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Originally published In Press as doi:10.1074/jbc.M111269200 on December 26, 2001

J. Biol. Chem., Vol. 277, Issue 10, 8260-8266, March 8, 2002
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Activation of Human MutS Homologs by 8-Oxo-guanine DNA Damage*

Anthony Mazurek, Mark Berardini, and Richard FishelDagger

From the Genetics and Molecular Biology Program, Department of Microbiology and Immunology, Kimmel Cancer Center, Thomas Jefferson University, Philadelphia, Pennsylvania 19107

Received for publication, November 27, 2001, and in revised form, December 14, 2001

    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The DNA lesion 8-oxo-guanine (8-oxo-G) is a highly mutagenic product of the interaction between reactive oxygen species and DNA. To maintain genomic integrity, cells have evolved mechanisms capable of removing this frequently arising oxidative lesion. Mismatch repair (MMR) appears to be one pathway associated with the repair of 8-oxo-G lesions (DeWeese, T. L., Shipman, J. M., Larrier, N. A., Buckley, N. M., Kidd, L. R., Groopman, J. D., Cutler, R. G., te Riele, H., and Nelson, W. G. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 11915-11920; Ni, T. T., Marsischky, G. T., and Kolodner, R. D. (1999) Mol. Cell 4, 439-444). Here we report the effect of double-stranded DNA oligonucleotides containing a single 8-oxo-G on the DNA binding affinity, ATPase, and ADP right-arrow ATP exchange activities of hMSH2-hMSH6 and hMSH2-hMSH3. We found that hMSH2-hMSH6 binds the oligonucleotide DNA substrates with the following affinities: 8-oxo-G/T > 8-oxo-G/G > 8-oxo-G/A > 8-oxo-G/C approx  G/C. A similar trend was observed for DNA-stimulated ATPase and ADP right-arrow ATP exchange activities of hMSH2-hMSH6. In contrast, hMSH2-hMSH3 did not appear to bind any of the 8-oxo-G containing DNA substrates nor was there enhanced ATPase or ADP right-arrow ATP exchange activities. These results suggest that only hMSH2-hMSH6 is activated by recognition of 8-oxo-G lesions. Our data are consistent with the notion that post-replication MMR only participates in the repair of mismatched 8-oxo-G lesions.

    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Oxidative DNA damage presents a serious challenge to the cells ability to maintain genomic integrity. Such damage results from a variety of interactions between reactive oxygen species with DNA (for review, see Ref. 3). Sources of these reactive oxygen species have been well documented and include chronic inflammation, by-products of cellular metabolism, by-products of peroxisome activity, and a considerable number of environmental factors (4, 5). Although there are many forms of oxidative modifications to DNA, 8-oxo-guanine (8-oxo-G)1 represents one of the most abundant and widely studied lesions (6, 7). The replication DNA polymerases alpha  and delta  often incorporate adenine opposite 8-oxo-G (8). Following a subsequent round of DNA replication, the erroneous adenine on the template strand will be paired with thymine, resulting in a signature G to T transversion. In addition, 8-oxo-G that is available from nucleotide pools may be incorporated into DNA during replication (9).

Bacteria, yeast, and human cells contain overlapping and redundant 8-oxo-G recognition systems (for review, see Ref. 3). For example, Escherichia coli has at least four 8-oxo-G processing glycosylases: 1) the MutM/fpg glycosylase removes global genomic 8-oxo-G lesions except those that occur in the context of an 8-oxo-G/A mismatch; 2) the MutY glycosylase presumably removes the A opposite 8-oxo-G in the nascent DNA strand following replication; 3) the Nei(EndoVIII) glycosylase presumably removes 8-oxo-G opposite A in the nascent DNA strand following replication; and 4) MutT acts as an 8-oxo-GTPase that effectively removes oxidative damage from the nucleotide pools. Examination of the Saccharomyces cerevisiae genome sequence as well as biochemical studies have suggested MutM/fpg (designated OGG1 in yeast), and Nei(EndoVIII) (designated OGG-2/Ntg1 in yeast) are functionally conserved (10-12). Interestingly, there does not appear to be a MutY or a MutT homolog in yeast. In contrast, human cells appear to have conserved functional and sequence homologs of all the bacterial oxidative damage glycosylases (hOGG1, hOGG2), a human MutY or hMYH, and perhaps two human MutT homologs or hMTH (13-17).

