB.
The activity of GST isoenzymes could also be further evaluated by
measuring microsomal GST (cytsolic GST was measured in these
investigations). Specificity for the cytosolic or microsomal GST isozyme
involved in catalyzing the conjugation reaction could be increased by utilizing
CEB as the substrate in an in vitro high performance liqiud chromatography
assay for measuring GST activity. Additionally, it should be noted that a
compound which acts as a substrate for GST does not necessarily act as an
inducing agent.
The activities of cytochrome P450 1A and 2B enzymes also deserve
further attention. Measurement of mRNA levels of P450 1A and 2B enzymes
would determine the effect of CEB on transcriptional activation or stabilization
of the mRNA. Alterations in P450 mRNA levels may occur with or without
corresponding responses in enzyme activity.
The activity of GST and P450 enzymes at different concentrations of
CEB should be investigated since high doses of CEB may saturate these
enzymes and inhibit their activity. Many enzymes follow an inverted
hyperbolic curve of activity with respect to substrate concentrations so that low
concentrations may induce activity while high concentrations inhibit their
activity.
Since these investigations suggest that the reactive thiol is responsible
for the nephrotoxicity, isolation of protein-thiol adducts using radiolabelled
CEB and affinity chromatography could identify and isolate the toxic
metabolite. Soft nucleophiles, such as the CEB-derived thiol, readily form
195
protein adducts while hard nucleophiles react with hard electrophiles like
those found in DNA.
Finally, it is important to establish the nephrotoxic and the GSHenhancing effects of CEB in renal cell culture in rats. Once the effects of CEB
metabolism are established for in vitro studies in the rat, human cell lines may
be investigated to assess the validity of extrapolation from rats to humans.
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261
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APPENDIX
Confirmation of the Structure of l-Cyano-3,4-epithiobutane and
Demonstration of the Purity of the Synthesized Product by Nuclear
Magnetic Resonance (NMR) and Gas Chromatography (GC)
262
263
i-C Y A N O -3 . 4 - E P I T H K BUTANE
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FIGURE A.1
NMR (200 MHz) of the distilled product. NMR spectrum
is consistent with the structure of l-cyano-3,4-epithiobutane.
PPM
l-CYANO-3. 4-EPITHIOBUTANE
FIGURE A.2
Enlarged NMR (200 MHz) spectrum of the distilled 1cyano-3,4-epithiobutane. Spectrum depicts increased detail consistent with the
structure of l-cyano-3,4-epithiobutane.
265
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FIGURES A.3 a and b
GC traces demonstrating the purity of the
synthesized l-cyano-3,4-epithiobutane: a) GC trace of the reaction at
completion and b) GC trace of the distilled l-cyano-3,4-epithiobutane.
*X
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i:
VITA
Judith S. Prescott-Mathews was born in Derby, England on March 5,
1965 to Roger and Margaret Ann Prescott. Her family immigrated to the
United States in January, 1967. She graduated from Science Hill High School
in May, 1983 and became a naturalized citizen of the United States on April 8,
1986. Judith received a Bachelor of Science degree in animal science from the
University of Tennessee on June 12,1987. On December 22,1988, Judith
married George A. Mathews, Jr. in Oak Ridge, Tennessee. She graduated
from the School of Veterinary Medicine at Louisiana State University with a
Doctor of Veterinary Medicine in May, 1991. In July, 1991 Judith enrolled in
the Department of Veterinary Pathology as a graduate assistant and resident
in veterinary clinical pathology. She completed the Doctor of Philosophy
degree with a major in Veterinary Medical Sciences in August, 1995.
266
DOCTORAL EXAMINATION AND DISSERTATION REPORT
Candidate:
Major Field:
Judith S. Prescott-Mathews
V et er i n a r y M e d i c a l Sciences
Title of Dissertation:
Elucidation and Modul at ion of CEB Metabo lis m
in Fischer 344 Rats:
Possible Mechanisms of To xi ci ty and
An ti ca rc in oge ni cit y
Approved:
or Professor and Chairman
EXAMINING COMMITTEE:
Date of Examination:
June 28,
1995