From the Section of Genetics and Development, Cornell University, Ithaca, New York 14853-2703
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ABSTRACT |
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In yeast, MSH2 interacts with MSH6 to repair base
pair mismatches and single nucleotide insertion/deletion mismatches and with MSH3 to recognize small loop insertion/deletion mismatches. We
identified a msh6 mutation (msh6-F337A) that
when overexpressed in wild type strains conferred a defect in both
MSH2-MSH6- and MSH2-MSH3-dependent mismatch repair
pathways. Genetic analysis suggested that this phenotype was due to
msh6-F337A sequestering MSH2 and preventing it from interacting with
MSH3 and MSH6. In UV cross-linking, filter binding, and gel retardation
assays, the MSH2-msh6-F337A complex displayed a mismatch recognition
defect. These observations, in conjunction with ATPase and dissociation rate analysis, suggested that MSH2-msh6-F337A formed an unproductive complex that was unable to stably bind to mismatch DNA.
DNA mismatches can arise through DNA replication errors, physical
damage, and heteroduplex formation during genetic recombination. If
left unrepaired, these mismatches become fixed in the genome as
mutations. The best understood mismatch repair system is the Escherichia coli mutHLS long-patch repair pathway (1). A
model for the initiation of mismatch repair by MutH, MutL, and MutS immediately after passage of a DNA replication fork has been developed based upon in vivo studies and an in vitro
mismatch repair system reconstituted from purified components (1-6).
In this model, a mismatch is first recognized and bound by a dimer of
MutS that displays an intrinsic ATPase activity. In a reaction that
requires ATP, a dimer of MutL binds to MutS and then activates the
methylation sensitive endonuclease MutH. Activation of MutH results in
cleavage of the unmethylated DNA strand at hemimethylated d(GATC) sites that are transiently present after replication fork passage, providing an entry point for excision and replication proteins to remove the
mismatch and repair the resulting DNA gap using the parental DNA strand
as a template.
The ability of MutS to both recognize base pair mismatches and trigger
downstream events that can be located several kilobases away from a
mismatch site suggests that it can bind to DNA in at least two
different modes. The first mode allows mismatch recognition, and the
second mode allows MutS protein to use the energy of ATP hydrolysis to
translocate along DNA with MutL so that it can activate MutH at GATC
sites (1, 7, 8). Support for the presence of multiple MutS DNA binding
modes was initially obtained from DNA binding assays showing that MutS
protein can specifically recognize base pair mismatches in the absence
of ATP and that the addition of ATP resulted in the loss of mismatch
binding specificity (8). An elegant electron microscopy analysis
performed by the Griffith laboratory (7) and the Modrich laboratory (9)
provides further evidence for this idea. They showed that MutS can form ATP-dependent loop structures on DNA substrates that
contain a mismatch site. The size of loop was dependent on incubation
time and the presence of a mismatch site. They hypothesized that MutS can bind to a mismatch substrate in an ATP independent step. After recognition, a second binding mode is activated through an
ATP-dependent conformational change in MutS resulting in
the loss of its mismatch recognition functions and a shift into a mode
that allows it to bidirectionally translocate along DNA away from a
mismatch site. This activity results in the formation of loop
structures that are thought to serve as topological intermediates for
strand excision and allow MutS to scan along DNA until it identifies
and activates MutH bound at GATC sites.
At present, little is known about which domains in MutS are important
for mismatch recognition. Recently, a DNA cross-linking analysis
performed by Malkov et al. (10) revealed that the
phenylalanine 39 residue in Thermus aquaticus MutS was
critical for photocross-linking of MutS to a mismatch substrate. They
also showed that a mutant derivative of MutS, mutS-F39A, displayed a
reduced affinity for mismatch substrate. Although this study identified
a domain that is important for mismatch recognition, it did not address
whether this domain functions in general DNA binding and/or in mismatch binding and whether it functions during one or more DNA binding modes.
Whereas studies in bacteria have greatly increased our understanding of
mismatch repair, analogous repair systems in eukaryotes appear to be
more complex (1, 11, 12). Several homologs of mutS and
mutL have been identified in eukaryotic organisms. A feature
of the MutS homologs is they all contain a highly conserved ATP binding
domain. In the yeast Saccharomyces cerevisiae, six mutS homologs and four mutL homologs have been
identified, and the gene products of the MSH2, MSH3, MSH6,
MLH1, and PMS1 genes have been identified as components
of nuclear mismatch repair. Several studies have suggested that during
mismatch recognition, MSH2 acts a scaffold for mismatch binding,
whereas MSH6 and MSH3 act as specificity factors (13-17). This
hypothesis was based on genetic studies showing that msh2
mutants are defective in the repair of both base pair and loop
insertion/deletion mismatches, whereas msh3 mutants are
principally defective in the repair of 2-4-nucleotide loop mismatches
and msh6 mutants are principally defective in the repair of
base-base and single nucleotide insertion/deletion mismatches (14, 16,
18). Interestingly, MSH3 and MSH6 functions have been shown to overlap,
as msh3 and msh6 single mutants do not display as
strong a defect in mismatch repair in their respective mismatch repair
assays as do msh2 mutants. msh3 msh6 double
mutants, however, display a mutator phenotype that is identical to that observed in msh2 mutants (14, 16). Subsequent biochemical studies have shown that MSH2 can interact with MSH3 and MSH6 to form
distinct complexes that bind subsets of mismatches predicted by the
genetic studies and that an MLH1-PMS1 heterodimer can interact with
these complexes when they are bound to a mismatch site (13, 17,
19-22).
In this study, we used genetic and biochemical assays to investigate
the mismatch recognition properties of MSH2-MSH6. Based on the
pioneering studies performed by Malkov et al. (10), we created site-specific mutations in putative DNA binding domains in the
MSH2 and MSH6 subunits and examined the effect of these mutations in UV
cross-linking and DNA binding studies. Our analysis, described below,
is consistent with MSH2 acting primarily as a scaffold for interactions
with MSH6 and MSH3 that confer specificity to mismatch recognition.
Interestingly, a mutation in MSH6 (msh6-F337A) was
identified that conferred a dominant negative phenotype that was
consistent with msh6-F337A sequestering MSH2 from functioning in
mismatch recognition. This analysis also suggested that subunit interactions, mismatch recognition functions, and ATPase activities are
coordinated to allow for the recognition and repair of DNA mismatches.
Strains and Genetic Procedures--
The E. coli
strain RDK2290 (F', Media, Reagents, and Chemicals--
E. coli strains
were grown in LB broth or on LB agar that was supplemented with 100 µg/ml ampicillin when required (29). Yeast strains were grown in
either YPD or minimal selective media (30). 2% glucose, 2% sucrose,
3% glycerol, 2% lactate, and 2% galactose were included as carbon
sources as indicated. When required, canavanine (Sigma) was included in
minimal selective media lacking arginine at 60 mg/liter. 5-FOA (U. S.
Biological, San Antonio, TX) plates were prepared as described (30).
Polyclonal antibodies raised against MSH2 and MSH6 were obtained as
described previously (15).
Nucleic Acid Techniques--
Site-directed mutagenesis of
MSH2 and MSH6 to create the msh2-Y42A,
msh6-F337A, msh6-G335D, and msh6D343A alleles was performed using
overlapping polymerase chain reaction mutagenesis (31). Oligonucleotide
synthesis and double-stranded DNA sequencing of the entire subcloned
fragments used to make the msh2 and msh6 mutant
alleles were performed at the Cornell Biotechnology
Analytical-Synthesis Facility (Ithaca, NY). All restriction
endonucleases, T4 polynucleotide kinase, T4 DNA ligase, T4 DNA
polymerase, and Vent polymerase were from New England Biolabs (Beverly,
MA) and used according to the manufacturers' specifications. Plasmid
DNA was isolated by alkaline lysis, and all DNA manipulations were
performed as described previously (32).
Oligonucleotides Used In DNA Binding Studies--
Duplex
oligonucleotide DNA substrates used in filter binding, ATPase and UV
cross-linking studies were created from the following oligonucleotides:
1) 5'-ATGTGAATCAGTATGGTTCCTATCTGCTGAAGGAAAT-3', 2)
5'-ATTTCCTTCAGCAGATAGGAACCATACTGATTCACAT-3', 3)
5'-ATGTGAATCAGTATGAGTTCCTATCTGCTGAAGGAAAT-3', 4)
5'-ATGTGAATCAGTATGGTTCXCTATCTGCTGAAGGAAAT-3', 5)
5'-ATTTCCTTCAGCAGATAGAGAACCATACTGATTCACAT-3', and 6)
5'-ATTTCCTTCAGCAGATAGGAACCATACTGATTCACAT-3'. All oligonucleotides were synthesized by Operon (Alameda, CA). In oligonucleotide 4, X refers to the position of an iododeoxyuridine residue.
Iododeoxyuridine (5-IdUrd) substituted derivatives were synthesized by
Operon using 5-I-dU-CE phosphoramidite (Glen Research Corp.). 5-IdUrd
oligonucleotides were shielded from light and handled according to
Operon specifications. Homoduplex and +1 mismatch DNA substrates (13)
used in the filter binding and ATPase studies were created by annealing
oligonucleotides 1 and 2 and oligonucleotides 3 and 2, respectively
(32). In the UV cross-linking studies, the AT-I homoduplex substrate
was formed by annealing oligonucleotides 4 and 5, and the Biochemical Techniques--
Overexpression and purification of
MSH2-MSH6 and msh2-Y42A-MSH6 was performed as described previously (13,
15). MSH2-msh6-F337A and MSH2-msh6-F337A/G987D were purified using the
same procedure with the exception that these purifications required
loading and washing the ssDNA cellulose column at 100 mM
NaCl Buffer A (25 mM Tris, pH 7.5, 1 mM EDTA,
10 mM UV Cross-linking--
In the UV cross-linking studies, 120 nM (1.5 µg) MSH2-MSH6 or MSH2-msh6-F337A was incubated in
50-µl reactions for 15 min at 30 °C with 20 nM
32P-labeled duplex DNA substrate in buffer containing 25 mM Tris, pH 7.5, 0.01 mM EDTA, 0.1 mM dithiothreitol, and 2.0 mM
MgCl2. The immunoprecipitation reactions presented in Fig.
1B and the cross-linking reactions presented in Fig.
4A were performed in the absence of MgCl2.
Samples were then transferred to polystyrene tubes and irradiated in a
Rayonet Photochemical reactor, model RPR100 (Branford, CT) equipped
with 350 nm bulbs for 0-60 min in the absence of visible light. After
UV irradiation, samples were immediately boiled for 3 min in SDS-PAGE
sample buffer and loaded onto an 8% SDS-PAGE gel for analysis by gel
electrophoresis. Following electrophoresis, the SDS-PAGE gel was
stained with Coomassie Blue, dried, and visualized using a
phosphorimager screen. Immunoprecipitations of denatured cross-linked
samples were carried out using methods described by Iaccarino et
al. (17). 10 µl of cross-linked protein-DNA samples were boiled
for 5 min in SDS sample buffer (New England Biolabs) and then added at
a 10% final concentration to reactions containing anti-MSH2,
anti-MSH6, or no antibody suspended in 0.1 M NaCl Buffer A. After a 45-min incubation at 4 °C, 30 µl of protein A-Sepharose
were added to each reaction followed by another 45-min incubation.
Protein A beads were spun down at 2500 rpm for 45 s. Supernatant
was removed from the beads, which were then washed with 200 µl of 0.1 M NaCl Buffer A. Beads were spun again, supernatant removed, and the beads were resuspended in SDS sample buffer, boiled
for 5 min, and loaded onto SDS-PAGE gels. All gels were visualized
using a phosphorimager screen (Molecular Dynamics), and bands were
quantified using Mac IQ Imagequant software.
DNA Binding, Dissociation, and ATPase Assays--
DNA filter
binding assays were performed as described previously (25). The
standard buffer for the DNA binding assay contained 25 mM
Tris, pH 7.5, 0.1 mM dithiothreitol, 0.01 mM
EDTA, and 40 µg/ml bovine serum albumin. When indicated, ATP was
added to a final concentration of 1.5-2.0 mM. Binding was
performed at 30 °C for 15 min in a 60-µl reaction. This time point
was chosen because maximal binding for MSH2-MSH6 to oligonucleotide
substrate was observed at 15 min, and no change in binding was observed in incubations that ranged from 15 to 90 min. Each reaction contained 16.7 nM 32P-labeled +1 substrate, 0-333
nM unlabeled +1 or homoduplex competitor, and 0.15-1.5
µg (10-100 nM) of MSH2-MSH6, msh2-Y42A-MSH6, or
MSH2-msh6-F337A. Following incubation, samples were analyzed by filter
binding to KOH-treated nitrocellulose filters (Schleicher and Schuell, Keene, NH) (34) as described by Chi and Kolodner (35). Dissociation of
wild type and mutant MSH2-MSH6-DNA complexes was examined in the
standard DNA binding buffer conditions as described in the text.
Wild type and mutant MSH2-MSH6-DNA complexes were also examined in gel
retardation assays. Binding was performed at 30 °C for 15 min in
10-µl reactions containing 360 nM (0.9 µg) MSH2-MSH6 or
MSH2-msh6-F337A, 10 nM 32P-labeled +1
substrate, 25 mM Tris, pH 7.5, and 0.1 mM
dithiothreitol. Unlabeled +1 and homoduplex substrate were added as
competitor as indicated in Fig. 3. After incubation, an equal volume of
buffer containing 50 mM Tris, pH 7.5, 3 mM
EDTA, and 50% glycerol was added to the binding reaction, and the
samples were loaded onto a 4% polyacrylamide-bis (29:1) gel. The gel
was run at room temperature in 1× Tris-Borate EDTA buffer (31) at 200 V, and the extent of binding was determined after the gel was dried
using a phosphorimager (Molecular Dynamics) and analyzed with Mac IQ
Imagequant software.
The affinity of MSH2-MSH6, msh2-Y42A-MSH6, and MSH2-msh6-F337A for
ssDNA cellulose was measured using a minicolumn protein retention
assay. Approximately 30 µg of wild type and mutant complex were
applied to a 100-µl bed volume ssDNA cellulose column in Buffer A
containing 0.1 M NaCl. Each of these columns was eluted with successive 10 column volumes of 1× buffer containing 0.1, 0.25, 0.30, and 1.0 M NaCl buffer. The amount of protein eluted with each wash was determined using Bradford assays (36).
ATPase assays were performed in 60-µl reactions containing 20 nM (0.3 µg) MSH2-MSH6, msh2-Y42A-MSH6, MSH2-msh6-F337A,
or MSH2-msh6-F337A/G987D, 1.2-100 µM
[ UV Cross-linking Indicates That Both MSH2 and MSH6 Are Cross-linked
to a Duplex Oligonucleotide at the Site of a Mismatch--
Previous
studies have led to the proposal that the MSH6 subunit of the S. cerevisiae MSH2-MSH6 complex, which is principally involved in
repairing base-base and single insertion/deletion mismatches, acts as a
mismatch recognition specificity factor for MSH2 (13-15). A similar
proposal for the human mismatch repair proteins, based on UV
cross-linking analysis of hMSH2-hMSH6 to mismatch substrates, was made
by Iaccarino et al. (17), in which it was found that only
the hMSH6 subunit could be cross-linked to a mismatch substrate.
Recently, Malkov et al. (10) demonstrated that mismatch DNA
substrates containing a iododeoxyuridine residue (5-IdUrd) at the site
of the mismatch could be specifically cross-linked to T. aquaticus MutS at phenylalanine 39, which is located in a region
that is highly conserved among MutS homologs. Malkov et al.
(10) also showed that a substitution of alanine at this position
(mutS-F39A) abolished binding of MutS to mismatch
substrates. We performed a similar analysis to identify residues
involved in yeast MSH2-MSH6 mismatch recognition. 38-mer homoduplex
(AT-I) and single nucleotide loop substrates (
UV-dependent cross-linking of mismatch and homoduplex
substrates to the MSH2-MSH6 complex was observed by incubating
32P-labeled
Immunoprecipitation analysis was performed on denatured cross-linked
species to determine whether Bands A-C contained MSH2 or MSH6. As
shown in Fig. 1B, Band B was recognized and
immunoprecipitated only by MSH6 antibody (lane 2), and Band
C was immunoprecipitated only by MSH2 antibody (lane 3).
Band A was recognized by both MSH2- and MSH6-specific antibodies,
suggesting that this species contained cross-links that physically
joined all three components of the MSH2-MSH6- The msh6-F337A Mutation Conferred a Mismatch Repair Defect--
We
constructed alanine substitutions at residue 42 in MSH2
(msh2-Y42A) and residue 337 in MSH6 (msh6-F337A)
based on the fact that these residues can be aligned with the T. aquaticus MutS Phe-39 residue (Fig.
2A) (10). The
msh2-Y42A and msh6-F337A alleles were cloned into
GAL10, 2 µ overexpression plasmids; galactose induction
studies indicated that each mutant was overexpressed to levels similar
to that observed for the corresponding wild type proteins (Fig.
2B and data not shown). The effect of these mutations was
then assessed for defects in mismatch repair.
The msh2-Y42A and msh6-F337A alleles were tested
to determine whether they can complement the mutator phenotype
exhibited by msh2 The msh6-F337A Mutation Confers a Dominant Negative Mutator
Phenotype That Is Similar to the Mutator Phenotype Observed in msh2 MSH2 Overexpression Suppresses the Dominant Negative Phenotype
Conferred by Overexpressing msh6-F337A--
As described above, the
msh6-F337A mutation conferred a dominant negative phenotype
that was similar to the mutator phenotype observed in
msh2 Analysis of msh2-Y42A-MSH6 and MSH2-msh6-F337A
Complexes--
MSH2-msh6-F337A and msh2-Y42A-MSH6 complexes were
purified as described in Fig. 2B and under "Experimental
Procedures." The complexes each contained a subunit stoichiometry
that was similar to the wild type complex, suggesting that the
site-specific mutations did not compromise interactions between
subunits or disrupt the overall structure of the mutant proteins.
Approximately the same yield of purified protein per unit of weight of
induced cells was observed for each of the wild type and mutant
heterodimers, and coimmunoprecipitation and gel filtration analysis
indicated that the mutant complexes behaved similarly to wild type in
these assays and formed stable heterodimers (data not shown) (13, 15).
In ATPase assays performed in the absence of DNA, the ATPase activities
of MSH2-msh6-F337A and MSH2-MSH6 were identical (see Fig. 5) (15)
(Km and Vmax were identical;
data not shown). The msh2-Y42A-MSH6 complex was indistinguishable from wild type in all genetic and biochemical assays described in this report (data not shown).
We obtained evidence that the MSH2-msh6-F337A complex was defective in
DNA binding during the purification of the complex. Under conditions
where the wild type complex was retained on a ssDNA cellulose column
(200 mM NaCl), the MSH2-msh6-F337A complex was not
retained. The purification procedure for the MSH2-msh6-F337A complex
was then modified by loading MSH2-msh6-F337A fractions onto the ssDNA
column at 100 mM NaCl (see under "Experimental Procedures"). To better quantify the binding of MSH2-MSH6 and MSH2-msh6-F337A complexes to DNA containing an insertion mismatch or
homoduplex DNA, filter binding assays were performed with +1 and
homoduplex DNA substrates (Fig. 2C; Refs. 13 and 33; see under "Experimental Procedures"). The 37-mer homoduplex substrate is identical to the +1 substrate with the exception that the +1 substrate contains an adenine insertion after base pair 15. In this
assay, increasing amounts of protein were added to a constant amount
(16.7 nM) of 32P-labeled +1 or homoduplex DNA.
As seen in Fig. 2C, MSH2-MSH6 binding to +1 substrate was
maximal at 3.6 pmol (60 nM) of protein with nearly 50% of
total counts bound. In contrast, less than 10% of total counts were
bound in a binding reaction containing 60 nM
MSH2-msh6-F337A complex and either homoduplex or +1 substrate. In
addition to a general DNA binding defect, the MSH2-msh6-F337A complex
appeared to display a defect in discriminating between +1 and
homoduplex substrates. At 0.6 pmol (10 nM) of complex, 33 and 17% of +1 and homoduplex substrate, respectively, were bound to
MSH2-MSH6 complex. In contrast, 8.5 and 7.5% of +1 and homoduplex
substrate, respectively, were bound by 3.6 pmol (60 nM) of
MSH2-msh6-F337A complex (Fig. 2C). It is important to note that DNA binding in all of the reactions was inhibited at high protein
concentrations because the NaCl present in the protein preparations
inhibited DNA binding when present at final concentrations greater than
50 mM; this is illustrated by the fact that the percentage of binding for all of the reactions reached a plateau at protein levels
greater than 3.6 pmol and started to decrease at levels greater than
7.2 pmol (Fig. 2C and data not shown).
MSH2-msh6-F337A Is Defective in Both Mismatch Binding and in
General DNA Binding--
To further explore the mismatch recognition
properties of wild type and mutant complexes, the DNA binding activity
of MSH2-MSH6 and MSH2-msh6-F337A were analyzed in filter binding and
gel retardation competition assays. In the filter binding assays (Fig.
3, A and B) 20 nM (0.3 µg) MSH2-MSH6 or MSH2-msh6-F337A complex was
incubated with 16.7 nM 32P-labeled +1 substrate
in the presence and absence of various concentrations of unlabeled +1
or homoduplex competitor. In these reactions the molar ratio of protein
to DNA substrate was approximately 1:1 and was chosen based on the
binding titrations shown in Fig. 2C. Mismatch binding
between the two competitors was measured by determining the maximal
horizontal separation between the binding curves (13, 33). As shown for
the MSH2-MSH6 complex in Fig. 3A, the +1 substrate is an
approximately 8-fold more effective competitor than homoduplex
substrate; this result is consistent with that found previously (13,
15). In competitions involving the MSH2-msh6-F337A complex (Fig.
3B), the +1 and homoduplex competitions were similar and
suggested a dramatic decrease in mismatch binding specificity compared
with the MSH2-MSH6 complex. A loss in mismatch binding specificity was
also observed in MSH2-MSH6 binding experiments performed in the
presence of 2.0 mM ATP; ATP addition was previously shown
to eliminate mismatch recognition (13) (data not shown).
In gel retardation assays, 360 nM (0.9 µg) MSH2-MSH6 or
MSH2-msh6-F337A complex was incubated with 10 nM
32P-labeled +1 substrate in the presence and absence of
various concentrations of unlabeled +1 or homoduplex competitor
substrates. As shown in Fig. 3, D and E, in the
absence of unlabeled competitor, MSH2-MSH6 completely retarded the
mobility of the 32P-labeled +1 substrate; in competition
reactions, the +1 unlabeled substrate was an approximately 20-fold more
effective competitor than homoduplex substrate. In reactions involving
MSH2-msh6-F337A, a discrete gel shift was not observed; instead, a
diffuse gel shift pattern was observed that spanned from the position
of unbound 32P-labeled +1 substrate to the position of the
MSH2-MSH6-+1 complex (Fig. 3D).
ATP Enhances UV Cross-linking and Binding of MSH2-msh6-F337A to
DNA--
UV DNA cross-linking studies were performed to examine in
greater detail the mismatch binding properties of the wild type and
MSH2-msh6-F337A complexes. These experiments were carried out as
described under "Experimental Procedures." 24 nM (0.3 µg) MSH2-MSH6 or MSH2-msh6-F337A was incubated with 20 nM
32P-labeled
In filter binding assays, we compared the effect of 1.5 mM
ATP on the binding of +1 and homoduplex DNA by the MSH2-MSH6 and MSH2-msh6-F337A complexes (Fig. 4B). In reactions containing
16.7 nM +1 substrate and 20 nM (0.3 µg)
MSH2-MSH6, 34% of total 32P-labeled +1 substrate was
bound; in identical reactions containing MSH2-msh6-F337A, 7% total
binding was observed. The addition of ATP to the MSH2-MSH6 +1 binding
reaction resulted in a significant reduction in binding from 34 to 25%
(Fig. 4B). In reactions involving MSH2-msh6-F337A, the
addition of ATP resulted in an increase in +1 substrate binding from 7 to 14%. In binding reactions involving homoduplex DNA that were
performed under the same conditions, the addition of ATP to either the
MSH2-MSH6 or MSH2-msh6-F337A reactions resulted in both cases in an
increase in homoduplex binding (Fig. 4B). These experiments
were performed at several different ATP concentrations with
qualitatively similar results (data not shown). Taken together, the
filter binding results are consistent with those obtained in the
UV-cross-linking experiments.
Mismatch Substrates Specifically Stimulate the ATPase Activity of
MSH2-msh6-F337A While Inhibiting the ATPase Activity of
MSH2-MSH6--
Previous analysis of the yeast MSH2-MSH6 complex
indicated that under low salt conditions (<50 mM NaCl),
the ATPase activity of MSH2-MSH6 was reduced in the presence of
homoduplex DNA and reduced even further in the presence of +1 substrate
(15, 39) (Fig. 5). Genetic analysis of
the msh6-F337A mutation and UV cross-linking analysis of the
MSH2-msh6-F337A complex encouraged us to examine the effect of DNA
substrate on the ATPase activity of the MSH2-msh6-F337A complex. As
described earlier, in the absence of DNA, the ATPase activity of
MSH2-msh6-F337A was identical to that observed for the MSH2-MSH6
complex (Fig. 5B). However, unlike the MSH2-MSH6 complex, +1
substrate addition (167 nM) to the MSH2-msh6-F337A ATPase
reaction resulted in a nearly 4-fold increase in its ATPase activity
(Fig. 5). The addition at 167 nM single-stranded
oligonucleotide or a mismatch substrate (+2) that is not recognized by
the MSH2-MSH6 complex (13) did not affect the ATPase activity of the
MSH2-msh6-F337A complex, suggesting that the enhanced ATPase activity
was not the result of a contaminating activity such as that conferred by a helicase (data not shown).
The stimulation of the MSH2-msh6-F337A ATPase activity by the +1
substrate was further investigated in experiments performed at a
constant concentration of ATP (33 µM) and varying amounts of +1 or homoduplex substrate. As shown in Fig. 5C, at all
concentrations tested, the +1 substrate displayed a stronger inhibition
of MSH2-MSH6 ATPase activity than the homoduplex substrate, and
increasing concentrations of either DNA substrate resulted in a gradual
reduction in ATPase activity (Fig. 5). In contrast, increasing
concentrations of homoduplex DNA had no effect on the ATPase activity
of the MSH2-msh6-F337A complex; however, the +1 substrate displayed a concentration-dependent effect on ATPase activity. At low
+1 substrate concentrations, the +1 substrate stimulated the ATPase
activity of the MSH2-msh6-F337A complex with maximal stimulation
observed at 167 nM +1 substrate. At +1 substrate
concentrations greater than 167 nM, the ATPase activity
progressively decreased; at 660 nM +1 substrate, the ATPase
activity of the MSH2-msh6-F337A complex was inhibited to a level below
that observed with homoduplex substrate.
Studamire et al. (15) showed that the ATPase activity of
complexes defective in the MSH2 ATPase was still modulated by mismatch and homoduplex substrates; this modulation, however, was not observed in complexes deficient in the MSH6 ATPase activity. To test whether the
enhancement of the MSH2-msh6-F337A ATPase activity by +1 substrate was
due to the ATPase activity of the msh6-F337A subunit, we purified a
complex containing both the msh6-F337A mutation and a
msh6 mutation (msh6-G987D) that was shown
previously to severely inhibit the MSH6 ATPase activity (15).
MSH2-msh6-F337A/G987D complex was purified as was described for
MSH2-msh6-F337A. A similar yield, purity, and subunit ratio were
obtained for purified MSH2-msh6-F337A/G987D compared with MSH2-MSH6 and
MSH2-msh6-F337A (data not shown). We also found that the ratio of MSH2
to MSH6 in immunoprecipitations involving MSH2-specific antibody and
crude extracts was indistinguishable for wild type and
MSH2-msh6-F337A/G987D complexes (13) (data not shown). In DNA binding
assays, the MSH2-msh6-F337A/G987D complex displayed a defect in
mismatch binding that was indistinguishable from that observed for the
MSH2-msh6-F337A complex (Fig. 3C).
The ATPase activity of the MSH2-msh6-F337A/G987D complex was tested in
the presence and absence of DNA substrate (Fig.
6). In the absence of DNA substrate, the
ATPase activity of the MSH2-msh6-F337A/G987D complex was less than half
of that observed for the MSH2-msh6-F337A complex but was similar to
that observed for the MSH2-msh6-G987D complex characterized previously
(15). The ATPase activity of the MSH2-msh6-F337A/G987D complex was
unaffected by the presence of either homoduplex or +1 substrate; this
result is consistent with the MSH6 ATPase playing an important role in
the enhancement of the MSH2-msh6-F337A ATPase by +1 substrate and also
supports the idea that the MSH6 ATPase plays a critical role in
signaling mismatch recognition (15).
Dissociation of the MSH2-MSH6 and MSH2-msh6-F337A-Oligonucleotide
Complexes--
The finding that MSH2-msh6-F337A was defective in
mismatch binding but was capable of detecting mismatch substrates as
demonstrated by mismatch-dependent changes in its ATPase
activity suggested that it was incapable of forming stable complexes
with mismatch substrates. We tested this idea by performing the
dissociation experiments presented in Fig.
7. In these studies, 5.7 nM
(0.3 µg) MSH2-MSH6 and MSH2-msh6-F337A were preincubated with either 1.4 nM 32P-labeled homoduplex or +1 substrate
for 15 min, after which a 300-fold excess of unlabeled competitor was
added. The stability of the original protein-DNA complexes was then
measured as a function of time in filter binding assays.
For the MSH2-MSH6-homoduplex complex, a rapid decay was observed
immediately after unlabeled homoduplex substrate was added. As shown in
Fig. 7A, approximately 70% of the MSH2-MSH6-homoduplex complexes dissociated within 1 min after unlabeled homoduplex was
added. For the MSH2-MSH6-+1 complex, the addition of unlabeled +1
substrate resulted in a biphasic decay, indicating the presence of two
types of complexes. One complex was unstable, with a
t1/2 of ~ 5 min, and the other was stable,
with a t1/2 of ~ 80 min (Fig. 7A).
These results indicated that ~40% of the MSH2-MSH6-+1 complexes
formed stable complexes. In experiments involving MSH2-msh6-F337A,
similar dissociation profiles were observed for the
MSH2-msh6-F337A-homoduplex and MSH2-msh6-F337A-+1 complexes, and the
dissociation of these complexes was more rapid than was observed for
the MSH2-MSH6-homoduplex complex (Fig. 7B). The inclusion of
2.0 mM ATP during the MSH2-MSH6-+1 preincubation phase
resulted in a dissociation profile that was similar to that found for
the MSH2-MSH6-homoduplex preincubated in the presence or absence of ATP
(Fig. 7C). The addition of ATP during the formation of
MSH2-msh6-F337A-homoduplex and MSH2-msh6-F337A-+1 complexes had no
effect on the dissociation of these complexes (data not shown).
The finding that MSH2-msh6-F337A complexed with +1 or homoduplex
substrate displayed a dissociation profile that was even more rapid
than was observed for MSH2-MSH6 provides an explanation for why we were
able to detect MSH2-msh6-F337A-+1 complexes in filter binding but were
unable to observe discrete complexes in the gel shift assay. We
hypothesize that the MSH2-msh6-F337A-+1 complexes were detected in
filter binding because these unstable complexes have a greater chance
to be recovered on nitrocellulose filters, as the filter binding step
was rapid and was performed in standard DNA binding buffer (see under
"Experimental Procedures"). We believe that these unstable
complexes were not observed in the gel shift assay because the gel
electrophoresis step provides an extended time period to allow for
dissociation of the complex. This argument can also explain why a
greater mismatch binding specificity for MSH2-MSH6 was observed in the
gel shift assay compared with the filter binding assay. An important
caveat in our hypothesis is that unlike gel retardation analysis, in
which specific protein-DNA complexes can be identified, it is difficult to determine in filter binding whether the complexes that were formed
resulted from specific protein-DNA interactions or resulted from
nonspecific protein-DNA aggregates.
The msh6-F337A Mutation Confers a Mismatch Recognition
Defect--
We analyzed mutations in MSH2
(msh2-Y42A) and MSH6 (msh6-F337A) that
are analogous to a mutation in T. aquaticus MutS that disrupts binding to mismatch DNA (10). Genetic analysis indicated that
the msh6-F337A but not the msh2-Y42A mutation
conferred a mismatch repair defect. In UV cross-linking and DNA binding
assays that involved titration analysis and substrate competitions,
MSH2-msh6-F337A displayed defects in both general and mismatch-specific
DNA binding (Figs. 2 and 3).
Additional support for the idea that the msh6-F337A mutation
conferred a mismatch binding defect came from experiments that examined
the effect of ATP on the mismatch binding activity of wild type and
MSH2-msh6-F337A complexes. In one set of experiments that involved
measuring the ATPase activity of complexes in the presence of DNA
substrate, we observed that in contrast to wild type complex, the
ATPase activity of MSH2-msh6-F337A was specifically stimulated by +1
substrate (Fig. 5). However, as the concentration of +1 substrate was
raised above 167 nM, the ATPase activity of the
MSH2-msh6-F337A complex decreased and eventually reached a level that
was below that observed in the absence of +1 substrate. Homoduplex DNA
did not affect ATPase activity, suggesting that msh6-F337A confers
specific defects in mismatch recognition that can be overcome in the
presence of a sufficiently high concentration of +1 substrate, leading
to a wild type response (decrease) in ATPase activity.
In a second set of experiments, the binding of MSH2-msh6-F337A to
single nucleotide insertion/deletion (+1, A Model for MSH2-MSH6 Binding to Base Pair Mismatches--
The
observation that MSH2-MSH6 forms unstable and stable complexes with DNA
provides an explanation for how the modest selectivity of the complex
for mismatches observed in filter binding assays could result in
efficient repair. Based on the off rate studies presented in Fig. 7, we
propose that MSH2-MSH6 initially binds to DNA and forms an unstable and
nonspecific complex that rapidly dissociates, allowing a new cycle of
binding to occur. A proportion of the complexes that form are stable,
and the formation of these stable complexes requires the presence of
mispaired DNA in the substrate. The stability of the MSH2-MSH6-+1
complex in dissociation experiments suggests that MSH2-MSH6 undergoes a
change in binding conformation when it is bound to a mismatch
substrate; this results in the stable binding of MSH2-MSH6 to a
mismatch and provides a recognition signal for interactions with repair
proteins that are involved in subsequent steps. Such a mechanism could
amplify the selectivity of the complex for mispaired bases (24, 33). A
similar binding mode has been demonstrated for the binding of MSH1,
MSH2, and hMSH2-hMSH6 to mismatch DNA (9, 24, 33). We propose that
because the MSH2-msh6-F337A complex is unable to form a stable complex
on mismatch DNA, it fails to provide a mismatch recognition signal for
interactions with mismatch repair proteins and instead undergoes futile
cycles of mismatch binding and release that involves ATP hydrolysis by
the MSH6 subunit.
The above model for mismatch binding does not address the genetic data
presented in Table I that showed that strains overexpressing msh6-F337A
displayed a strong dominant negative phenotype in both the mutator and
DNA slippage assays. Overexpression of MSH6, on the other hand, did not
dramatically affect mismatch repair in either assay. As shown in the
biochemical assays presented in this paper, the MSH2-msh6-F337A complex
was unable to stably bind to mismatch DNA substrates. A hypothesis that
can account for these genetic and biochemical observations is that in
wild type cells, MSH2 is able to switch between MSH3 and MSH6 subunits
when present in a mismatch binding competent complex. In such a
scenario, the presence of a high level of one MSH partner would not
prevent interactions with a partner present at lower levels. Based on this idea, we suggest that the inability of MSH2-msh6-F337A to stably
bind to DNA prevents MSH2 within the mutant complex from switching
between MSH3 and MSH6 subunits and thus withdraws MSH2 from pools
available for repair. Experiments to test this idea are in progress.
Does MSH2 Contribute to Mismatch Binding?--
The observation
that the MSH2-msh6-F337A complex is defective in mismatch binding
whereas its ATPase activity responds to the presence of a mismatch
suggests that additional residues in the MSH2-MSH6 complex are involved
in mismatch recognition; this idea is also supported by the finding
that the mutant complex showed residual cross-linking to the +1
substrate (Fig. 4). Does the entire mismatch recognition activity of
the MSH2-MSH6 complex reside in the MSH6 subunit, or is MSH2 also
involved? Previous analysis of MSH2 and MSH6 purified individually
indicated that neither polypeptide demonstrated the mismatch binding
specificity displayed by the complex (13, 25). In cross-linking
analysis, we observed a Band C at lower intensity that was shown by
immunoprecipitation analysis to contain MSH2 (Figs. 1, 4). The
intensity of this band could be increased in binding reactions that
omitted magnesium chloride or in reactions that involved incubating
MSH2-MSH6 complex with DNA substrates containing 5-IdUrd substitutions
at different distances away from the mismatch
site.3 At present, these data
cannot distinguish whether MSH2 is playing a role in general DNA
binding or in recognition. In an initial attempt to investigate the
putative mismatch binding domain shown in Fig. 2A, we made
two substitutions in MSH6 (msh6-G335D and msh6-D343A) that map to residues that are in the vicinity of
Phe-337 and are highly conserved in MutS homologs. Unfortunately, these substitutions resulted in the destabilization of the mutant msh6 proteins and prevented their purification (data not shown). We are
currently analyzing the contribution of the MSH2 and MSH6 subunits in
mismatch recognition by examining a set of msh2 and msh6 mutants that display genetic phenotypes consistent with
a defect in mismatch binding.
The MSH6 ATPase Acts as a Signaling Factor in Mismatch
Recognition--
Previously, Studamire et al. (15)
individually mutagenized the ATP binding domains of MSH2 and MSH6 and
tested the mutant complexes for ATPase activity in the presence and
absence of mismatch substrate. Their results supported a role for MSH6
as a mismatch recognition signaling factor, as complexes containing
only a functional MSH6 ATPase could be modulated by mismatch binding in
a manner qualitatively similar to that observed for the MSH2-MSH6
complex (Fig. 5). The ATPase activity of complexes that contained only a functional MSH2 ATPase was unchanged by the presence of DNA substrate
(15). The data presented in this paper provide further support for this
idea, as biochemical analysis showed that the enhanced ATPase activity
of the MSH2-msh6-F337A complex in the presence of a mismatch was
eliminated if a second mutation was introduced into the MSH6 subunit
that inactivated the MSH6 ATPase (MSH2-msh6-F337A/G987D). We
hypothesize that the MSH6-dependent mismatch signaling
activity plays an important role in recruiting downstream mismatch
repair factors, such as the MLH1-PMS1 complex (15). Recent data from
the Prakash laboratory (22) are consistent with this idea, as they
observed that ATP was required for the assembly of a ternary complex
consisting of a DNA mismatch and the yeast MSH2-MSH6 and MLH1-PMS1 complexes.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
80dlacZ
M15,
(lacZYA-argF)U169, endA1, rec A1,
hsdR17(rK ,
mK+), deoR, thi-1,
supE44,
-gyrA96, relA1) was kindly
provided by R. Kolodner and was used to amplify and manipulate all
plasmids described in this paper. Dominance, complementation, and
suppression studies were performed using the S. cerevisiae
strain FY23 (MATa, ura3-52, leu2
1, trp1
63) (23) and
msh2 (EAY281-MATa, ura3-52, leu2
1, trp1
63,
msh2
::hisG) and msh6 (EAY338-MATa,
ura3-52, leu2
1, trp1
63, msh6
::hisG)
derivatives. Strain were transformed using the lithium acetate method
(24) with the following episomal vectors individually or in
combination: pEAE51 (GAL10-MSH6, TRP1, 2 µ) (13), pEAE84
(GAL10-msh6-G987D, TRP1, 2 µ) (15), pEAE20 (GAL10-MSH2, URA3, 2 µ) (25), pEAE86 (GAL10-MSH2,
TRP1, 2 µ) (15), pEAE95 (GAL10-MSH6, URA3, 2 µ;
this paper), pEAE82 (GAL10-MSH3, LEU2, 2 µ; this paper),
pEAE88 (GAL10-msh6-F337A, TRP1, 2 µ; this paper), pEAE89
(GAL10-msh6-F337A/G987D, TRP1, 2 µ; this paper), pAP2
(GAL10, URA3, 2 µ; kindly provided by Arlen Johnson),
pRS424 (TRP1, 2 µ) (26), and pEAE91
(GAL10-msh2-Y42A, URA3, 2 µ; this paper). The S. cerevisiae strain BJ5464 (MAT
, ura3-52, trp1, leu2
1,
his3
200, pep4::HIS3, prb1
1.6R, can1, GAL) was
obtained from the Yeast Genetic Stock Center and was used for the
overexpression and purification of MSH2-MSH6 and the mutant derivative
complexes. MSH2-MSH6 complex was purified from BJ5464 transformed with
pEAE9 and pEAE51, MSH2-msh6-F337A was from BJ5464 transformed with
pEAE9 and pEAE88, and msh2-Y42A-MSH6 was from BJ5464 transformed with pEAE90 and pEAE51. Mutation rates were obtained after determining the
median frequency (of 11 colonies) of forward mutations to canavanine
resistance and DNA slippage rates were obtained by determining the
median frequency of frameshift events (of 11 colonies) that resulted in
resistance to 5-fluoroorotic acid
(5-FOA)1 in FY23-derived
strains containing pEAA69
((TG)16T-URA3, ARSH4, CEN6,
LEU2) (15, 27, 28).
1-I
mismatch substrate was formed by annealing oligonucleotides 4 and 6. Duplex DNA substrates were 32P-labeled by 5'-end labeling
oligonucleotides with [
-32P]ATP and T4 polynucleotide
kinase prior to the annealing reaction.
-mercaptoethanol, 1 mM
phenylmethylsulfonyl fluoride) instead of the 200 mM salt
conditions used to load and wash the MSH2-MSH6 complex. In cell
extracts obtained from overexpression strains, the levels of soluble
wild type and mutant MSH2-MSH6 proteins were indistinguishable and were
found to be equivalent to 3% of total protein. This was determined in
SDS-PAGE by directly comparing the intensity of MSH2 and MSH6 bands in
crude extracts to known amounts of purified complex. Through Western
blot analysis, we determined that the number of MSH2 monomers per wild
type mid-log cell was approximately 400; a 700-fold increase in this
level was observed in strains that overexpressed MSH2 using the
GAL10 promoter.2
The purity of protein preparations was monitored by SDS-PAGE (8% gels)
(33) (Fig. 3A). The approximate molecular weight of the
MSH2-MSH6 and MSH2-msh6-F337A was determined by measuring the time
required to elute polypeptides from a Superose 6HR gel filtration
column as described previously (13). This analysis was kindly performed
by William Enslow at the Cornell Biotechnology Analytical-Synthesis Facility.
-32P]ATP, 25 mM Tris, pH 7.5, 2.0 mM MgCl2, 0.1 mM dithiothreitol, 0.01 mM EDTA, and 40 µg/ml bovine serum albumin as
described previously (15, 37). +1 and homoduplex DNA substrate was
included in the reactions as indicated.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
1-I) were synthesized that contained single 5-IdUrd substitutions at the indicated thymine residues (see under "Experimental Procedures"). In the
1-I
substrate, the 5-IdUrd substitution constitutes an extrahelical loop of
one nucleotide (see under "Experimental Procedures"). In filter
binding competition assays (see below), MSH2-MSH6 displayed a 10-fold binding specificity for the
1-I substrate compared with the AT-I substrate: this specificity was similar to that observed with +1 and
homoduplex substrates tested previously (see Fig. 3A and data not shown).
1-I or AT-I substrate with MSH2-MSH6 under
standard binding conditions (Fig. 1; see
under "Experimental Procedures"). After incubation, samples were
irradiated with UV light for 0-60 min, resolved by SDS-PAGE, and then
exposed to film to detect radioactively labeled protein-DNA complexes.
As shown in Fig. 1A, in binding reactions involving either
the
1-I or the AT-I substrates, a major band migrating at ~180 kDa
(Band B) was observed that was dependent on MSH2-MSH6 and UV exposure
and migrated in SDS-PAGE slightly above the position of MSH6 (142 kDa).
The size of the labeled complex was consistent with a
1-I or AT-I
substrate (~30 kDa) cross-linked to MSH6 (142 kDa). At UV exposures
15 min and longer, a band migrating at greater than 205 kDa (Band A) was observed, as well as a faint band migrating at ~116 kDa (Band C).
At 30 min of UV exposure, a cross-linking efficiency of 17% (as
measured by densitometry) was observed in the
1-I reactions; this
cross-linking efficiency was 7-fold higher than that found in reactions
involving the AT-I substrate.
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Fig. 1.
MSH6 specifically cross-links to a
1-I mismatch substrate containing an
iodouracil residue at the site of the mismatch. A,
SDS-PAGE analysis of MSH2-MSH6 complexes cross-linked to mismatch
substrates. 20 nM 32P-labeled
1-I and AT-I
homoduplex substrates were incubated in the presence or absence of 100 nM MSH2-MSH6 complex for 15 min at 30 °C as described
under "Experimental Procedures." Samples were irradiated with 350 nM UV light as indicated, subjected to SDS-PAGE (8% gels),
and visualized using a phosphorimager. Lanes 1-5 display
the indicated control reactions. Lanes 6 and 7 indicate the position of Coomassie Blue-stained MSH2-MSH6 complex and
molecular weight standards (205, 116, 97, 66, 43 kD), respectively.
MSH2 and MSH6 are 109 and 142 kDa, respectively. Lanes 8-15
display a UV irradiation time course for MSH2-MSH6 binding to the
1-I (lanes 8-11) and AT-I (lanes 12-15)
substrates. B, immunoprecipitation analysis.
32P-Labeled
1-I heteroduplex was incubated with
MSH2-MSH6 as above and irradiated with 350 nM UV light for
15 min. UV-irradiated samples were denatured and immunoprecipitated
with MSH6-specific (lane 2) or MSH2-specific (lane
3) antibodies, and the immunoprecipitate was subjected to SDS-PAGE
(8% gels) (see under "Experimental Procedures"). Lane 1 displays the autoradiogram of a cross-linked sample that was directly
loaded onto SDS-PAGE gels, and lane 4 displays the results
of an immunoprecipitation reaction performed with cross-linked sample
in the absence of antibody. Bands A-C are described in the
text.
1-I complex. It is
important to note that in addition to Bands A-C, lower intensity bands
were observed in the cross-linking reactions shown in Fig.
1A; based on immunoprecipitation analysis and the mobility
of these bands relative to Bands A-C in SDS-PAGE, we hypothesize that
they represent complexes containing photocleavage products of the
5-IdUrd DNA substrates.
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Fig. 2.
Purification of MSH2-MSH6 and
MSH2-msh6-F337A. A, amino acid alignment of MutS
homologues in a region shown by UV cross-linking studies to be involved
in binding of T. aquaticus MutS to DNA mismatch substrates
(10). Highly conserved residues are shown with diamonds. The
msh2-Y42A and msh6-F337A, msh6-G335D,
and msh6-D343A substitutions described in this study are
indicated. B, purification of MSH2-MSH6, msh2-Y42A-MSH6, and
MSH2-msh6-F337A. Coomassie Blue staining of 0.6 µg of MSH2-MSH6
(lane 1), MSH2-msh6-F337A (lane 2), and
msh2-Y42A-MSH6 (lane 3) complexes subjected to SDS-PAGE (8%
gel). Molecular weights (in thousands) of standards are provided.
C, filter binding assay. 16.7 nM
32P-labeled +1 or homoduplex substrate were incubated with
increasing amounts of MSH2-MSH6 or MSH2-msh6-F337A in 60-µl reactions
(see under "Experimental Procedures"), and the amount of DNA bound
to protein was determined in filter binding assays (see under
"Experimental Procedures"). Open squares, MSH2-MSH6
binding to +1; filled squares, MSH2-MSH6 binding to
homoduplex; open circles, MSH2-msh6-F337A binding to +1;
filled circles, MSH2-msh6-F337A to homoduplex.
and msh6
strains,
respectively (14). In an assay that assesses mutation rate (principally
base-base and single nucleotide insertion/deletion mutations) through
the acquisition of resistance to the arginine analog canavanine,
msh2
strains exhibit a 13-20-fold higher mutation rate
than either wild type strains or msh2
strains that
overexpress MSH2 (14, 38, 39). The msh2-Y42A allele fully
complemented the msh2
mutator phenotype when its protein product was expressed in both GAL10 promoter inducing
(galactose) and repressing (glucose) conditions (Table
I) (10) (data not shown). These data
indicated that the msh2-Y42A mutation does not disrupt
mismatch repair. In the same assay, the msh6-F337A allele
failed to complement the modest mutator phenotype exhibited in
msh6
strains and instead conferred a mutation rate
equivalent to that observed in msh2
strains (a 20-fold
increase for a msh6
strain containing
pGAL10-msh6-F337A versus a 3.8-fold increase for
a msh6
strain containing a dummy plasmid).
Average rates of spontaneous mutations and DNA slippage events in
msh2, msh6
, and wild type strains bearing the msh6-Y42A,
msh6-F337A, and msh6-G987D alleles on GAL10, 2µ plasmids
(EAY 281), msh6
(EAY338), and wild
type (FY23) strains were transformed individually or in pairs with
pEAE51 (pGAL10-MSH6), pEAE20 or pEAE86
(pGAL10-MSH2), pEAE91 (pGAL10-msh2-Y42A), pEAE84
(pGAL10-msh6-G987D), and pEAE88
(pGAL10-msh6-F337A) and tested for forward mutations to
canr. These strains were also transformed with pEAA69
((TG)16T-URA3, ARS, CEN) and, where
indicated, tested in the (TG)16T tract alteration assay
described under "Experimental Procedures." All assays presented
were performed using media containing 2% galactose, 2% sucrose as a
carbon source. None refers to pRS control plasmids (26) that lacked
MSH2 or MSH6 sequences. The median frequency was
determined from 11 independent colonies for each experiment, and the
mutation rate per cell generation was determined as described by Lea
and Coulson (28). The average rate of two to six independent
experiments for each strain is presented. Note that overexpressed
MSH6 confers a weak dominant negative phenotype in the
canavanine resistance assay and only partially complements the
msh6
mutator phenotype (15). This partial complementation
and weak dominance phenotype was not observed in strains that expressed
the GAL10-MSH6 plasmid at lower levels (15).
Strains--
Recently, Studamire et al. (15) analyzed a
mutation in the MSH6 ATP binding domain (msh6-G987D) that
severely inhibited the ATPase activity of the MSH2-MSH6 complex. When
overexpressed in wild type strains, the msh6-G987D allele
conferred a weak dominant negative mutator phenotype that was similar
to the phenotype found in msh6
strains, as it was limited
to defects in the MSH2-MSH6 repair pathway (Table I) (15).
In contrast, when the msh6-F337A allele was similarly
overexpressed in wild type strains, a mutator phenotype was observed
that was similar to that observed in msh2
strains. In the
canavanine assay, wild type strains overexpressing msh6-F337A displayed
a 21-fold higher mutation rate than strains lacking this plasmid and a
5-fold higher rate than msh6
strains or wild type strains
that overexpressed msh6-G987D (Table I). In DNA slippage assays (26)
that detect 2-4 insertion/deletion mismatches formed in an in-frame
(TG)16T cassette placed within the URA3 gene,
msh2
strains exhibited an approximately 17-fold higher
rate of slippage events, as measured by resistance to 5-fluoroorotic acid, than wild type. msh6
strains or a wild type strain
overexpressing MSH6 displayed DNA slippage frequencies that were
indistinguishable from wild type. However, strains overexpressing
msh6-F337A displayed a DNA slippage rate that was similar to that
observed in msh2
strains (Table I). It is important to
note that the overexpression of all msh6 single and double mutant
protein combinations in this study was similar to that observed for
overexpression of wild type MSH6 (data not shown).
strains. We hypothesized that this resulted from
msh6-F337A sequestering MSH2 in an inactive complex that was prevented
from interacting with either MSH6 or MSH3 subunits. We tested this idea
by performing canavanine mutator assays on wild type strains
overexpressing the msh6-F337A, MSH2, and MSH6 subunits individually or
in combination. As shown in Table I, co-overexpression of MSH2 or MSH6
with msh6-F337A fully suppressed the dominant negative phenotype
observed in strains that only overexpressed msh6-F337A, consistent with
the idea that MSH2 was sequestered from interacting with MSH3 or MSH6.
This experiment, however, did not rule out the possibility that excess
MSH2 suppressed the dominant negative phenotype conferred by the
msh6-F337A allele by suppressing its mismatch repair defect.
To test this, mutator assays were performed in a msh6
strain that overexpressed both MSH2 and msh6-F337A. As shown in Table
I, this strain displayed a mutation rate that was similar to that
observed in msh6
strains, indicating that overexpression
of MSH2 cannot suppress the msh6-F337A mismatch repair
defect to achieve wild type mismatch repair function.
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Fig. 3.
Filter binding and gel retardation
competition assays performed with MSH2-MSH6 and MSH2-msh6-F337A
complexes. Filter binding reactions were performed with 20 nM (0.3 µg) MSH2-MSH6 (A), MSH2-msh6-F337A
(B), or MSH2-msh6-F337A/G987D complex (C) as
described under "Experimental Procedures." 32P-Labeled
+1 substrate was present at 16.7 nM in the absence or
presence of the indicated unlabeled homoduplex or +1 competitor. The
percentage of input 32P-labeled +1 bound for each of the
complexes in the absence of competitor was 21.2% for MSH2-MSH6, 8.2%
for MSH2-msh6-F337A, and 9.3% for MSH2-msh6-F337A/G987D. D,
gel retardation assays were performed with 10 nM
32P-labeled +1 substrate and 360 nM (0.9 µg)
MSH2-msh6-F337A (lane 2) or MSH2-MSH6 (lanes
3-16) as described under "Experimental Procedures."
Lane 1, no protein control. Unlabeled homoduplex was present
at 0, 250, 500, 1000, 2000, and 4000 nM in lanes
3-8, respectively, and unlabeled +1 substrate was present at 0, 16, 32, 64, 125, 250, 500, and 1000 nM in lanes
9-16, respectively. E, the gel shift assays in
D are presented graphically. In A, B, C, and
E, filled squares represent homoduplex
competitor, and open squares represent +1 competitor.
1-I DNA at 30 °C for 15 min,
UV-irradiated for 15 min, electrophoresed on 8% SDS-PAGE gels, and
then quantified by using a phosphorimager (see under "Experimental
Procedures"). As shown in Fig.
4A, the MSH2-msh6-F337A
complex displayed a 6-fold lower level of cross-linking to the
1-I
substrate compared with MSH2-MSH6 (Fig. 4A, lanes 1 and
2). When 1.5 mM ATP was included in the binding
reaction prior to UV cross-linking, a 2.7-fold reduction in the level
of MSH2-MSH6 cross-linking to the
1-I substrate was observed
(lane 4). However, in the MSH2-msh6-F337A reactions, ATP
enhanced cross-linking of the complex to the
1-I substrate by
2.5-fold (Fig. 4A, lanes 1 and 3).
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Fig. 4.
UV cross-linking and filter binding assays of
MSH2-MSH6 and MSH2-msh6-F337A performed in the presence and absence of
ATP. A, cross-linking reactions were performed with 24 nM (0.3 µg) MSH2-MSH6 or MSH2-msh6-F337A complex and 20 nM 32P-labeled 1-I. The reactions were
incubated at 30 °C for 15 min, UV-irradiated for 15 min, submitted
to SDS-PAGE (8% gels), and quantified using a phosphorimager.
Lanes 1 and 3, MSH2-msh6-F337A cross-linking
reactions performed in the absence (lane 1) and presence
(lane 3) of 1.5 mM ATP. Lanes 2, 4, and 5, MSH2-MSH6 cross-linking reactions performed in the
absence (lane 2) and presence (lane 4) of 1.5 mM ATP and in the absence of UV irradiation (lane
5). Bands A-C are described in the text. B,
filter binding analysis of MSH2-MSH6 (2-6) and
MSH2-msh6-F337A (2-6FA) complexes incubated in the presence
(hatched columns) or absence (filled columns) of
1.5 mM ATP. DNA binding assays were performed by filter
binding as described under "Experimental Procedures" with 16.7 nM 32P-labeled +1 substrate and 20 nM MSH2-MSH6 or MSH2-msh6-F337A. Duplicate experiments were
performed, and the difference between the two values is shown with
error bars.
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Fig. 5.
The effect of +1 and homoduplex substrate on
the ATPase activity of MSH2-MSH6 and MSH2-msh6-F337A. In
A and B, 20 nM (0.3 µg) MSH2-MSH6
(A) or MSH2-msh6-F337A (B) was assayed for ATPase
activity as described under "Experimental Procedures" in the
presence of 167 nM +1 or homoduplex substrate. The amount
of ATP hydrolyzed was determined after 15 min at 30 °C for duplicate
reactions, and the differences between these two values are shown with
error bars. In C and D, 20 nM (0.3 µg) MSH2-MSH6 (C), or MSH2-msh6-F337A
(D) was incubated in the presence of the 33.3 µM [ -32P]ATP, and the indicated
concentrations of +1 or homoduplex substrate and the amount of ATP
hydrolyzed were determined after a 15-min incubation. A-D:
open triangles, no DNA substrate; filled squares,
homoduplex substrate; open squares, +1 substrate.
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Fig. 6.
The effect of +1 and homoduplex substrate on
the ATPase activity of MSH2-msh6-F337A/G987D and MSH2-msh6-F337A.
20 nM (0.3 µg) MSH2-msh6-F337A/G987D or MSH2-msh6-F337A
was assayed for ATPase activity as described under "Experimental
Procedures" in the presence of 100 µM
[ -32P]ATP and 167 nM +1 or homoduplex
substrate (homo). The amount of ATP hydrolyzed was
determined after 15 min at 30 °C in duplicate reactions, and the
differences between the duplicates are shown with error
bars.
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Fig. 7.
Dissociation studies revealed slow and fast
off rates for MSH2-MSH6 binding to +1 substrate. 5.7 nM (0.3 µg) MSH2-MSH6 (A) or MSH2-msh6-F337A
(B) was preincubated for 15 min with either 1.4 nM 32P-labeled homoduplex (filled
squares) or +1 substrate (open squares) in a 210-µl
standard DNA binding reaction (see under "Experimental
Procedures"). In C, MSH2-MSH6 was preincubated with DNA
substrate in the presence of 2.0 mM ATP. After
preincubation, a 300-fold excess of the respective unlabeled substrate
was added, and at various time points a 30-µl aliquot was removed and
the amount of 32P-labeled substrate that remained bound was
measured by filter binding. The percentage of binding is normalized to
a control reaction in which unlabeled competitor was not added. The
percentages of binding in the absence of competitor were as follows:
MSH2-MSH6, 43% to +1 and 26% to homoduplex; MSH2-msh6-F337A, 8.9% to
+1 and 9.3% to homoduplex; and MSH2-MSH6 in the presence of 2.0 mM ATP, 27% to +1 and 30% to homoduplex. Background
levels of binding were observed for both +1 and homoduplex reactions
when unlabeled competitor was included in the preincubation
reaction.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
1-I) and homoduplex substrates was measured in UV cross-linking and in filter binding assays performed in the presence and absence of ATP (Fig. 4). Previous
studies indicated that the specific binding of MutS homolog complexes
to mismatch substrates in vitro was abolished by ATP (8, 13,
19, 41, 42). This loss of mismatch binding specificity in the presence
of ATP most likely reflects a switch from a recognition mode to a
translocation mode (7, 9). In contrast, the binding of MSH2-msh6-F337A
complex to both homoduplex and single nucleotide insertion/deletion
substrates was improved in the presence of ATP, and approached one-half
of the level observed for wild type MSH2-MSH6 (Fig. 4). Based on the
two binding mode model for MutS described in the Introduction, we
propose that the mutant complex is defective in mismatch recognition
but can still function in a second mode that allows it to bind and/or translocate along DNA in an ATP-dependent fashion. It is
important to note that in the presence of ATP, the MSH2-msh6-F337A
complex did not reach the level of binding to homoduplex DNA that was observed for MSH2-MSH6. This finding can be reconciled in the context
of a two binding mode model if the initial loading of the MSH2-MSH6
complex onto DNA is accomplished primarily through the first binding
mode, the disruption of which would be expected to reduce both general
DNA binding and mismatch recognition. At present it is unclear whether
the ATP-dependent binding of the MSH2-msh6-F337A complex is
mediated by residues in the conserved domain presented in Fig.
2A or is located elsewhere.
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ACKNOWLEDGEMENTS |
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We thank Elizabeth Evans, Richard Kolodner, Chris Roberts, Jeff Roberts, Barbara Studamire, and Yali Xie for technical advice and helpful discussions and Elizabeth Evans, Jeff Roberts, and members of the Alani laboratory for their insightful comments on the manuscript.
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FOOTNOTES |
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* 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.
These authors contributed equally to this work. Supported by
National Institutes of Health predoctoral training grants.
§ Supported by an undergraduate summer research fellowship from the Howard Hughes Medical Institute awarded to Cornell University.
¶ Supported by National Institutes of Health Grant GM53085 and United States Department of Agriculture Hatch Grant NYC-186424. To whom correspondence should be addressed: Section of Genetics and Development, Cornell University, 459 Biotechnology Bldg., Ithaca, NY 14853-2703. Tel.: 607-254-4811; Fax: 607-255-6249; E-mail: eea3{at}cornell.edu.
2 T. Sokolsky and E. Alani, unpublished data.
3 T. Quach, J. Bowers, and E. Alani, unpublished data.
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ABBREVIATIONS |
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The abbreviations used are: 5-FOA, 5-fluoroorotic acid; hMSH, human MSH; PAGE, polyacrylamide gel electrophoresis; 5-IdUrd, iododeoxyuridine; ssDNA, single-stranded DNA.
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