DNA mismatches and insertion/deletion loops (IDL) which arise from errors during DNA replication represent another challenge to genomic integrity. Mismatch repair (MMR) is the predominant pathway for the repair of these misincorporation errors. The human MutS homologs hMSH2, hMSH3, and hMSH6 play a fundamental role in the recognition and signaling of MMR (for review, see Refs. 18-20). The hMSH2-hMSH6 heterodimer primarily recognizes DNA base/base mismatches and single nucleotide IDL mismatches (21). The hMSH2-hMSH3 heterodimer also recognizes single nucleotide IDLs as well as larger IDLs (21).

The eukaryotic MutS homologs have been extensively studied for their role in the recognition and repair of DNA mismatches and IDLs, but they also appear to recognize other abnormalities in DNA. For example, hMSH2-hMSH6 has been shown to bind DNA oligonucleotides containing cisplatin and O6-methylguanine lesions (22-24). Genetic and biochemical evidence also suggests a role for MSH2 and hMSH6 in the repair of 8-oxo-G lesions (25). Compared with wild type cells, Msh2-deficient mouse embryonic stem cells appeared resistant to prolonged low level ionizing radiation and accumulated a significantly greater number of 8-oxo-G in their genomic DNA (1). In addition, cell lines deficient for hMSH2 have been found to be deficient for transcription-coupled DNA repair (TCR) of oxidative lesions (26). Moreover, the combination of S. cerevisiae msh2 or msh6 (but not msh3) mutations with a mutation of ogg-1 resulted in a synergistic increase in G to T transversions (2). Consistent with these genetic results, purified scMsh2-scMsh6 was shown to bind DNA oligonucleotides containing a single 8-oxo-G lesion by gel-shift analysis and to be released from these substrates in the presence of ATP but not ADP (2). Taken together, these latter results suggest that MSH2 and MSH6 (presumably as a heterodimer) recognize and are activated by 8-oxo-G lesions.

The mechanism by which the mismatch repair heterodimers perform their role in the repair of DNA mismatches, as well as other abnormalities in DNA structure, continues to be an area of debate. A model that is consistent with the available biochemical and structural data suggests that both hMSH2-hMSH6 and hMSH2-hMSH3 heterodimers behave as molecular switches that are regulated by binding ADP or ATP (19, 20, 24, 27-31). In the ADP-bound form, rapid high affinity DNA binding is observed between the heterodimers and oligonucleotides containing a mismatch or an IDL. Recognition of the mismatch results in ADP release by a mechanism that appears similar to GDP release by guanine nucleotide exchange factors.2 In the presence of physiological levels of ATP, these MutS homologs undergo rapid ADP right-arrow ATP nucleotide exchange (24, 27-29, 31). As with G proteins undergoing GDP right-arrow GTP exchange, ADP right-arrow ATP exchange by MutS homologs is accompanied by a large conformational transition. In the case of MutS homologs this conformational transition results in the formation of a hydrolysis-independent sliding clamp associated with the duplex DNA adjacent to the mismatch (28, 31). Diffusion of the MutS homologs away from the mismatch allows the loading of multiple redundant sliding clamps that propagate a mismatch-recognition signal to downstream protein machinery such as the MutL homologs (MLH); ultimately culminating in a mismatch repair event. One of the fundamental predictions of this model is that bona fide mismatch and/or lesion recognition is determined by whether the abnormality elicits ADP right-arrow ATP nucleotide exchange. In addition, the ability of a mismatch and/or lesion to elicit nucleotide exchange is likely to be specific for the MSH heterodimers. For example, oligonucleotides containing an O6-methylguanine adduct were found to activate nucleotide exchange by hMSH2-hMSH6 but not hMSH2-hMSH3 (24). These biochemical observations were confirmed by the observation that hMSH2- or hMSH6-deficient cells were resistant to the O6-guanine alkylating agent MNNG, but hMSH3-deficient cells remained sensitive to MNNG (32). In addition to MNNG, MMR-deficient cell lines have been found to be resistant to cisplatin, 5-fluorouracil, and temozolomide (33-37). These and other results have suggested that the MMR proteins may also function as sensors connecting DNA damage to apoptosis (20, 30, 35).

