|
|
|
|||||||
|
|||||||||
J. Biol. Chem., Vol. 278, Issue 37, 35584-35591, September 12, 2003
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
The Phosphorylation Domain of the 32-kDa Subunit of Replication Protein A (RPA) Modulates RPA-DNA Interactions
EVIDENCE FOR AN INTERSUBUNIT INTERACTION*
||
From the
Department of Biochemistry, University of
Iowa College of Medicine, Iowa City, Iowa 52242-1109 and
¶Pacific Northwest National Laboratory, Richland,
Washington 99352
Received for publication, May 22, 2003
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
Replication protein A (RPA) is a heterotrimeric (subunits of 70, 32, and 14 kDa) single-stranded DNA-binding protein that is required for DNA replication, recombination, and repair. The 40-residue N-terminal domain of the 32-kDa subunit of RPA (RPA32) becomes phosphorylated during S-phase and after DNA damage. Recently it has been shown that phosphorylation or the addition of negative charges to this N-terminal phosphorylation domain modulates RPA-protein interactions and increases cell sensitivity to DNA damage. We found that addition of multiple negative charges to the N-terminal phosphorylation domain also caused a significant decrease in the ability of a mutant form of RPA to destabilize double-stranded (ds) DNA. Kinetic studies suggested that the addition of negative charges to the N-terminal phosphorylation domain caused defects in both complex formation (nucleation) and subsequent destabilization of dsDNA by RPA. We conclude that the N-terminal phosphorylation domain modulates RPA interactions with dsDNA. Similar changes in DNA interactions were observed with a mutant form of RPA in which the N-terminal domain of the 70-kDa subunit was deleted. This suggested a functional link between the N-terminal domains of the 70- and 32-kDa subunits of RPA. NMR experiments provided evidence for a direct interaction between the N-terminal domain of the 70-kDa subunit and the negatively charged N-terminal phosphorylation domain of RPA32. These findings suggest that phosphorylation causes a conformational change in the RPA complex that regulates RPA function.
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
Replication protein A (RPA)1 is a heterotrimeric (70-, 32-, and 14-kDa subunit) single-stranded DNA-binding protein that is required for DNA replication, recombination, and repair (1, 2). RPA was first identified as a factor necessary for SV40 replication (3–5). Subsequently homologues have been identified in all eukaryotic cells examined (1, 6). Human RPA binds single-stranded (ss) DNA with high affinity (7) but low specificity and cooperativity (8, 9). RPA has also been shown to specifically interact with multiple proteins involved in DNA metabolism (1, 2).
RPA is composed of three subunits with multiple structurally related,
functional domains (Fig. 1).
The primary ssDNA binding activity is localized to the central region of the
70-kDa subunit (RPA70) (10).
This region is composed of two structural domains called DNA binding domains
(DBDs) A and B (Fig.
1A) (11,
12). Both domains form an
oligosaccharide/oligonucleotide binding fold (OB-fold) that contains five
-strands arranged into a Greek-key -barrel capped by an
-helix between the third and fourth strands
(13). The N- and C-terminal
domains of RPA70 are also composed of OB-folds. Structural studies by NMR have
shown that the N-terminal 168 residues of RPA70 form two distinct domains
(14). The first 108 residues
form an OB-fold, defined as DBD F based on structural homology and DNA binding
ability (15). The remaining 60
residues are a flexible linker to DNA binding domains A and B in RPA70. In
addition to binding DNA, DBD F has also been shown to interact with various
proteins involved in DNA metabolism
(1). The C-terminal region of
RPA70, DBD C, exhibits weak DNA binding and is necessary for heterotrimeric
complex formation (16,
17). The OB-fold of DBD C has
two unique characteristics, a three helical cap and a zinc ribbon motif
(18). Two additional OB-fold
domains are located in the 32- and 14-kDa subunits. The 32-kDa subunit has a
central OB-fold domain, DBD D, that has weak DNA binding, flanked by the
N-terminal phosphorylation domain and a C-terminal winged helix domain
(19). The latter has been
shown to interact with proteins involved in DNA repair
(19). The 14-kDa subunit
(RPA14) is also composed of an OB-fold that has been defined as DBD E although
there is currently no evidence that this domain interacts with DNA. RPA14 is
necessary for RPA complex formation
(1,
2).
