Sulfotransferases and Acetyltransferases in Mutagenicity Testing: Technical Aspects

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[13] Sulfotransferases and Acetyltransferases in Mutagenicity Testing: Technical Aspects By HANSRUEDI GLATT and WALTER MEINL Abstract

Sulfotransferases (SULTs) and N‐acetyltransferases (NATs) mediate the terminal activation step of various mutagens and carcinogens. Target cells of standard in vitro mutagenicity tests do not express any endogenous SULTs. NATs are expressed in some cells, but may not reflect the substrate specificity of human NATs. External activating systems usually lack the cofactors for these enzymes. Upon addition of the cofactor, the ultimate mutagen may be formed, but especially sulfo conjugates—anions—may not reliably penetrate into the target cells. This chapter presents methods used to incorporate these enzyme systems into in vitro mutagenicity test systems and to identify the critical human forms. The method of choice is direct expression of the enzymes in target cells. We present procedures on how this can be reached in bacteria and in mammalian cell lines in culture. Furthermore, genetically manipulated mouse models are a very promising perspective for answering open questions.

Introduction

Mutagenicity is an important toxicological effect, as it can lead to inheritable damage to the progeny, the initiation and progression of neoplasias, and degenerative changes in tissues. Because the structure and processing of DNA are highly conserved, mutagenicity studies conducted in any biological systems may be useful for detecting possible genotoxic hazards to human. Indeed, in vitro investigations using bacterial or mammalian target cells form the dominating part in standard programs for studying the genotoxicity of new compounds. Chemical damage to DNA is by far the most common mechanism underlying chemical mutagenesis. It involves reactive molecules, which usually are formed within the organism, either as side products of endogenous metabolism or during the biotransformation of xenobiotics. Biotransformation can vary enormously among species, genotypes, tissues, and physiological states. This variation tremendously complicates the design of appropriate genotoxicity studies in vitro and in vivo and the assessment of risks for humans. It is often assumed that phase I metabolism is particularly prone METHODS IN ENZYMOLOGY, VOL. 400 Copyright 2005, Elsevier Inc. All rights reserved.

0076-6879/05 $35.00 DOI: 10.1016/S0076-6879(05)00013-3

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to the formation of reactive intermediates. Therefore, hepatic microsomal or postmitochondrial (‘‘S9’’) fractions, supplemented with NADPH (the cofactor of cytochromes P450) or an NADPH‐generating system, are used frequently in in vitro genotoxicity assays. Cofactors for phase II enzymes are normally not added to activating systems, as many toxicologists associate phase II metabolism primarily with detoxification rather than toxification. We are convinced that this view is one‐sided and that involvements of phase II enzymes in toxification are rather frequent. However, we recognize another reason for omitting cofactors for phase II enzymes in activating systems. The primary function of typical phase II enzymes is the introduction of an anionic group into the acceptor molecule to prevent the passive penetration of cell membranes, the property needed for its vectorial transport and effective excretion. Therefore, many reactive (and often short‐lived) phase II metabolites formed externally may not reach the DNA of the target cell. For this reason, other strategies are required for the reliable detection of genotoxicants formed by phase II enzymes. A phase II metabolite, 2‐acetylaminofluorene N‐sulfate, was the first ultimate carcinogen, formed from a procarcinogen, to be discovered (DeBaun et al., 1968; King and Phillips, 1968). Subsequently, it was found that sulfotransferases (SULTs) are also involved in the bioactivation of other carcinogens, e.g., many aromatic amines (Miller, 1994). Using SULT‐ proficient target cells we then demonstrated that a large number of compounds belonging to several chemical classes could be activated to mutagens by SULTs (Glatt, 2000, 2005). The high chemical reactivity of many sulfuric acid esters, formed via sulfo conjugation of hydroxylated acceptor molecules, can be explained by the fact that sulfate is a good leaving group in certain chemical linkages. For example, heterolytic cleavage is facilitated if the resulting cation is resonance stabilized, as is the case with sulfuric acid esters derived from aromatic amines (Fig. 1). O‐Acetylation is another metabolic reaction creating a potential leaving group, acetate, in xenobiotics. The enzymes catalyzing this reaction in vertebrates are termed N‐acetyltransferases (NAT) because they appear to transfer the acetyl group exclusively to nitrogen atoms and N‐centered hydroxyl groups. Therefore, NAT‐mediated activation has been observed primarily with aromatic hydroxylamines (usually formed from amino‐ and nitroarenes). Various aromatic hydroxylamines may be activated by SULTs and NATs via analogous mechanisms (Fig. 1). In both cases, the same nitrenium/carbonium ion is transferred to the DNA (and other nucleophilic acceptors). However, in agreement with the fact that sulfuric acid (pKa
3 and 1.92 for the first and second proton, respectively) is a more potent acid than acetic acid (pKa 4.74), sulfuric acid esters are usually more reactive than the corresponding acetic acid esters. Moreover, acetylation is an

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FIG. 1. Activation of aromatic amines (1) and nitro compounds (2) via hydroxylamines (3) to reactive esters (4,5), which may react spontaneously with DNA and other nucleophiles via a resonance‐stabilized nitrenium/carbonium ion (6). Sulfuric acid esters (4) differ from acetic acid esters (5) by their negative charge and higher reactivity.

atypical conjugation reaction, as it involves the transfer of a neutral group, which may not constrict the penetration of membranes. Finally, the substrate specificity of the enzymes may be another factor determining which pathway is important for a given substrate. In the mammalian species investigated, usually 2 members of the NAT superfamily (three in the mouse) and approximately 12 members of the SULT superfamily have been detected. The activation of various promutagens by NATs (Hein, 2000) and SULTs (Glatt, 2000, 2005) has been reviewed elsewhere. This chapter focuses on methodological aspects—how to detect the involvement of these enzyme classes in the activation of a compound and how to identify the enzyme forms involved. The dominating part deals with the

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expression of these enzymes in standard target cells of mutagenicity assays, using gene‐technical approaches, and with the usage of the systems. Natural Expression of SULTs and NATs in Target Cells of Test Systems

SULTs are expressed in vivo with high tissue and cell‐type selectivity (Glatt, 2002). In human, some SULTs are highly expressed in liver, whereas other forms are absent or very low in this tissue, but high at specific extrahepatic sites. In rats and mice, the expression of most SULTs is much more focused to the liver than in human. The expression of the mRNA of SULTs is completely lost (or is reduced for SULT1A1) in primary cultures of rat hepatocytes within a few hours (Liu et al., 1996). Likewise, SULT activity levels are much lower in hepatic and other epithelial cell lines in culture than in liver tissue (Glatt et al., 1994). In general, fast‐growing mesenchymal cell lines, such as Chinese hamster V79 and CHO and mouse L5178Y cells, are used for mutagenicity experiments. Using numerous substrates and antisera (cross‐reacting with rat, mouse, and human SULTs), we have not detected any SULT expression in V79 cells, the cell line widely employed in our laboratory for genetic engineering and genotoxicity studies (Glatt et al., 1994; and many results that have not been published systematically). Sporadic investigations in CHO and L5178Y cells consistently gave negative results. Furthermore, we have not detected SULT activity toward any xenobiotics in Ames’s Salmonella typhimurium strains (Glatt et al., 1994; and many results that have not been published systematically), the most widely used target cells in mutagenicity research. The V79 cells maintained in our laboratory for many years (termed V79‐MZ) do not express any NAT proteins or activities (Glatt et al., 2004). The situation may be different in V79 sublines used in other laboratories. In particular, the subline V79‐NH demonstrated some endogenous NAT activity (Perchermeier et al., 1994). However, Chinese hamster NATs have not yet been characterized on a molecular level, and thus it is not known which forms are expressed in V79‐NH cells and whether they are similar to human forms. Ames’s S. typhimurium strains express an endogenous acetyltransferase. Knockout of this enzyme (in strains marked with the suffix ‘‘DNP’’ or ‘‘1,8‐DNP’’) leads to a drastic decrease in the mutagenicity and cytotoxicity of many nitro‐ and aminoarenes, such as 1,8‐dinitropyrene (1,8‐DNP) (Rosenkranz and Mermelstein, 1983). This bacterial enzyme differs in its substrate specificity from mammalian enzymes, despite some overlaps. For example, mammalian NATs, unlike Salmonella acetyltransferase, show N‐hydroxyacetamide N,O‐transacylase activity, and thus

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can activate the N‐hydroxy metabolites of various aromatic amides (Ando et al., 1993). SULT and NAT Activity in Subcellular Activating Systems

Xenobiotic‐metabolizing SULTs and NATs are soluble proteins normally localized in the cytoplasm. However, some SULTs were detected in the nuclei rather than the cytoplasm in some tissues (He et al., 2004). This localization may require a modification of some protocols for subcellular fractionation. Furthermore, each SULT and NAT form is not expressed in each tissue and ontogenetic stage. In adult rat liver, several SULT forms are only present in females or males (reviewed by Glatt, 2002). Apart from these complications, a role of SULTs or NATs in a biotransformation (e.g., activation) reaction in a subcellular system can be demonstrated readily by conducting incubations in the presence and absence of their characteristic cofactors, 30 ‐phosphoadenosine‐50 ‐phosphosulfate (PAPS) and acetyl coenzyme A (acetyl‐CoA), respectively. Identification of the critical form may be strived for by comparing the effect of preparations from tissues differing in the forms expressed or by using selective inhibitors. Likewise, individual cDNA‐expressed and/or purified enzymes may be employed (Glatt et al., 1995). Tissue levels of PAPS and acetyl‐CoA are low. PAPS levels of 3.6 to 76.8 nmol per gram tissue have been reported for various tissues from different species (Klaassen and Boles, 1997). In liver of the guinea pig and the rat, acetyl‐CoA was detected at levels of 3 and 14 nmol per gram, respectively (Erfle and Sauer, 1967; Kato, 1978). Thus, their endogenous concentration is negligible in the strongly diluted subcellular preparations commonly used in biotransformation studies, and normally they have not to be removed (e.g., by dialysis) for the negative controls. For the addition of the cofactor, the following information may be useful. Apparent Km values of SULTs for PAPS can range from <1 to 155
M depending on the enzyme form and experimental conditions used (reviewed by Glatt, 2002). PAPS, unlike its desulfonated form, PAP, does not inhibit SULTs when its concentration is increased. Therefore, we recommend a generous supply of PAPS, especially if the sulfo acceptor substrate is used at a high concentration and a high turnover is strived for. Commercially available PAPS usually contains high levels of PAP, which inhibits SULTs. Fortunately, complete cytosolic fractions from tissues and cells contain PAP‐degrading enzymes and thus reduce this inhibition. When accurate quantification of reaction parameters is aimed at or when a purified SULT is studied, it is, however, advised to use high‐quality PAPS (synthesized in some laboratories working with SULTs).

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PAPS (which is relatively expensive) may be replaced in some situations by a PAPS‐generating system. The physiological synthesis of PAPS is conducted by PAPS synthetases from inorganic sulfate and ATP and requires the presence of Mg2þ. PAPS synthetase is found in the cytosolic fraction of most tissues, including liver. Therefore, it is sufficient to supplement such preparations with ATP (e.g., 10 mM) and MgSO4 (e.g., 10 mM). An alternative is the ‘‘transfer assay,’’ which can lead to particularly high sulfo conjugation rates and involves the transfer of the sulfo group from a conjugate (usually 4‐nitrophenylsulfate, used at a high concentration, e.g., 5 mM) to SULT‐bound PAP and from there to the actual acceptor substrate (Frame et al., 2000; Duffel et al., 2001). Although this approach can be very efficient, it should only been used with well‐characterized and appropriate enzyme systems. In particular, it is necessary that a SULT is present that accepts 4‐nitrophenol/4‐nitrophenylsulfate as a substrate, and ideally the same enzyme mediates the sulfo conjugation of the final substrate. Primarily, human SULT1A1 is appropriate for this reaction (Frame et al., 2000). This enzyme shows particularly broad substrate tolerance toward promutagens (Glatt, 2000, 2005). However, the use of PAPS‐generating systems is not possible for determining accurate kinetic data. In general, we prefer the use of highly pure PAPS in subcellular enzyme systems. Some SULTs are strongly inhibited at excessive substrate concentrations. For example, human liver thermostable phenol sulfotransferase (SULT1A1) showed an apparent Km of 0.94
M for the standard substrate 4‐nitrophenol; the conjugation rate was maximal at 4
M and then decreased rapidly with increasing substrate concentrations (Campbell et al., 1987). This effect was also observed with various other substrates and has to be taken into account when studying possible bioactivation reactions. The use of high substrate concentrations may lead to an underestimation of the activation potential. We address this point here, as conventional activity measurements with subcellular preparations are usually conducted under enzyme‐limiting, substrate‐saturated conditions, whereas toxicological experiments with cells are normally done under substrate‐limiting conditions in the presence of high enzyme levels. This difference may be the reason for various conflicting results in the literature. A concentration of 200
M acetyl‐CoA is appropriate for studying NAT activities; it is recommended to add acetyl phosphate (5 mM) and phosphotransacetylase (4.5 U/ml) to recycle CoA, which is an inhibitor of NATs (Andres et al., 1985). Others have used 2 mM of acetyl‐CoA in the absence of a regenerating system (Arlt et al., 2005). Subcellular preparations have been incorporated in cell‐free genotoxicity test systems using the formation of adducts to DNA and other biomolecules as the end point. This was how the SULT‐dependent activation of