In this study, we examined recognition of oligonucleotides containing a single 8-oxo-G lesion by hMSH2-hMSH6 and/or hMSH2-hMSH3. Three criteria were established for bona fide recognition and activation of MSHs by 8-oxo-G: 1) affinity for a DNA oligonucleotide substrate containing a well defined 8-oxo-G; 2) the ability to enhance ATPase activity; and 3) the ability to enhance ADP right-arrow ATP nucleotide exchange activity. We found that only the hMSH2-hMSH6 heterodimer was capable of being activated by oligonucleotides containing 8-oxo-G and only when they were mismatched with non-complementary nucleotides. Our results are consistent with the notion of overlapping and redundant repair systems for oxidative damage.

    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Protein Purification-- The overexpression and purification of hMSH2-hMSH6 was performed as previously described (27), with the exception that PBE chromatography was followed by Mono Q chromatography. Buffers used for this additional step were identical to those used for PBE purification, and hMSH2-hMSH6 eluted from the Mono Q resin at ~675 mM NaCl. Overexpression and purification of hMSH2-hMSH3 was also performed as previously described (29) except for the following modifications. The hMSH2-hMSH3 heterodimer was overexpressed in Hi5 insect cells (Invitrogen) and purified as previously described with the exception that an additional Mono Q chromatography step was performed following PBE/heparin-Sepharose (29). hMSH2-hMSH3 eluted at ~550 mM NaCl from Mono Q.

DNA Substrate Preparation-- All single-stranded 41-bp oligonucleotides were synthesized and purified by reverse phase high performance liquid chromatography (Midland, TX). Double stranded (ds) DNA substrates used for ATPase and ADP right-arrow ATP exchange assays were prepared as follows. Oligonucleotide substrates were annealed in equimolar concentrations in TNE (10 mM Tris, pH 8.1, 1 mM EDTA, 100 mM NaCl). Residual single-stranded (ss) DNA was purified away from dsDNA using benzoylated naphthoylated DEAE-cellulose (Sigma). DsDNA was then concentrated using a YM-10 microcon centrifugal filter (Amicon Bioseparations). Purified dsDNA substrates were stored at -20 °C in TNE. DsDNA substrates used for IAsys total internal reflectance (TIR) studies were prepared by annealing 20 µg of 3'-biotinylated 41-bp ssDNA oligonucleotides to 60 µg of their respective complimentary non-biotinylated 41-bp ssDNA oligonucleotides. The resultant ds/ss DNA mixture was stored at -20 °C in TNE.

IAsys Total Internal Reflectance-- TIR using the IAsys Auto Plus system (Affinity Sensors) was done to measure the binding of hMSH2-hMSH6 to DNA substrates. The IASYS Auto Plus microcuvettes (Affinity Sensors) contained reaction cells coated with biotin and were subsequently bound with excess streptavidin (Prozyme). Unbound streptavidin was removed from the cuvette with PBST buffer. 2 µg of the appropriate ds/ss-annealed DNA substrate containing a single 3'-biotin, was added to the streptavidin-coated reaction cells, and excess unbound ssDNA removed with phosphate-buffered saline + 0.05% Tween 20 (PBST). Reaction cells were equilibrated with 50 µl of B-buffer (25 mM Hepes pH 8.1, 100 mM NaCl, 2 mM MgCl2, 1 mM dithiothreitol) plus 2% glycerol. Purified recombinant hMSH2-hMSH6 was added to the equilibrated cells to final concentrations of 12.5, 25, 50, 75, 100, 125, 150, or 175 nM. Binding was allowed to continue until the proteins reached equilibrium (saturation) binding with the DNA. Following each experiment, the protein was removed from the chips and the DNA bound to the surface regenerated by washing with 3 M NaCl for 2 min. Grafit 4 software (Affinity Sensors) was used to calculate kinetic constants (kassociation, kdissociation, and KD).