|
In addition to binding ssDNA and proteins involved in DNA metabolism RPA can destabilize double-stranded DNA (dsDNA) (20, 21). Although RPA binds dsDNA with low affinity, it is able to promote helix destabilization by stabilizing single-strand regions in dsDNA (20, 22). RPA destabilization of duplex DNA is functionally distinct from that of helicases, because ATP and Mg2+ are not required (20). Linearized SV40 dsDNA is preferentially denatured by RPA at internal regions rich in A-T pairs and at both ends (21). Although other ssDNA-binding proteins are capable of destabilizing dsDNA, RPA is more efficient than the ssDNA-binding proteins tested: Escherichia coli SSB, herpes simplex virus ICP8 protein, bacteriophage T4 gp32, and bacteriophage T7 gp2.5 (20, 21, 23). Association and helix destabilization reactions of RPA with duplex DNA and duplex DNA containing a central eight-nt ss bubble indicated that the denaturation of dsDNA occurs in at least two steps (16, 23). The two steps are (i) the formation of stable RPA/ssDNA with a small region of ssDNA (nucleation), followed by (ii) efficient strand separation. The destabilizing activity of RPA may play a role in the initiation of replication, as well as the denaturation of damaged DNA, in nucleotide excision repair (16).
Helix destabilization by RPA requires the core ssDNA binding region (DBDs A
and B) and is stimulated by the presence of DBD F
(22). DBD F contains a basic
cleft in the -sheet-rich OB-fold
(Fig. 2) and interacts with the
acidic domains of the transcriptional activators Gal-4, VP16, and p53
(24,
25). NMR data indicate that
residues within the basic cleft of DBD F interact with ssDNA
(15). A direct competition
between ssDNA and the acidic activation domains of Gal-4, p53, and VP16 has
been proposed to regulate DBD F function
(15).
|
RPA is phosphorylated in a cell cycle-dependent manner (during S and
G2) (26,
27) and in response to DNA
damage (28,
29). Hyperphosphorylation
occurs in response to DNA-damaging agents, such as UV or ionizing radiation
(30,
31), hydroxyurea or
camptothecin treatment (31,
32), and cellular apoptosis
(33). Phosphorylation occurs
primarily within the N-terminal 33 residues of RPA32
(28,
34). Proteolytic and NMR
analyses indicated that this domain, termed the N-terminal phosphorylation
domain, exists in an extended, flexible conformation
(19,
35). Phosphorylation or
mutations that add multiple negative charges to the N-terminal phosphorylation
domain of RPA32 cause altered interactions with p53, T antigen, and DNA
polymerase /DNA primase
(37).2
Thus the N-terminal phosphorylation modulates RPA activity.
The effect of phosphorylation on RPA interactions with DNA is not well understood. Independent studies have reported that RPA phosphorylation causes stimulation of RPA helix destabilization activity (38), dramatic decreases in DNA binding (39), and dissociation of the RPA complex (40). Although these reports have not been reproduced in subsequent studies (41, 42, 47),3 it seems likely that phosphorylation affects DNA interactions. To test this hypothesis, we examined the interactions between ssDNA or dsDNA and mutant forms of RPA with modifications in the N-terminal phosphorylation domain of RPA32. We found that whereas all forms of RPA examined had equal affinity for short oligonucleotides, a mutant form containing serine/threonine to aspartic acid mutations in the N-terminal phosphorylation domain of RPA32 had a significant defect in helix destabilization activity. A similar defect was observed with a mutant form of RPA that had the N-terminal domain of RPA70 (DBD F) deleted, suggesting a functional linkage between the N-terminal domains of RPA70 and RPA32. Helix destabilization assays with different substrates and time course experiments demonstrated that the observed decreases in helix destabilization activity were caused by deficiencies in both the nucleation and strand separation steps. Finally, we present evidence of a direct interaction between a negatively charged N-terminal phosphorylation domain of RPA32 and the basic cleft of DBD F of RPA70 using heteronuclear single quantum correlation NMR (HSQC). These findings suggest that there is a conformational change in RPA upon phosphorylation involving an intersubunit interaction between DBD F in RPA70 and the N-terminal phosphorylation domain of RPA32. This conformational change appears to regulate RPA function through modulation of RPA interactions with both proteins and DNA.