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2‐acetylaminofluorene in the presence of hepatic preparations was detected (DeBaun et al., 1968; King and Phillips, 1968). The approach has also been employed with cDNA‐expressed SULTs and NATs (e.g., Arlt et al., 2005). More often, subcellular preparations are combined with cellular genotoxicity test systems, such as the NADPH‐fortified hepatic S9 fraction in standard Salmonella and mammalian cell mutagenicity tests (Bradley et al., 1981; Maron and Ames, 1983). It appears that the cofactor acetyl‐CoA has not been fortified in such studies, probably because the most popular target cells (S. typhimurium and Escherichia coli) express their own acetyltransferase and produce the necessary cofactor. PAPS‐ fortified activating systems have been used with varying success. The major problem involves penetration of the active metabolite into the target cells, which depends on the individual sulfo conjugate, the target cell, and the incubation conditions. We noticed that some benzylic sulfuric acid esters readily exchange the anionic leaving‐group sulfate against a neutral leaving group (such as chloride) in a medium containing the corresponding component and that the resulting secondary reactive species can readily penetrate into the target cells (Enders et al., 1993). Thus, we have used the bioactivation of 1‐hydroxymethylpyrene to a bacterial mutagen as an assay for monitoring the active fractions in the purification of a SULT (Czich et al., 1994). Other classes of reactive sulfuric acid esters, including those derived from aromatic amines, are usually not detected as mutagens when generated extracellularly. For example, chemically synthesized 2‐acetylaminofluorene N‐sulfate showed negligible mutagenicity to S. typhimurium (Smith et al., 1986). Likewise, the addition of PAPS to the hepatic‐metabolizing system decreased the mutagenic activity of N‐hydroxy‐2‐acetylaminofluorene (Mulder et al., 1977), whereas intracellular (cDNA‐expressed) SULT strongly enhanced its effect in isogenic bacterial strains (next section). Recombinant Systems

Historical Background Watanabe et al. (1990) cloned the acetyltransferase of S. typhimurium into the pBR322 vector. This plasmid was introduced into the standard S. typhimurium strains TA98 and TA100, yielding strains YG1024 and YG1029, respectively. This manipulation led to an approximately 100‐fold increase in acetyltransferase activity and to strongly enhanced mutagenic activity of various N‐hydroxylamino‐ and nitroarenes (tested directly) and aminoarenes (in the presence of a hepatic S9 preparation). Grant et al. (1992) were the first to express human enzymes directly in target cells of a bacterial mutagenicity test system. They cloned human NAT1 and NAT2

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into the phagemid vector pKEN2 and then transformed strain TA1538/1,8‐ DNP (deficient in endogenous acetyltransferase). The resulting strains, DJ400 and DJ460, respectively, were much more sensitive to the mutagenic action of 2‐aminofluorene, benzidine, and 2‐amino‐3,4‐dimethylimidazo‐ 4,5‐f ]quinoline than the parental strain TA1538/1,8‐DNP. We constructed analogous strains (named TA1538‐DNP‐hNAT1 and ‐hNAT2) using the pKK223–3 vector (Table I) (Muckel et al., 2002). The responsiveness of these strains was similar to those of strains DJ400 and DJ460, respectively. E. coli and S. typhimurium strains coexpressing a human cytochrome P450, together with high levels of Salmonella acetyltransferase, have also been constructed for detection of the mutagenicity of various aromatic amines without the need of external activating systems (Josephy et al., 1998; Suzuki et al., 1998). However, analogous strains with human NATs are not yet available. We have used vector pKK233‐2 (Table I) for expressing a total

TABLE I VECTORS USED FOR THE EXPRESSION OF SULTS AND NATS IN TARGET CELLS OF MUTAGENICITY TESTS

Vector

Unique restriction sites within multiple cloning site

Expression mode

Selection marker

trc

Constitutive

Ampicillin

trc

Constitutive

Ampicillin

tac

Constitutive

Ampicillin

trc Two adjacent tac motifs

Constitutive Induciblea

Neomycin Ampicillin

Constitutive

Noneb

Constitutive

Noneb

Promoter

Expression in S. typhimurium pKK233‐2 pKKnew

pKK223‐3 pKNeo pCWmodI

NcoI, PstI, HindIII NcoI, SacI, SmaI, XbaI, PstI, HindIII EcoRI, SmaI, PstI, HindIII see main text NdeI, XbaI, SalI, HindIII

Expression in V79 cells pMPSVEH

EcoRI, SalI, BamHI, HindIII

pSI

EcoRI, XbaI, SalI, SmaI, NotI a

Long terminal repeat of myeloproliferative sarcoma virus SV40 early promoter

Vector contains lac Iq repressor (absent in the S. typhimurium LT2 genome), induction by isopropylthio‐‐D‐1‐galactopyranoside. b Usually cotransfected with a separate vector (pBSpac
p) conferring resistance to puromycin.

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of 40 different mammalian SULTs in S. typhimurium TA1538 (e.g., Glatt et al., 1996; Meinl et al., 2002). Doehmer et al. (1988) were the first to employ gene transfer techniques for expressing a xenobiotic‐metabolizing enzyme, a cytochrome P450, in mammalian target cells (V79) to be used in gene mutation tests. We adopted this technology for expressing SULTs and NATs in V79 cells (Czich et al., 1994; Teubner et al., 2002). However, we selected other vectors and resistance markers to make possible the expression of these conjugating enzymes in cells already engineered for a cytochrome P450 (Glatt et al., 1996, 2004). The following sections present technical aspects of our experience with the expression of SULTs and NATs. Vectors cDNAs are normally cloned in E. coli, e.g., in strain XL blue‐1 (Stratagene, Heidelberg, Germany). Plasmids are adapted to the restriction enzymes of Ames’s S. typhimurium strains (derivatives of LT2) by passaging through the restriction‐deficient, but methylation‐proficient S. typhimurium strain LB5000 (Bullas and Ryu, 1983). All bacterial expression vectors used contain a trc or tac promoter (Table I), leading to repression in common E. coli strains (containing an endogenous lac Iq) in the absence of an inducer (such as isopropylthio‐‐D‐1‐galactopyranoside). However, these promoters lead to constitutive expression in S. typhimurium due to the absence of endogenous lac Iq unless such a repressor is contained on the plasmid. For the expression of SULTs in bacteria, we generally use pKK233‐2 (Amann and Brosius, 1985) (GenBank accession number X70478; the vector was obtained from Pharmacia, Freiburg, Germany, but is now delisted). The NcoI restriction site used for inserting the 50 end of the cDNA contains an ATG translation initiation codon followed by a G. Fortunately, the second codon of most SULTs starts with this base. In other cases (primarily SULT2A forms), the base was belatedly corrected by site‐directed mutagenesis. To enhance the practicality of this vector, we introduced additional unique restriction sites within the multiple coding region (Meinl et al., 2002); in the meantime we named this vector pKKnew. These pKK vectors contain an ampicillin resistance marker. Such a resistance factor is already present in standard S. typhimurium strains containing plasmid pKM101 (e.g., strains TA98 and TA100). For the transformation of these strains, we exchanged the ampicillin resistance marker in pKK233‐2 against a neomycin resistance marker to give vector pKNeo (Glatt and Meinl, 2004). This vector contains additional NcoI restriction sites. Therefore, cDNAs

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were first cloned in pKK233‐2 or pKKnew and then transferred into pKNeo using SalI (pKKnew) or SalI/HindIII fragments. Similar levels of SULT proteins were expressed from vectors with the different resistance markers in strains TA1538 or TA1537 (not containing pKM101). Nevertheless, the activities of SULT‐dependent and ‐independent mutagens were stronger (in general by a factor of nearly 2) with the original vector than with the pKNeo vector, indicating additional influences of the resistance markers. Vector pKK223‐3 (M77749, from Pharmacia, delisted) differs primarily from pKK233‐2 in the restriction sites in the multiple cloning region (Table I). It was more appropriate than the latter vector for the insertion of NATs. Vector pCWmodI (a generous gift of Dr. T. Friedberg, University of Dundee, Scotland) was generated from pCW (Gegner and Dahlquist, 1991) by deleting the CheW gene. The NdeI restriction site contains the ATG translation initiation codon (but no additional bases reaching into the next codon). pCWmodI encodes a lac Iq repressor sequence. For the stable expression in V79 cells, we initially used pMPSV (Artelt et al., 1988), carrying the long terminal repeat promoter of the myeloproliferative sarcoma virus (Czich et al., 1994; Teubner et al., 2002). Later, we also explored other vectors. In particular, pSI (U47121, purchased from Promega, Mannheim, Germany) appeared to give somewhat higher expression levels than pMPSV, although a robust comparison is difficult, as the expression in each clone is affected by chance (resulting from the number of copies integrated and the integration sites). Expression from pSI is driven by the SV40 early promoter. Characteristics and Levels of Expressed Proteins The heterologous enzyme proteins in recombinant cells are analyzed by electrophoresis in polyacrylamide gels under denaturing conditions with subsequent immunoblotting. With most SULT and NAT expression vectors, a single polypeptide band is observed, which comigrates with the corresponding protein naturally expressed in mammalian tissues. The only exception among the 13 human SULTs and NATs (reference sequences) and their allelic variants was found with SULT2B1b. It produced a single band in E. coli and V79 cells, but an additional, weaker band, representing a smaller polypeptide, in S. typhimurium. Site‐directed mutagenesis of an internal ATG abolished the occurrence of this second band, suggesting that this ATG was used as an additional translation initiation codon. Two double bands, which were just separated by electrophoresis, were also observed with mouse and rat Sult1e1 (containing ATG as codon 4) expressed in S. typhimurium. Dog SULT1D1 (Tsoi et al., 2001) cDNA produced

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two immunoreactive bands differing by nearly 7 kDa in E. coli, with the larger band comigrating with the authentic protein. S. typhimurium only exhibited the shortened polypeptide. We have not studied its origin, but SULT1D1 contains an ATG codon in position 60. If used for starting the translation it would create a polypeptide with the approximate size of the shortened product found in bacteria. For the estimation of expression levels, purified enzyme protein or— nearly homogeneous, but enzymatically inactive—inclusion bodies were used as standards in immunoblot analyses. Expressed from the natural cDNA sequence in pKK and pKNeo vectors in S. typhimurium, the level of SULTs usually was in the range of 0.5 to 10% of the soluble protein fraction (Meinl et al., 2005). Human SULT2B1b was the most notable exception, with a very low expression under these conditions (see later). Using the same approach for quantification, we found SULT1A1 levels of 0.07 to 0.3% in the cytosolic fraction of three liver samples concurrently tested, the highest value being one‐third of that observed in the standard S. typhimurium strain TA1538‐hSULT1A1 (Meinl et al., 2005). Expression levels for NATs were lower in S. typhimurium in absolute terms than for SULTs, but exceeded hepatic levels by factors of 100. In some situations, one would like to vary the level of the activating enzyme in a mutagenicity study. This is simple with subcellular preparations (Glatt et al., 1994, 1995), but difficult with enzymes expressed within the target cells. For example, when we compared the activation potential between alloenzymes of human SULT1A2, we noticed that they were expressed at different levels in the S. typhimurium strains constructed (Meinl et al., 2002). We then analyzed several separately transformed clones, but only found negligible variation in expression levels and promutagen activation. We also varied the concentration of ampicillin, the selection marker, from zero to a high level (100
g/ml) when growing the strains; this modification did not affect the activation of promutagens either. In an attempt to reduce the level of an enzyme, we introduced synonymous low‐ usage codons in the 50 ‐terminal region of its cDNA. To our surprise, this modification enhanced, rather than decreased, the expression (Meinl et al., 2002). Consequentially, we then employed this method with various poorly expressed cDNA—consistently with success. For example, the level of SULT2B1b protein in E. coli and S. typhimurium was enhanced from barely detectable with the natural cDNA sequence to nearly 3% of the soluble protein after the introduction of several synonymous low‐usage codons (M. Osterloh‐Quiroz, W. Meinl, and H. R. Glatt, manuscript in preparation). We have not yet clarified whether the low‐usage codons as such were decisive or accidentally associated changes in the secondary structure of the mRNA.

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The other approach for modulating expression levels in S. typhimurium involved the use of an inducible promoter (pCWmodI, Table I). However, the repression of this vector in S. typhimurium is incomplete, as shown for the activation of N‐hydroxy‐2‐amino‐3‐methylimidazo[4,5‐f ]quinoline in a strain engineered for expression of rat NAT1 (Fig. 2). The activation was only enhanced two‐ to threefold when the inducer, isopropylthio‐‐D‐1‐ galactopyranoside, was added to the growth medium. Effects of other test compounds and with other NAT forms were enhanced to similar extents. Finally, it was possible to modify the expression of various NATs and SULTs by changing the temperature at which the cultures were grown. However, this modification may also alter the expression of endogenous factors involved in the handling of the test compound or the primary damage. Thus, the method is not appropriate for elucidating the influence of varying levels of a heterologous enzyme on the mutagenicity of a chemical. Unlike in the bacterial model, the transfection of expression vectors into V79 cells created transformed cell clones that differed strongly in the level and stability of the heterologous protein. This is due to the fact that

FIG. 2. Mutagenicity (reversion to histidine prototrophy) of N‐hydroxy‐2‐amino‐3‐ methylimidazo[4,5‐f ]quinoline (N‐OH‐IQ) in S. typhimurium strain TA1538‐1,8‐DNP‐rNAT1 grown in the presence ( ) or absence (□) of isopropylthio‐‐D‐1‐galactopyranoside (1 mM, added to the overnight culture for the last 2 h before bacteria were used in the mutagenicity tests). This strain contains rat NAT1 cDNA inserted into vector pCWmodI. The NAT‐ deficient recipient strain TA1538–1,8‐DNP (○) was used as a control. The mutagenicity experiment was conducted as described by Muckel et al. (2002). Values are means and SE of three plates. The initial slopes of the dose–response curves amounted to 3000, 1200, and 10 revertants per picomole test compound, respectively. Unpublished data from E. Muckel and H. R. Glatt.

▪

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the transfected DNA has to be integrated into the host genome. The number of copies integrated and the integration sites are accidental with the vectors used. With SULT, we usually selected strains with expression levels in the high natural range in tissues, e.g., liver for human SULT1A1 and SULT2A1 and female liver for rat hydroxysteroid sulfotransferase a (a member of the SULT2A subfamily) (Glatt et al., 1998; Meinl et al., 2005). Expression of NATs in our recombinant cells lines is markedly higher than in tissues such as liver (Glatt et al., 2004). Various transformed V79 clones showed stable expression levels for 100 or more population‐doubling times, even in the absence of a selecting agent, which we only use for the initial picking of transfectants (Czich et al., 1994; Teubner et al., 2002). Cofactor Supply The cofactors PAPS and acetyl‐CoA can be added directly to subcellular‐metabolizing systems, as described in a preceding section. However, intact cells would not take up these cofactors and thus have to synthesize them. In practice, the synthesis has to occur during the incubation period, as the tissue and cell levels of these cofactors are low. The regeneration of acetyl‐CoA requires common intermediates from the endogenous metabolism and thus appears unproblematic—changes in the exposure media did not materially affect the responses with NAT‐dependent mutagens in S. typhimurium and V79 cells. The situation is different for PAPS, which is synthesized from inorganic sulfate (and ATP). Mulder and Jakoby (1990) observed apparent Km values of 0.3 to 0.5 mM for sulfate in the sulfonation of various substrates in isolated hepatocytes. Most cell culture media contain approximately 0.8 mM sulfate. We increased this concentration up to 5 mM when studying SULT‐dependent mutagenicity in recombinant V79 cells, but did not find any influence of this modification. In these experiments, we conducted the exposure in regular medium (Dulbecco’s modified Eagle’s minimum essential medium). It is also possible, and not uncommon, to use phosphate‐buffered saline for the exposure to the mutagen; it would be imperative to supplement sulfate when using this protocol with SULT‐dependent mutagens. Salmonella typhimurium does not appear to express any endogenous SULTs for transforming xenobiotics, but it produces PAPS for endogenous reactions. This is demonstrated by the fact that we could activate hundreds of chemicals to mutagens by mammalian SULTs expressed in these bacteria. There are two common exposure protocols for the Ames test: the plate incorporation assay (bacteria and test compound are mixed with soft agar and then immediately poured on agar plates) and the liquid preincubation assay (involving incubation of bacteria and test compound at 37 for some

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time before adding the top agar) (Maron and Ames, 1983). The second method is much more efficient than the former one for SULT‐dependent mutagens. Omission of sulfate in the exposure medium completely abrogates SULT‐dependent mutagenicity. We recommend using a concentration of at least 10 mM MgSO4 and a liquid preincubation time of at least 20 min. In general we now expose the bacteria to the test compound in 100 mM MgSO4 for 60 min (Meinl et al., 2002), based on our findings from various optimization experiments. Positive Controls The expression of SULTs and NATs usually had little effect on the number of spontaneous revertants in S. typhimurium TA1538 (and other strains without plasmid pKM101). We frequently use SULT‐independent, in addition to SULT‐dependent, mutagens as positive controls. The mutagenic activity of benzo[a]pyrene 4,5‐oxide (nearly 15,000 revertants per nanomole under our conditions) was usually unchanged after transformation with SULT‐ or NAT‐encoding vectors. However, during this work we noticed that our stock culture of TA1538 contained a subpopulation of cells with reduced response toward this compound (usually by a factor of four). Since some previous recombinant strains were derived from this population, we made new strains for the corresponding SULTs and observed enhanced mutagenicity not only toward benzo[a]pyrene 4,5‐oxide, but also some SULT‐dependent mutagens. Transformation of strain TA100 (containing pKM101) frequently led to a decrease in the number of spontaneous and benzo[a]pyrene 4,5‐oxide‐induced revertants. The effect was enhanced when the same protein was expressed at a higher level (by synonymous base exchange in the cDNA). 1‐Hydroxymethylpyrene (available from Aldrich, Taufkirchen, Germany) is particularly useful as a SULT‐dependent positive control compound because it is not mutagenic in SULT‐deficient recipient strains, but reverts most standard strains (TA1538, TA1537, TA98, and TA100, but not TA1535) after sulfo conjugation. The parental compound shows low bacteriotoxicity and thus can be used at high dose levels (up to 500 nmol per plate). It was activated to a mutagen by many different SULT forms. However, the appropriate dose levels varied substantially between different SULT forms, as shown in Fig. 3 for human SULTs. Activation of 1‐ hydroxymethylpyrene was weak (but unambiguous) with human SULT1C3 (Fig. 3, lower right) and absent (or marginal) with human SULT1C1, 2B1a, 2B1b, and 4A1. 6‐Hydroxymethyanthathrene is a more potent positive control compound to SULT1C3, but shows some mutagenic activity in

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FIG. 3. Mutagenicity (reversion to histidine prototrophy) of 1‐hydroxymethylpyrene to S. typhimurium TA1538 strains expressing the indicated human SULT. Strains TA1538‐ SULT1A1*1, *2, and *3 express allelic variants (involving amino acid substitutions) of SULT1A1 from their natural cDNA sequences. Expression levels were similar for all three alloenzymes. Synonymous codon exchanges were used to enhance the expression of SULT1A1 in strain TA1538‐SULT1A1Y. The mutagenicity experiment was conducted as described by Meinl et al. (2002). Values are means and SE of three plates.

the parental strain (10% of the activity observed with the SULT1C3‐ expressing strain) (Fig. 4, left). We have not yet found satisfactory positive control compounds activated by human SULT1C1, 2B1a, and 2B1b. However, cytosolic preparations of these strains demonstrated SULT activity

[13]

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FIG. 4. Positive control compounds activated to mutagens by human SULT1C3 in TA1538‐derived strains (left) and by human NAT1 and NAT2 (reference‐type alleles 1*4 and 2*4, respectively) in TA1538‐DNP‐derived strains (right). The mutagenicity experiment was conducted as described by Meinl et al. (2002). Values are means and SE of three plates.

toward conventional substrates. Although SULT4A1 is a member of the SULT family based on its amino acid sequence, no substrates have been found for this form up to date (Falany et al., 2000). We now use this strain as a (negative) vector control instead of, or in addition to, the parental strains (e.g., Figs. 3 and 4). Human and rat NATs, expressed in Salmonella, activate numerous aromatic hydroxylamines to mutagens. However, most of these compounds also show substantial mutagenic activity in acetyltransferase‐deficient strains when used at high dose levels. This background activity is particularly low with N‐acetyl‐2‐hydroxylaminophenanthrene, making it a convenient positive control compound (Fig. 4, right). Positive control compounds in recombinant V79 cells have to be adjusted to the needs of individual cell lines. 1‐Sulfooxymethylpyrene is mutagenic in these cells, but its metabolic precursor, 1‐hydroxymethylpyrene, is cytotoxic at relatively low concentrations even in SULT‐deficient control cells, limiting the concentrations that can be used. Its mutagenicity can be demonstrated in V79‐derived cells expressing human SULT1A1 or SULT1E1, forms that are particularly active in the Salmonella system. 2‐Nitropropane (used at a high concentration, 1 to 10 mM) is a convenient alternative for cell lines expressing human SULT1A1. Other positive control compounds are found in publications on the individual cell lines (Czich et al., 1994; Glatt et al., 1996, 2004; Teubner et al., 2002).

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Perspectives

Genotoxicity studies with recombinant bacteria and mammalian cell lines have demonstrated marked differences in the substrate specificity toward promutagens between orthologous SULT and NAT forms from human and rodent species normally used in carcinogenicity studies. Moreover, it is known that the tissue distribution of SULTs can vary strongly between different species. Based on these findings, one might suspect enhanced or reduced susceptibility of human toward standard rat and mouse models. Moreover, whereas SULT‐dependent carcinogenicity in these models usually was targeted to the liver, one might expect a higher level of extrahepatic effects in human. To test these hypotheses, animal models are required that express enzymes with human‐like substrate specificity and tissue distribution. Initial results from our laboratory indicate that this is possible for some SULTs by introducing human genomic sequences (rather than cDNAs) with long flanking regions into mouse oocytes. Acknowledgment Our current work on SULTs is financially supported by European Union (FP6–506820) and Philip Morris Incorporated.

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[14] A Comparative Molecular Field Analysis‐Based Approach to Prediction of Sulfotransferase Catalytic Specificity By VYAS SHARMA and MICHAEL W. DUFFEL Abstract

Understanding the catalytic function and substrate specificity of cytosolic sulfotransferases (SULTs) involved in drug metabolism is essential for predicting the metabolic outcomes of many xenobiotics. Although multiple isoforms of cytosolic SULTs have been identified and characterized in humans and other species, relatively little is known about the specific molecular interactions that govern their selectivity for substrates. The use of three‐dimensional quantitative structure–activity relationship METHODS IN ENZYMOLOGY, VOL. 400 Copyright 2005, Elsevier Inc. All rights reserved.

0076-6879/05 $35.00 DOI: 10.1016/S0076-6879(05)00014-5