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INTRODUCTION |
Bases in DNA suffer oxidative damages from normal cellular
metabolism even in the absence of exogenous oxidative stress (1, 2).
The resulting alterations in base-pairing properties (3) lead to
increased mutational frequencies at these damaged bases during
subsequent replication (4) with wide-ranging pathological consequences
(5). The accumulation of oxidative lesions in DNA have been associated
with aging (6-10) and are also strongly correlated to cancers
resulting from known exposure to oxidative environmental carcinogens
(1, 11-15).
The removal of oxidative lesions and the correction of mutagenic
mispairing at these lesions, therefore, represent two essential but
distinct lines of defense against genome degradations. The prevalent
oxidatively damaged base, 8-oxo-guanine, is excised from DNA when
base-paired to cytosine by MutM (16-20) and its eukaryotic homolog,
OGG1 (21-26). Misincorporation of adenines by DNA polymerase opposite
8-oxo-guanines that escape MutM-mediated removal are subsequently
repaired by the adenine glycosylase, MutY (20, 27-33), and its
homolog, MYH (16, 34-36). Additionally, MutT (16, 20, 37-40), a
nucleoside triphosphate pyrophosphohydrolase, removes oxidatively
damaged dGTPs, thereby preventing their incorporation into DNA.
MutY, therefore, functions physiologically as a mismatch repair enzyme,
as originally suggested (41, 42), albeit targeted toward adenines
misincorporated by DNA polymerase at template 8-oxo-guanines. As such,
the accuracy of repair depends on the exclusive removal of adenines
from the newly synthesized, daughter strand of DNA. By contrast, MutY
repair activity at a mispair where the 8-oxo-guanine has been
misincorporated opposite a template adenine would be mutagenic. The
observation (43) that A:T
C:G transversion frequencies in
MutT
/MutY+
Escherichia coli is increased relative to
MutT
/MutY
strains,
whereas the high rate of C:G
A:T mutations in
MutY
strains is unaffected by the status of
MutT, corroborates the mutagenic effects of such
MutY-mediated removal of adenines from the template strand.
However, the generally accepted view of MutY as a prototypical
mono-functional base excision glycosylase provides no mechanistic basis
for daughter versus template strand selectivity.
Interestingly, the human MutY homolog, hMYH, has recently been
shown to interact with the human mismatch repair enzyme, hMSH
(44),
providing the first insight into the mechanistic possibility that
nascent strand recognition may result from cooperative recruitment of the mismatch repair system. Even so, such cooperative interaction between different repair systems does not mitigate the need for a
regulatory mechanism intrinsic to MutY itself to forestall
indiscriminate excision prior to recruitment. Toward this end, we have
reexamined the kinetic mechanism of MutY using presteady-state active
site titrations to reveal a polysteric and allosteric basis for such a
molecular switch.
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EXPERIMENTAL PROCEDURES |
Buffers and Reagents--
Except as noted, all buffers were made
with reagent grade chemicals and Milli-Q Plus (Millipore, Bedford, MA)
purified distilled-deionized water. MutY Storage Buffer contained 50 mM HEPES-NaOH (pH 7.5), 500 mM NaCl, and 50%
spectroscopic grade glycerol. MutY Assay Buffer contained 5 mM HEPES-NaOH (pH 7.5), 20 mM Tris-HCl (pH 7.5), 50 mM NaCl, and 5% spectroscopic grade glycerol. All
buffer stock solutions were filtered through 0.2-µm polyethersulfone filters (Nalgene, Rochester, NY).
Enzymes and Oligonucleotides--
The E. coli MutY
was purified from E. coli BL21(DE3) harboring the
overproducing plasmid pET24a/MutY-8 as described previously (53). The
MutY concentration was determined spectrophotometrically using an
extinction coefficient of
280 = 77,510 M
1cm
1. Purified MutY had an
A400 to A280 ratio
between 0.15 and 0.19 and was stored in MutY Storage Buffer at
80 °C. Synthetic oligodeoxyribonucleotides were synthesized and
further purified by urea-polyacrylamide gel electrophoresis and
electro-elution as described previously (61). Duplex substrates were
constructed by annealing 5'-32P-labeled *t-AAT
(5'-CATCAGAACAATTCATCGTTA) with its complement, b-AOT
(5'-TAACGATGAAOTGTTCTGATG, O = 8-oxo-guanine) as
described (61), placing a single adenine:8-oxoguanine mismatch at the center (position 11) of the sequence as denoted by the underscore. Concentrations were determined spectrophotometrically using extinction coefficients calculated according to Cantor et al.