|
EXPERIMENTAL PROCEDURES |
|---|
|
TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
Materials—[
-32P]ATP (6000 µCi/mmol) was
purchased from Amersham Biosciences. The 35-residue peptides, RPA32-WT and
RPA32-Asp, corresponding to the wild-type or aspartic acid-substituted
N-terminal phosphorylation domain of RPA32, respectively
(Fig. 1), were from the Howard
Hughes Medical Institute Biopolymer/Keck Foundation Biotechnology Resource
Laboratory at Yale University. Biotinylated dT30 for surface plasmon resonance
was purchased through the University of Iowa DNA Core (Sigma Genosys). Buffers—HI buffer contained 30 mM Hepes (diluted from 1 M stock at pH 7.8), 1 mM dithiothreitol, 0.25 mM EDTA, 0.5% (w/v) inositol, and 0.01% (v/v) Nonidet P-40. 1x Tris borate/EDTA (1x TBE) gel buffer contained 89 mM Tris, 89 mM boric acid, and 2 mM EDTA.
DNA Templates and Manipulation—A 40-bp dsDNA fragment of the
early palindromic SV40 origin was made for helix destabilizing studies by
labeling an oligonucleotide (SV40 top,
5'-CTCCAAAAAAGCCTCCTCACTACTTCTGGAATAGCTCAGA-3') with
[-32P]ATP by T4 polynucleotide kinase (New England
Biosystems) following the manufacturer's recommendations. The labeled DNA was
separated from free ATP with a P-30 Tris chromatography column (Bio-Rad)
following the manufacturer's specifications and annealed to equal molar
concentrations of the complementary sequence (SV40 bottom,
5'-TCTGAGCTATTCCAGAAGTAGTGAGGAGGCTTTTTTGGAG-3') to make a 40-nt
double-stranded DNA substrate for helix destabilization assays. A
double-stranded 40-nt substrate containing an eight-nt bubble was made by
annealing SV40 top to SV40 bottomB
(5'-TCTGAGCTATTCCAGAGACGACAGGGAGGCTTTTTTGGAG-3'). The
underlined portion indicates a region of non-complementary bases. Annealing
reactions (10 mM Tris-HCl, pH 7.5, 20 mM
MgCl2, and 50 mM NaCl) were placed in a PCR machine at
95 °C for 3 min ramped to 22 °C by a rate of 0.01 °C/s with a hold
temperature of 4 °C. Annealing was monitored by 15% polyacrylamide gel
electrophoresis (1x TBE). In all experiments, greater than 95% of
labeled DNA was in double-stranded form.
Proteins—Wild-type RPA was purified as described
(43). A form of human RPA that
could not be phosphorylated, RPA·32 S4, 8, 11–13, 23, 29, 33,
39A,T21A (RPA·32Ala10), and one that mimics the constitutively
phosphorylated form, RPA·32 S8, 11–13, 23, 29, 33D, T21D
(RPA·32Asp8), were created and purified as described.2 The
deletions of DBD F (RPA70
1–168 (RPA·70N168))
(17), DBD C
(RPA70442–616 (RPA70C442))
(Fig. 1), and the
phosphorylation domain (RPA·32 1–33
(RPA·3233)) (34)
were purified as described previously.
The 15N-labeled RPA7169–616 (DBD F) was made by
transforming vector p11d-tRPA70 169–616
(44) into DE3 cells. The cells
were grown in 5 g/liter Celtone®-N medium (Martek Biosciences Corp.)
supplemented with 1 g/liter glucose, 1.8 g/liter K2HPO4,
1.4 g/liter KH2PO4, 1 g/liter
MgSO4·7H2O, and 11 mg/liter
CaCl2·2H2O
(14). The protein was purified
by standard RPA purification procedures as described previously
(43).
