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J. Biol. Chem., Vol. 276, Issue 35, 33233-33240, August 31, 2001

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DNA Chain Length Dependence of Formation and Dynamics of hMutS·hMutL·Heteroduplex Complexes^*

Leonard J. Blackwell, Shuntai Wang^§, and Paul Modrich^§¶

From the  Department of Biochemistry and ^§ Howard Hughes Medical Institute, Duke University Medical Center, Durham, North Carolina 27710

Received for publication, June 3, 2001, and in revised form, July 3, 2001

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Formation of a ternary complex between human MutS, MutL, and heteroduplex DNA has been demonstrated by surface plasmon resonance spectroscopy and electrophoretic gel shift methods. Formation of the hMutL·hMutS·heteroduplex complex requires a mismatch and ATP hydrolysis, and depends on DNA chain length. Ternary complex formation was supported by a 200-base pair G-T heteroduplex, a 100-base pair substrate was somewhat less effective, and a 41-base pair heteroduplex was inactive. As judged by surface plasmon resonance spectroscopy, ternary complexes produced with the 200-base pair G-T DNA contained ~0.8 mol of hMutL/mol of heteroduplex-bound hMutS. Although the steady-state levels of the hMutL·hMutS· heteroduplex were substantial, this complex was found to turn over, as judged by surface plasmon resonance spectroscopy and electrophoretic gel shift analysis. With the former method, the majority of the complexes dissociated rapidly upon termination of protein flow, and dissociation occurred in the latter case upon challenge with competitor DNA. However, ternary complex dissociation as monitored by gel shift assay was prevented if both ends of the heteroduplex were physically blocked with streptavidin·biotin complexes. This observation suggests that, like hMutS, the hMutL·hMutS complex can migrate along the helix contour to dissociate at DNA ends.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

The MutS and MutL homologs, which are required for the initiation of mismatch repair, have been implicated in the correction of DNA biosynthetic errors, the transcription-coupled repair of DNA damage, and the fidelity of genetic recombination (1-6). In mammalian cells, MutS (MSH2·MSH6 heterodimer), MutS (MSH2·MSH3 heterodimer), and MutL (MLH1·PMS2 heterodimer) are also thought to function as lesion sensors for certain types of DNA damage that kill by activating apoptosis (2, 6, 7).

Repair in the Escherichia coli system is initiated by the binding of MutS to a mismatch (8-10). Formation of a MutL·MutS·heteroduplex complex has been demonstrated by DNase I footprint analysis (11), electron microscopy (12), and surface plasmon resonance spectroscopy (SPRS)¹ (13), with assembly of this ternary complex being ATP-dependent. Several lines of evidence indicate that assembly of the ternary complex is required for subsequent steps in mismatch repair. Both MutS and MutL are required for the mismatch-dependent activation of the d(GATC) endonuclease activity of MutH, which cleaves the unmethylated strand of a hemimethylated d(GATC) site, with the ensuing strand break serving to direct repair to the unmethylated DNA strand (14). MutS and MutL are also required for the mismatch-dependent activation of DNA helicase II, which enters the helix at the strand break and initiates the excision step of repair (15).

Formation of a MutL·MutS·heteroduplex complex has been demonstrated by electrophoretic gel shift analysis with yeast mismatch repair proteins using synthetic heteroduplexes of ~50 base pairs (bp) in size (16, 17). However, using gel shift methods and surface plasmon resonance spectroscopy, we have consistently been unable to demonstrate the corresponding ternary complex between E. coli MutS and MutL or human MutS and MutL and synthetic heteroduplexes of similar size.² This is despite the fact, as noted above, that footprint analysis, electron microscopy, and SPRS has indicated formation of a MutL·MutS·heteroduplex complex with the bacterial proteins (11-13). Since the latter experiments utilized heteroduplexes 140 bp or longer, we have examined the effect of DNA chain length on ternary complex formation using surface plasmon resonance spectroscopy and electrophoretic gel shift. We show here that efficient formation of the MutL·MutS·heteroduplex ternary complex is dependent on DNA chain length. We also show that ternary complex formation with the human proteins requires ATP, and probably its hydrolysis, and that these complexes turn over rapidly with respect to binding and release from the DNA. However, dissociation can be prevented and ternary complexes kinetically stabilized by placement of streptavidin blocks at both ends of a linear heteroduplex.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

hMutS and hMutL Preparations-- hMutS and hMutL were prepared from baculovirus constructs expressing the appropriate human cDNAs in SF9 cells. The two subunits of hMutS were expressed from a single virus constructed using the Dual Bac system (Life Technologies, Inc.). Briefly, hMSH2 cDNA (provided by Bert Vogelstein, Johns Hopkins Oncology Center, Baltimore, MD) was inserted into the NcoI site just downstream of the p10 promoter, while hMSH6 cDNA (a gift from Rick Fishel, Thomas Jefferson University, Philadelphia, PA) was expressed from the polyhedrin promoter by insertion between the BamHI and SalI sites. cDNAs for human MLH1 and PMS2 (generously provided by Mike Liskay, Oregon Health Sciences University, Portland, OR) were expressed from individual viral constructs prepared using the pFastBac I system (Life Technologies, Inc.). hMLH1was expressed from the polyhedrin promoter by insertion between the BamHI and NotI sites. hPMS2 was also expressed from the polyhedrin promoter after insertion between BamHI and XbaI sites.

Infected SF9 cells for hMutS isolation were grown by Kemp Biotechnologies, Inc. (Frederick, MD). Frozen cell pellets were thawed and suspended (10 ml/g of cells) in 25 mM HEPES-KOH, pH 7.5, 0.1 mM EDTA, 1 mM dithiothreitol containing 1 µg/ml each of aprotinin, leupeptin, Pefabloc (Roche Molecular Biochemicals), and E64 (Peptides International). Cells were lysed with five strokes of a Dounce B pestle, and the extract supplemented with KCl to 200 mM. After centrifugation (30,000 × g, 15 min), hMutS was isolated from the supernatant by minor modifications of the method used previously for isolation of the HeLa cell activity (18). Recombinant hMutS preparations obtained in this manner had a purity of 98% or better, were characterized by a 1:1 subunit stoichiometry, and were fully active in mismatch repair as judged by in vitro complementation of nuclear extracts derived from MSH2-deficient human cells.

For hMutL purification, SF9 cells were continuously cultured in 800 ml of serum-free HyQ-SFX medium (HyClone, Inc.) at 27 °C in 2.8-liter Fernbach flasks. When culture density reached 1 × 10⁶/ml, cells were co-infected with hMLH1 and hPMS2 baculovirus constructs at a multiplicity of infection of 5. Infected cells were harvested 60 h later by centrifugation at 4,000 rpm for 10 min in a Sorvall RC-3B centrifuge. Cell pellets were suspended in 120 ml of 20 mM KPO₄, pH 7.6, 5 mM KCl, and 1 mM MgCl₂ containing 0.1% phenylmethylsulfonyl fluoride (relative to a saturated solution in isopropanol) and 1 µg/ml each of aprotinin, leupeptin, and E64. After incubation on ice for 10 min, cells were lysed with 20 strokes using a Dounce B pestle, the suspension adjusted to 100 mM KCl, and then clarified by centrifugation at 20,000 × g for 10 min. The supernatant was frozen in liquid N₂ and stored at 80 °C (fraction I). Forty ml of fraction I was thawed and loaded at 4 °C onto a 5-ml heparin HiTrap column (Amersham Pharmacia Biotech) equilibrated with 25 mM HEPES-KOH, pH 7.5, 100 mM KCl, 1 mM EDTA, and 10% (v/v) glycerol at flow rate of 1.5 ml/min. After wash with starting buffer, the column was eluted with a 60-ml gradient of KCl (100-450 mM) in 25 mM HEPES-KOH, pH 7.5, 1 mM EDTA, and 10% (v/v) glycerol. hMutL fractions, which eluted at ~230 mM KCl, were diluted to 80 mM KCl with 20 mM KPO₄, pH 7.4, 0.1 mM EDTA, 10% (v/v) glycerol and loaded onto a 1-ml Mono Q column (Amersham Pharmacia Biotech) equilibrated with 20 mM KPO₄, pH 7.4, 0.1 mM EDTA, 10% (v/v) glycerol (buffer A) containing 80 mM KCl at a flow rate of 0.5 ml/min. After wash with starting buffer, the column was eluted with a 20 ml gradient of KCl (80-380 mM) in buffer A. hMutL fractions, which eluted at ~220 mM KCl, were diluted to 80 mM KCl with buffer A and loaded onto a Mono S column (Amersham Pharmacia Biotech) equilibrated with buffer A containing 80 mM KCl at a flow rate of 0.5 ml/min. After wash with starting buffer, the column was eluted with a 20-ml KCl gradient (80-380 mM) in buffer A. hMutL fractions, which eluted at ~150 mM KCl, were pooled, and aliquots frozen in liquid N₂ and stored at 80 °C. Purification from 40 ml of extract yielded 1.6 mg of hMutL with an MLH1:PMS2 subunit ratio of 1:1 and an estimated purity of 98%. Activity of such preparations are comparable to that of hMutL isolated from HeLa cells (19), as judged by in vitro complementation of nuclear extracts derived from cells deficient in hMLH1.

DNAs-- Oligodeoxyribonucleotides were purchased from Oligos Etc. (Wilsonville, OR), and when indicated were radiolabeled at the 5'-terminus with T4 polynucleotide kinase and [-³²P]ATP (3000 Ci/mmol, PerkinElmer Life Sciences) to a specific activity of 1 × 10⁶ cpm/pmol. Poly d(I)·d(C) was purchased from Amersham Pharmacia Biotech.

A 41-bp G-T heteroduplex used for gel shift analysis was prepared by mixing, in a 100-µl volume, 80 nM top strand 5-³²P-d(AGCCGAATTTTTAGACTCGATAGCTTGCTAGCAATTCGGCG) with 120 nM unlabeled bottom strand 5'-biotin-d(CGCCGAATTGCTAGCAAGCTGTCGAGTCTAAAAATTCGGCT). Strands were annealed in 10 mM Tris-HCl, pH 8.0, 1 mM EDTA, and 150 mM NaCl by heating at 99 °C in a PerkinElmer Life Sciences Gene Amp 9600 thermocycler for 2 min and cooling to 25 °C over a period of 90 min. The corresponding A·T homoduplex was prepared in a similar manner by annealing 5'-biotin-d(CGCCGAATTGCTAGCAAGCTATCGAGTCTAAAAATTCGGCT) with the top strand above. Identical 41-bp substrates were prepared for SPRS by annealing 1 µM each of top and bottom strands described above in a 100-µl volume. These 41-bp DNAs correspond to coordinates 5612-5652 of the f1MR phage DNAs used for preparation of in vitro mismatch repair substrates (9).

PCR-derived 200-bp G-T heteroduplex and homoduplex DNAs were prepared after strand separation by denaturing HPLC as described previously (20). Briefly, a biotinylated viral strand sequence obtained by strand separation of a 200-bp PCR product derived from phage f1MR1 (0.3 µM) was annealed in 100 µl with 0.2 µM 200-nucleotide, ³²P-labeled complementary strand sequence derived by PCR from f1MR1 or f1MR3 (9). Duplexes were annealed as described above. Non-radioactive 200-bp homoduplex and heteroduplex DNAs for SPRS were prepared in a similar manner by annealing 0.5 µM 5'-biotinylated f1MR1 viral strand sequence with 0.5 µM complementary strand sequence derived from f1MR1 or f1MR3.

100-bp homoduplex and G-T heteroduplexes used for gel shift analysis were prepared by denaturing HPLC strand separation of a 150-bp PCR product derived from coordinates 5582-5732 of bacteriophages f1MR1 and f1MR3 (9). Forward and reverse primers for PCR were d(CGCTTTCTTCCCTTCCTTTCTCG) and d(AAGTTTTTTGGGGTCGAGGT). The ³²P-labeled, 150-residue viral strand sequence from f1MR1 was combined with the complementary sequence (0.45 µM each in 100 µl) prepared from f1MR1 or f1MR3 to yield homoduplex or G-T heteroduplex, respectively. DNAs were annealed as described above in 20 mM Tris acetate, pH 7.9, 10 mM magnesium acetate, 50 mM potassium acetate, and 1 mM dithiothreitol. Resulting duplexes were cleaved with 20 units of BanII for 1 h at 37 °C. After dilution to 400 µl with buffer B (10 mM Tris-HCl, pH 8.0, 1 mM EDTA) containing 300 mM NaCl, the DNA was loaded onto a Gen-Pak Fax column (4.6 × 100 mm) equilibrated with this buffer at a flow rate of 0.55 ml/min. After washing 5 min with buffer B containing 0.3 M NaCl, the DNA was eluted with a gradient of NaCl (0.3-1 M) in buffer B over a 35-min period. 5'-Biotinylated 110-bp DNAs for surface plasmon resonance spectroscopy were prepared in a similar manner from PCR products derived from region 5570-5732 of f1MR1 and f1MR3 using the same reverse primer described above and 5'-biotin-d(GCCCGCTCCTTTCGCTTTCT) as forward primer. The biotinylated viral strand sequence from f1MR1 was annealed with the complementary strand sequence (1 µM each in 100 µl) prepared from f1MR1 or f1MR3, subjected to BanII cleavage, and the 110-bp heteroduplex and homoduplex purified as described above.

3'-³²P-Labeled 200 bp homoduplex and heteroduplex DNAs that were tagged with 5'-biotin at both ends were prepared as described previously (20).

Gel Shift Analysis-- DNA binding reactions (20 µl) contained 20 mM Tris-HCl, pH 7.6, 1 mM dithiothreitol, 50 µg/ml bovine serum albumin, 5 mM MgCl₂, 100 mM KCl, 1 nM [³²P]homoduplex or [³²P]heteroduplex DNA (as indicated), 25 µg/ml BstEII digest of bacteriophage  DNA (New England Biolabs), and ATP as indicated. Solutions were prewarmed to 25 °C for 10 min and reactions initiated by addition of hMutS and hMutL as indicated. After incubation for 10 min at 25 °C, reactions were stopped by addition of 2 µl of 50% (v/v) glycerol, 0.05% xylene cyanol, 0.05% bromphenol blue, and 20 mM EDTA, placed on ice, and then loaded onto a 4% native polyacrylamide gel (acrylamide-bisacrylamide, 37.5:1) in 6.7 mM Tris acetate, pH 7.5, and 1 mM EDTA. Gels were electrophoresed at room temperature at 11.4 V/cm in this buffer. ³²P-Labeled complexes were visualized by autoradiography after drying.

Western and Southern Blotting of Gel-shifted Complexes-- A nitrocellulose membrane was placed on polyacrylamide gels and a NA45 DEAE-cellulose membrane (Schleicher & Schuell) placed on top of the nitrocellulose. Protein and DNA were then electrophoretically transferred in 50 mM Tris, 376 mM glycine, and 20% methanol for 1 h at 100V. Under these conditions proteins are retained by the nitrocellulose, but DNA passes through to be retained on the DEAE membrane. Radiolabeled DNA was visualized by autoradiography of the DEAE membrane. Mismatch repair polypeptides were identified by Western blot using monoclonal anti-hMLH1 (PharMingen), monoclonal anti-hPMS2 (PharMingen), monoclonal anti-hMSH2 (Calbiochem), or goat anti-MSH6 (N-20, Santa Cruz Biotechnology Inc.). Nitrocellulose membranes were incubated in blocking buffer (10 mM Tris-HCl, pH 8.0, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, 5% (w/v) nonfat dried milk) for 30 min, followed by a 1-h incubation with the appropriate antibody diluted 1/500 into blocking buffer. After three 10-min washes with blocking buffer, membranes were incubated for 30 min with a 1/500 dilution of anti-mouse Ig horseradish peroxidase conjugate (Amersham Pharmacia Biotech) or anti-goat IgG peroxidase conjugate (Sigma) as appropriate. Immune complexes were detected by ECL. When indicated nitrocellulose membranes were stripped by incubating a 65 °C for 4 h in 62.5 mM Tris-HCl, pH 6.8, 100 mM 2-mercaptoethanol, 2% (w/v) sodium dodecyl sulfate. After washing three times for 10 min with blocking buffer, the membrane was probed with a second antibody.

Surface Plasmon Resonance Spectroscopy-- Surface plasmon resonance measurements were performed on a BIAcore 2000. Streptavidin sensor chips were derivatized with the 41-, 110-, and 200-bp homoduplex or G-T heteroduplex DNAs described above, in which one strand was tagged with a 5'-terminal biotin. Solutions contained 20 mM Tris-HCl, pH 7.6, 1 mM dithiothreitol, 0.005% surfactant P20, 5 mM MgCl₂, 100 mM KCl and ATP, and hMutS and hMutL as indicated. Measurements were performed at 25 °C at a flow rate of 20 µl/min, and samples were maintained at 4 °C prior to injection. Dissociation kinetics were monitored by coinjecting 120 µl of ATP supplemented reaction buffer immediately following protein association. SA chips were regenerated by a 20-µl injection of 0.5% sodium dodecyl sulfate.

DNA-bound protein monitored by SPRS is expressed as equivalents of MutS (M_r = 258,000; Refs. 18 and 21-24) bound/mol of chip-bound DNA. This value was calculated from the mass ratio, which is given by (RU_exp)/(0.79 × RU_DNA), where RU_exp is the output signal due to protein binding and RU_DNA corresponds to the amount of DNA bound to the chip in resonance units. The factor 0.79 corrects for the fact that the refractive index increment for a typical protein is 79% of that obtained with an equivalent mass of DNA (25). Mass_ratio values were converted to molar ratios using the molecular weights of the DNA and protein in question. Since this method is based on several assumptions (for example, that the relative refractive index increment cited above is generally valid and that all chip-bound DNA is equally accessible to protein flow), it can only be regarded as approximate. However, we have also used this method to estimate relative binding stoichiometries of hMutS and hMutL in ternary complexes with DNA, and these values should be quite accurate.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Mismatch-, ATP-, and Chain Length-dependent Formation of a hMutL·hMutS·Heteroduplex Complex-- Ternary complexes of yeast MutS and MutL with synthetic heteroduplexes of ~50 bp in size have been demonstrated by electrophoretic gel shift (16, 17). However, we have been unable to reproducibly detect ternary complexes involving bacterial MutS and MutL, or human MutS and MutL, utilizing synthetic heteroduplexes 41 bp in length. Since there is abundant evidence for ternary complex formation between the bacterial mismatch repair proteins and heteroduplexes of 143 bp or longer (11-13), we reasoned that this problem might be due to the small size of synthetic heteroduplexes.

Using surface plasmon resonance spectroscopy, specific binding of hMutS to a 200-bp G-T heteroduplex (see "Experimental Procedures") was evident in the presence of ATP·Mg²⁺, with the heteroduplex signal approximately 3 times that observed with an otherwise identical A·T homoduplex (Fig. 1). Although hMutL did not bind detectably to the 200-bp heteroduplex under these conditions, passage of a mixture of hMutS and hMutL over the chip resulted in a substantial enhancement of the mass of heteroduplex-bound protein, as compared with that observed with hMutS alone. This mass enhancement is mismatch-dependent since it was not observed with the A·T homoduplex control, and experiments described below demonstrate that it is not a simple consquence of increased hMutS binding due to presence of hMutL. This effect requires the simultaneous presence of hMutS and hMutL since no enhancement of heteroduplex-bound protein was observed in experiments in which the two proteins were passed over the chip in a sequential manner (data not shown).

View larger version (11K): [in this window] [in a new window]   Fig. 1.   Ternary complex formation requires a mismatch and hMutS. SPRS analysis was performed as described under "Experimental Procedures" using a SA sensor chip derivatized with 143 RU of a 200-bp G-T heteroduplex (blue lines) and 146 RU of an otherwise identical 200-bp A·T homoduplex (red lines). Protein solutions contained 20 mM Tris, pH 7.6, 100 mM KCl, 1 mM dithiothreitol, 5 mM MgCl2, 0.005% surfactant P20, 100 µM ATP, 200 nM hMutS alone (red and blue solid lines), 200 nM hMutL alone (hyphenated blue line), or 200 nM of hMutS and 200 nM hMutL (dashed red and blue lines). Chip-bound protein is expressed as mass equivalents of hMutS (see "Experimental Procedures"), where one mass equivalent corresponds to 258 kDa.

The hMutL-dependent mass enhancement was only observed in the presence of ATP, where it displayed a strong dependence on DNA chain length (Fig. 2). In the absence of ATP (lower panel), ~1 hMutS heterodimer was bound/41-bp G-T heteroduplex, and this value increased to approximately two equivalents with the 200-bp heteroduplex. Under these conditions in the absence of ATP, the protein mass bound by 41-, 100-, and 200-bp heteroduplexes was unaffected by inclusion of hMutL along with hMutS, as compared with that observed with hMutS alone.

View larger version (16K): [in this window] [in a new window]   Fig. 2.   Ternary complex formation between hMutS, hMutL, and a heteroduplex requires ATP and is dependent on DNA chain length. Ternary complex formation between hMutS and hMutL and 41-, 110-, and 200-bp G-T heteroduplex DNA substrates (183, 165, and 166 RU, respectively) was monitored by SPRS as described under "Experimental Procedures" and in the legend to Fig. 1. hMutS (200 nM), hMutL (200 nM), and ATP (1 mM) were present as indicated. Progress curves are shown for 41-bp (blue), 110-bp (green), and 200-bp (red) G-T heteroduplexes. Solutions contained hMutS only (solid lines) or both hMutS and hMutL (dashed lines). The solid and dashed lines are essentially coincident with the 41-bp heteroduplex (blue) in the presence or absence of ATP.

As observed previously (18, 20, 26), the presence of ATP resulted in a dramatic reduction in hMutS binding to the 41-bp heteroduplex, and the presence of hMutL was without effect (Fig. 2, compare upper and lower panels). However, the extent of hMutS binding to 100- and 200-bp heteroduplexes in the presence of ATP was similar to that observed in the absence of nucleotide, and the presence of hMutL resulted in a substantial enhancement of protein mass bound by these two heteroduplex DNAs. Based on experiments described below, and previous observations with bacterial and yeast mismatch repair proteins (11-13, 16, 17), we have concluded that this enhancement of heteroduplex-bound protein mass reflects mismatch-, ATP-, and hMutS-dependent presence of hMutL in a nucleoprotein complex with heteroduplex DNA. The stoichiometries of formation of this hMutL·hMutS·heteroduplex ternary complex will be considered below.

Ternary complex formation was demonstrable in the presence of ATP·Mg²⁺, but we have been unable to detect a hMutL-dependent increase in the mass of heteroduplex-bound protein in the presence of AMPPNP·Mg²⁺ (data not shown). This finding, which is similar to previous observations with bacterial MutS and MutL (13), strongly suggests that ternary complex formation is dependent upon ATP hydrolysis by one or both proteins.

Apparent hMutS and hMutL Affinities and Stoichiometry of Ternary Complex Formation-- Ternary complex formation requires ATP·Mg²⁺, conditions that reduce the affinity of hMutS for a mispair (18, 20, 26). The apparent specific affinity of hMutS for the 200-bp G-T heteroduplex and A·T homoduplex in the presence of ATP was estimated by SPRS (Fig. 3, upper panel). Under these conditions, hMutS·heteroduplex and hMutS·homoduplex formation was hyperbolic, with apparent K_d values of 205 and 420 nM, respectively. These data were also evaluated after subtraction of the hMutS·homoduplex values from those observed with the heteroduplex in order to correct interactions with heteroduplex for mismatch-independent binding. Correction in this manner also yielded an excellent hyperbolic fit with an apparent K_d of 140 nM and an asymptotic value of 3 equivalents of the MSH2·MSH6 heterodimer bound per DNA at saturation (Fig. 3, upper panel). Although the stoichiometry of hMutS·heteroduplex formation calculated from SPRS can be regarded as only approximate (see "Experimental Procedures"), the finding that the limiting stoichiometry of heteroduplex binding exceeds unity, even after correction for homoduplex effects, may seem surprising given that the DNA contains a single mismatch. There are several possible explanations for this effect. For example, multiple copies of hMutS may oligomerize on a heteroduplex in a mismatch-dependent reaction. A second possibility is based on the observation that hMutS can leave a mismatch in the presence of ATP by movement along the helix contour to dissociate at DNA termini (20, 27, 28). Such a mechanism would account for the loading of multiple copies of hMutS onto a heteroduplex, provided that the migrating species fail to reach a DNA terminus before another hMutS binds to the mispair (27).

View larger version (17K): [in this window] [in a new window]   Fig. 3.   Apparent Kd values for assembly of hMutS·hetero- duplex and hMutL·hMutS·heteroduplex complexes. Upper panel, binding of hMutS to a 200-bp G-T heteroduplex (175 RU) and an A·T homoduplex (174 RU) was determined in the presence of 1 mM ATP. Plateau values of SPRS sensorgrams obtained as a function of hMutS concentration are shown as a function of the concentration of the protein. Data were fit to a square hyperbola using a nonlinear least squares routine. Apparent Kd values for the G-T heteroduplex () and the A·T homoduplex () are 205 and 420 nM, respectively. The data are also plotted after subtraction of homoduplex values from those obtained with heteroduplex in order to correct heteroduplex binding for nonspecific effects (). Hyperbolic fit of this corrected data yielded an apparent Kd of 140 nM and an asymptotic value of 3 equivalents of hMutS/200-bp heteroduplex. Lower panel, the SA chip was derivatized with 143 RU of the 200-bp G-T heteroduplex. The hMutS concentration was 200 nM, ATP was 100 µM, and hMutL concentration varied as shown. The mass change, beyond that produced by hMutS alone, was fit to a square hyperbola, yielding an apparent Kd of 70 nM. The binding maxima from the fit corresponded to a relative, hMutL-dependent mass increase of 201 kDa, or 1.1 equivalents of the MLH1·PMS2 heterodimer (180 kDa; Refs. 19 and 35-37).

Like hMutS binding to the 200 bp G-T heteroduplex in the presence of ATP, ternary complex formation was a hyperbolic function of hMutL concentration (Fig. 3, lower panel), characterized by an apparent K_d of 70 nM in the presence of a hMutS concentration of 200 nM. At this MutS concentration, which is approximately equal to the K_d for binding to heteroduplex DNA (above), the SPRS results shown in Figs. 1-3 indicate a relative ternary complex stoichiometry of ~0.6 and 0.8 mol of hMutL/mol of hMutS for the 100- and 200-bp heteroduplexes, respectively. This calculation is based on the extent of hMutS binding to heteroduplex DNA after correction for nonspecific complexes formed with homoduplex controls (Figs. 1 and 3). It is noteworthy that a similar hMutL-dependent enhancement of heteroduplex-bound mass was observed at 800 nM hMutS (4 times the K_d), providing additional evidence that the observed mass increase is due to hMutL binding rather than increased hMutS binding in the presence of the MutL homolog. It is important to note that we have consistently found the 110-bp heteroduplex to be somewhat less effective than the 200-bp DNA in supporting ternary complex formation, as judged by the additional mass enhancement observed in the presence of hMutL.

hMutS·hMutL·Heteroduplex Complexes Are Dynamic-- The kinetic lifetimes of hMutL·hMutS·heteroduplex complexes were monitored with SPRS by terminating protein flow and continuing wash with reaction buffer containing ATP and Mg²⁺ (Fig. 4, upper curve). Dissociation of the ternary complexes from the 200-bp G-T heteroduplex was multiphasic, with decay curves fitting well to a sum of two exponentials. The major amplitude (60% of the complexes) dissociated rapidly (t_1/2 ~ 1 s), a second component (20%) dissociated more slowly (t_1/2 ~ 10 s), and the residual (20%) dissociated so slowly that a rate could not be estimated. Complexes prepared with homoduplex DNA in the presence of hMutS and hMutL displayed similar multiphasic dissociation kinetics (Fig. 4 lower curve). The major species (60%) dissociated rapidly with a t_1/2 of ~1 s, the second component (16%) dissociated more slowly (t_1/2 ~ 19 s), with the residual (24%) dissociating too slowly to permit an estimate of lifetime.

View larger version (15K): [in this window] [in a new window]   Fig. 4.   The hMutS·hMutL· heteroduplex ternary complex turns over rapidly in the presence of ATP. Ternary complex formation between 200 nM hMutS, 200 nM hMutL, and a 200-bp G-T heteroduplex was monitored by SPRS as described in the legend to Fig. 1. Protein flow was terminated at the discontinuity, and wash with reaction buffer (20 mM Tris-HCl, pH 7.6, 100 mM KCl, 1 mM dithiothreitol, 5 mM MgCl2, 0.005% surfactant P20, and 1 mM ATP) continued. These results are shown in the upper curve (). The lower curve () presents an otherwise identical experiment performed with a 200-bp A·T homoduplex. The nature of the dissociation curves, which were multiphasic in both cases, is discussed in the text.

The relative extents of binding to heteroduplex and homoduplex DNAs (Figs. 1, 3, and 4) indicate that 60-70% of the protein mass bound to the heteroduplex under these conditions is dependent on the presence of a single mismatch. We therefore think it likely that the more rapidly dissociating heteroduplex species (t_1/2 values of 1 and 10 s, 80% by mass) correspond to several classes of mismatch-dependent ternary complexes. This implies that the levels of these species observed by SPRS prior to termination of protein flow correspond to a dynamic steady state, i.e. the complexes are turning over rapidly via dissociation and reassociation. It is important to emphasize that the multiphasic dissociation kinetics observed with both heteroduplex and homoduplex DNAs imply the existence of several distinct types of specific and nonspecific complexes.

hMutL·hMutS·Heteroduplex Ternary Complexes by Gel Shift Analysis-- The chain length dependence of formation and the nature of hMutL·hMutS·heteroduplex ternary complexes was also examined by electrophoretic gel shift assay. As observed by SPRS, electrophoretic assay indicated that hMutL did not bind detectably to 41-, 100-, or 200-bp G-T heteroduplexes (Fig. 5). In the presence of hMutS, specific complexes were evident with each of these heteroduplexes, and the presence of both proteins led to production of one or more supershifted species. A hMutL-dependent supershifted complex was produced with the 41-bp heteroduplex, as well as its homoduplex control. Although production of this species required presence of hMutS, the lack of a mismatch requirement indicates that it is nonspecific in nature. Three supershifted species were observed with 100- and 200-bp DNAs in the presence of hMutS and hMutL (these were in addition to the hMutS·heteroduplex binary complex, which was evident at a low level with the 200-bp substrate). Two of these (Fig. 5, asterisks) were produced with both homoduplex and heteroduplex, and as observed with 41-bp DNAs, production of these nonspecific complexes was dependent on the presence of both hMutS and hMutL. However, with both 100- and 200-bp DNAs, a heteroduplex-specific, supershifted complex was also produced (Fig. 5, arrows). Combined Southern and Western blot analysis confirmed the presence of both hMutS and hMutL in specific and nonspecific ternary complexes produced with the 100-bp G-T heteroduplex (Fig. 6).

View larger version (36K): [in this window] [in a new window]   Fig. 5.   DNA chain length dependence of hMutL and hMutS ternary complex formation determined by gel shift analysis. Electrophoretic gel shift analysis of protein-DNA complexes was performed as described under "Experimental Procedures" with 41-, 100-, and 200-bp G-T heteroduplexes, and otherwise identical A·T homoduplex control DNAs. Reactions contained 200 nM hMutS, 200 nM hMutL, and 0.5 mM ATP as indicated. Specific complexes are indicated by arrows, and nonspecific complexes by asterisks.

View larger version (50K): [in this window] [in a new window]   Fig. 6.   Specific and nonspecific ternary complexes contain hMutS and hMutL. Gel shift reactions (see "Experimental Procedures") contained 200 nM hMutS, 200 nM hMutL, 32P-labeled 100-bp G-T heteroduplex (lanes 1 and 3 of each experiment) or A·T homoduplex (lane 2 of each experiment). ATP (0.5 mM) was present in reactions shown in lanes 2 and 3, but omitted from samples shown in lane 1. After electrophoresis, protein and DNA were electrotransferred to nitrocellulose and DEAE membranes (see "Experimental Procedures"). DNA bound to the DEAE membrane was visualized by autoradiography and nitrocellulose-bound mismatch repair polypeptides identified by Western blot. In the experiment shown at the top, the nitrocellulose membrane was probed as indicated with anti-MLH1, and then with anti-MSH6 after stripping. The experiments shown in the center and at the bottom were performed in a similar manner except that stripping was not used; parallel gels were examined individually for DNA and MSH2, or for DNA and PMS2. Specific complexes are indicated by arrows and nonspecific complexes by asterisks. In the experiment shown at the top, the nonspecific complex that runs more slowly than the specific component is barely visible in the DNA and MSH6 panels, but is evident in the MLH1 panel. The faster migrating species in lane 1 of each DNA panel corresponds to the hMutS·DNA complex. That portion of the gel where free DNA runs is not shown.

These observations confirm the chain length dependence of specific ternary complex formation observed in SPRS experiments. By contrast, nonspecific complexes of the sort observed by gel shift assay were not detected as a mass enhancement in SPRS experiments with homoduplex DNA (Fig. 1). The reason for this is not clear, but as noted above, the multiphasic dissociation kinetics observed by SPRS with homoduplex DNA could be indicative of several classes of nonspecific complex.

The stability of ternary complexes was evaluated by challenge of preformed complexes with polyd(I)·d(C). As shown in Fig. 7, polyd(I)·d(C) challenge resulted in a dramatic reduction in ternary complex formation with the 200-bp G-T (lanes 4 and 5). The yield of binary hMutS·heteroduplex complexes was also extremely low under these conditions when ATP was present in the reaction. However, distinct results were obtained when the two ends of the duplex were blocked with biotin-streptavidin complexes. The presence of end blocks at both heteroduplex termini stabilized binary hMutS·heteroduplex complexes in the presence of ATP (compare lanes 2 and 3 with lanes 8 and 9), confirming previous observations in this respect (20, 27). The biterminal end block also stabilized hMutL·hMutS·heteroduplex ternary complexes, but a single end block did not; that fraction of the heteroduplex that contained only one end block was recovered as free DNA after polyd(I)·d(C) challenge, whereas heteroduplexes with streptavidin-biotin blocks at both duplex termini were not (compare lanes 10 and 11). These observations are consistent with the conclusion discussed above that ternary complexes are dynamic in nature.

View larger version (79K): [in this window] [in a new window]   Fig. 7.   The hMutL·hMutS·heteroduplex ternary complex is dynamic. Gel shift analysis was performed as described under "Experimental Procedures" in the presence or absence of 0.5 mM ATP using 3'-32P-labeled 200-bp G-T heteroduplex (lanes 1-5 and 7-11) or a control 200-bp A·T homoduplex (lanes 6 and 12). Both DNAs contained a 5'-terminal biotin on each DNA strand. Reactions in the right panel contained 0.5 mg/ml streptavidin and were preincubated 10 min prior to addition of mismatch repair proteins to allow conjugation of biotin. Reactions were initiated by addition of hMutS and hMutL as indicated, to a final concentration of 200 nM. After 10 min at room temperature, a poly(dI)·d(C) competitor was added to a final concentration of 25 µg/ml. Reactions were terminated after an additional 5-min incubation subjected to polyacrylamide gel electrophoresis (see "Experimental Procedures"). The location of the specific ternary complex formed in the absence of streptavidin is shown by an arrow. Mobilities of free DNA with 0, 1, or 2 bound streptavidin molecules are also indicated.

Due to the size of the heteroduplex and the presence of streptavidin end blocks, the supershifted complex observed in the presence of hMutS and hMutL was not resolved into specific and nonspecific components (lanes 10 and 11, compare with lane 9). However, a surprising effect of streptavidin-biotin end blocks became evident upon examination of nonspecific interactions with the 200-bp homoduplex. Two classes of nonspecific complex are produced with homoduplex DNA in the presence of hMutS, hMutL, and ATP (Fig. 5, lane 8; Fig. 7, lane 6). Unexpectedly, the corresponding nonspecific complexes were not detectable under conditions where the homoduplex molecules were blocked at both ends with streptavidin-biotin complexes (lane 12). These observations suggest that presence of free duplex DNA termini have an important role in the production of nonspecific complexes that are observed by gel shift assay.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Although MutS homologs bind readily to small synthetic heteroduplexes (10, 18, 23, 24, 29-31), the experiments described here indicate that formation of the hMutL·hMutS·heteroduplex requires ATP (and probably its hydrolysis) and depends on DNA chain length. The 200-bp heteroduplex used in this report supports efficient ternary complex formation, 100- and 110-bp heteroduplexes appear to be somewhat less effective in this regard, and we have been unable to detect mismatch-dependent ternary complex formation with a 41-bp heteroduplex substrate. We have also obtained similar results with E. coli MutS and MutL.³

The SPRS and gel shift analyses described here show that the hMutL·hMutS·heteroduplex ternary complex is dynamic, undergoing rapid dissociation and reassociation in the presence of ATP·Mg²⁺ at near physiological ionic strength. In fact, the kinetics of dissociation of ternary complexes are similar to those observed with the binary hMutS·heteroduplex in the presence of ATP. As observed for ternary complexes (Fig. 4), dissociation of binary complexes as monitored by SPRS was multiphasic (data not shown); the major amplitude dissociated rapidly (60% of complexes, t_1/2 = 3 s), a second species dissociated more slowly (26%, t_1/2 = 50 s), and a third component (14%) dissociated too slowly to determine an accurate rate. These observations are similar to those obtained previously by Galio et al. (13) in SPRS studies of bacterial MutS and MutL. As described here for hMutS and hMutL, these earlier studies led to the conclusion that ternary complexes of MutS, and MutL, with a 149-bp heteroduplex turn over rapidly as compared with the lifetime of MutS·heteroduplex complexes. In the case of the human system, we have also shown that hMutL· hMutS·heteroduplex complexes can be kinetically stabilized by placement of a physical block at each end of a linear DNA. The simplest interpretation of this finding is that turnover of the ternary complex depends on movement of one or both mismatch repair proteins along the helix with dissociation occurring at free ends. By contrast, the recent work of Hsieh and colleagues (32) has led to the conclusion that bacterial MutL stabilizes mismatch-bound MutS in the presence of ATP, resulting in a much longer lifetime for the MutL·MutS·heteroduplex ternary complex as compared with that of the MutS·heteroduplex (32). The basis of these differing conclusions is uncertain, although different methods were used to monitor dissociation kinetics. Our conclusions and those of Galio et al. (13) are based on real time analysis using SPRS, whereas Schofield et al. (32) monitored dissocation kinetics by gel shift assay after addition of a heteroduplex trap.

The molecular basis of the chain length dependence for ternary complex formation is not clear, but there are a number of potential explanations for this effect. One possibility is that the presence of flanking homoduplex is necessary to accommodate both hMutS and hMutL. The formation of nucleoprotein complexes containing hMutL that we have detected require hMutS; however, the ATPase of bacterial MutL is known to be activated in the absence of MutS by single strands and to a lesser extent by duplex DNA (33, 34), implying presence of a DNA binding center. MutL and hMutL are large asymmetric proteins (Stokes radii of 61 and 74 Å, respectively (Refs. 11 and 19)) and are potentially capable of occluding a substantial segment of helix. It is also possible that ternary complex formation involves oligomerization (or polymerization along the helix) of hMutS or hMutL. Indeed, we have concluded that ternary complexes with 200-bp heteroduplex DNA contain several copies of each heterodimer, but as discussed above, the presence of multiple protein copies can also be explained by a mechanism that invokes movement of repair protein complexes along the helix contour. A third interesting possibility is that the chain length requirement is indicative of a major DNA conformational transition associated with ternary complex formation, e.g. the opening of a significant length of helix or the introduction of a substantial bend, perhaps due to a partial wrapping of DNA about one of the repair activities.

	ACKNOWLEDGEMENTS

We thank Keith Bjornson for many useful discussions during the course of this work and for comments on the manuscript. We also thank Elisabeth Penland for preparation of baculovirus-infected SF9 cells.

	Addendum

While this manuscript was in preparation, we learned of a paper in press by Hsieh and colleagues (32) demonstrating chain length-dependent effects with respect to bacterial MutL modulation of MutS·heteroduplex interaction. The DNA chain length effects observed in the two systems are not surprising since early work with the bacterial proteins demonstrated that the MutS footprint in the presence of DNase I expands dramatically from 20 bp to ~100 bp in the presence of MutL and ATP (11).

	FOOTNOTES

^* This work was supported in part by Grant GM45190 from the NIGMS, National Institutes of Health.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.

^¶ Investigator of the Howard Hughes Medical Institute. To whom correspondence should be addressed. Tel.: 919-684-2775; Fax: 919-681-7874; E-mail: modrich@biochem.duke.edu.

Published, JBC Papers in Press, July 3, 2001, DOI 10.1074/jbc.M105076200

² D. Chandrasekhar, D. Allen, L. Blackwell, and P. Modrich, unpublished observations.

³ L. J. Blackwell and P. Modrich, unpublished experiments.

	ABBREVIATIONS

The abbreviations used are: SPRS, surface plasmon resonance spectroscopy; bp, base pair(s); PCR, polymerase chain reaction; HPLC, high performance liquid chromatography; RU, response units.

	REFERENCES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

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