Elucidation and Modulation of CEB Metabolism in Fischer 344 Rats: Possible Mechanisms of Toxicity and Anticarcinogenicity
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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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Recommended Citation 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. https://digitalcommons.lsu.edu/gradschool_disstheses/6045
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ELUCIDATION AND 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 me 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...... 1 Introduction ...... 1 Influence of Dietary Constituents on Carcinogenesis 2 Cruciferous Vegetables ...... 26 Glucosinolates ...... 31 Functions of Glutathione ...... 40 Glutathione Metabolism and Transport ...... 48 Nitriles ...... 73 Objectives ...... 83
CHAPTER 2 PROTECTIVE EFFECT OF AMINOOXYACETIC ACID ON CEB-INDUCED NEPHROTOXICITY IN FISCHER 344 RATS...... 87 Introduction ...... 87 Materials and Methods ...... 90 Animal Maintenance and Treatment ...... 90 Materials ...... 91 Tissue Preparation ...... 93 Biochemical Assays ...... 93 Histological and Histomorphometric Examination.... 94 Statistical Analysis ...... 95 Results...... 95 Renal and Hepatic GSH ...... 95 Renal and Hepatic GCS Activity ...... 98 Histological Evaluation ...... 101 Karyometry ...... 101 Discussion ...... 116
CHAPTER 3 EFFECT OF CEB ON RENAL AND HEPATIC XENOBIOTIC METABOLIZING ENZYMES IN FISCHER 344 RATS 125 Introduction ...... 125 Materials and Methods ...... 131 Animal Maintenance and Treatment ...... 131
v Materials ...... 132 Tissue Preparation ...... 134 Biochemical Assays ...... 135 Results ...... 138 Renal and Hepatic GCS ...... 138 Renal and Hepatic GST ...... 138 Renal and Hepatic Cytochrome P-450, EROD, PROD, andECOD ...... 138 Renal and Hepatic Epoxide Hydrolase ...... 143 Discussion ...... 143
CHAPTER 4 IDENTIFICATION OF A CEB-DERIVED URINARY MERCAPTURIC ACID IN FISCHER 344 RATS...... 152 Introduction ...... 152 Materials and Methods ...... 157 Animal Maintenance and Treatment ...... 157 Materials ...... 158 Biochemical Assays ...... 160 Gas Chromatography-Mass Spectrometry ...... 161 Sample Preparation ...... 161 Gas Chromatography-Mass Spectrometry 161 Statistical Analysis ...... 162 Results...... 162 Urinary Total Nonprotein Thiols ...... 162 Urinary Thioethers ...... 164 Gas Chromatography-Mass Spectrometry ...... 164 Discussion ...... 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 with the exception of AOAA pretreated rats at 24 hours. A predominant urinary metabolite of CEB was identified as the mercapturate, N-acetyl-S- (4- cyano-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-glutamyl- cysteine 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 probenecid- pretreated 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 compounds derived from cruciferous plants have demonstrated toxic effects when 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 well as the effect of CEB on xenobiotic metabolizing enzymes.
The anticarcinogenic potential of various dietary constituents, the influence of these compounds on xenobiotic metabolizing enzymes, and the effect of these enzymes on anticarcinogenicity will be reviewed.
Additionally, the functions, metabolism, and transport of GSH and the
GSH-enhancing effect and toxicity of cyanoepithioalkanes similar to CEB will be described in detail.
1 2
Influence of 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 consumption of anticarcinogens may directly or indirectly counteract the effects of carcinogens in the human diet (Ferguson, 1994). More than 500 chemical compounds 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 well- defined phenomenon referring to the induction of a permanent transmissible alteration in the genetic code (Cotran et al, 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 synonymous 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 vivo 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 non- genetic 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
(Negeshi 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 well 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 compounds 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 most important and, certainly, the most well-known 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 mixed 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 enzyme
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 molecule 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 al, 1990; Lephart et al, 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 and/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 enzymes 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 al, 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 enzymes following administration of phenobarbitol (PB) or 3- methylcholanthrene (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 enzyme 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 following 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 enzyme 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. DBA/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 simple 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 well 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 with 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 shown 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 shown to elevate
P450 activity (Kimura et al., 1986).
The mechanism of induction of the P450 2B subfamily as well 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 enzymes 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 within 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, when 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 well 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 was testis>liver>lung>skin>kidney>intestine (Oesch et al., 1977).
Similarly, cEH, also referred to as 'EH 2 ' (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 wide 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 shown 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 5,6- oxide hydrolase, 'chEH' (Guenther, 1990), 'mEHch' (Oesch, et ah, 1984), or 'mCE' (Finley and Hammock, 1988), catalyzes the hydration of steroid 5,6- oxides. 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 enzyme (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 al, 1987) and a lack of cross-reactivity between chEH and mEH-specific antibodes (Levin et al, 1983; Oesch et al, 1984). A unique cytosolic epoxide hydrolase involved in leukotriene biosynthesis has additionally been characterized (Evans et al, 1985; McGee and Fitzpatrick, 1985; Ohishi et al, 1987). Leukotriene A 4 (LTA4 ) hydrolase or LTA 4 H hydrolyzes the oxirane ring of LTA 4 to form LTB4. LTA 4 H is located exclusively in the cytosol and differs catalytically from both cEH and mEH in that the addition of water by LTA 4 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 LTA 4 and closely related analogs of LTA 4 act as substrates (Evans et al, 1985; McGee and Fitzpatrick, 1985; Ohishi et al, 1987). LTA 4 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, LTA 4 (McGee and Fitzpatrick, 1985; Ohishi et al, 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). LTA 4 H was also present in rodent
(Evans et al., 1985; Haeggstrom et al, 1985; Pace-Asciak et al, 1985; Medina et al, 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 molecules 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 enzymes 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 isoenzymes 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 enzymes 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 non- catalytic covalent binding (Jakoby, 1978; Listowsky et al., 1988). This has been proposed as a 'suicidal' defense mechanism since the enzyme molecule is often inactivated in the process (van Bladeren and van
Qmmen, 1991; Pabst et al., 1974; Corrigall et al., 1989; Adams 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, immunological 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 isoenzyme GST £ which is immunologically 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 isozymes, 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 isoenzymes 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 isoenzymes 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 endoplasmic reticulum and the outer mitochondrial membrane from both humans and rats (Kraus, 1975; Morgenstem and DePierre, 1988; Wahllander et al., 1979;
Morgenstern 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 enzyme 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 enzymes (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 y- glutamylcysteine (Sugimoto et al, 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 wide 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-2,4-
dinitrobenzene 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 al, 1982; Coles et al, 1988).
Reactions of GSH with 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-9,10-
oxide, 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 al, 1975;
Hernanandez et al, 1980; Plummer et al, 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 N- acetylbenzoquinone 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 isoenzymes which is characteristic for the specific tissue. The isoenzyme pattern of distribution as well as the isoenzyme 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-4,5- oxide 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 isoenzyme 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 mouse 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 al, 1993).
GST isoenzyme 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 well 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;
Ploemen 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 compounds 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 isoenzymes 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), 3- methylcholanthrene (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.
Numerous 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 etal, 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 DNA binding of food-borne carcinogens (Lin et al., 1994).
Similarly, the dietary antioxidants, 2(3)-ferf-butyl-4-hydroxyanisole and 1,2- 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 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 and/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 ( Amoracia rusticana) and the roots (radish, turnip), stems (kohl-rabi), flower buds
(cauliflower, broccoli), and leaves (cabbage, Brussels sprouts, Takana,
Nozawana, 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 development of cancer at various sites including the colon, rectum, bladder, lung, thyroid, pancreas, skin, stomach, and mesothelium (Graham et al., 1978; Graham et al, 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)anthracene- treated 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 - methylimidazo[4,5-f]quinoline, 2-amino-3,4-dimethylimidazo[4,5-f]- quinoline, 2-amino-3,8-dimethylimidazo[4,5-f]quinoxaline in the Ames 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 al, 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;
Wortelboer 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 shown 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 shown 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 al, 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 (indolylmethyl-
glucosinolate) 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 p-
D-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\ NOSO3
Figure 1 . 1 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 2-
propenyl 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;
Seimiya 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 al, 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 al, 1993; Sugie et al., 1993; Jiaro et al., 1994; Kojima et al., 1994; Stresser et al, 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 al, 1988; Nugon-Baudon et al, 1990; Smith et al., 1990; Rao et al, 1991; Steinmetz and
Potter, 1991; Baldwin and LeBlanc, 1992; Loft et al, 1992; Ferguson, 1994;
Stresser et al, 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 enzymes may diminish the formation of toxic metabolites thereby reducing DNA-adduct formation and carcinogenesis (Ishizaki et al, 1990;
Smith et al, 1990; Zhang and Talalay, 1994).
In tandem with the inhibition/ activation of phase I enzymes, 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 DNA 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, DNA 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 al, 1988).
Arachadonic acid, found within cellular membranes, is converted to 5- hydroperoxy-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, LTC 4, which is cleaved at its y-glutamyl linkage to yield LTD 4 . The cysteinyl-glycine linkage of LTD 4 is then cleaved to produce LTE 4 . 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 al, 1988; Hidaka et al, 1990; Buckley et al, 1991; Nejad et al, 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 al, 1988; Hidaka et al., 1990; Nejad et al., 1991). Increases in GSH appear to stimulate
the synthesis of PGE 2 relative to other prostaglandins in the urinary bladder
epithelium (Mimata et ah, 1988) and renal medullary homogenates (Nejad et al, 1991).
The effect of GSH on prostaglandin biosynthesis may be mediated
through GSH conjugation. Glutathione S-transferases have been shown to
reduce PGH2 to PGF 20U presumably due to GSH peroxidase activity (Burgess
et al, 1987) and may be involved in isomerization of PGH 2 to PGE 2 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-aminoazobenzene (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 S- transferase 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
(Swenson 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 S- transferases (Degen and Neumann, 1978; Emerole et al., 1979).
Glutathione Metabolism and Transport GSH possesses two structurally characteristic features: 1) the y-
glutamyl 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 enzyme 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)
y - GLUTAMYL AA y - GLUTAMYL CYSTEINE (Y-GLUTAMYL A CYCLO- CH2SH A D P+P: TRANSFERASE I 2 H2NCHCOOH (Y-GLUTAMYL RCHCOOH CYSTEINE CYSTEINE (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, cysteinyl- glycine 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 ATP- dependent 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 shown 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 monophosphate- dependent 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 al, 1988; Garcia-Ruiz et al., 1992) although Inoue et al. 52 (1984) characterized two kinetically distinct, Na+-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 widely 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 hepatoblastoma- derived 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 Na+-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, Na+-independent, saturable, carrier- mediated 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 C 4 . 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 al, 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 al, 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 well (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 Na+-K+ pump (Ozaki et al, 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 2 0 % 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 AT- 125, 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 compounds 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 Na+-dependent (alveolar type II epithelial cells and enterocytes) or Na+-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; Gopalan- Kriczky 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 y- glutamyltranspeptidase 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 widely 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 DNA 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-bromo- propyl)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~3- chloropropane (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 well as the cysteine conjugate (S-pentachlorobuta-1,3- dienylcysteine) and mercapturic acid [N-acetyl-(S-l,l,2,3,4- pentachloro- butadienyl)-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-(2- chloro-l,l,2-trifluoroethyl)-L-cysteine, is metabolized by 0-lyase to 2-chloro- 1,1,2-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 p- elimination 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 compounds 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 - g l y (Y - GLUTAMYL TRANSPEPTIDASE) glutamate- CYS - GLY I X (DIPEPTIDASE) glycini AOAA CY! (B - LYASE) (N - ACETYL - TRANSFERASE) PYRUVATE N - ACETYL - CYS I NH, X A REACTIVE THIOL (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; / / \ \ ' n SO —( ' '> —COOH 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 5 2 - 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 well 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 widely 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 PROBENECID GS-R NAC-CSY-R BILE PLASMA CYS-R GSH + R ENTEROHEPATIC CIRCULATION NAC-CSY-R NAC-CSY-R GS-R NAC-CSY-R CYS-R PROBENICID CYS-R EXCRETION EPITHELIUM INTESTINE BASO- KIDNEY LUMENAL LATERAL Figure 1.7 Diagram depicting the inhibitory effect of probenecid (p-di-n- propylsulfamoyl 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 COOH I * H2N— CH H2N - CH HC— 0N ^2 H r i 4 CH2 o I I Cl n h 2 AT-125 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 f RENAL PROXIMAL TUBULE y GLU-CYS-GLY R-SH MA GSH AT-125 \ / CY^-GLY- CYS-R GLY AA(CYS; YGLU-AA yGLU BASOLATERAL BRUSH-BORDER MEMBRANE 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 y- glutamyltranspeptidase 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 vivo (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-2- hydroxy-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 mg/kg 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 mg/kg 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 (GST) (GSH REDUCTASE) GLUTATHIONE amino acu (GSH) ADP+Pj (Y - GLUTAMYL (GSH TRANS - SYNTHETASE) CYSTEINYL-GLYCINE PEPTIDASE) ATP (DIPEPTIDASE) Y - GLUTAMYL AA y - GLUTAMYL CYSTEINE GLYCINE (Y-GLUTAMYL CYCLO» ,/ADP+Pi TRANSFERASE) 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-5- isoxazoleacetic 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 GSH- enhancing effect. Enhanced renal GSH may prove anticarcinogenic by conjugating and detoxifying other electrophiles which may be potential 85 mutagens and/or carcinogens. Additionally, elevated renal GSH levels may scavenge free radicals associated with oxidative stress and free radical- induced 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 end- products 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 metabolizing 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-2,4- dinitrobenzene (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 y- glutamylcysteine synthetase (GCS) and GSH-depleting agent (VanSteenhouse, etal, 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-5- isoxazoleacetic 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 cysteine- thioester 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 GSH- enhancing 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 AOAA/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, New 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-(2- nitrobenzoic 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 double- beam UV/Vis 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 Units/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 2ATP, 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 5- 10 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 GCS/hr/mg protein. Total protein levels were determined by the Pierce Procedure 23225X- BCA 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 UV/Vis 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 hours S 4 8 h o u rs Control AOAA Probenecid AT-125 CEB 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 ours □ 48 H ours Control AOAA Probenecid AT-125 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 hours H 48 h o u rs 10 - Q, CD E >_ 6 - JC w o 4 -- o o 2 - E 5. C ontrol AOAA Probenecid AT-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 hours H 48 h o u rs Control AOAA Probenecid AT-125 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. No 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 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 jum 2 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 //m2 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 ;«m 2 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 Control AOAA Probenecid AT-125 CEB 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 ♦- ■ • Control • — AOAA + CEB *—-CEB Nuclear Area (jim2) Figure 2.15 Frequency distribution of proximal tubule epithelial cell nuclear size with 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 - ■ ■ ■ - C ontrol - * — AOAA + CEB 0.4 -- A CEB or 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 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 /mi 2 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 etal, 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 etal, 1977), N-(DL-2-amino-2-carboxyethyl)-L- lysine (Reyniers etal., 1974), lead acetate (Choie and Richter, 1972), N-(4'- fluoro-4-biphenylyl)acetimide (Dees etal., 1980a, b), 2S-l-cyano-2-hydroxy-3,4- epithiobutane (Gould etal., 1980), aflatoxin Bi (Butler, 1964), trichloroethylene (Maltoni etal., 1988), cisplatin (Goudie etal., 1994), l-(2-chloroethyl-4- methylcyclohexyl)-l-nitrosourea (MeCCNU) and chlorozotocin (Kramer etal., 120 1986) and nephrotoxic mycotoxins such as ochratoxin A (Huff, 1991; Mantle et al, 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 al, 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 al, 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 Icy stine (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 etal, 1990). Additionally, cystine transport into human erythrocytes was increased in the presence of extracellular l-chloro-2,4- dinitrobenzene (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-glutamyl- cysteine. 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-3,4- 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 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 and/or carcinogens and scavenge free radicals associated with oxidative stress and free radical-induced mutagenesis and tumorigenesis (Feig etal., 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 3- methylcholanthrene (3MC). Antibody inhibition experiments indicated that ethoxyresorufin O-deethylase (EROD) was catalyzed mainly by P450 1 Al 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 O- depentylase (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 P450- dependent 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, New 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 monoperoxy- phthalate 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 1- 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). 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 2ATP), 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, 7- hydroxycoumarin, 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 (MgCl 2 )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 RC- 58 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 EDTA/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 Na 2 EDTA, 100 //I of 5 mM Na 2ATP, 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 5- 10 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 UV/Vis 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 (EX 5 3 0 ) and an emission of 580 nm (EM 5 8 0 ) for EROD and PROD activities and at EX5 3 0 and EM 4 5 2 for ECOD activity. The activities of these enzymes were expressed in pmole/min/mg 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 UV/Vis spectrophotometer. Total protein levels were determined by the Pierce Procedure 23225X- BCA Protein Assay Reagent method. The absorbance of the samples was recorded at 562 nm. Total protein levels were expressed in mg protein/ml. 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 U 6 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 ml/ 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 CEB- treated 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 □ Control □ 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 4,12,24,36 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. No significant differences were observed between control and CEB treated groups at any time period evaluated. 141 0.30 -r □ C ontrol c 0.25 SCEB ‘3 2 0.20 a . 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 4,12,24,36 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. 142 □ Control 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 No 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 4,12,24,36 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 □ Control HCEB 4 12 24 36 48 Time (Hours) Figure 3.6 Comparison of hepatic ECOD activity at 4,12,24,36 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 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. 