ATPase-- ATPase experiments were performed as previously described (24). The assays were carried out in 20-µl reactions containing B-buffer plus 15% glycerol and 240 nM of the respective 41-bp dsDNA substrate. Unlabeled ATP was added into each reaction for final concentrations of 10, 15, 20, 30, 40, 80, 160, and 250 µM for hMSH2-hMSH6 studies and 0.5, 1, 2, 4, 8, 12, 16, 24, and 32 µM for hMSH2-hMSH3 studies. The final concentration of [gamma -32P]ATP for each reaction was fixed at 17 nM. The concentrations of purified recombinant heterodimers were varied (10-50 nM hMSH2-hMSH6 and 10-40 nM hMSH2-hMSH3) in the experiments such that no more than 20% of the total [gamma -32P]ATP was hydrolyzed. Assays were carried out for 30 min at 37 °C, stopped with 0.4 ml of 10% (w/v) charcoal, 1 mM EDTA and placed on ice for 30 min. The samples were centrifuged in a table top microcentrifuge at 14,000 × g for 10 min. 100 µl of the supernatant was collected and counted in a RackBeta 1209 liquid scintillation counter (Amersham Biosciences, Inc.). The data was then fitted to the Michaelis-Menton equation to generate values for Km and Vmax (kcat).

ADP-ATP Exchange-- ADP exchange experiments were performed in 20-µl reactions containing B-buffer plus 15% glycerol, 2.3 µM [3H]ADP, 60 nM of the respective 41-mer dsDNA substrate, and 60 nM of either hMSH2-hMSH3 or hMSH2-hMSH6. Purified recombinant heterodimers were preincubated with [3H]ADP for 15 min at 25 °C. The respective DNA substrate was added and the mixture incubated 5 min at 25 °C. These reactions were then placed on ice. To begin the ADP right-arrow ATP exchange, reactions were taken off ice and ATP was added for a final concentration of 25 µM. Each exchange reaction was allowed to proceed at 25 °C until the noted time points. Reactions were stopped by dilution to 4 ml with ice-cold B-buffer minus dithiothreitol and plus 15% glycerol and then immediate vacuum filtered through a prewet nitrocellulose filter (Millipore HAWP02500, 25 mm, 0.45 µm). The filters were washed with 4 ml of ice-cold B-buffer minus dithiothreitol and plus 15% glycerol. Each filter was allowed to dry at room temperature. 3 ml of scintillation fluid (Amersham Biosciences, Inc.) was added before counting. ATP was not added to samples used to determine 100% [3H]ADP bound to the heterodimers.

    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

IAsys Total Internal Reflectance-- Surface mass measurements were performed by examining TIR using IAsys (Affinity Sensors, Franklin, MA). The binding affinities of the hMSH2-hMSH6 heterodimer with dsDNA oligonucleotide substrates were determined for well defined dsDNA sequences (Fig. 1). Unlike surface mass measurements by surface plasmon resonance using Biacore (Amersham Biosciences, Inc.), TIR maintains continuous equilibrium in a closed cuvette that is then displayed as real-time binding isotherms. A closed system reduces the possibility of flow-driven non-equilibrium binding plates that may artificially induce a binding mass.


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  		Fig. 1.   Sequences of double-stranded oligonucleotide substrates tested. The 3' biotin modification on the noted DNA strands was added only for the DNA substrates used for total internal reflectance experiments. Differences in substrates are indicated in bold and the substrate designation indicates nucleotide pair at position 21.

IAsys binding studies with hMSH2-hMSH3 suggested that the affinity of this heterodimer for 8-oxo-G containing DNA was not significantly elevated when compared with homoduplex DNA (data not shown). We observed no significant difference in the equilibrium binding of hMSH2-hMSH3 or hMSH2-hMSH6 to consensus mispair recognition substrates in the absence of adenosine nucleotide or in the presence of ADP (data not shown). In addition, no significant increase in equilibrium binding was observed in the presence of ATP when compared with the homoduplex DNA substrate (data not shown). These results stand in contrast to those of Blackwell et al. (38) and may reflect the equilibrium distinctions between the different detection systems (IAsys/TIR versus Biacore/surface plasmon resonance).

Once appropriate dsDNA substrates were bound to the surface of an IAsys cuvette, the binding isotherms at a variety of hMSH2-hMSH6 concentrations were determined (representative isotherms shown in Fig. 2). Analysis of this binding isotherm data using Grafit 4 software (Affinity Sensors, Franklin, MA) results in calculated kassociation, kdissociation, and KD values (Table I). We found the binding of hMSH2-hMSH6 to the 8-oxo-G/A substrate (KD approx  78.0 × 10-9 M) was roughly 2-fold better than that observed with homoduplex DNA (KD approx  146.6 × 10-9 M). The binding of hMSH2-hMSH6 to a G/A mismatch (KD approx  58.7 × 10-9 M) was nearly 2-fold better than to an 8-oxo-G/A mismatch (KD approx  78.0 × 10-9 M). There was no difference in the binding of homoduplex DNA (G/C) (KD approx  146.6 × 10-9 M) versus homoduplex DNA containing 8-oxo-G (8-oxo-G/C) (KD approx  143.3 × 10-9 M). The binding of hMSH2-hMSH6 to DNA substrates containing either a G/T (KD approx  5.5 × 10-9 M) or G/G (KD approx  9.2 × 10-9 M) mismatch was significantly greater (~8-fold) than the binding to the G/A mismatch (KD approx  58.7 × 10-9 M). This observation supports previous reports suggesting G/T and G/G mismatches are more efficiently repaired than the G/A mismatch by hMSH2-hMSH6 (39-41).


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  		Fig. 2.   Binding of hMSH2-hMSH6 to various DNA substrates assessed by IAsys total internal reflectance. Biotinylated DNA substrates were immobilized on the surface of streptavidin-coated reaction cells. The representative binding isotherms shown were obtained with 75 nM hMSH2-hMSH6 and the DNA substrates described in Fig. 1. KD values (Table I) were determined by hMSH2-hMSH6 protein titration and curve fitting using Grafit software. Binding isotherms are color coded to indicate the oligonucleotide substrate shown in the box.

                              
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Table I
Affinity constants for the interaction of hMSH2-hMSH6 with the noted 41-mer double-stranded oligonucleotide substrates
Each value represents the mean derived from three independent experiments.

ATPase Activity-- There is considerable controversy surrounding the role of the conserved ATPase associated with MutS homologs. Accumulating evidence suggests that the efficiency for repair of an individual mismatch is linked to the mismatch-dependent ATPase activity (31). We examined the ATPase activity of hMSH2-hMSH3 and hMSH2-hMSH6 in the presence of the DNA substrates containing 8-oxo-G by the Norit method (Fig. 3; Table II). No significant stimulation of the hMSH2- hMSH3 ATPase was observed for substrates containing 8-oxo-G (kcat approx  1.0-1.3 min-1; Km approx  1.1-1.9 × 10-6 M; Fig. 3A; Table II). These observations should be compared with the stimulation of the hMSH2-hMSH3 ATPase by a consensus mispair DNA substrate +(CA) (kcat = 2.7 min-1; Km = 7.6 × 10-6 M).


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  		Fig. 3.   ATPase activity of hMSH2-hMSH6 and hMSH2-hMSH3 in the presence of 8-oxo-G DNA substrates. hMSH2-hMSH3 (A) or hMSH2-hMSH6 (B) were incubated in the presence of 240 nM DNA substrate and increasing concentrations of ATP. ATP hydrolysis was measure by the release of gamma -32P by the Norit method. Data from at least three independent experiments were fitted to the Michaelis-Menton equation to calculate the kinetic parameters with standard deviation (error bars) shown and listed in Table II.

                              
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Table II
Stimulation of hMSH2-hMSH6 and hMSH2-hMSH3 ATPase activity by various DNA substrates
Each value represents the mean derived from three independent experiments.

The hMSH2-hMSH6 ATPase is significantly stimulated by DNA substrates containing mismatched nucleotides and mismatched 8-oxo-G nucleotides (Fig. 3B; Table II). The hierarchy of mismatch-stimulated catalytic rate (kcat) was similar to our previous report: G/T (kcat = 20.3 min-1; Km = 50.4 × 10-6 M) > G/G (kcat = 17.8 min-1; Km = 51.4 × 10-6 M) > G/A (kcat = 13.8 min-1; Km = 52.2 × 10-6 M) > G/C (kcat = 6.4 min-1; Km = 29.1 × 10-6 M) (31). Furthermore, the hierarchy and the magnitude of ATPase stimulation is conserved when the DNA substrates contain 8-oxo-G: 8-oxo-G/T (kcat = 19.2 min-1; Km = 53.3 × 10-6 M) > 8-oxo-G/G (kcat = 17.3 min-1; Km = 48.2 × 10-6 M) > 8-oxo-G/A (kcat = 10.2 min-1; Km = 36.2 × 10-6 M) > 8-oxo-G/C (kcat = 4.4 min-1; Km = 18.8 × 10-6 M). No difference in the stimulation of the hMSH2-hMSH6 ATPase was observed when homoduplex G/C DNA was compared with 8-oxo-G/C DNA. These data suggest that only hMSH2-hMSH6 can be activated by 8-oxo-G containing DNA. Moreover, this activation appears to require mismatched 8-oxo-G.

