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Oncogene (2002) 21, 8895 – 8904 ª 2002 Nature Publishing Group All rights reserved 0950 – 9232/02 $25.00 www.nature.com/onc Oxidative nucleotide damage: consequences and prevention Mutsuo Sekiguchi*,1 and Teruhisa Tsuzuki2 1 Biomolecular Engineering Research Institute, Suita, Osaka 565-0874, Japan; 2Department of Medical Biophysics and Radiation Biology, Faculty of Medical Sciences, Kyushu University, Fukuoka 812-8582, Japan 8-Oxoguanine (8-oxo-7,8-dihydroguanine) is produced in DNA, as well as in nucleotide pools of cells, by reactive oxygen species normally formed during cellular metabolic processes. 8-Oxoguanine nucleotide can pair with cytosine and adenine nucleotides with an almost equal efficiency, then transversion mutation ensues. MutT protein of Escherichia coli and related mammalian protein MTH1 specifically degrade 8-oxo-dGTP to 8oxo-dGMP, thereby preventing misincorporation of 8oxoguanine into DNA. The bacterial and mammalian enzymes are close in their size and share a highly conserved region consisting of 23 residues with 14 identical amino acids. Following saturation mutagenesis of this region, most of these residues proved to be essential to exert 8-oxo-dGTPase activity. Gene targeting was done to establish MTH1-deficient cell lines and mice for study. When examined 18 months after birth, a greater number of tumors were formed in the lungs, livers, and stomachs of MTH17/7 mice, as compared with findings in wild-type mice. These proteins protect genetic information from untoward effects of threats of endogenous oxygen. Oncogene (2002) 21, 8895 – 8904. doi:10.1038/sj.onc. 1206023 Keywords: DNA replication fidelity; oxygen radicals; oxidized guanine base; mutagenesis; carcinogenesis; genomic stability Introduction Reactive oxygen species, such as superoxide, hydrogen peroxide, hydroxyl radical and singlet oxygen, are produced through normal cellular metabolism, and formation of such radicals is further enhanced by ionizing radiations and by various chemicals (Ames and Gold, 1991; Henle and Linn, 1997). Nucleic acids exposed to oxygen radicals generate various modified bases, and more than 20 different types of oxidatively altered purines and pyrimidines have been detected (Gajewski et al., 1990; Demple and Harrison, 1994). Among them, 8-oxo-7, 8-dihydroguanine (8-oxoguanine) is the most abundant, and seems to play critical roles in mutagenesis and in carcinogenesis (Kasai and *Correspondence: M Sekiguchi; E-mail: [email protected] Nishimura, 1984; Fraga et al., 1990). Unlike other oxidative DNA damage, such as thymine glycol and 5’, 8-purine cyclodeoxynucleoside (Evans et al., 1993; Brooks et al., 2000; Kuraoka et al., 2000), 8oxoguanine does not block DNA synthesis, rather it induces base mispairing. 8-Oxoguanine can pair with both cytosine and adenine during DNA synthesis, and this mispairing could contribute significantly to spontaneous mutations in genomic DNA (Shibutani et al., 1991; Smith, 1992). Studies on Escherichia coli mutator mutants revealed that cells possess elaborate mechanisms that can prevent mutations caused by oxidation of guanine residues of DNA. 8-Oxoguanine residues in DNA can be removed by an enzyme that is coded by the mutM gene (Cabrera et al., 1988; Chung et al., 1991; Michaels et al., 1991; Bessho et al., 1992), while MutY protein removes adenine from adenine : 8oxoguanine pairs, which are produced as a result of misincorporation of adenine opposite to 8-oxoguanine (Au et al., 1988, 1989; Nghiem et al., 1988; Michaels et al., 1992). Thus, two proteins, MutM and MutY, act consecutively at the site of the oxidized guanine residue in DNA to prevent the occurrence of specific transversion mutation (Tchou and Grollman, 1993). In higher organisms, similar enzyme activities have been detected, and may account for the rapid elimination of 8-oxoguanine from chromosomal DNA. These include two types of MutM-related proteins, OGG1 and AtMMH, which carry 8-oxoguanine-DNA glycosylase activity (van der Kemp et al., 1996; Nash et al., 1996; Ohtsubo et al., 1998; Boiteux and Radicella, 1999), and MYH protein that excises adenine paired with 8oxoguanine in DNA (Yeh et al., 1991; Slupska et al., 1996). Oxidation of guanine proceeds also in the cellular nucleotide pool, and 8-oxo-dGTP, the oxidized form of dGTP, is the mutagenic substrate for DNA synthesis. It can be incorporated opposite adenine or cytosine residues of template DNA, the result being A : T to C : G and G : C to T : A transversions (Maki and Sekiguchi, 1992; Cheng et al., 1992). However, in normally growing cells, the frequency of these types of mutations remains low, owing to the action of enzymes degrading such mutagenic substrates (Schaaper et al., 1986; Mo et al., 1991). The MutT protein of E. coli hydrolyzes 8-oxo-dGTP to 8-oxo-dGMP, thereby preventing misincorporation of 8-oxoguanine into DNA (Maki and Sekiguchi, 1992). A similar enzyme Oxidative nucleotide damage M Sekiguchi and T Tsuzuki 8896 activity has been detected in mammalian cells, and the protein responsible was named MTH1 (Mo et al., 1992; Sakumi et al., 1993; Furuichi et al., 1994). It has been proposed that one early step in the progression of human cancers is elevation of the rate of spontaneous mutation, that is, acquisition of a mutator phenotype (Ionov et al., 1993; Jackson and Loeb, 1998; Stoler et al., 1999). If changes in spontaneous mutation rates are indeed involved in carcinogenesis, it is expected that defectiveness in the MTH1 or related genes would increase the likelihood of tumor occurrence. To address this question, targeted disruption of the MTH1 as well as the OGG1 gene has been done (Klungland et al., 1999; Minowa et al., 2000; Tsuzuki et al., 2001). These studies are significantly important in terms of multiple defence mechanisms against induction and progression of tumors in animal bodies. MutT-related error avoidance mechanism for DNA synthesis There are at least three steps required to prevent errors during DNA replication; (1) selection of a nucleotide complementary to the template by DNA polymerase, (2) removal of a misincorporated non-complementary nucleotide by an editing nuclease, associated with DNA polymerase, and (3) correction of a misincorporated nucleotide by the post-replicational mismatch repair system (Kornberg and Baker, 1992). Recently, another process for preventing replication errors was revealed. This error-avoiding mechanism functions at the pre-replicational step by degrading a naturally occurring mutagenic substrate for DNA synthesis (Maki and Sekiguchi, 1992). Here, MutT protein plays a major role while MutM and MutY participate in the process for providing a unique mutational specificity to the mutT7 mutant (Tajiri et al., 1995). Among the many mutators found in E. coli, mutT has drawn particular attention. MutT is one of the first mutators found in organisms (Treffers et al., 1954) and specifically induces A : T to C : G transversion mutations (Yanofsky et al., 1966). As a consequence of this unidirectional mutator activity, mutT7 cells have increased GC content in the chromosomal DNA (Cox and Yanofsky, 1967). Akiyama et al. (1987) cloned the mutT gene and, based on sequence analysis, identified a protein with 129 amino acid residues. The MutT protein was purified to physical homogeneity and was shown to have nucleoside triphosphatase activity (Bhatnagar and Bessman, 1988). Using an in vitro DNA synthesis system, Akiyama et al. (1989) demonstrated that the MutT protein specifically prevented misincorporation of dGMP onto the poly(dA)/oligo(dT)20 template-primer. Subsequently Maki and Sekiguchi (1992) found that the nucleotide misincorporated opposite the adenine residue of the template is not dGMP but rather its oxidized form, 8oxo-dGMP. Although the guanine base assumes almost exclusively anti conformation in DNA, in the case of 8oxoguanine, there exists almost the same quantity of Oncogene anti and syn forms, the former pairing with cytosine and the latter pairing with adenine. When 8-oxo-dGTP was added to an in vitro DNA replication system, 8oxo-dGMP was indeed incorporated opposite cytosine and adenine residues of the template, with almost equal frequencies. The MutT protein possesses enzyme activity which hydrolyzes 8-oxo-dGTP to 8-oxodGMP, thereby preventing the misincorporation of 8oxoguanine into DNA. Here, one must emphasize the importance of eliminating the oxidized form of guanine base from DNA. 8Oxoguanine was first described to be a minor modified base found in DNA treated with heated glucose (Kasai and Nishimura, 1984). This modified base was later revealed to be present in untreated DNA, albeit at levels no higher than approximately three molecules of 8oxoguanine/106 guanine residues in the chromosomal DNA (Tajiri et al., 1995). It thus seems that the level of reactive oxygen species produced by cellular metabolic intermediates may be sufficient to oxidize the guanine base of the nucleotide pool as well as that of DNA, even in normally growing cells. Spontaneous mutagenesis caused by the oxidation of guanine nucleotides can be separated into two pathways: one starting from oxidation of guanine in DNA and the other from oxidation of guanine nucleotide in the precursor pool (Figure 1). The oxidation of DNA forms the 8-oxoG : C pair which would induce a G : C?T : A transversion if the 8-oxoG is not corrected by MutM (Fpg) protein. When 8-oxoG remains unrepaired until DNA polymerase arrives at the lesion, dAMP would be inserted opposite the mutagenic lesion. However, in most cases, the resulting 8-oxoG : A pair is reversed back to 8-oxoG : C by the action MutY protein which removes the adenine base from the 8oxoG : A mispair. These dual defence mechanisms against mutagenesis were first proposed by Michaels et al. (1992), based on findings that the combination of mutM and mutY mutations resulted in synergistic mutator effects specific for G : C?T : A transversion and that a multi-copy plasmid carrying the mutM gene suppressed the mutator effect of the mutY7 mutant. The high rate of G : C to T : A mutation in the mutM mutY double mutator strain likely reflects the high level of 8-oxoG content in the DNA (Tajiri et al., 1995). While the mutation caused by the oxidation of DNA is unidirectional, incorporation of 8-oxo-dTP into DNA would result in base substitutions in two directions. In E. coli cells, the MutT protein seems to prevent both A : T to C : G and G : C to T : A transversions by eliminating 8-oxo-dGTP from the nucleotide pool. For the spontaneous mutagenesis caused by 8-oxo-dGTP, MutM and MutY proteins also have a role, but their functions in