Helix Destabilization Assay—The proteins were dialyzed against HI-30 (30 mM KCl). The extent of dialysis was monitored by solution conductivity. Helix destabilization assays were carried out as described previously (22). Briefly, 15-µl reactions with HI buffer containing 30 mM KCl, 2 fmol radiolabeled DNA, increasing amounts of RPA (0–3160 fmol), and 50 µg/ml bovine serum albumin were incubated for 20 min at 25 °C. Reactions were terminated by adding SDS to a final concentration of 0.2% (to disrupt RPA·DNA complexes), followed by 4% glycerol and 0.01% bromphenol blue. The reaction products were separated on a 15% polyacrylamide gel (1x TBE) at 200 volts for 2.5 h. The gels were dried on Whatman 3-mm chromatography paper, and the radioactive bands were visualized by autoradiography. The radioactivity in each band was quantified using a Packard Instant Imager. The amount of dsDNA (%) was plotted against the concentration of RPA and then analyzed by non-linear least squares fitting to a Langmuir binding equation using Nonlin (22, 45). Although the destabilization of DNA by RPA is not a simple bimolecular binding reaction, the Langmuir binding equation was used to precisely determine the midpoint of the transition between dsDNA and ssDNA. The midpoint (given with units of M–1) of the fitted curves was used as a value for comparing the activities of RPA forms in this assay.
Time Course of Helix Destabilization—Saturating amounts of
RPA (562 fmol for wild-type RPA, RPA·32Ala10, and
RPA·3233 and 3160 fmol for RPA·32Asp8,
RPA·70N168, and RPA70C442) were combined with HI-30 to
create a 120-µl reaction mixture. The DNA mixture consisted of 60 µl
containing 50 µg/ml bovine serum albumin and 120 fmol of radiolabeled
double-stranded 40-nt substrate or a double-stranded 40-nt substrate with an
eight-nt bubble. The reaction mixture was incubated with the DNA mixture.
Samples (15 µl) were removed at the indicated time points and immediately
placed in microcentrifuge tubes with SDS (0.2% final concentration) to
dissociate RPA·DNA complexes. The zero-min time point was made by
adding the saturating amount of RPA to a mixture containing 2 fmol of DNA, 50
µg/ml bovine serum albumin, and 0.2% SDS in HI-30. The products were
separated on a 15% polyacrylamide gel (1x TBE). The amount of ssDNA at
each time point was quantitated by the Packard Instant Imager and plotted as a
function of time.
RPA70169–616 Interactions with
Peptides—The NMR data collection was performed as described
previously (14). Lyophilized
RPA32-WT peptide and RPA32-Asp peptide were dissolved in the same buffer as
[15N]RPA70169–616. Two-dimensional
1H-15N correlation maps were acquired on two identical
[15N]RPA70169–616 samples after addition
stoichiometric and superstoichiometric amounts of either RPA32-WT or RPA32-Asp
peptide to each RPA sample. Final spectra contained nearly a 20-fold excess of
peptide relative to protein, and spectral changes were minimal in final
spectra with RPA32-Asp, indicating an approach to saturation of peptide
binding. Spectra were acquired at 750 MHz on a Varian Inova console, using
water flip-back HSQC. We used a spectral width of 3000 Hz, 128 increments in
the indirect dimension. Temperature was 25 °C.
Surface Plasmon Resonance—Interaction of RPA with ssDNA was monitored using a surface plasmon resonance biosensor instrument, BIAcore 3000. The streptavidin biosensor surface was prepared by manually injecting 5'-biotinylated dT30 diluted to 0.5 nM in 10 mM sodium acetate, pH 4.8, and 1.0 M NaCl into the desired flow cell. Proteins were diluted in HBS-EP buffer (10 mM HEPES, pH 7.4, 150 mM NaCl, and 0.005% polysorbate-20) from BIAcore brought to 1 mM dithiothreitol. Protein (20 µl of 4 nM solution) was loaded by the kinject option with the dissociation time of 500 s and a flow rate of 10 µl/min. Each experiment was repeated at least twice. Data were analyzed with the BIA Evaluation program and fit to a bimolecular Langmuir binding curve.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
Role of N-terminal Phosphorylation Domain in RPA-DNA Interactions—RPA is composed of a high affinity DNA binding core (DBDs A and B) and three additional domains that can interact with DNA (DBDs F, C, and D; see Fig. 1). Mutations that decrease ssDNA binding also show defects in double-stranded DNA binding and helix destabilization activities (see Table I and Ref. 22). There have been several reports of phosphorylation modulating DNA interactions, but no consistent models have emerged for how RPA phosphorylation affects DNA interactions (see the Introduction). Our initial studies with RPA phosphorylated with CDK2 family kinases or DNA-dependent protein kinase (DNA-PK) suggested that phosphorylation had minimal effects on RPA binding to single-stranded oligonucleotides.4 To extend these observations we compared the ssDNA binding activity of various mutant forms of RPA including N-terminal phosphorylation domain mutants (Fig. 1). We have shown that addition of negative charges to the N-terminal phosphorylation domain of RPA32 modulates RPA-protein interactions and that a mutant form, which has aspartic acid substitutions in the N-terminal phosphorylation domain of RPA32 (RPA·32Asp), has properties similar to those of phosphorylated RPA.2 Binding of the mutant forms of RPA to a 30-residue oligonucleotide was determined by surface plasmon resonance. The apparent binding constants for the phosphorylation domain mutants were the same as the wild-type RPA complex (Table I). Deletion or introduction of multiple point mutations of the N-terminal phosphorylation domain had no effect on the binding constant with oligo(dT)30. This is consistent with previous deletion analysis demonstrating that the core DNA binding domain of RPA70 (DBDs A and B) is necessary and sufficient for high affinity binding to oligonucleotides (10, 22).
|
Single-stranded oligonucleotides are the most simple form of DNA to which
RPA can bind. In the cell most single-stranded DNA is part of partially duplex
structures. Therefore, we next examined the ability of the phosphorylation
domain mutants to destabilize a short DNA duplex. In this assay increasing
concentrations of RPA are incubated with radiolabeled duplex DNA, and the
relative amount of dsDNA and ssDNA is determined at each protein
concentration. A radiolabeled 40-nt oligonucleotide from the SV40 early
palindromic region annealed to its complementary strand was used as the
substrate. This region of DNA has been shown to require only T antigen and RPA
for denaturation during the early steps of SV40 initiation
(46). Protein·DNA
complexes were dissociated with SDS, and the products were separated on a 15%
polyacrylamide gel. The amount of ssDNA and dsDNA in each reaction was
quantitated, and the midpoint of the transition was determined. Deletion of
the N-terminal phosphorylation domain (RPA·3233) or introduction
of multiple alanine mutations (RPA·32Ala10) had no effect on helix
destabilization (Fig.
3A). In contrast, introduction of multiple aspartic
acidic residues in the N-terminal phosphorylation domain caused a 4-fold
increase in the amount of protein needed to destabilized 50% of the DNA
template (Fig. 3A).
This decrease in helix destabilization activity is unusual. Other forms of RPA
have been shown to have decreased helix destabilization activity. For example,
forms of RPA with mutations or deletions in the core DNA binding domain (DBDs
A and B) or in the conserved zinc finger domain (DBD C) have decreased
oligonucleotide binding and helix destabilization activity
(22). However, in all but one
case, these mutants also had decreased affinity for oligonucleotides
(Table I; see also Ref.
22). The one exception to this
general rule is RPA·70N168, which has wild-type affinity for
oligonucleotides but also shows an activity in helix destabilization
one-quarter that of wild-type RPA (see
Table I and
Fig. 3A). We conclude
that both RPA·32Asp8 and RPA·70N168 have altered
interactions with double-stranded or partially duplex DNA substrates that are
not caused by differences in ssDNA binding activity.
|
Helix-destabilizing Activities of RPA with an Eight-nt Bubble
Substrate—Helix destabilization by RPA appears to be a multi-step
process in which there is first the formation of an RPA·DNA complex at
a small region of ssDNA (nucleation) followed by efficient strand separation
coupled to binding of additional RPA molecules. A pseudo-origin substrate with
an eight-nt bubble in the center has been used to investigate the steps that
are effected in these RPA mutants
(46). The rate-limiting step
is hypothesized to be the generation of a small region of ssDNA, because RPA
binds rapidly and with high affinity to ssDNA. The decreased helix
destabilization of RPA·32Asp8 and RPA·70N168 could be
because of deficiencies in either nucleation or subsequent strand separation
steps. If the decreased helix destabilization activity of RPA·32Asp8
and RPA·70N168 could be rescued by a substrate with an eight-nt
bubble, it would suggest a defect in the nucleation step.
When helix destabilization was examined using an eight-nt bubble substrate,
two levels of helix destabilization activity were observed. Both
RPA·3233 and RPA·32Ala10 had activity similar to
wild-type RPA, although RPA·32Ala10 consistently had a slightly lower
activity (Fig. 3B). In
contrast, RPA·32Asp8 and RPA·70N168 had one-half the
destabilization activity with the eight-nt bubble substrate compared with
wild-type RPA (Fig.
3B). