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ATP binding causes the mispair-bound Msh2-Msh6 mismatch recognition complex to slide along the DNA away from the mismatch, and ATP is required for the mispair-dependent interaction between Msh2-Msh6 and Mlh1-Pms1. It has been inferred from these observations that ATP induces conformational changes in Msh2-Msh6; however, the nature of these conformational changes and their requirement in mismatch repair are poorly understood. Here we show that ATP induces a conformational change within the C-terminal region of Msh6 that protects the trypsin cleavage site after Msh6 residue Arg1124. An engineered disulfide bond within this region prevented the ATP-driven conformational change and resulted in an Msh2-Msh6 complex that bound mispaired bases but could not form sliding clamps or bind Mlh1-Pms1. The engineered disulfide bond also reduced mismatch repair efficiency in vivo, indicating that this ATP-driven conformational change plays a role in mismatch repair.
Mismatch repair (MMR)2 identifies and repairs mispaired bases that occur in DNA as the result of errors during DNA replication, the formation of recombination intermediates, or chemical damage to DNA (1–5). As MMR normally functions to suppress the accumulation of mutations, defects in MMR genes underlie both inherited and spontaneous cancers that are driven by high rates of accumulating mutations (6–9). In Escherichia coli, the MutS homodimer recognizes mispairs (2, 3, 10–13), whereas in Saccharomyces cerevisiae and humans, one of two heterodimers of MutS homologs, Msh2-Msh3 or Msh2-Msh6, recognizes mispairs (4, 14–17). Mispair recognition by the MutS family of proteins is the first step of MMR that leads to the recruitment of either the bacterial MutL homodimer or the eukaryotic MutL homolog heterodimers, Mlh1-Pms1 or Mlh1-Mlh3, to mediate downstream steps, which lead to excision and resynthesis of the DNA strand that contained the misincorporated bases (18–26).
MutS and its homologs contain ABC ATPase domains, and multiple lines of evidence indicate that ATP plays crucial roles in the function of these proteins (27–36). In the absence of nucleotide or the presence of ADP, these complexes can recognize mispairs; however, these pre-bound complexes form “sliding clamps” that dissociate from the mispair and slide along the DNA when challenged with ATP or nonhydrolyzable ATP analogs (18, 19, 28, 30). These sliding clamps can be trapped on DNA fragments with sterically blocked ends (18, 19, 24, 30, 37). ATP binding by MutS and its homologs is also required for the ability of these proteins to interact with MutL or the eukaryotic MutL homologs, respectively, and thereby mediate downstream events in MMR (18–20, 24). Together these biochemical results are consistent with the presence of an ATP-induced conformational change in MutS, Msh2-Msh3, and Msh2-Msh6 that propagates from the ATPase domain to other parts of the molecule.
To date, the details of ATP-driven conformational changes in MutS, Msh2-Msh3, and Msh2-Msh6 have proven elusive (38). Crystal structures of bacterial MutS, human Msh2-Msh3, and Msh2-Msh6 bound to mispairs have been solved and have provided important details of the ADP- and mispair-bound states of these proteins (2, 10, 39, 40). This state has been called the open form (41, 42). In contrast, the conformation of the ATP-bound forms of these proteins is unknown. Based on structural homology to ATP-bound Rad50 (43), this form of the MutS family of proteins has been called the closed form (41, 42). Consistent with these inferred states, deuterium exchange studies with S. cerevisiae Msh2-Msh6 demonstrated that ATP binding protects the Msh2-Msh6 dimer interface between the ATPase domains and exposes the mispair-binding domains (42). ATP binding moderately increases deuterium exchange in the connector domains of MutS and Msh2 (42), which interact with MutL or Mlh1-Pms1, respectively (44, 45). These data are consistent with a model in which ATP binding releases the mispair-binding domain from the mispair and exposes the connector domain to recruit MutL/Mlh1-Pms1. However, the details of and the requirement for these conformational changes in MMR remain poorly characterized. In the studies presented here, we have used a combination of genetic and biochemical approaches to identify a region undergoing an ATP-induced conformational change and demonstrate its functional involvement in MMR.
S. cerevisiae strains were grown in standard medium, including either yeast extract/peptone/dextrose or complete supplement mixture medium (U.S. Biological Corp.) lacking specific amino acids to select for plasmid markers and/or Thr+ revertants. E. coli strains used for the propagation of plasmids were grown in LB medium containing 100 μg/ml of ampicillin to select for plasmids. All S. cerevisiae strains used for genetic tests were isogenic derivatives of the same parental S288C strain (46). The strain used for the complementation studies was RDKY4234 (MATα ura3–52 leu2Δ1 trp1Δ63 his3Δ200 hom3–10 lys2::InsE-A10 msh3Δ::hisG msh6Δ::hisG). Plasmid pRDK439 was from our laboratory collection and was constructed by cloning a 6-kb fragment of S. cerevisiae genomic DNA containing MSH6 into the ARS CEN vector pRS315. The dual Msh2-Msh6 E. coli expression plasmid pET11a-Msh2-Msh6 was from Dr. Manju Hingorani (47). Derivatives of pRDK439 and pET11a-Msh2-Msh6 containing the indicated MSH6 alleles were made by site-directed mutagenesis and verified by sequencing the complete MSH6 gene (Table 1). The msh6 truncation alleles 1125stop, 1135stop, 1152stop, 1172stop, 1182stop, 1191stop, 1201stop, 1211stop, 1221stop, and 1241stop were engineered by inserting two stop codons at the indicated positions. The msh6 truncation alleles 1231stop and 1236stop were engineered by deleting codons 1231–1242 and 1236–1242, respectively. All of the genetic methods used in the described studies, including those used for site-directed mutagenesis, verification by DNA sequencing, and evaluation of mutator phenotypes by patch tests and fluctuation analysis, were as described previously (15, 35, 48, 49).
Wild-type and mutant Msh2-Msh6 complexes were purified from E. coli as described (24, 47, 49, 50). Wild-type Msh2-Msh6 and Msh2-Msh6 L1129C/S1137C were purified in buffer A (25 mm Tris, pH 8.0, 314 mm NaCl, 1 mm EDTA, 10% glycerol, 5 mm DTT, 0.02% Igepal, 1 mm phenylmethylsulfonyl fluoride (PMSF), 1 mm benzamidine, 0.5 mg/liter of bestatin, and 1 mg/liter each of chymostatin, pepstatin A, aprotinin, and leupeptin). Reduced wild-type Msh2-Msh6 and reduced Msh2-Msh6 L1129C/S1137C were maintained in buffer A. Wild-type Msh2-Msh6 and Msh2-Msh6 L1129C/S1137C were oxidized by overnight dialysis at 4 °C against buffer A lacking DTT. ATP-supplemented oxidized Msh2-Msh6 L1129C/S1137C was generated by overnight dialysis at 4 °C against buffer A lacking DTT and containing 100 μm ATP in the absence of Mg2+. Mlh1-Pms1 was purified from S. cerevisiae as described (24, 41). LacI protein was generously provided by Kathleen Matthews (Rice University).
To determine which region of Msh6 was undergoing the ATPγS-induced conformational change, ~300 ng each of full-length Msh6 and the 90-, 80-, and 75-kDa Msh6 tryptic fragments were separated by SDS-PAGE on a 4–15% gradient gel. After staining and destaining, the gel was incubated in water overnight, and the bands of interest were each excised from the gel with a clean razor blade. In gel slice reduction, alkylation, and trypsin digestion was performed by the Proteomics Mass Spectrometry Laboratory at the Scripps Research Institute, La Jolla, CA. Briefly, the proteins were reduced in 10 mm DTT (Sigma) for 1 h, then alkylated with 55 mm iodoacetamide (Sigma) for 30 min in the dark, and finally digested for 18 h with trypsin at 37 °C using a 1:30 (w/w) enzyme to substrate ratio. The digestion reaction was stopped by lowering the pH of the reaction to 2.0. Tryptic peptides were analyzed by reverse phase chromatography using Zorbax SB-C18 stationary phase (Agilent). The data-dependent MS/MS data were obtained on a LTQ linear ion trap mass spectrometer (Thermo Scientific). Protein identification was performed using Mascot (Matrix Science, version 2.1.04). Tandem mass spectra were extracted by the Xcalibur software, and all MS/MS samples were analyzed using Mascot. Scaffold (Proteome Software Inc.) was used to additionally validate MS/MS-based peptide and protein identifications.
To demonstrate the presence of the disulfide bond, 20 μg of oxidized and reduced Msh2-Msh6 L1129C/S1137C were each treated with 2% iodoethanol (Sigma) for 1 h at 37 °C in the dark. The protein was then precipitated by addition of a mixture of 50% ethanol, 50% acetone containing 0.1% acetic acid for 1 h on ice. Then the proteins were denatured with 8 m urea, reduced with 10 mm DTT for 10 min at 42 °C, alkylated with 30 mm iodoacetamide (Sigma) for 30 min at room temperature in the dark, digested with 0.2 μg of sequencing grade trypsin (Promega) at 37 °C overnight, and the digestion reaction was stopped with 0.2% trifluoroacetic acid. Peptide cleanup was then performed using a 50-mg Sep-Pak C18 column (Waters) that had been equilibrated with 0.1% acetic acid. Bound peptides were eluted from the column with 300 μl of 80% acetonitrile, 0.1% acetic acid; the eluate was lyophilized; and the resulting peptides were resuspended in 0.1% trifluoroacetic acid. The samples were then analyzed by LC-MS/MS using an LTQ Orbitrap Discovery XL mass spectrometer (Thermo Scientific). Data dependent MS/MS data collected were searched using SorcererTM-SEQUEST (51) with a semi-tryptic modified database designed for the mutant Msh6. Variable modifications for cysteine were applied for carbamidomethyl (+57 Da) and 2-hydroxyethyl (+44 Da) with a peptide mass tolerance of 20 ppm. MS/MS peptide modification and protein identification were additionally validated by manual inspection.
To monitor the ATPγS-induced conformational change in Msh2-Msh6, 4 μg of wild-type Msh2-Msh6 or Msh2-Msh6 L1129C/S1137C was incubated with 0–500 ng of trypsin with or without 100 μm ATPγS in reaction buffer (25 mm Tris, pH 8.0, 110 mm NaCl, 4 mm MgCl2, 0.01% Igepal, 2 mm dithiothreitol, 2% glycerol) in a final volume of 10 μl for 1 h at room temperature (23 °C). Reactions were stopped by the addition of PMSF to a final concentration of 10 mm. The proteolysis products were separated by SDS-PAGE on a 4–15% gel (Bio-Rad), and the resulting gel was silver-stained.
Experiments analyzing the interaction between the Msh2-Msh6 complex and mispaired DNA as well as the formation of the Msh2-Msh6-Mlh1-Pms1 ternary complex on mispaired DNA were performed with a Biacore T100 instrument (GE Healthcare) essentially as described previously (41, 44). Note that the buffers used in these experiments contained Mg2+.
Nucleotide binding was measured by UV cross-linking essentially as described previously (28). Briefly, 20-μl reactions containing 10 pmol of reduced wild-type Msh2-Msh6, 30 pmol of oxidized wild-type Msh2-Msh6, 30 pmol of reduced Msh2-Msh6 L1129C/S1137C, 50 pmol of oxidized Msh2-Msh6 L1129C/S1137C, or 50 pmol of oxidized Msh2-Msh6 L1129C/S1137C that had been dialyzed overnight against buffer containing ATP without Mg2+, were each performed in binding buffer that contained 50 mm Tris, pH 8.0, 110 mm NaCl, 100 μg/ml of BSA, 0.5 mm EDTA, 5% glycerol. The reactions with reduced protein also contained 2 mm dithiothreitol. The absence of magnesium prevented ATP hydrolysis. Proteins were mixed with 25 Ci/mmol of [γ-32P]ATP and incubated on ice for 10 min. Samples were then subjected to UV cross-linking (Stratalinker) for 20 min followed immediately by fractionation by SDS-PAGE using 4–15% gradient gels. The gels were exposed to a PhosphorImager screen overnight, and the screen was developed using a PhosphorImager (GE Healthcare).
We previously demonstrated that ATP binding protected Msh2-Msh6 from digestion with trypsin (Fig. 1A), consistent with a conformational change (41). To determine which part of Msh2-Msh6 undergoes the ATP-induced conformational change, bands corresponding to full-length Msh6 and the 90-, 80-, and 75-kDa trypsin fragments were excised from a gel, digested to completion with trypsin, and analyzed by mass spectrometry (MS; Fig. 1, B–E). Consistent with Western blots (41), MS confirmed that the 90-, 80-, and 75-kDa trypsin fragments were derived from Msh6.
The 90-, 80-, and 75-kDa fragments yielded the same pattern and proportions of N-terminal tryptic peptides, and therefore appeared to have the same N terminus. For each trypsin fragment, MS failed to detect significant amounts of any peptides N-terminal to the trypsin cleavage site at Arg441-Glu442 (Fig. 1, C–E), unlike full-length Msh6 from which substantial amounts of peptides starting with the first residue were detected (Fig. 1B). This suggests that the N terminus of the 90-, 80-, and 75-kDa fragments was residue Glu442. Residue Glu442 lies in the N terminus of the connector domain of Msh6, C-terminal to a loop connecting the connector domain with the mispair-binding domain and appears to be exposed. Thus, the 90-, 80-, and 75-kDa fragments lack the mispair-binding domain and the N-terminal tether that contains the PCNA interaction site (Fig. 1, C–E).
MS analysis indicated the 90-, 80-, and 75-kDa fragments had different C termini, which were consistent with their molecular weights estimated by SDS-PAGE. As for intact Msh6, significant amounts of peptides from the 90-kDa fragment sample were observed up to and including the C-terminal peptide ending in residue Ser1242 (Fig. 1C). An Msh6 Glu442-Ser1242 fragment has a calculated molecular mass of 90.4 kDa, consistent with the estimated size of the fragment analyzed. In contrast, the 80-kDa fragment had reduced levels of tryptic peptides between residues Arg1171 and Ser1242 (Fig. 1D). Residue Arg1171 is in the last α-helix of the ATPase domain of Msh6, as predicted by homology to the human Msh2-Msh6 structure (Fig. 1, F and G), and an Msh6 Glu442-Arg1171 fragment has a calculated molecular mass of 82.7 kDa, consistent with the estimated size of the fragment analyzed. MS analysis of the 75-kDa fragment was indicative of an even smaller fragment; significant levels of tryptic peptides were not identified C-terminal to residue Arg1124 (Fig. 1E). Residue Arg1124 is between the last two β strands of the ATPase domain, as predicted by homology to the human Msh2-Msh6 structure (Fig. 1, F and G), and an Msh6 Glu442-Arg1124 fragment has a calculated molecular mass of 77.5 kDa, consistent with the estimated size of the fragment analyzed. These results support the idea that ATP binding induces a conformational change in the region containing Msh6 residue Arg1124, which protects the Arg1124-Asn1125 trypsin site from cleavage.
To determine whether the C-terminal region of Msh6 that is affected by ATP binding is required for MMR, we constructed a series of MSH6 mutations that eliminated amino acids from the C terminus of Msh6. These mutant genes were expressed on low copy number plasmids and tested for their ability to complement the high frameshift mutation rate of an msh3Δ msh6Δ strain. The mutant lacking the last two codons of MSH6 (1241stop) fully complemented the high frameshift mutation rate of an msh3Δ msh6Δ strain in patch tests, like a wild-type MSH6 plasmid (Fig. 2A), and had a similar mutation rate to that of the wild-type MSH6 plasmid in quantitative mutation rate tests (Table 2). The mutant lacking the last seven codons of MSH6 (1236stop) had an intermediate mutation rate, which was 8-fold higher than seen with the wild-type MSH6 plasmid (Table 2). In contrast, deletion of 12 or more amino acids from the C terminus (1125stop to 1231stop) resulted in plasmids that failed to complement the high frameshift mutation rate of an msh3Δ msh6Δ strain in patch tests, like the empty vector control plasmid (Fig. 2A). The strains containing plasmids with stop codons at 1221 and 1231 had mutation rates that were 199- and 149-fold higher, respectively, than that obtained with a plasmid containing wild-type MSH6, and were similar to the rates obtained with the empty vector control plasmid (Table 2). These results indicate that virtually the entire C-terminal region of Msh6 is required for MMR, including regions not ordered in the human Msh2-Msh6 structure (Fig. 2B) (39), and would be consistent with a C-terminal Msh2-Msh6 dimerization domain equivalent to the human Msh2-Msh3 C-terminal domain (40).
We next analyzed the C-terminal region of Msh6 near the ATP-protected Arg1124-Asn1125 trypsin site for residues that could be changed to cysteines that would then have the potential to generate a disulfide bond under oxidizing conditions. Residues 1129 and 1137 are in close proximity and are in a region undergoing ATP-induced changes (42), but they are not ideally positioned for disulfide bond formation in the crystal structure of human Msh2-Msh6 (39). We reasoned that if conformational changes allowed for disulfide bond formation in the mutant protein, then disulfide bond formation might in turn affect conformational changes in Msh2-Msh6.
We then created single and double cysteine mutations in a plasmid-encoded MSH6 gene and tested for their effect on MMR (Fig. 3A). The msh6-L1129C and msh6-S1137C mutations individually appeared to be weak mutator mutations in patch tests as did alternative mutations affecting these residues including msh6-L1129D and msh6-S1137A. In contrast, the msh6-L1129C/S1137C pair of mutations, which we predicted to form a disulfide bond, caused a significantly stronger mutator phenotype in patch tests, unlike the msh6-L1129D/S1137A pair. Other potential cysteine pairs in this region were investigated but not studied further as the other double cysteine mutants did not have increased mutation rates over the single cysteine mutants. Quantitative mutation rate tests confirmed that the msh6-L1129C/S1137C allele caused a significant increase in mutation rate although not as large as a complete deletion of MSH6 (Table 2). In contrast, the mutation rates caused by the msh6-L1129C and msh6-S1137C mutations were not significantly different from the wild-type mutation rate (Table 2). Thus, the MMR defect occurred only in the double cysteine mutant.
To demonstrate the formation of a disulfide bond, Msh2-Msh6 L1129C/S1137C was overexpressed and purified. An oxidized sample was generated by overnight dialysis against buffer A lacking DTT to enable any potential disulfide bonds to form. Both oxidized and reduced Msh2-Msh6 L1129C/S1137C were treated with iodoethanol, denatured, reduced, and treated with iodoacetamide. In this experiment, exposed cysteines not in disulfide bonds will react with iodoethanol to become 2-hydroxyethyl-modified and have a total mass of 147 Da. Cysteines in disulfide bonds or buried within the protein will react with iodoacetamide after denaturation and reduction to become carbamidomethyl-modified and have a total mass of 160 Da (Fig. 4A). Thus, exposed cysteines in disulfide bonds should be 2-hydroxyethyl-modified in the reduced sample and carbamidomethyl-modified in the oxidized sample. In contrast, cysteines not in disufide bonds should be either 2-hydroxyethyl-modified if they are exposed or carbamidomethyl-modified if they are buried regardless of whether the sample is oxidized or reduced.
After this treatment, S1137C was observed as carbamidomethyl-modified 73% of the time in the oxidized sample and 31% in the reduced sample (Fig. 4B). L1129C was not observed in the oxidized sample and was only observed once in the reduced sample. L1129C may not have been detected for a variety of technical reasons including vaporization and/or time of flight. Thus no significant conclusions can be drawn about L1129C. However, it should be noted that this analysis method allows inference of disulfide bond status even when both cysteine-containing peptides are not recovered as disulfide formation is detected through analysis of the properties of the modified cysteines. Eleven additional Msh6 cysteines were observed more than once in both the oxidized and reduced samples (Fig. 4C), but only the modification of S1137C was significantly affected by the presence of DTT (p = 0.003; Fisher's exact test). Thus, at least 73% of S1137C in oxidized Msh2-Msh6 L1129C/S1137C was disulfide bonded. Possible explanations for the observation of 27% 2-hydroxyethyl-modified S1137C are incomplete disulfide bond formation due to protein dynamics in this region of the protein or that dialysis treatment was insufficient to oxidize 100% of the cysteine residues.
To determine how the engineered disulfide bond affected the ATP-induced Msh2-Msh6 conformational change, we repeated the partial proteolysis analysis using reduced and oxidized Msh2-Msh6 L1129C/S1137C. As for both reduced and oxidized wild-type Msh2-Msh6, ATPγS prevented trypsin from cleaving the reduced Msh6 L1129C/S1137C into the 75-kDa fragment (Fig. 5). In contrast, trypsin generated the 75-kDa fragment from oxidized Msh2-Msh6 L1129C/S1137C regardless of whether or not ATPγS was present, consistent with the possibility that disulfide bond formation trapped Msh2-Msh6 L1129C/S1137C in the ATP-unbound conformation. When the oxidized Msh2-Msh6 L1129C/S1137C was dialyzed overnight against buffer lacking both DTT and Mg2+ and containing 100 μm ATP at 4 °C, trypsin digestion was unable to generate the 75-kDa fragment, regardless of whether or not ATPγS was in the proteolysis reaction (Fig. 5), consistent with slow binding of ATP and subsequent conformational changes during dialysis. Together, these results suggest that oxidized Msh2-Msh6 L1129C/S1137C can bind ATP and undergo ATP-driven conformational changes but on an extremely slow time scale.
Wild-type Msh2-Msh6 in either the nucleotide free or ADP-bound state can bind to mispairs. Challenging these complexes with ATP converts wild-type Msh2-Msh6 to a sliding clamp that slides away from the mispair along the DNA (28). Sliding clamp formation by Msh2-Msh6 can be monitored by surface plasmon resonance by following the accumulation of multiple Msh2-Msh6 molecules on an immobilized DNA molecule containing a single mispair under conditions in which the DNA ends are blocked (24). To do this, a 236-base pair oligonucleotide with a central GT mispair, a biotin at one end, and a lac operator sequence at the other end was bound to a streptavidin-coated Biacore chip at its biotinylated end. Addition of LacI, which binds the lac operator and can be removed by addition of isopropyl 1-thio-β-d-galactopyranoside, sterically blocks the DNA end and prevents Msh2-Msh6 from sliding off of the DNA.