the A : T to C : G pathway differ from those in the G : C to T : A pathway. The repair enzymes, especially the MutY protein, promote the fixation of A : T to C : G transversion whereas the MutM and MutY proteins cooperate to suppress the G : C to T : A transversion, in the same manner as they do for oxidative DNA, as illustrated in Figure 1c. Oxidative nucleotide damage M Sekiguchi and T Tsuzuki 8897 Figure 1 8-Oxoguanine-related mutagenesis. (a) Base substitution mutations, classified into transition and transversion according to orientations of the base changes. Among two types of transition and four types of transversion mutations, mutT deficiency causes A : T?C : G transversion while deficiencies in mutM and mutY induce G : C?T : A transversion. (b) Distribution of types of mutations, as measured in the rpsL gene of E. coli (Tajiri et al., 1995). Blue bars represent G : C?T : A transversion and yellow bars represent A : T?C : G transversion. All six types of base substitutions appearing though the mutation frequency of wild-type cells is 100 times less than those for mutant strains (Mo et al., 1991). (c) 8-Oxoguanine-related mutagenesis pathways leading to induction of specific types of mutations. The guanine residue in DNA as well as dGTP in the nucleotide pool can be oxidized to 8-oxoguanine. As shown in the left side surrounded by a blue line, 8-oxoguanine produced in DNA induces G : C?T : A transversion, which is prevented by the actions of MutM and MutY proteins. Incorporation of 8-oxoguanine-containing nucleotide into DNA could induce A : T?C : G and G : C?T : A transversions, both of which can be prevented by MutT protein, by hydrolyzing 8oxo-dGTP. In mutT7 mutant cells, occurrence of G : C?T : A transversion is specifically prevented by functions of MutM and MutY proteins, resulting in a preferential occurrence of A : T?C : G transversion, as surrounded by a yellow line Oncogene Oxidative nucleotide damage M Sekiguchi and T Tsuzuki 8898 Mammalian MTH1 protein with 8-oxo-dGTPase activity In search of an activity similar to E. coli MutT in mammalian cells, Jurkat cells (a human T-cell leukemia cell line) were found to contain a high level of such activity. Taking advantage of this high level of activity, the enzyme was purified to apparent physical homogeneity (Mo et al., 1992; Sakumi et al., 1993). Substrate specificity of the enzyme was examined using a-32Plabeled dNTPs. Although dGTP and dATP were also hydrolyzed to the corresponding nucleoside monophosphates, product yields were about 5% of those with 8oxo-dGTP. Neither TTP nor dCTP was hydrolyzed by the enzyme. The apparent Km for hydrolysis of 8-oxodGTP was 70 times lower than that for the degradation of dGTP, whereas the maximal reaction rates observed with both substrates were similar. Based on the partial amino acid sequence determined with the purified human 8-oxo-dGTPase protein, a cDNA for human enzyme was cloned and the nucleotide sequence determined (Sakumi et al., 1993). The molecular mass of the protein, as calculated from the predicted amino acid sequence, was 17.9 kDa, a value close to that estimated from analysis of SDS – PAGE. By placing the cDNA under control of the lac promoter, a high level of 8-oxo-dGTPase activity was induced in the presence of isopropyl-1-thio-b-Dgalactopyranoside. On expression of cDNA in E. coli mutT7 cells the increased level of spontaneous mutation frequency was considerably reduced. Similar but more striking suppressive effects were observed when mouse or rat cDNA was expressed in the mutT7 cells (Kakuma et al., 1995; Cai et al., 1995). Thus, mammalian 8-oxo-dGTPase functions in E. coli cells to prevent mutations caused by the accumulation of 8oxo-dGTP in the nucleotide pool. The mammalian gene for 8-oxo-dGTPase has been named MTH1 for mutT homolog 1. Transfection of human MTH1 cDNA brought about a significant reduction in 8-oxoguanine content of DNA in mouse embryonic fibroblasts as well as in tumor cells, with or without H2O2 treatment (Colussi et al., 2002). As MTH1 protein decreases both steady-state and oxidant-induced 8-oxoguanine levels in DNA, endogenous oxidation of the deoxynucleotide pool is a definite source of DNA damage and the deoxynucleotide pool is a significant target for exogenous oxidative damage. To elucidate structure of the gene, human genomic libraries were screened using cDNA as a probe (Furuichi et al., 1994). The gene which spans approximately 8 kb consists of five exons, and the coding sequence resides on exons 2, 3, 4 and 5 (Oda et al., 1997, 1999). As will be described below, some of the exons are composed of differentially processed segments. Using fragments of genomic DNA as probes, chromosal assignment of the gene was made. These fragments derived from different regions, consistently hybridized to a single locus on the short arm of chromosome 7 at p22. Thus, the MTH1 gene is located on human chromosome 7p22 (Furuichi et al., 1994). Oncogene Northern blot analysis revealed that MTH1 mRNA is present in almost all tissues of adult mouse, among which expression levels are exceedingly high in thymus and testis (Oda et al., 1997). Fetal tissues (brain and liver) contained large amounts of MTH1 mRNA. In peripheral blood lymphocytes, the level of MTH1 mRNA was significantly increased after concomitant treatment with phytohemagglutinin