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 pmole/min/mg protein and 25 pmole/min/mg protein, respectively, while hepatic and renal PROD activity were 5 pmole/ min/ mg protein and 1 pmole/min/mg protein, respectively. EROD activity was 30 pmole/min/mg protein and 2 pm 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 1- aminobenzotriazole 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 GST- catalyzed 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 peroxisome- proliferating 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 (Fenwick etal., 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 n h 3 o c h 2 O* I II I C- CH— CH2- CH2- C N H - C H - C - N H - CH2 -O II O Figure 4.2 Structure of glutathione (GSH). 154 c h 3 I c h 2 o o 1 II II CH2- CH,- S = N - P— O' I c h 2 o - c h 2 I HC— N H ,+ 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 etal, 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 CEB- associated 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/kg 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-n- propylsulfamoyl 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 AOAA/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, New 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 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, N- nitroso-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 GC/M S/M S/DS from Finnigan MAT (San Jose, CA, USA), respectively. A Hitachi U-2000 double-beam UV/Vis 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 UV/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 GC/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) B 12 h o u rs C 'E 1.0 + □ 24 hours 148 h o u rs CO ® 0.8 + o 2 0.6 + O 0 .4 + jE Jjj 0.2 ^ 0.0 4 - C ontrol 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 post- CEB 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. No 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 4.8- 4.13). The metabolite was also present in the urine of rats pretreated with 165 0 .3 5 t S 12 hours 0.30 - □ 24 hours ■ 48 hours $ 0 . 2 5 + 0.20 - 3L 0 .1 5 -■ 0.10 - 0 .0 5 - Control AOAA Probenecid AT-125 Treatment Figure 4.5 Urinary nonprotein thiols excreted at 12,24, and 48 hours post- treatment 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 a> j= 0 .8 -r c Q 12 h o u rs *3 □ 24 h o u rs (0 0) ■ 48 h o u rs 0.6 - o 0.5 - 5. w 0.4 - 0) £«-< 09 O 0.2 - !E h" 0.1 - s 3. Control AOAA Probenecid AT-125 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 -- □ 24 h o u rs 48 h o u rs 0.25 -- ® 0.15 0.1 -- 0.05 - Control AOAA Probenecid 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 40.0-650.0 amu. CEB CONTROL #44 JSPM TIC ££0 £40 £80 300 Q 100000 - 9.5 10.0 10.5 11.0 11.5 1£.0 TIME 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 4Q.Q-65Q.0 amu. N0RMRLF344 URINE “ JSPM STC TIC ££0 £40 £80 300 w 400000- 5 100000 - 9.0 9.5 10.0 10.5 11.0 11.5 1 £. 0 TIME 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 F ile >JM406 CEB CONTROL #44 JSPM l-'4/'95 URINE S c a n 2 6 8 Bpk fib 7767. SUB ADO CLP DVC 1 0 . 6 4 m in . 110 4 3 8000- 100 7 0 0 0 - ■90 8 0 U9 u 7 0 5QQ0- 3 7 6 0 Q 2 D 4 0 0 0 - 5 0 CD < 4 0 3Q 00- 1 7 7 3 0 7 9 2 0 1 2 0 10QO- 112 1 5 2 2 4 5 10 -0 8 0 1 6 0 200 2 4 0 2 8 0 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. 170 177 .00 CH- 123 .00 87 .00 5000; Q z 0- p F 1 1 a >JM4Q5 4 0 .0 -6 5 0 .0 amu. CEB CONTROL #44 258 254 256 258 260 262 264 266 268 270 80000 40000; O 1 0 .3 0 1 0 .4 0 10 '.50 10 .6 0 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 .00 C0Ntl?0L #27 F344 UR JSPtl TIC 260 264 200000- ’ - '" T • ’ ■ I I " ■ ' • I I ...... I"' ■ ' ’ I • ■ • ■ I ' ■ 10.30 10.40 10.50 10.60 10.70 TIME 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 (*). 100-1 127 r 149240 00 177 144 201 102 5 0 .0 - 74 53 134 152 80 112 104 47 160 117 93 1SS £ 215 257 w 245 2Ul l- M/Z 50 100 150 2 50 2 100 r 149248 5 0 .0 - 269 279 269 301 314 328 343 356 405 475 M/Z 3 00 3 50 4 00 450 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-(4- cyano-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]. 174 100.0 366 r 3 8 9 5 2 9 0 267 208 235 £ CO z Ui 164 H Z 317 179 302 133 289 385 328 115 RIC 415 445 468 100 150 200 250 300 350 400 450 500 SCAN 3:20 5:00 6:40 8:20 10:00 11:40 13:20 15:00 16:40 TIME 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 2 8 4 2 6 2 0 234 222 255 374 302 162 195 332 351 305 119 361 RIC 414 461 490 100 150 200 2 50 300 350 400 450 5 00 SCAN 3:20 5:00 6:40 8:20 10:00 11:40 13:20 15:00 16:40 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 5 0 .0 - 305.091 ±0.5 £ CO 302 385 z(U H Z 368 r-3895290 267 ABUNDANCE 208 235 RIC “ 164 317 179 3B5 415 445 486 1 00 15 0 200 2 5 0 300 350 400 450 5 00 SCAN 3 :20 5:00 6 :40 8:20 10:00 11:40 13:20 15:00 16:40 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. Figure 4.18 Mass spectrum of scan # 367 from the chemical ionization chemical the from 367 # scan of spectrum chromatogram. Mass 4.18 Figure % % INTENSITY 0- .0 0 5 0- .0 0 5 100 100 M/Z M/z 100 333 lull 1 114 1 50 35 128 355 144 150 7 37 9 40 1 42 4 458 449 432 419 410 397 387 378 160 450 0 0 4 176 190 200 7 9 54 1 57 540 527 516 504 493 475 245 250 257 263 ■ 500 K - 273 i ■ i ■ 00 30 305 550 r-196352 6352 9 1 r ABUNDANCE 177 M + 41 (345) ions consistent with a molecular weight of 304. of weight molecular a with and consistent (333), the 29 ions + (345) M from 1367 (305), # 41 + + M M the scan of reveals spectrum mass chromatogram the of ionization view closer chemical A 4.19 Figure % % INTENSITY 100 0- .0 0 5 M /Z /Z M . 0-1 0 0 3 301 305 310 1 343935 2 331 328 325 319 314 311 20 3 330 333 4 350 340 4 351339343 345 146432 r ABUNDANCE 178 179 RELATIVE QUANTIFICATION OF URINARY MERCAPTURATE 25 00000 — ■ — CEB — • -“ AT-125 2000000 “ A - AOAA 15 00000 1000000 5 0 00 00 0 0 4 12 24 36 48 Time (Hours) 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 200000 - 1 2 7 . 0 0 8 0 0 0 0 - Q 4 0 0 0 0 - File >TJUDU 40.0-650.0 amu. URiNE TREATED TIC 50 100 150 , 200 250 300 350 400 . i . ■ ■ ■ i.. ■ ■ I.. ■ ... i .. .. i-.... .i ■. i l.U.i »„liui Ixtt.Jn .1 I i »-> ■ I l 4 0 0 0 0 0 0 2000000 CH 4 I 0 ' s'.O 6.0 7.0 816 olo lo'.o 11.0 12.0 13.0 14.0 15.0 TIME b) 128 .00 20000 0 - ..i. .A . .. __ 1 2 7 . 0 0 1000001 Q 5 0 0 0 0 2 So ch File >J0 8 2 3 40.0-650.0 a m u . CONTROL # 2 7 F 3 4 4 UR JSPM TIC 50 100 150 200 250 300 350 400 4 0 0 0 0 0 0 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-glutamyl 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 N- acetyltransferase 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 (Seutter- Berlage 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)-L- cysteine (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-7- guanyl]ethy!)-L-cysteine (Kim and Guengerich, 1989). Ethylene oxide, a 186 potential human carcinogen, similarly forms N-acetyl-S-(2-hydroxyethyl)-L- cysteine 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 al, 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-glutamyl- cysteine 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 O- deethylase (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/kg body w eight) 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. 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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 is consistent with the structure of l-cyano-3,4-epithiobutane. of structure the with consistent is FIGURE A.1 NMR (200 MHz) of the distilled product. NMR spectrum NMR product. distilled the of MHz) (200 NMR A.1 FIGURE PPM "I -- ■ | ’ " - “T_l - | " • ’ ' | • ■ ■ ■“""I -'- O 4.0 3.0 2.0 tO 0.0 0 t.O 0 . 2 0 . 3 0 . 4 .O S 0 . 6 0 . 7 -1",- | • . , '. ' - - - j - • | - I-'11" , gtietefektitidStisHK - 3 4- THK BUTANE K H IT P -E 4 -3. O N A Y i-C PPM I ' I T-^— Jl 263 structure of l-cyano-3,4-epithiobutane. of structure FIGURE A.2 Enlarged NMR (200 MHz) spectrum of the distilled 1- distilled the of spectrum MHz) (200 NMR the Enlarged with consistent detail increased depicts Spectrum cyano-3,4-epithiobutane. A.2 FIGURE PPM l-CYANO-3. 4-EPITHIOBUTANE 265 a) « B »»■ ^T IsSs.S! X M ■# ♦ M •* - J N - 3 ) ' a » — — —. n ce ?•» n *■ «» s j » jt* \ S M7* aJ Sm >* t* f •* S..-, £.M 5S i ,-i «p Jj 9 9 < r s n .*) « S 9 9 15 b) MdS,*)4f V S IS g «N»54V.'4V.»— Q>.rj 15 3 m s s - i s l s s a 8 3 ’ ‘ ^ ££££3>£3i£S£3 sv^r' ^ jj.Ni r* S’ s * X -»»9?»f/)rt \ ye . 3*SSSZ2R5SSS *fi u Li I i i: * =S* &cvW 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. 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: Judith S. Prescott-Mathews Major Field: Veterinary Medical Sciences Title of Dissertation: Elucidation and Modulation of CEB Metabolism in Fischer 344 Rats: Possible Mechanisms of Toxicity and Anticarcinogenicity Approved: or Professor and Chairman EXAMINING COMMITTEE: Date of Examination: June 28, 1995