Both oxidized and reduced wild-type Msh2-Msh6 bound mispaired DNA in the presence of ATP and Mg2+ (Fig. 6, A and C, red dashed line). Under these conditions, free Msh2-Msh6 hydrolyzes ATP to yield the ADP-bound state in which it binds the mispair, exchanges ADP for ATP, and then slides off the DNA end, resulting in the steady-state level of binding seen (24). Addition of LacI increases the number of molecules bound to the DNA substrate (Fig. 6, red solid line), and release of LacI with isopropyl 1-thio-β-d-galactopyranoside returns binding back to the steady-state level (Fig. 6, green dashed line). This pattern of LacI-dependent accumulation of Msh2-Msh6 on the DNA substrate demonstrated that both oxidized and reduced wild-type Msh2-Msh6 can form sliding clamps (24). Reduced Msh2-Msh6 L1129C/S1137C had the same binding properties as wild-type Msh2-Msh6, indicating that the mutant can form sliding clamps under reduced conditions (Fig. 6B). In contrast, oxidized Msh2-Msh6 L1129C/S1137C (Fig. 6D) and oxidized Msh2-Msh6 L1129C/S1137C that had been dialyzed overnight against buffer containing ATP (Fig. 6E) were able to bind mispaired bases (Fig. 7), but appeared to be defective for the formation of sliding clamps as they showed no increase in binding to substrates with blocked ends versus unblocked ends, and there was little to no decrease in binding to the end-blocked substrate upon addition of isopropyl 1-thio-β-d-galactopyranoside. It should be noted that ATP-supplemented oxidized Msh2-Msh6 L1129C/S1137C can bind to the mispaired base because the Biacore running buffer contains Mg2+, which allows the bound ATP to be hydrolyzed, creating a protein state that is permissible for mispair binding (24, 28). Therefore, the inability of oxidized Msh2-Msh6 L1129C/S1137C and ATP-supplemented oxidized Msh2-Msh6 L1129C/S1137C to form a sliding clamp suggests that disulfide bond formation completely or partially inhibits ATP-induced conformational changes.
Surface plasmon resonance was also used to assess the ability of Msh2-Msh6 to assemble ternary complexes with Mlh1-Pms1 in the presence of ATP. In the presence of ATP, reduced or oxidized wild-type Msh2-Msh6 interacted with Mlh1-Pms1 and formed ternary complexes resulting in increased binding to the end-blocked mispaired substrate (Fig. 8, A and C, solid red line) compared with binding of Msh2-Msh6 alone (Fig. 8, A and C, dashed red line). Reduced Msh2-Msh6 L1129C/S1137C also mediated the formation of ternary complexes with Mlh1-Pms1, albeit to a lesser extent than wild-type Msh2-Msh6 (Fig. 8B). In contrast, oxidized Msh2-Msh6 L1129C/S1137C (Fig. 8D) and oxidized Msh2-Msh6 L1129C/S1137C that had been dialyzed overnight against buffer containing ATP (Fig. 8E) did not support the binding of Mlh1-Pms1. These results suggest that the ability of the Msh6 C-terminal region to switch between the two conformations is required for interaction between Msh2-Msh6 and Mlh1-Pms1.
We previously demonstrated that ATP binding to both the Msh2 and Msh6 subunits as well as communication between the subunits is required for sliding clamp formation, and that ATP binding to only the Msh6 subunit is required for ternary complex formation (41). We used ATP cross-linking to determine whether the inability of oxidized Msh2-Msh6 L1129C/S1137C to form sliding clamps or ternary complexes with Mlh1-Pms1 might result from a reduced affinity for ATP. The absence of Mg2+ in the cross-linking reaction buffer prevented ATP hydrolysis but could have had a minor influence on the nucleotide binding characteristics of Msh2-Msh6 (28, 52). Reduced and oxidized wild-type Msh2-Msh6 had similar affinities for ATP. Consistent with the results of previous studies (28), 50% saturation of ATP (S0.5) in wild-type Msh6 for ATP was achieved at concentrations below 1 μm, and the S0.5 of Msh2 for ATP was between 20 and 50 μm (Fig. 9). In reduced Msh2-Msh6 L1129C/S1137C, the S0.5 of Msh2 and Msh6 for ATP were similar to each other and were between 20 and 50 μm, similar to the affinity of Msh2 for ATP in wild-type Msh2-Msh6. This indicates that in reduced Msh2-Msh6 L1129C/S1137C, the Msh6 subunit has a lower affinity for ATP relative to that in wild-type Msh2-Msh6; however, reduced Msh2-Msh6 L1129C/S1137C is likely proficient for sliding clamp formation and the formation of ternary complexes because the assays for these features were performed in buffer containing 250 μm ATP, a concentration significantly higher than the S0.5 for ATP binding to each subunit.