and interleukin-2. Analyses of the 5’ regions of the MTH1 transcripts revealed that seven types of MTH1 mRNA, which may be produced by transcription initiation at different sites and/or alternative splicing (Figure 2). In most or all of human cells and tissues, type 1 mRNA which lacks the exon 2 sequence is present as a major form. More than one ATG initiation codons in-frame were found in the 5’ regions of some of the MTH1 mRNAs and, moreover, there is a polymorphic alteration at the 5’ splicing site located in exon 2. Thus, regulation of expression of the human MTH1 gene is complex and Figure 2 Genomic structure and alternative splicing of the human MTH1 gene (Oda et al., 1997). In the upper part, the overall structure of the gene is shown, in which exons 1 to 5 are indicated by filled boxes. In the lower part, seven types of MTH1 mRNA are shown, which can be produced by transcription initiation at different sites and alternative splicing. Hatched boxes represent exons Oxidative nucleotide damage M Sekiguchi and T Tsuzuki 8899 may be subjected to cell type-specific modification (Oda et al., 1999). In eukaryotic cells, a pool of dNTP for nuclear DNA replication is present mainly in the cytosol (Bestwick et al., 1982). Mitochondria, which preserve a pool of dNTP for mitochondrial DNA synthesis, consist of more than 10% of the total intracellular dNTP. The mitochondrial respiratory chain located on inner membranes is a major site for the initiation of lipid peroxidation, which can lead to oxidation of the guanine to 8-oxoguanine. In addition, the mitochondrial respiratory chain produces superoxide, which can be converted to hydroxyl radicals via hydrogen peroxide. The hydroxyl radical is the main species of reactive oxygen that attacks the guanine base. Thus, DNA and dNTP in the mitochondrial pool may be exposed to a greater oxidative stress than is the case in the nucleus. In this context, the intracellular location of 8-oxo-dGTPase was determined (Kang et al., 1995). In human Jurkat cells, most of the enzyme activity was found in cytosolic and the mitochondrial fractions. The specific activity of 8-oxo-dGTPase in the mitochondrial fraction was about 17% of that in the cytosolic fraction; almost no enzyme activity was found in nuclear and microsomal fractions. Electron microscopic immunocytochemistry, using a specific antibody against MTH1 protein, showed that MTH1 protein was localized in the mitochondrial matrix. An identical molecular form of MTH1 protein appears to be present in the cytosol and in the mitochondria. Structural features of MTH1-related proteins Human MTH1 and E. coli MutT proteins are similar in size and there is a certain degree of sequence homology in these proteins. Genes for analogous functions were isolated from Proteus vulgaris and Streptococcus pneumoniae, bacteria distantly related to E. coli (Kamath and Yanofsky, 1993; Bullions et al., 1994). The products of the latter two genes carry enzyme activity specifically degrading dGTP to dGMP and are structurally and functionally related to the E. coli MutT protein. Most of the identical residues are in a region corresponding to the 23 residues from Gly37 to Gly59 of E. coli MutT, known as the MutT signature (Bessman et al., 1996). Homologs of human MTH1 protein were identified in mice and rats, based on isolated cDNAs (Kakuma et al., 1995; Cai et al., 1995). Both proteins comprise 156 amino acid residues, as was the case for the human MTH1 protein, and amino acid sequences are highly conserved. Alignment of the sequences of these six proteins shows that all carry a highly conserved sequence in nearly the same region, corresponding to amino acids 36 to 58 for human MTH1 (Figure 3a). Ten of 23 amino acid residues in this region are identical, hence this probably constitutes an active center for the enzyme. The secondary structure of MutT protein has been determined by heteronuclear multi-dimensional NMR (Abeygunawardana et al., 1993, 1995; Weber et al., 1993, Frick et al., 1995) and the conserved region was found to consist of a loop – helix – loop motif which may contribute to nucleotide binding and nucleotide pyrophosphohydrolase activity. NMR studies were further extended to determine the roles of conserved amino acid residues in the possible active site of the MutT protein, in conjunction with site-directed mutagenesis analyses (Lin et al., 1996, 1997). Recently, NMR analysis of human MTH1 protein has been made (Mishima et al., unpublished result). Structure of MutT protein, both free and Mn-associated forms, was resolved by X-ray crystallographic analysis (Yamagata et al., unpublished result). The 23-residue sequence is a sole conserved sequence among all MutT and MTH1 homologs with 8-oxodGTPase, and of the many other proteins with the MutT signature so far identified, some hydrolysing various nucleotide derivatives, such as dATP, diadenosine oligophosphates, NADH, ADP-ribose, and GDP-mannose (O’Handley et al., 1996, 1998; Sheikh et al., 1998). Furthermore, a diphosphoinositol polyphosphate phosphohydrolase that hydrolyzes a nonrelated polyphosphate also contains the 23-residue (Safrany et al., 1998). A chimeric protein MTH1-Ec, in which the 23-residue sequence of MTH1 was replaced with that of MutT, retains its capacity to hydrolyze 8-oxo-dGTP (Fujii et al., 1999), thereby indicating that the 23-residue sequences of MTH1 and MutT are functionally and structurally equivalent and constitute functional modules (Figure 3b). Among these 23 residues in the putative active site region, 10 amino acids are conserved through bacteria to human. To establish the functional significance of these residues, saturation mutagenesis of the 10 conserved residues of MutT and MTH1 proteins were done, in combination with negative and positive mutant screening, using mutagenesis primers that contain a completely degenerated codon for each residue (Figure 3c). Shimokawa et al. (2000) found that Gly37, Gly38, Glu44, Arg52, Glu53, and Glu57 in MutT protein could not be replaced by any other residue without losing activity. Replacement of the corresponding residues of MTH1 protein also caused loss of function (Fujii et al., 1999). In the latter case two additional residues were found to be essential, though there is the possibility that formation of the human protein in heterologous cell (E. coli) might cause a more severe effect on expression of the function. It seems that these essential amino acid residues interact with the substrate nucleotide and Mg++ to exert catalytic functions. Tumorigenesis and mutagenesis in mice lacking MTH1 To obtain insight into the role of MTH1 protein in terms of spontaneous tumorigenesis as well as mutagenesis caused by the oxygen-induced DNA damage, it is necessary to establish cell and mouse lines defective in the MTH1 gene. This was achieved recently using gene targeting techniques. Oncogene Oxidative nucleotide damage M Sekiguchi and T Tsuzuki 8900 Figure 3 Comparison of structures of MutT family proteins. (a) The 23-residue modules of MutT-related proteins from various organisms, as shown by hatched boxes. Conserved amino acid residues are indicated by bold letters. (b) Secondary structures of E. coli MutT, human MTH1 and a chimeric protein MTH1-Ec. In the chimeric MTH1 protein, the 23-residue module is replaced with that of E. coli MutT protein. (c) Targeted mutagenesis analyses of E. coli MutT and human MTH1 protein. Amino acid residues conserved through the six MutT-related proteins are indicated by bold letters and those that could not be replaced by any other residue without losing function are placed in the yellow boxes. The data were taken from Fujii et al. (1999) and Shimokawa et al. (2000) The mouse MTH1 gene is composed of five exons and spans about 10 kb (Igarashi et al., 1997). Using the isogenic genomic DNA fragment, the entire area of the third exon containing the initiation codon and the adjacent intron regions were replaced with a neo cassette (Tsuzuki et al., 2001). The resulting construct was electroporated into ES cells, and cells showing resistance to both G418 and ganciclovir were selected to obtain MTH1+/7 cells. MTH17/7 cells were generated from the MTH1+/7 cells by growing in the presence of 1.5 mg/ml of G418, a concentration which is six times higher than that used for the isolation of MTH1+/7 cells. MTH17/7 cells thus obtained were used to examine the effect of MTH1 deficiency on spontaneous mutagenesis. Mutations in the Hprt gene, located on the X chromosome in the mouse genome, render cells resistance to 6-thioguanine and, in this forward mutation assay, two independently isolated MTH17/7 cell lines exhibited an approximately twofold higher mutation rate, compared with the value of MTH1+/+ cells. Thus, MTH1 may have the potential to prevent the occurrence of mutations under normal growth conditions. It should be noted, however, that degree of increase in spontaneous mutation frequency, due to the loss of Oncogene MutT-related functions, differs considerably in mouse and E. coli cells. Increases in Hprt mutations detected in mouse MTH17/7 cells is twofold compared with that in MTH1+/+ cells, whereas the value of mutation frequency for E. coli mutT7 cells is 100 times over that in wild-type cells (Tajiri et al., 1995). Several hypotheses to explain this difference may be considered, among which the most plausible is that mammalian cells may possess an enzyme(s) capable of degrading 8-oxo-dGTP in addition to MTH1. This notion was strengthened by the recent finding that even E. coli, despite its predominant MutT activity, possesses an additional enzyme activity that degrades 8-oxo-dGTP. GTP cyclohydrase II, encoded by the ribA gene of E. coli, can hydrolyze 8-oxo-dGTP to the corresponding nucleoside monophosphate. In the mutT7 background, ribA7 cells showed two times higher spontaneous mutation frequencies as compared with ribA+ cells (Kobayashi et al., 1998). It is unlikely that a GTP cyclohydrase II-type enzyme is present in mammalian cells because the biosynthesis of riboflavin, in which this type of enzyme functions, does not take place in animals. Recently, a mouse cDNA clone that considerably suppresses the high mutability of E. coli mutT7 cells was isolated (Cai et al., unpublished result). The protein encoded by this cDNA carried Oxidative nucleotide damage M Sekiguchi and T Tsuzuki 8901 activity to degrade 8-oxo-dGTP, and could act as an MTH1 redundancy factor. MTH1 homozygous mutant mice were generated by microinjecting MTH1+/7 ES cell clones into blastocysts and subsequently crossing the resulting chimera and heterozygous mice. MTH17/7 mice have a normal physical appearance, but have