(62). T4 polynucleotide kinase was obtained from USB (Cleveland,
OH), and [
32P]ATP (3000 Ci/mmol) was obtained from
Amersham Biosciences.
Presteady-State Excision Assays--
Single-turnover experiments
were carried out using a KinTek RFQ-3 rapid chemical quench (KinTek
Instruments, State College, PA) instrument thermostatically maintained
at 37 °C with a Neslab RTE-111 refrigerated water bath. Reactions in
MutY Assay Buffer were initiated by rapidly mixing 15 µl of MutY with
15 µl of 5'-32P-labeled duplex DNA substrate and were
chemically quenched with 90 µl of 0.2 M NaOH. The abasic
site-containing excision products were then heated to 90 °C for 8 min to cleave the phospho-ribose backbone of the DNA at the abasic site
to generate 10-nucleotide products, which were separated from the
21-nucleotide substrates by electrophoresis on a 20% acrylamide, 8 M urea gel. Following visualization with a
PhosphorImager (Amersham Biosciences), the intensities of DNA substrate
and product bands were quantified using ImageQuant software (Amersham
Biosciences) as described (47, 61). Single-turnover time courses with
enzyme present in excess over substrate were fitted to a single
exponential function, [P]t = A1(1
e
kobst).
First turnover time courses in excess substrate were also fitted to
single exponential functions for time courses less than 2 min because
of the undetectable contribution of the linear steady-state phase
within this time frame.
First turnover burst amplitudes were determined by performing excision
assay using time points in excess of 4 min. Accurate data could be
obtained using a minimal set of three time points at 4, 8, and 12 min;
however, time courses were typically linear for up to 2 h. Burst
amplitudes were obtained as the y-intercept values returned by the
linear regression best fit of the data. Active site titrations
consisted of a series of first turnover burst amplitude determinations
obtained over a serially diluted range of MutY concentrations at fixed
DNA concentrations.
Fit to Statistical Thermodynamic Model--
The statistical
thermodynamic model of Fig. 2 has been described previously in detail
(48, 49). Fits to active site titrations were modeled after the species
fraction of the Y2D complex as given by Equation 1.
Nonlinear fits were obtained for all data sets using an
adaptive nonlinear regression algorithm (63) as implemented in the
program NLREG (Phillip H. Sherrod, Brentwood, TN). Fractions of DNA
found in the Y2D2 and YD subpopulations were
simulated according to Equations 2 and 3.
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(Eq. 2)
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(Eq. 3)
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Simulated curves shown were generated using KaleidaGraph v.3.51
(Synergy Software, PA) as described (48, 49).
Footprint Titrations--
Hydroxyl-radical (Fe-EDTA)
footprint titrations were adapted from published protocols (50, 52,
64). Duplex tAAT:bAOT (100 nM), radiolabeled either in the
adenine, *tAAT, or 8-oxo-guanine strand, *bAOT, were incubated with
MutY at 37 °C in Assay Buffer containing 2% glycerol for 30 s
prior to the addition of 25 µM ferrous ammonium sulfate,
50 µM EDTA, 5 mM sodium ascorbate, and 0.24%
H2O2 followed by further incubation at room
temperature for 8 min. Reactions were quenched by transfer to
prechilled tubes containing ice-cold 25% glycerol, 4 M
thiourea, and 4 M urea. Elevated ascorbate and
H2O2 concentrations used were optimally calibrated to account for the presence of the 2% glycerol. Conversely, inclusion of the usual 5% glycerol in the assay buffer inhibited DNA
cleavage. Footprints were visualized following separation by denaturing
PAGE in 8 M urea and Tris-borate-EDTA (TBE) on an Amersham
Biosciences PhosphorImager. Intensities of bands corresponding to
protected regions, designated as If,i, were
quantified using ImageQuant software and compared with the intensities
of reference bands outside of the footprint region, designated as Ib,i, to correct for background variability between
lanes. The corrected intensity at each ith
concentration of MutY was further normalized against the corrected intensity in the absence of MutY to obtain the normalized intensity, Inorm,i, according to Equation 4,
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(Eq. 4)
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where If,0 and
Ib,0 represent the intensities of the
footprint and reference bands, respectively, in the absence of MutY.
The extent of binding at each MutY concentration,
i, was calculated for each concentration of MutY
by normalizing against the maximum extent of protection observed, Inorm,
, by Equation 5,
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(Eq. 5)
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where Inorm,
was obtained by averaging
normalized intensities at saturation.
Gel Mobility Shift Assays--
Gel mobility shift assays were
performed in MutY Assay Buffer without glycerol at 6, 8, 10, and 12%
acrylamide using a Bio-Rad Mini-PROTEAN 3 apparatus. Using a 10-well
comb, samples were prepared in MutY Assay Buffer with glycerol and
loaded into the central two wells of each gel. Sample buffer containing
bromphenol blue was loaded into the remaining wells. Electrophoresis
was carried out at 70 V for 40 min at room temperature. Due to the poor
buffering capacity of the MutY Assay Buffer, the pH in the outer lanes
would drop during the course of the run as indicated by a color change of the bromphenol blue tracking dye from blue to yellow. Care was,
therefore, taken to calibrate the time of electrophoresis to maintain
blue coloring in the tracking dye of the lanes immediately adjacent to
the innermost, sample-containing lanes. Inclusion of glycerol in the
gel buffer as well as electrophoresis at reduced temperatures both
exacerbated the pH artifact.
 |
RESULTS |
First Turnover Kinetics and Active Site Titrations--
For
presteady-state assay of MutY activity, we radiolabeled the
adenine-containing strand of a 21-bp duplex substrate,
*tAAT:bAOT, which contained a centrally positioned
adenine:8-oxo-guanine mispair. The base-labile abasic product was
cleaved in 0.2 N NaOH at 80 °C to generate a shortened
10-mer, which was separated from the 21-mer substrate by denaturing
polyacrylamide gel electrophoresis. In a typical single-turnover assay
performed with 4-fold molar excess of MutY (200 nM) over
substrate (50 nM), we observed rapid, single-exponential
excision of 98 ± 2% of the mispaired adenine at 0.24 ± 0.01 s
1, (Fig.
1A, closed
circles). In experiments performed with excess substrate (200 nM) over enzyme (50 nM), however, only 20 ± 0.4% of the substrates were turned over in the exponential phase
but with the identical rate constant of 0.22 ± 0.01 s
1 (open circles). Data obtained at longer
times (Fig. 1B) showed that a presteady-state burst
of product was formed in the first turnover followed by steady-state
turnovers at 2 × 10
6 s
1 corresponding
to the slow rate of one or more product release step(s) that limited
the regeneration of free active sites. The amplitude of the burst
phase, therefore, provided a direct measure of the molar amount of
productively bound substrate during the initial turnover (45-47).

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Fig. 1.
MutY excision assays.
As shown in A, presteady-state excision of adenine from an
adenine:8-oxo-guanine mismatch obtained under single-turnover
conditions ( , 200 nM MutY, 50 nM DNA)
yielded a single-exponential best-fit amplitude of 0.98 ± 0.02 and a rate constant of 0.24 ± 0.01 s 1. Under burst
or steady-state conditions ( , 50 nM MutY, 200 nM DNA), only 0.2 ± 0.004 of the adenines were
excised in the first turnover; however, the rate constant of 0.22 ± 0.01 s 1 for excision in the burst phase was within
error identical to the rate constant measured with excess substrate. As
shown in B, steady-state turnover following the initial
burst phase, measured for 2 h at 50 nM MutY and 200 nM DNA, was extremely slow at 2.1 ± 0.2 × 10 6 s 1.
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In active site titrations, burst amplitudes were measure at different
MutY and DNA concentrations. The resulting titration curves, plotted as
a function of MutY concentration (Fig.
2A), were sigmoidal at eight
different DNA concentrations ranging from 10 to 600 nM. At
saturating DNA concentrations (>200 nM), complete binding
of substrate required a 2-fold excess of MutY, limiting the reaction
stoichiometry to
2. However, the nonlinear response at lower MutY
concentrations ruled out a monomeric oligomeric species with a single
binding site (48, 49).

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Fig. 2.
Active site titrations. As shown in
A, for active site titrations, first turnover amplitudes
were obtained as the y-intercept values extrapolated from a linear fit
to time points at 4, 6, and 8 min. These amplitudes, representing the
fraction of DNA bound in productive complexes, were plotted as a
function of MutY concentration at 10 ( ), 25 ( ), 50 ( ), 100 ( ), 200 ( ), 300 ([triao]), 400 ( ), and 600 nM
( ) *tAAT:bAOT. As shown in B, titration data from
panel A were replotted as fractional occupancy of MutY
active sites as a function of DNA concentration at 71 ( ), 95 ( ),
127 ( ), 169 ( ), 225 ( ), 301 ( ), and 401 nM
( ) MutY. Solid lines represent simulated species
population according the model in Fig. 3A using best-fit
parameters listed in Table I as described in the text.
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To further clarify the apparent reaction stoichiometry, we
reconstructed the titration data to reflect the fractional occupancy of
total available active sites as a function of increasing DNA concentrations at various fixed concentrations of MutY (Fig.
2B). The titration curves were linearly dependent on DNA
concentration up to ~45% saturation of active site corresponding to
a reaction stoichiometry of two MutY monomers per substrate molecule.
Addition of DNA beyond this point, however, resulted in decreased burst amplitudes. As mass action dictated that the addition of DNA ligand must lead to additional total binding, the decline in burst amplitude demonstrated allosteric inhibition of the initial population of two-to-one active complexes by the binding of a second equivalent of
substrate. Furthermore, the reduction in burst amplitudes was not
accompanied by any change in exponential time course of the first
turnover (Fig. 1A); therefore, the binding of the second substrate abolished, rather than attenuated, the excision activity of
the dimer. Our results demonstrate that the active form of MutY is a
half-saturated dimeric, Y2D, whose excision activity is
allosterically inhibited upon binding a second substrate to form the
fully saturated dimer-DNA complex, Y2D2.
Identical results were obtained using MutY purified by other protocols
including enzyme refolded from guanidine-solubilized inclusion bodies
(data not shown).
Statistical Thermodynamic Model--
To quantitatively model this
allosteric regulation, we used a general statistical thermodynamic
scheme (Fig. 3A) developed previously to describe the protein assembly and DNA binding properties of dimeric proteins (48, 49). The model required four interaction parameters to describe the dynamic equilibrium between two monomeric states, Y and YD, and three dimeric states, Y2,
Y2D, and Y2D2. K0 describes the protein-protein dimerization
constant in the absence of bound DNA. K11
describes the DNA binding affinity of the monomer for DNA.
K1 describes the dimerization constant between a
DNA-bound monomer and a free monomer to form the half-saturated dimer,
Y2D. Lastly, K22 describes the DNA
affinity of Y2D for a second DNA molecule to form the fully
saturated dimer, Y2D2. The two remaining
equilibrium constants, K2 and
K21, were constrained by thermodynamic linkage
to the values of K0, K1,
K11, and K22. Lastly, the
value of K0, the dimerization constant in the
absence of substrate, was arbitrarily fixed at 10 mM
1 as gel filtration chromatography
experiments showed that MutY was monomeric in the absence of DNA even
at micromolar concentrations (data not shown).

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Fig. 3.
Dimeric model. As shown in A,
the active site titration data were modeled using the thermodynamic
linkage cycle of Wong and Lohman (48). Y, D, and
YD represent unbound MutY monomers, unbound
adenine:8-oxo-guanine-containing oligonucleotide substrates, and
monomeric MutY-substrate complexes, respectively.
Y2, Y2D and
Y2D2 represent MutY dimers bound
with zero, one, and two substrates. Of the six equilibrium association
constants shown, only four (K0,
K11, K1, and
K22) were needed to describe the linkage scheme.
Additionally, K0 was constrained at 1 × 104 M 1 based on the absence of
dimerization in the absence of DNA. Global best fit of all active site
titrations was obtained based on Y2D being the catalytic
species. As shown in B, simulated species fractions of
Y2D, Y2D2, and YD showed that
activity correlated only with Y2D. Additionally, the
apparent sigmoidal behavior stemmed from DNA binding to form
nonproductive species, Y2D2 and YD, at low
molar ratios of MutY:DNA.
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Previously, we have shown that titration data obtained at four
different DNA concentrations were sufficient to yield satisfactory resolution of the fitted interaction parameters in global nonlinear regression analysis using this model (48). Nonlinear best fit of the
active site titrations obtained at eight DNA concentrations to this
model showed excellent global agreement between the burst amplitudes
and the species population of Y2D predicted by the model (Fig. 2, solid lines) at all DNA and MutY
concentrations, yielding the values for K1,
K11, and K22 shown in
Table I. Simulations were insensitive to
values of K0 < 40 mM
1, confirming our assumption that the
unliganded dimer is sparsely populated and validating the value of
K0 chosen for the fitting. DNA binding to the
monomer was relatively weak (K11 = 6 µM
1) but was strongly linked to protein
dimerization with a 5 order of magnitude increase in dimerization
constant (K1 = 3 nM
1)
upon binding DNA, placing the affinity for the DNA by Y2,
the unliganded dimer, at K21 > 450 nM
1 (Kd, 21 < 2.2 pM). This positive polysteric linkage between DNA binding
and protein oligomerization thermodynamically guaranteed near
stoichiometric binding of DNA by a MutY dimer despite the weak affinity
of the monomer for DNA. The active half-saturated dimer,
Y2D, however, bound the second equivalent of DNA much more weakly, with K22 = 1.5 µM
1. This negative cooperativity of binding
by Y2D coupled with the negative allosteric inhibition of
excision activity by the second bound lesion together accounted for the
apparent gradual decline of burst amplitude observed at higher DNA
concentrations.
Fig. 3B shows the population distribution of all DNA-bound
species for an active site titration at 300 nM DNA. As
expected, the data correlated well with the sigmoidal accumulation of
Y2D with increasing MutY added. At low MutY concentrations,
however, the high ratio of DNA to MutY favored the formation of species with a one-to-one binding stoichiometry. Accordingly, the fully saturated species, Y2D2, was maximally
populated at equimolar concentrations of MutY and DNA (300 nM) but disproportionated to form Y2D at higher
MutY concentrations. Because Y2D2 lacked excision activity, the adenine:8-oxo-guanine mismatches of DNA bound in
this configuration were silent in active site titrations, thereby
giving rise to the sigmoidal behavior observed. Because the DNA-bound
monomer, YD, accounted for only ~2% of the DNA bound at this DNA
concentration due to the strong positive polysteric linkage, we are
less certain that YD also lacked excision activity. However, fits to
any model with full excision activity by YD tended to systematically
lessen the sigmoidality of the resulting titration curves at all DNA
concentrations, compelling us to hypothesize that the monomer is inactive.
Footprint Titrations--
Because the model proposes the existence
of a silent Y2D2 state, we performed
hydroxyl-radical DNA footprint (50) titrations to directly monitor all
site-specific DNA interactions (51, 52). Fig.
4A shows sequencing gels of
footprint titrations performed in parallel using duplex substrate DNA
radiolabeled either on the adenine strand, *tAAT (left), or
on the 8-oxo-guanine strand, *bAOT (right). The adventitious
lyase activity of MutY resulted in partial strand cleavage at the
position of the scissile adenine, which was visible in the *tAAT
footprint. Additionally, a distinct, MutY-dependent
footprint covering 2-3 nucleotides was observed immediately 5' of the
scissile adenine. The footprint on the 8-oxo-guanine strand, *bAOT,
covered 4-5 nucleotides and appeared to be centered about the
8-oxo-guanine.

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Fig. 4.
Hydroxyl-radical DNA footprint
titrations. In A, sequencing gels show MutY
concentration-dependent protection of substrate DNA,
tAAT:bAOT (100 nM) from hydroxyl-radical-induced strand
cleavage. Experiments performed with the radiolabel on *tAAT
(left) showed protection on the adenine-bearing strand
(indicated by bold bar marked If,i)
several bases 5' of the scissile adenine. The position of the scissile
adenine is marked by major cleavage product generated by the
adventitious lyase activity of MutY. Experiments using radiolabeled
*bAOT showed protection on the 8-oxo-guanine-bearing strand surrounding
the 8-oxo-guanine. The bold bars marked
Ib,i denote regions of unprotected DNA used for
normalization. In B, a plot of fractional protection as a
function of MutY concentration showed that protection on the adenine
strand ( ) was sigmoidal and co-titrated with the active site
titration data ( ). The solid line represents the
simulated Y2D population. In contrast, the protection on
the 8-oxo-guanine strand ( ) was not sigmoidal and correlated well
with the modeled total fraction of DNA bound.