ADP right-arrow ATP Exchange-- We have previously shown that ADP right-arrow ATP nucleotide exchange is the rate-limiting step for the hMSH2-hMSH6 ATPase and its transition to a hydrolysis-independent sliding clamp (27, 28). Mismatch-dependent ADP right-arrow ATP exchange has also been shown to be rate-limiting for hMSH2-hMSH3 (29), the yeast scMSH2-scMSH6,3 and bacterial MutS.4 These results suggest that bona fide recognition and activation of MutS homologs requires mismatch/lesion provoked ADP right-arrow ATP exchange.

We examined the ability of DNA substrates to provoke ADP right-arrow ATP exchange by hMSH2-hMSH3 and hMSH2-hMSH6 (Fig. 4). None of the 8-oxo-G DNA substrates stimulated ADP right-arrow ATP exchange by hMSH2-hMSH3 above the background for homoduplex DNA (Fig. 4A). In contrast, the +(CA) IDL efficiently provoked ADP right-arrow ATP exchange by hMSH2-hMSH3. These results are consistent with the IAsys binding studies and ATPase activity and are consistent with the notion that hMSH2-hMSH3 is incapable of activation by 8-oxo-G DNA damage.


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  		Fig. 4.   Rate of ADP-ATP exchange by hMSH2-hMSH3 and hMSH2-hMSH6 induced by various DNA substrates. DNA substrates (60 nM) were incubated with 60 nM of either hMSH2-hMSH3 (A) or with 60 nM hMSH2-hMSH6 (B) in the presence of 2.3 µM [3H]ADP. At time 0, ATP (25 µM) was added and the exchange reaction was allowed to proceed until the noted time points. The amount of bound [3H]ADP was determined by filter binding. Each symbol point represents an average of at least three independent experiments ± S.D. (error bars).

The stimulation of hMSH2-hMSH6 ADP right-arrow ATP exchange by DNA substrates containing 8-oxo-G/G or 8-oxo-G/T appeared similar to the corresponding G/G and G/T mismatches and significantly more rapid than ADP right-arrow ATP exchange in the absence of DNA or in the presence of homoduplex DNA (Fig. 4B). The 8-oxo-G/A containing DNA substrate did not appear to induce ADP right-arrow ATP exchange by hMSH2-hMSH6 at a rate similar to the substrate containing a G/A mismatch. However, the accelerated rate of hMSH2-hMSH6 ADP right-arrow ATP exchange provoked by the 8-oxo-G/A DNA substrate was significantly greater than that observed for homoduplex DNA (G/C) or the non-mismatched 8-oxo-G/C DNA substrate. These data are consistent with the IAsys DNA binding data and ATPase activity and support the conclusion that hMSH2-hMSH6 is only activated by mismatch 8-oxo-G containing DNA substrates. Taken as a whole, our studies suggest that 8-oxo-G alone is insufficient to activate hMSH2-hMSH6 and does not appear to be treated as a mispair by the human mismatch recognition system.

    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Oxidative damage incurred by reactive oxygen appears to be the most common lesion of DNA (42). Biology has adapted to the continual assault of reactive oxygen by evolving overlapping and redundant mechanisms for the repair of oxidative DNA damage (3). Most of these repair systems have been functionally or structurally conserved from bacteria to man. However, modest differences in the conservation of these repair systems may explain both genetic and biochemical observations.

Previous studies have suggested an involvement of eukaryotic MSH2 in the repair of oxidative DNA damage (1, 2). Both msh2 or msh6 mutant yeast strains display a significant increase in the signature of 8-oxo-G damage, G to T transversions. However, a synergistic increase in G to T transversions was observed in double mutant combinations carrying ogg1msh2 or ogg1msh6 (2). This genetic evidence indicated that the OGG1 base excision repair glycosylase and the MMR machinery function in different although redundant oxidative damage repair pathways. These same studies suggested that the scMsh2-scMsh6 heterodimer was capable of significantly binding oligonucleotide substrates that contained 8-oxo-G/A or 8-oxo-G/C (2). Our studies demonstrate that only the hMSH2-hMSH6 heterodimer specifically binds DNA substrates containing a mismatched 8-oxo-G lesion and that this binding activated its ATPase and ADP right-arrow ATP exchange activities. Conversely, hMSH2-hMSH3 does not appear to bind any of the 8-oxo-G DNA substrates nor do these 8-oxo-G DNA substrates activate either the ATPase or ADP right-arrow ATP exchange activities. We conclude that the hMSH2-hMSH6 heterodimer, but not the hMSH2-hMSH3 heterodimer, is capable of being activated by mismatched 8-oxo-G lesions. In addition, we found that hMSH2-hMSH6 was incapable of significant binding to non-mismatched 8-oxo-G/C DNA, nor was the ATPase or ADP right-arrow ATP exchange activities appreciably activated by this DNA substrate. This observation appears to represent a noteworthy difference between the yeast and human MSH2-MSH6 heterodimers (2).

Differences between the bacterial, yeast, and human recognition and processing of oxidative damage might be traced to differences in overlapping and redundant recognition systems (for review, see Ref. 3). The lack of a MutT or a MutY homolog in yeast is consistent with reduced redundancy and suggests that the post-replication MMR system is likely to play an expanded role in the repair of 8-oxo-G DNA damage. Thus, both replication misincorporation of adenosine opposite 8-oxo-G that has escaped the OGG-1 global genomic oxidative damage repair system as well as the misincorporation of 8-oxo-G into nascent DNA from nucleotide pools is likely processed by the MMR system in yeast (Fig. 5). Thus, it is not surprising that mutation of both MMR (msh2 or msh6) plus OGG1 (ogg1) in yeast would result in a synergistic increase in spontaneous G to T transversions. In contrast, such a large synergistic increase in G to T transversions would not be predicted in bacteria, mouse, or human cells where MutT and MutY homologs have been identified. Such studies have yet to appear in the literature.


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  		Fig. 5.   Model for the pathways of oxidative damage repair.

A major puzzle in the mechanism of oxidative DNA damage glycosylases includes the appropriate contextual recognition of mismatched 8-oxo-G lesions. For example, OGG-1 mediated inappropriate removal of mismatched 8-oxo-G from the template strand following replication could result in G to A transition mutations. Conversely, MutY-mediated removal of adenosine from the template strand following misincorporation of an 8-oxo-G from nucleotide pools could result in an A to C transversion. Since mismatched 8-oxo-G can only arise from replication errors, it is possible the glycosylases that might interfere with appropriate non-mutagenic recognition of oxidative damage may be excluded and/or regulated during S-phase. Indeed, nuclear expression of hMYH appears to be induced during S-phase (43).

Our studies suggest a significant role for hMSH2-hMSH6 in the non-mutagenic processing of 8-oxo-G lesions that remain in the DNA through S-phase. Because hMSH2-hMSH6 is only activated by mismatched 8-oxo-G, cyclic MMR excision-resynthesis would be expected until a C is inserted opposite the 8-oxo-G (Fig. 5). An 8-oxo-G/C pair does not appear to activate hMSH2-hMSH6, effectively disengaging MMR. This process thus retains the status quo that existed prior to replication and provides a DNA substrate for non-mutagenic repair by OGG1. The lack of binding and activation of hMSH2-hMSH6 by an 8-oxo-G/C DNA substrate appears distinct from the results of Ni et al. (2). We consider two possibilities: 1) the recognition of 8-oxo-G/C by the yeast scMSH2-scMSH6 is different from hMSH2-hMSH6; or 2) the binding of scMSH2-scMSH6 may not provoke requisite activation that is capable of inciting MMR. We regard the latter possibility as unlikely since the yeast scMSH2-scMSH6 appears to be equally displaced from the 8-oxo-G/C and 8-oxo-G/A substrates. It should also be noted that nascent strand misincorporation of 8-oxo-G from nucleotide pools may be recognized and removed by MMR-dependent excision-resynthesis.

hMSH2 appears to be involved in TCR of ionizing radiation and peroxide-induced oxidative DNA damage (26). A model for TCR has been proposed in which a transcription bubble is impeded by oxidative damage on the transcribed strand (44). TCR requires components of the nucleotide excision repair machinery that may be initiated by hMSH2 lesion recognition (45, 46). Yet, neither of the hMSH2 heterodimers (hMSH2-hMSH3 or hMSH2-hMSH6) appear to be activated by homoduplex 8-oxo-G. This observation places significant constraints on any model for TCR that requires hMSH2. It is possible that another structure associated with a stalled transcription fork may activate one of the hMSH2 heterodimers (46). Alternatively, it is also possible that hMSH2 may not participate in the TCR of 8-oxo-G damage. For example, only the TCR of peroxide-induced thymine glycol lesions has been directly demonstrated to require hMSH2 (26). Because a wide variety of oxidative DNA lesions may be induced by ionizing radiation, it is unclear which of these might be dependent on hMSH2 function (47, 48).

Finally, the activation of hMSH2-hMSH6 by lesions and mismatched substrates has begun to provide an accumulating picture of the initial recognition process. Comparison of the recognition and activation of hMSH2-hMSH6 by otherwise identical G/T, O6-methyl-G/T and 8-oxo-G/T DNA substrates, suggests that displacement of both the T and G away from the helical axis (49, 50) causes reduced stacking which then contributes to efficient activation of hMSH2-hMSH6. For example, the O6-methyl-G/T mismatch forms 1-imino and 2-amino hydrogen bonds on the G with the 3-imino and 2-oxo groups of the T, respectively (49). This base pairing interaction appears qualitatively similar to an A/T or G/C base pair. Thus, the T and G are only marginally displaced from a normal helix stack. Comparison of DNA binding, ATPase, and ADP right-arrow ATP exchange activities of hMSH2-hMSH6 in the presence of a O6-methyl-G/T substrate suggests only modest activation over an A/T or G/C homoduplex substrate (24). In contrast, the G/T mispair, and by projection the 8-oxo-G/T mispair, induce a shift from the central helical axis to form 1-imino and 6-oxo hydrogen bonds with the 2-oxo and 3-imino groups of the T, respectively. The overall effect is to further displace the 5-methly group of the T into the major groove as well as the 2-amino group of the G into the minor groove. The DNA binding, ATPase, and ADP right-arrow ATP exchange activities of hMSH2-hMSH6 in the presence of a G/T or 8-oxo-G/T mispair is 2-3-fold better than an O6-methyl-G/T and 5-8-fold better than homoduplex DNA. It is tempting to speculate that the shift in helix stacking contributes to localized flexibility of the DNA that is subsequently captured by the MutS homolog clamp (28, 52).

A significant accumulation of 8-oxo-G in irradiated mouse Msh2-/- embryonic stem cells supports the notion that oxidative stress may contribute to HNPCC (1). Our observation that the hMSH2-hMSH6 heterodimer is uniquely responsible for 8-oxo-G recognition among the MMR proteins also suggests that patients with altered hMSH6 may be susceptible to similar oxidative stress. Consideration of the types of lesions recognized by the human MMR machinery may be useful in understanding the process(es) of carcinogenesis as well as appropriate therapeutics for HNPCC-related tumors.

    ACKNOWLEDGEMENTS

We thank Samir Acharya, Christoph Schmutte, and Kristine Yoder for helpful discussions and extensive editorial comments.

    Note Added in Proof

We thank Dr. John Hays (Oregon State University) for pointing out an apparent inconsistency in our representation of the exponential values for the kinetic parameters shown in Table II. We have clarified this as a footnote to Table II.

    FOOTNOTES

* This work was supported by National Institutes of Health Grants CA56542 and CA67007.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Dagger To whom correspondence should be addressed: Kimmel Cancer Center, BLSB933, 233 S. 10th St., Philadelphia, PA 19107. Tel.: 215-503-1345; E-mail: rfishel@lac.jci.tju.edu.

Published, JBC Papers in Press, December 26, 2001, DOI 10.1074/jbc.M111269200

2 M. Berardini, C. Heinen, T. Wilson, and R. Fishel, unpublished results.

3 S. Acharya, M. Hess, R. Kolodner, and R. Fishel, unpublished results.

4 S. Acharya and R. Fishel, unpublished results.

    ABBREVIATIONS

The abbreviations used are: 8-oxo-G, 8-oxo-guanine; MMR, DNA mismatch repair; HNPCC, hereditary nonpolyposis colorectal cancer; MSH, MutS homolog; TIR, total internal reflectance; IDL, insertions/deletions loop-type; MNNG, N-methyl-N'-nitro-N-nitrosoguanidine; TCR, transcription-coupled repair; ds, double-stranded; ss, single-stranded.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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