a high susceptibility for spontaneous tumorigenesis (Tsuzuki et al., 2001). Around 18 months after birth, many tumors were found in lungs and livers of MTH17/7 mice, but there were few in MTH1+/+ mice. The elevated incidence of tumor formation in the liver of MTH17/7 mice correlated well with the highest content of MTH1 protein in this organ of the wild-type mouse (Kakuma et al., 1995). As generally observed in spontaneous and carcinogen-induced hepatocarcinogenesis in rodents (Poole and Drinkwater, 1996), there is a high susceptibility of male mice to liver tumorigenic events, compared with their female counterparts. It has been suggested that the hormonal environment of the host affects the development of many types of tumors, especially those in the liver. More tumors tend to form in the stomach of MTH17/7 mice, as compared with MTH1+/+ mice, although the statistical significance is weak. Indeed, as there are different profiles of antioxidant enzyme, such as superoxide dismutases and catalases, different susceptibility to tumorigenesis among organs may reflect the metabolic balance of oxidative stress, inducing lesions in DNA as well as in the nucleotide pool. Altogether, it can be concluded that the intracellular level of MTH1 protein is an important factor in determining susceptibility of mice to tumor induction by endogenous oxidative damage. To examine in vivo mutation events due to the MTH1-deficiency, a reporter gene, rpsL of E. coli, was introduced into MTH17/7 mice (Egashira et al., 2002). Interestingly, the net frequency of rpsL7 forward mutants showed no apparent increase in MTH17/7 mice as compared to findings in MTH1+/+ mice. However, there are differences between these two genotypes in class- and site-distributions of the rpsL7 mutations recovered from the mice. Unlike MutTdeficient E. coli, an increase in frequency of A : T?C : G transversion was not evident in MTH1 nullizygous mice. Nevertheless, the frequency of singlebase frameshifts at mononucleotide runs was 5.7-fold higher in spleens of MTH17/7 mice than in those of wild-type mice. Since the elevated incidence of singlebase frameshifts at mononucleotide runs is a hallmark of the defect in MSH2-dependent mismatch repair system, this weak site-specific mutator effect of MTH17/7 mice could be attributed to a partial sequestration of the mismatch repair function that may act to correct mispairs with the oxidized nucleotides. Studies of OGG1-defective mice would provide much more insight into the matter. OGG17/7 mice accumulate abnormally high levels of 8-oxoguanine in their genomes and exhibit a moderately elevated spontaneous mutation rates in their tissues (Klungland et al., 1999; Minowa et al., 2000). Nevertheless, there was no sign of development of malignancy during about 12 months after birth. Alternative mechanisms might function to minimize the effects of an increased load of 8-oxoguanine in mammalian genomes. Metabolism of oxidized nucleotides Figure 4 summarizes enzymatic reactions for interconversion of 8-oxoguanine-containing nucleotides in mammalian cells. Enzymatic conversion of ribonucleotides to deoxyribonucleotides occurs at the level of nucleoside diphosphate, and ribonucleotide reductase, the enzyme responsible, has a relatively broad substrate specificity. Four types of naturally occurring ribonucleotides, ADP, GDP, CDP and UDP, are converted to the corresponding deoxyribonucleotides by a single species of reductase enzyme (Thelander and Reichard, 1979). However, this enzyme is inactive on the 8oxoguanine-containing nucleotide, as revealed with mouse ribonucleotide reductase (Hayakawa et al., 1999). This implies that 8-oxoguanine-containing deoxyribonucleotides must be generated in the cellular deoxyribonucleotide pool. Human cells contain nucleoside diphosphate kinase, an enzyme activity which phosphorylates various nucleoside diphosphates to the corresponding nucleoside triphosphates (Kornberg and Baker, 1992). This enzyme can convert 8-oxo-dGDP to 8-oxo-dGTP, although the rate of phosphorylation of 8-oxo-dGDP was one-third that of dGDP (Hayakawa et al., 1995). Thus, in addition to direct oxidation of dGTP, 8-oxodGTP may be generated by the phosphorylation of 8oxo-dGDP. Once 8-oxo-dGTP is produced, this can be incorporated into DNA. Various DNA polymerases from eukaryotes and prokaryotes have the potential to utilize 8-oxo-dGTP as substrate (Cheng et al., 1992; Maki and Sekiguchi, 1992; Pavlov et al., 1994; Minnick et al., 1994). Thus, the action of 8-oxo-dGTPase is a pre-requisite for high fidelity of DNA replication. 8-Oxo-dGMP, produced by the action of 8-oxodGTPase, cannot be re-phosphorylated by cellular enzymes. Human guanylate kinase, which phosphorylates both GMP and dGMP to the corresponding nucleoside diphosphates, is totally inactive for 8-oxodGMP (Hayakawa et al., 1995). This would provide another basis for excluding this mutagenic substrate from the DNA precursor. 8-Oxo-dGMP is dephosphorylated to yield the corresponding nucleoside, 8-oxodeoxyguanosine. Nucleosides are readily transported through the cell membrane, and extracellular nucleosides can be excreted into the urine. Dephosphorylation of 8-oxodGMP, therefore, may be an essential step for excretion of 8-oxoguanine-containing compounds. The enzyme that catalyzes this reaction, 8-oxo-dGMPase, was partially purified from an extract of human Jurkat cells, and the mode of action was elucidated (Hayakawa et al., 1995). 8-Oxo-dGMP is the preferred substrate of the enzyme, and other nucleoside monoOncogene Oxidative nucleotide damage M Sekiguchi and T Tsuzuki 8902 Figure 4 Interconversion of 8-oxoguanine-containing nucleotides in mammalian cells. This scheme is based on the results of Hayakawa et al. (1995, 1999). (1) Ribonucleotide reductase; (2) nucleoside diphosphate kinase; (3) 8-oxo-dGTPase (MTH1); (4) guanylate kinase; (5) DNA polymerase; (6) 8-oxo-dGMPase. O. denotes oxidative reaction phosphates are cleaved albeit at significantly lower rates. As discussed above, MTH1 and MutT are key enzymes for sustaining high fidelity of DNA replication, in oxygenated stated of living systems. These enzymes of diverged origins share a common character to degrade 8-oxo-dGTP to 8-oxo-dGMP, thereby eliminating the mutagenic substrate from the DNA precursor pool. In spite of this common feature, they exhibit certain significant differences. First, MTH1 and MutT differ in the capacity to degrade 8-oxoguaninecontaining ribonucleotides. E. coli MutT protein cleaves 8-oxoGTP as efficiently as does 8-oxo-dGTP (Taddei et al., 1997). On the other hand, human MTH1 is hardly capable of hydrolyzing 8-oxoGTP; its potential to degrade 8-oxoGTP is only 2% that for 8oxo-dGTP (Hayakawa et al., 1999). These apparent differences in efficiency of degrading an error-evoking substrate may be counterbalanced by the capacity of RNA polymerases to discriminate the unfavorable substrate. E. coli RNA polymerase can utilize 8oxoGTP as substrate at a rate 10% that of GTP whereas mammalian RNA polymerase II incorporates little of 8-oxoguanine into RNA (Hayakawa et al., 1999). The second point is that the mammalian enzyme has a broader substrate specificity as compared with the bacterial enzyme. MTH1 hydrolyzes 8-oxo-dATP, 2hydroxy-dATP and 2-hydroxyATP as well as 8-oxodGTP (Sakai et al., 2002). This is in contrast with MutT, which acts on 8-oxoguanine-containing nucleotides alone. Recognition of 8-oxo-dGTP and 2hydroxy-dATP appears to depend on different residues of MTH1 protein. Transgenic mice expressing engineered MTH1 proteins lacking each one of the activities would be useful to reveal the biological significance of these two activities. Acknowledgements We thank Drs H Hayakawa, H Maki, Y Nakabeppu, Y Yamagata, Y Nakatsu and H Shimokawa for discussion and helpful advice, and M Ohara for useful comments on the manuscript. References Abeygunawardana C, Weber D, Frick DN, Bessman MJ and Mildvan AS. (1993). Biochemistry, 32, 13071 – 13080. Abeygunawardana C, Weber D, Gittis AG, Frick DN, Lin J, Miller A-F, Bessman MJ and Mildvan AS. (1995). Biochemistry, 34, 14997 – 15005. Oncogene Akiyama M, Horiuchi T and Sekiguchi M. (1987). Mol. Gen. Genet., 206, 9 – 16. Akiyama M, Maki H, Sekiguchi M and Horiuchi T. (1989). Proc. Natl. Acad. Sci. USA, 86, 3949 – 3952. Ames BN and Gold LS. (1991). Mutat. Res., 250, 3 – 16. Oxidative nucleotide damage M Sekiguchi and T Tsuzuki 8903 Au KG, Cabrera M, Miller JH and Modrich P. (1988). Proc. Natl. Acad. Sci. USA, 85, 9163 – 9166. Au KG, Clark S, Miller JH and Modrich P. (1989). Proc. Natl. Acad. Sci. USA, 86, 8877 – 8881. Bessho T, Tano K, Kasai H and Nishimura S. (1992). Biochem. Biophys. Res. Commun., 188, 372 – 378. Bessman MJ, Frick DN and O’Handley SF. (1996). J. Biol. Chem., 271, 25059 – 25062. Bestwick RK, Moffett GM and Mathews CK. (1982). J. Biol. Chem., 257, 9300 – 9304. Bhatnagar SK and Bessman MJ. (1988). J. Biol. Chem., 263, 8953 – 8957. Boiteux S and Radicella JP. (1999). Biochimie, 81, 59 – 67. Brooks PJ, Wise DS, Berry DA, Kosmoski JV, Smerdon MJ, Somers RL, Mackie H, Spoonde AY, Ackerman EJ, Coleman K, Tarone RE and Robbins JH. (2000). J. Biol. Chem., 275, 22355 – 22362. Bullions LC, Mejean V, Claverys JP and Bessman MJ. (1994). J. Biol. Chem., 269, 12339 – 12344. Cabrera M, Nghiem Y and Miller JH. (1988). J. Bacteriol., 170, 5405 – 5407. Cai JP, Kakuma T, Tsuzuki T and Sekiguchi M. (1995). Carcinogenesis, 16, 2343 – 2350. Cheng KC, Cahill DS, Kasai H, Nishimura S and Loeb LA. (1992). J. Biol. Chem., 267, 166 – 172. Chung MH, Kasai H, Jones DS, Inoue H, Ishikawa H, Ohtsuka E and Nishimura S. (1991). Mutat. Res., 254, 1 – 12. Colussi C, Parlanti E, Degan P, Aquilina G, Barnes D, Macpherson P, Karran P, Crescenzi M, Dogliotti E and Bignami M. (2002). Current Biol., 12, 912 – 918. Cox EC and Yanofsky C. (1967). Proc. Natl. Acad. Sci. USA, 58, 1895 – 1902. Demple B and Harrison L. (1994). Annu. Rev. Biochem., 63, 915 – 948. Egashira A, Yamauchi K, Yoshiyama K, Kawate H, Katsuki M, Sekiguchi M, Sugimachi K, Maki H and Tsuzuki T. (2002). DNA Repair, 75, 1 – 13. Evans J, Maccabee M, Hatahet Z, Courcelle J, Bockrath R, Ide H and Wallace S. (1993). Mutat. Res., 299, 147 – 156. Fraga CG, Shigenaga MK, Park JW, Degan P and Ames BN. (1990). Proc. Natl. Acad. Sci. USA, 87, 4533 – 4537. Frick DN, Weber DJ, Abeygunawardana C, Gittis AG, Bessman MJ and Mildvan AS. (1995). Biochemistry, 34, 5577 – 5586. Fujii Y, Shimokawa H, Sekiguchi M and Nakabeppu Y. (1999). J. Biol. Chem., 274, 38251 – 38259. Furuichi M, Yoshida MC, Oda H, Tajiri T, Nakabeppu Y, Tsuzuki T and Sekiguchi M. (1994). Genomics, 24, 485 – 490. Gajewski E, Rao G, Nackerdien Z and Dizdaroglu M. (1990). Biochemistry, 29, 7876 – 7882. Hayakawa H, Hofer A, Thelander L, Kitajima S, Cai Y, Oshiro S, Yakushiji H, Nakabeppu Y, Kuwano M and Sekiguchi M. (1999). Biochemistry, 38, 3610 – 3614. Hayakawa H, Taketomi A, Sakumi K, Kuwano M and Sekiguchi M. (1995). Biochemistry, 34, 89 – 95. Henle ES and Linn S. (1997). J. Biol. Chem., 272, 19095 – 19098. Igarashi H, Tsuzuki T, Kakuma T, Tominaga Y and Sekiguchi M. (1997). J. Biol. Chem., 272, 3766 – 3772. Ionov Y, Peinado MA, Malkhosyan S, Shibata D and Perucho M. (1993). Nature, 363, 558 – 561. Jackson AL and Loeb LA. (1998). Genetics, 148, 1483 – 1490. Kakuma T, Nishida J, Tsuzuki T and Sekiguchi M. (1995). J. Biol. Chem., 270, 25942 – 25948. Kang D, Nishida J, Iyama A, Nakabeppu Y, Furuichi M, Fujiwara T, Sekiguchi M and Takeshige K. (1995). J. Biol. Chem., 270, 14659 – 14665. Kamath AV and Yanofsky C. (1993). Gene, 134, 99 – 102. Kasai H and Nishimura S. (1984). Nucleic Acids Res., 12, 2137 – 2145. Klungland A, Rosewell I, Hollenbach S, Larsen E, Daly G, Epe B, Seeberg E, Lindahl T and Barns DE. (1999). Proc. Natl. Acad. Sci. USA, 96, 13300 – 13305. Kobayashi M, Ohara-Nemoto Y, Kaneko M, Hayakawa H, Sekiguchi M and Yamamoto K. (1998). J. Biol. Chem., 273, 26394 – 26399. Kornberg A and Baker TA. (1992). DNA Replication 2nd edn. New York: W. H. Freeman and Company,. Kuraoka I, Bender C, Romieu A, Cadet J, Wood RD and Lindahl T. (2000). Proc. Natl. Acad. Sci. USA, 97, 3832 – 3837. Lin J, Abeygunawardana C, Frick DN, Bessman MJ and Mildvan AS. (1996). Biochemistry, 35, 6715 – 6726. Lin J, Abeygunawardana C, Frick DN, Bessman MJ and Mildvan AS. (1997). Biochemistry, 36, 1199 – 1211. Maki H and Sekiguchi M. (1992). Nature, 355, 273 – 275. Michaels ML, Pham L, Cruz C and Miller JH. (1991). Nucleic Acids Res., 19, 3629 – 3632. Michaels ML, Cruz C, Grollman AP and Miller JH. (1992). Proc. Natl. Acad. Sci. USA, 89, 7022 – 7025. Minowa O, Arai T, Hirano M, Monden Y, Nakai S, Fukuda M, Itoh M, Takano H, Hippou Y, Aburatani H, Masumura K, Nohmi T, Nishimura S and Noda T. (2000). Proc. Natl. Acad. Sci. USA, 97, 4156 – 4161. Minnick DT, Pavlov YI and Kunkel TA. (1994). Nucleic Acids Res., 22, 5658 – 5664. Mo JY, Maki H and Sekiguchi M. (1991). J. Mol. Biol., 222, 925 – 936. Mo JY, Maki H and Sekiguchi M. (1992). Proc. Natl. Acad. Sci. USA, 89, 11021 – 11025. Nash HM, Bruner SD, Schärer OD, Kawate T, Addona TA, Spooner E, Lane WS and Verdine GL. (1996). Curr. Biol., 6, 968 – 980. Nghiem Y, Cabrera M, Cupples CG and Miller JH. (1988). Proc. Natl. Acad. Sci. USA, 85, 2709 – 2713. Oda H, Nakabeppu Y, Furuichi M and Sekiguchi M. (1997). J. Biol. Chem., 272, 17843 – 17850. Oda H, Taketomi A, Maruyama I, Ito R, Nishioka K, Yakushiji H, Suzuki T, Sekiguchi M and Nakabeppu Y. (1999). Nucleic Acids Res., 27, 4335 – 4343. O’Handley SF, Frick DN, Bullions LC, Mildvan AS and Bessman MJ. (1996). J. Biol. Chem., 271, 24649 – 24654. O’Handley SF, Frick DN, Dunn CA and Bessman MJ. (1998). J. Biol. Chem., 273, 3192 – 3197. Ohtsubo T, Matsuda O, Iba K, Terashima I, Sekiguchi M and Nakabeppu Y. (1998). Mol. Gen. Genet., 259, 577 – 590. Poole TM and Drinkwater NR. (1996). Carcinogenesis, 17, 191 – 196. Pavlov YI, Minnick DT, Izuta S and Kunkel TA. (1994). Biochemistry, 33, 4695 – 4701. Safrany ST, Caffrey JJ, Yang X, Bembenek ME, Moyer MB, Burkhart WA and Shears SB. (1998). EMBO J., 17, 6599 – 6607. Sakai Y, Furuichi M, Takahashi M, Mishima M, Iwai S, Shirakawa M and Nakabeppu Y. (2002). J. Biol. Chem., 277, 8579 – 8587. Sakumi K, Furuichi M, Tsuzuki T, Kakuma T, Kawabata S, Maki H and Sekiguchi M. (1993). J. Biol. Chem., 268, 23524 – 23530. Oncogene Oxidative nucleotide damage M Sekiguchi and T Tsuzuki 8904 Schaaper RM, Danforth BN and Glickman BW. (1986). J. Mol. Biol., 189, 273 – 284. Sheikh S, O’Handley SF, Dunn CA and Bessman MJ. (1998). J. Biol. Chem., 273, 20924 – 20928. Shibutani S, Takeshita M and Grollman AP. (1991). Nature, 349, 431 – 434. Shimokawa H, Fujii Y, Furuichi M, Sekiguchi M and Nakabeppu Y. (2000). Nucleic Acids Res., 28, 3240 – 3249. Slupska MM, Baikalov C, Luther WM, Chiang JH, Wei YF and Miller JH. (1996). J. Bacteriol., 178, 3885 – 3892. Smith KC. (1992). Mutat. Res., 277, 139 – 162. Stoler DL, Chen N, Basik M, Kahlenberg MS, RodriguezBigas MA, Petrelli NJ and Anderson GR. (1999). Proc. Natl. Acad. Sci. USA, 96, 15121 – 15126. Taddei F, Hayakawa H, Bouton M, Cirinesi A, Matic I, Sekiguchi M and Radman M. (1997). Science, 278, 128 – 130. Tajiri T, Maki H and Sekiguchi M. (1995). Mutat. Res., 336, 257 – 267. Oncogene Tchou J and Grollman AP. (1993). Mutat. Res., 299, 277 – 287. Thelander L and Reichard P. (1979). Annu. Rev. Biochem., 48, 133 – 158. Treffers HP, Spinelli V and Belser NO. (1954). Proc. Natl. Acad. Sci. USA, 40, 1064 – 1071. Tsuzuki T, Egashira A, Igarashi H, Iwakuma T, Nakatsuru Y, Tominaga Y, Kawate H, Nakao K, Nakamura K, Ide F, Kura S, Nakabeppu Y, Katsuki M, Ishikawa T and Sekiguchi M. (2001). Proc. Natl. Acad. Sci. USA, 98, 11456 – 11461. van der Kemp PA, Thomas D, Barbey R, de Oliveira R and Boiteux S. (1996). Proc. Natl. Acad. Sci. USA, 93, 5197 – 5202. Weber D, Abeygunawardana C, Bessman MJ and Mildvan AS. (1993). Biochemistry, 32, 13081 – 13088. Yanofsky C, Cox EC and Horn V. (1966). Proc. Natl. Acad. Sci. USA, 55, 274 – 281. Yeh YC, Chang DY, Masin J and Lu AL. (1991). J. Biol. Chem., 266, 6480 – 6484.