In contrast, the affinities of both Msh2 and Msh6 for ATP were significantly reduced in oxidized Msh2-Msh6 L1129C/S1137C, with the S0.5 for ATP for both Msh2 and Msh6 being between 50 and 250 μm (Fig. 9). This result supports the possibility that the inability of oxidized mispair-bound Msh2-Msh6 L1129C/S1137C to form either sliding clamps or ternary complexes with Mlh1-Pms1 is due to the drastically reduced affinities of both the Msh2 and Msh6 subunits for ATP. Interestingly, both subunits, particularly Msh6 L1129C/S1137C, of the oxidized Msh2-Msh6 double mutant, which had been dialyzed in ATP overnight in the absence of Mg2+ and had undergone the ATP-induced conformational change as evidenced by the partial proteolysis pattern (Fig. 5), had even weaker affinities for ATP than those of oxidized Msh2-Msh6 L1129C/S1137C. This is most likely due to stable binding of unlabeled ATP, which forced the mutant to undergo the conformational change, competing with the radioactively labeled ATP in the cross-linking reaction. The severely reduced affinity of ATP-supplemented oxidized Msh2-Msh6 L1129C/S1137C for ATP is consistent with its inability to form the sliding clamp or ternary complex.
The Msh2-Msh6 complex appears to exist in at least two biochemically distinct forms that are modulated by ATP binding and hydrolysis. The nucleotide free and ADP bound forms are proficient for mispair binding (19, 28). ATP binding prevents Msh2-Msh6 from binding mispairs, but converts the mispair bound form to a sliding clamp and a form that recruits Mlh1-Pms1 (18–20, 24, 28, 41). These changes in biochemical properties are also mirrored by changes in partial proteolysis (36, 41) and are consistent with results from electron microscopy (30), deuterium exchange (42), analytical ultracentrifugation (38), and single-molecule FRET (53). Despite this, the requirement for conformational changes in the MutS family of proteins during MMR is not well understood (30, 38, 41, 42). To understand the role this conformational change plays in the activities of Msh2-Msh6 and MMR, we sought to introduce a reversible disulfide cross-link to prevent Msh2-Msh6 from transitioning between states.
To model the effects of ATP binding on the ATPase domains of Msh2-Msh6, we compared the DNA-bound structures of MutS, Msh2-Msh6, and Msh2-Msh3, which are considered to be the open form of the ABC ATPases (2, 10, 39, 40, 54, 55), with the ATP-bound form of Rad50, which is considered to be the closed form of the ABC ATPases (43). This analysis indicated that the ATPase domains could readily switch between an open ATP-free conformation and a closed ATP-bound conformation, except for the final two ordered helices of both Msh2 and Msh6 in the DNA co-crystal structure (39). These helix pairs most likely move relative to the rest of the ATPase domain (Fig. 10A) due to steric collision with each other in the ATP-bound form as inferred from simple superposition of Msh2 and Msh6 onto Rad50 (Fig. 10A) or potentially by rigid separation of these helix pairs by a C-terminal dimerization domain equivalent to that of E. coli MutS (56) or human Msh2-Msh3 (40) that is disordered in the human Msh2-Msh6 crystal structure (39). The existence of this dimerization domain in S. cerevisiae Msh2-Msh6 and its requirement in MMR, as with Thermus aquaticus and E. coli MutS (56, 57), would explain why all of the Msh6 C-terminal residues, except for the last seven, are required for MMR (Fig. 2A). Conservation of this domain across MutS family members may be required to stabilize dimers so that the interface between the ATPase domains can change dynamically upon ATP binding. Regardless of the cause, movement of the final ordered helices of Msh2-Msh6 relative to the rest of the ATPase domain would move the loop containing Msh6 residues Leu1129 and Ser1137, and is consistent with ATP binding causing increased protection from deuterium exchange (Fig. 11) (42) and protection of the Arg1124-Asn1125 trypsin site from cleavage (Fig. 1).
Introduction of double cysteine amino acid substitutions, L1129C and S1137C, into the loop of Msh6 preceding the final ordered helices generated a protein that formed a disulfide cross-link upon oxidation (Fig. 4). Genetically, the double cysteine mutation reduced MMR proficiency in vivo albeit not to the same extent as caused by deleting MSH6 (Fig. 3A, Table 2). One potential reason why an MSH6 deletion causes a higher mutation rate is that the cytoplasm of S. cerevisiae is reducing (58) and hence disulfide bond formation may not go to completion in vivo. However, under reducing conditions in vitro, 31% of mutant protein contained a L1192C/S1137C disulfide bond (Fig. 4). Taken together, this suggests that a sufficient fraction of protein containing the disulfide bond can form in vivo to produce the elevated mutation rate observed in vivo.