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Quantitation of the extent of protection by MutY (Fig. 4B)
revealed different protein concentration dependences for the observed footprints on the two strands of DNA. The protection of the adenine strand was sigmoidal and correlated well with both the active site
titration data and the predicted population of Y2D. In
contrast, the protection observed on the 8-oxo-guanine strand was not
sigmoidal, correlating instead with the predicted fraction of
total bound DNA. This difference in concentration dependences between
the protection patterns on the two strands supported the presence of
multiple binding modes. More importantly, the stoichiometric titration
of the protection on the 8-oxo-guanine strand demonstrated that more
DNA was bound at low MutY concentrations than was apparent in the
excision-based active site titrations, thereby establishing the
existence of non-productive complexes. Furthermore, the sigmoidal titration of the scissile strand footprint provided evidence of structurally distinct protein conformations associated with productive versus non-productive binding. The correlation between
activity and localized protection immediately adjacent to the scissile base likely reflected the formation of intimate contacts with the
active site in the productive complex, Y2D, that are absent in the inhibited complexes, perhaps suggesting that the adenine base
may not be flipped out of the DNA duplex, as is required for excision
(53), in the inhibited Y2D2 state.
Direct Detection of Inhibited Complexes by Gel Mobility
Shift--
Gel mobility shift assays provided an alternate method of
separating different binding complexes based on differences in mass and
in charge. Previous reports using this technique to measure DNA binding
affinity of MutY, however, revealed only a single bound species with
bands corresponding to higher order aggregates appearing only at high
concentrations of MutY (54-56). Although simple to perform, gel
mobility shift assays have been shown by Engler et al. (57)
to be sensitive to buffer-dependent artifacts. In our own
attempts to reproduce assays under published conditions using
TBE buffer in the gel, we observed that the banding patterns as well as
the resultant titration curves changed with acrylamide concentration.
We, therefore, tested a variety of gel buffers other than TBE and found
that MutY Assay Buffer without glycerol reproducibly provided good results.
Fig. 5A shows typical results
obtained with 100 nM DNA at 1:1 and 2:1 MutY to DNA molar
ratios resolved on 6, 8, 10 and 12% polyacrylamide gels in this
buffer. In all cases, three distinct bands were visible. Assays run
without MutY (data not shown) verified that the fastest migrating
species, band III, corresponded to free DNA. Most of the DNA was
unbound at a 1:1 ratio of MutY to DNA. At this ratio, the bound DNA was
roughly equally divided between bands I and II with the faster
migrating band II showing slightly more DNA than band I. On increasing
the MutY:DNA ratio to 2:1, however, most of the DNA became bound as
band II. Additionally, the amount of DNA in band I, the slowest
migrating species, actually decreased at the higher MutY concentration.
Identical results were observed at all four acrylamide concentrations.
Fig. 5B, which compares identical samples resolved in 10%
polyacrylamide gels in TBE versus MutY Assay Buffer, shows
the absence of band I in TBE gels at both MutY:DNA molar ratios,
demonstrating that the ability to detect this species was gel
buffer-dependent.

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Fig. 5.
Gel mobility shift analysis. As shown in
A, gel mobility shift assays were performed at 100 nM DNA and either 100 (1:1) or 200 nM (2:1)
MutY and electrophoretically separated in 6, 8, 10, and 12% acrylamide
gels using MutY Assay Buffer. In addition to the free DNA (band
III), two slower migrating bands (I and II)
corresponding to complexes were detected. The faster migrating of the
two (band II) was preferentially populated at higher MutY
concentrations. As shown in B, samples subjected to
electrophoresis on a 10% gel using TBE instead of MutY Assay Buffer
failed to show band I. As shown in C, Ferguson analysis was
performed by plotting the logarithm of the mobility versus
acrylamide concentration of a band. The resulting linear correlations
showed that band I was more subject to both the sieving effect of the
matrix (steeper slope) and the applied electric field (larger
y-intercept) than band II. The inset shows that the DNA
extracted from band I and subjected to base cleavage separated as
substrate 21-mers, whereas the DNA from band II was 100% cleaved to
product 10-mers.
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The preferential appearance of band I at a low MutY:DNA ratio suggested
that band I may correspond to the inactive Y2D2
state, whereas band II represented the active half-saturated dimer,
Y2D. To verify this assignment, we constructed a Ferguson
plot (58) to analyze the relationship between the electrophoretic
mobility of the two bands and the acrylamide percentage in the gel. As electrophoresis at all four acrylamide concentrations was carried out
in parallel at a constant 70 V for a fixed 40 min, migration distances
of each band provided a direct measure of its electrophoretic mobility.
We, therefore, plotted the logarithm of migration distance as a
function of acrylamide concentration (Fig. 5C), which showed linear dependences for all three bands. Additionally, the line for band
I had a steeper slope and a larger y-intercept than the line for band
II. As larger complexes are more sensitive to sieving by the gel
matrix, the steeper slope, which indicated a greater dependence on
acrylamide concentration, showed that band I corresponded to a larger
complex than band II. Conversely, the larger y-intercept, which
revealed a higher sensitivity to the applied electric field in the
absence of sieving by the gel matrix, indicated that band I
corresponded to a complex with a higher charge. These results, therefore, confirm our assignment of band I to
Y2D2, which differed from Y2D by
one DNA substrate.
Lastly, we excised the two bands from the gel and placed them
separately into 0.2 N NaOH to inactivate MutY activity. The gel slices were then pulverized and heated to 90 °C for 10 min to
extract the DNA and to alkali-cleave abasic DNA products. Denaturing PAGE separation of the cleaved DNA extracts from each band (Fig. 5C, inset) revealed that primarily unreacted
substrate 21-mers were recovered from band I, whereas the DNA extracted
from band II was completely cleaved. These results identified band II
as being the active complex, Y2D, and confirmed the lack of
excision activity by the inactive Y2D2 complex
found in band I.
 |
DISCUSSION |
A New Mechanistic Model--
Using UV cross-linking and
stopped-flow fluorimetry, we have previously established a double
base-flipping reaction sequence requiring a minimum of three distinct
structural changes between the substrate binding and excision that
provided the mechanistic basis for the efficient screening and accurate
selection of both bases in the target adenine:8-oxo-guanine mispair
(53). In the current report, we have additionally demonstrated that the
glycosylase functions as a dimer with several salient mechanistic
features. Firstly, DNA binding induced a 105-fold increase
in the dimerization constant of MutY. This strongly positive
thermodynamic linkage couples free energies of protein dimerization and
substrate binding to provide overall tight binding of a single lesion
mispair by the dimer. Secondly, the resulting half-saturated dimer,
Y2D, is the functionally active form of the enzyme.
Thirdly, binding of a second mispair by the Y2D dimer is
negatively cooperative, i.e. with weak affinity, and
allosterically inhibitory such that the resulting fully saturated
dimer, Y2D2, whereas difficult to form, lacks
all excision activity.
Porello et al. (30) used active site titrations similar to
ours to determine the active concentration of MutY in their enzyme preparations. However, because they assumed a monomeric stoichiometry, their assays utilized only three enzyme concentrations within a limited
range and a single fixed DNA concentration to determine the fractional
activity of their enzyme preparation. In contrast, we obtained our
active site titration data over an extended range of enzyme and DNA
concentrations including conditions of high DNA:MutY ratios in which we
observed a DNA-dependent decline of excision activity. This
result cannot be rationalized by any explanation based on inactive
protein in the enzyme preparation. No decline in burst amplitude would
have been observed if the 45% maximum burst amplitude had resulted
from only 45% active MutY in the enzyme preparation as the binding of
DNA by the remaining 55% inactive MutY should have no additional
effect on the measured activity unless the two populations of proteins
interacted directly. Interestingly, Porello et al. (30)
reported typical yields of 25-60% percent active enzyme, a range that
is, within statistical error, compatible with our alternative induced
dimeric model.
Recruitment of MutS by Y2D2--
The lack
of excision activity by the Y2D2 complex is
initially surprising as the inhibition of repair in response to
elevated lesion concentrations would appear to be counterproductive.
However, the accumulation of 8-oxo-guanines in DNA has been estimated
at 2.5-6 × 10
6 per guanine even under oxidative
conditions of H2O2 challenge (1, 59), placing
these lesions, on average, 0.67-1.5 × 106 base pairs
apart within the genome. The probability of forming a
Y2D2 complex between two lesions is, therefore,
low in vivo in light of the negative cooperativity of
binding the second lesion (K22 = 1.5 µM
1). Consequently, although induced
dimerization guarantees recognition, tight binding, and excision of
rare lesions by the active Y2D dimer, the negative
cooperativity of binding to its second site ensures that the repair of
a single isolated lesion is not impaired.
This negative cooperativity, therefore, imparts a sensitivity to the
proximity between pairs of lesions by the dimer, forming a mechanistic
basis for detecting catastrophic damages in the DNA. This makes
physiological sense if we take into account the fact that the target
adenine:8-oxo-guanine mispair results from the accumulation of two
separate errors: the oxidative damage of a guanine and a subsequent but
independent polymerase error that base-paired it with an adenine
instead of cytosine. Thus the localized occurrence of numerous
adenine:8-oxoguanine lesions indicates repeated failures by the base
excision enzyme MutM to repair the original oxidatively damaged
8-oxo-guanine:cytosine base pairs prior to a replication event,
suggesting that the one-base-at-a-time approach of the base excision
repair system may not be the most efficient means of removing such
clusters of damages. In this context, the bidentate binding of multiple
lesions by the Y2D2 complex may serve the dual
function of blocking further rounds of replication (or transcription)
at these damage sites while signaling the recruitment of an alternate
repair system to ensure more efficient removal of all damages.
Several lines of evidence lead us to favor the recruitment of the
methyl-directed mismatch repair system in such instances. Firstly, Gu
et al. (44) have recently identified direct protein-protein interactions between the human mismatch repair homolog, hMSH
, and the human MutY homolog, hMYH. In that report, the interaction site
was mapped to conserved amino acid residues 232-254 of hMYH, which is
homologous to amino acids 148-170 in E. coli MutY. We have
also recently obtained preliminary evidence of a similar interaction between E. coli MutY and
MutS.1 Secondly, the
recruitment of MutS by MutY in response to locally clustered lesions
makes functional sense as MutS-initiated mismatch repair typically
results in replacement of hundreds of nucleotides per repair event and
would, therefore, provide for a more efficient, single-pass means of
eliminating multiple lesions. Thirdly, Mazurek et al. (60),
in demonstrating the ability of hMSH
to recognize and initiate
repair of 8-oxo-guanine-containing mismatches, reported a hMSH
binding affinity for an adenine:8-oxoguanine mispair that is only
2-fold tighter than its affinity for a correctly base-paired homoduplex
and 17-fold weaker than the affinity for a thymine:8-oxoguanine mispair. This apparently poor selectivity by hMSH
for the
adenine:8-oxo-guanine mispair, therefore, further suggests a
complementary role for MutY in facilitating the recognition of the
adenine:8-oxo-guanine mismatch by MutS.
Contextual Recognition by Y2D and
Y2D2--
Because an adenine:8-oxo-guanine
mismatch arises from a polymerase error (60), the accuracy of repair by
an adenine glycosylase such as MutY depends on its ability to recognize
the template versus nascent strand context of the adenine.
Repair without regard to contextual recognition is mutagenic when an
adenine in the template strand becomes indiscriminately removed. For
this reason, the recruitment of a nascent strand-specific repair system
makes functional sense. However, recruitment alone does not guarantee accurate repair, as MutY excision of adenines in the template strand
followed by MutS-initiated repair of the nascent strand would result in
the detrimental loss of genetic information from both strands of the
DNA. Thus the utilization of the strand specificity of mismatch repair
enzymes does not obviate the need for a mechanism of
context-dependent inhibition of MutY activity.
We hypothesize that the allosteric inhibition of excision activity
observed in Y2D2 coupled with the negative
cooperativity of binding the second lesion may together provide such a
mechanism. As noted , the negative cooperativity of binding confers a
sensitivity toward the proximity of lesions. Additionally, direct
random oxidative damages to the DNA genome would position 8-oxo-guanine
residues, on average, a million base pairs apart. Consequently,
adenines misincorporated into the nascent strand opposite template
strand 8-oxoguanines are likely to be found only in isolation. MutY
recognition and binding at these mismatches would, therefore, lead to
formation of the active Y2D dimer, resulting in the
appropriate removal of the nascent strand adenine.
On the other hand, mismatches with the adenine in the template strand
arise from the misinsertion of 8-oxo-guanosine-monophosphates by
polymerase into the nascent strand. As oxidatively damaged dGTPs are
enzymatically eliminated from the dGTP pool by MutT, the accumulation
of 8-oxo-guanosine-triphosphates available for misincorporation
following oxidative challenge would be transient. Replication during
such a transient rise in 8-oxo-guanosine-triphosphate concentration
would, therefore, lead to the appearance of 8-oxo-guanines in the
nascent DNA strand in localized patches. MutY binding within such a
patch would lead to formation of the allosterically inhibited Y2D2 complex due to the high local
concentration of mismatches. This negative regulation of excision
activity by proximity of adenine:8-oxo-guanine mismatches, therefore,
effectively constitutes contextual recognition and selection of nascent
versus template strand adenines.