The reduced Msh2-Msh6 L1129C/S1137C behaved like wild-type in the accessibility of the Arg1124-Asn1125 trypsin site in the open and closed conformations (Figs. 5 and and1010D), the formation of the sliding clamp (Fig. 6), and recruitment of Mlh1-Pms1 (Fig. 8). The greatest difference between the reduced mutant and wild-type proteins was that the mutant Msh6 had a reduced affinity for ATP, which was equivalent to Msh2 in wild-type Msh2-Msh6 and Msh2-Msh6 L1129C/S1137C. Sliding clamp formation and recruitment of Mlh1-Pms1 differ in their ATP requirements in that recruitment of Mlh1-Pms1 only requires ATP binding to the high affinity Msh6 site, whereas sliding clamp formation requires ATP binding to both the Msh2 and Msh6 sites (41); however, the concentration of ATP in the experiment, 250 μm, was sufficient to saturate the ATP binding sites of the Msh2 and Msh6 subunits in this mutant explaining how it can be proficient for sliding clamp formation and recruitment of Mlh1-Pms1. This contrasts with other mutants, such as Msh2-R730C-Msh6 and Msh2-Msh6-R1024C, which were unable to form sliding clamps or bind Mlh1-Pms1 even when the ATP concentration was above the measured S0.5 values for each subunit; Msh2-R730C-Msh6 and Msh2-Msh6-R1024C most likely have additional defects in communication between the ATP-binding sites (41).
The oxidized Msh2-Msh6 L1129C/S1137C, however, appeared to be defective for normal rapid rates of ATP binding as evidenced by the generation of the 75-kDa band in the presence of ATPγS (Figs. 5 and and1010E), its ability to bind mispairs (Fig. 7), its failure to form a sliding clamp or recruit Mlh1-Pms1 (Figs. 66–8), and its reduced affinity for ATP in both subunits (Fig. 9). Although these effects can be attributed to lack of ATP binding by Msh6, the L1129C/S1137C disulfide bond is not at the nucleotide-binding site; it is located in a region undergoing ATP-induced changes according to deuterium exchange data (42). Thus, inhibition of rapid ATP binding is more likely due to disruption of ATP-induced conformational changes that generate the high affinity ATP-binding site, rather than disruption of the nucleotide-binding site per se. The fact that extensive dialysis of oxidized Msh2-Msh6 L1129C/S1137C against buffer containing ATP without Mg2+ converted it to a state in which trypsin was unable to generate the 75-kDa fragment (Figs. 5 and and1010F) is consistent with a slowed ATP-induced transition and is at odds with models in which the disulfide bond directly disrupts the nucleotide-binding site. Thus, the formation of the disulfide bond appears to have generated an Msh2-Msh6 complex in which the ATP-induced conformational changes occur on the order of hours.
Supplementing oxidized Msh2-Msh6 L1129C/S1137C with ATP reduced the affinity of both its subunits for ATP relative to oxidized Msh2-Msh6 L1129C/S1137C. The unlabeled ATP from the dialysis most likely competed with the added labeled ATP in the UV cross-linking experiment (Fig. 9). The ATP-supplemented oxidized Msh6 L1129C/S1137C subunit appeared to retain more unlabeled ATP than Msh2, resulting in less cross-linked labeled ATP. This result suggests that the C-terminal region of Msh6 affects ATP binding to Msh2, consistent with previous results (28, 41, 52). Interestingly, this ATP-bound form of oxidized Msh2-Msh6 L1129C/S1137C was capable of binding mispaired bases (Fig. 7), but did not form the sliding clamp or recruit Mlh1-Pms1 (Figs. 6 and and8).8). This is most likely because the bound ATP was hydrolyzed to ADP due to the presence of Mg2+ in the Biacore running buffer, which converted it to a mispair-binding proficient form (19, 28), but the slow ATP binding prevented either sliding clamp formation or Mlh1-Pms1 binding.
The fact that in vitro oxidized Msh2-Msh6 L1129C/S1137C is in the open conformation but can be slowly converted to the closed conformation (Figs. 5 and and10)10) indicates that the disulfide bond slows ATP binding to Msh2 and Msh6, and hence the dynamic interconversion between the open and closed forms that would be predicted to occur during MMR. Therefore the C-terminal conformational change in Msh6 plays a role in ATP processing, specifically ATP hydrolysis or ADP exchange for ATP. This, in turn, disrupts the formation of both Msh2-Msh6 sliding clamps and ternary complexes between Msh2-Msh6 and Mlh1-Pms1 and results in an MMR defect.
We thank Kathleen Matthews for the generous gift of LacI protein. We thank Jason Liang and Huilin Zhou for their assistance in analyzing the disulfide bond formation by mass spectrometry.
*This work was supported, in whole or in part, by National Institutes of Health Grants GM50006 and CA092584.
2The abbreviations used are: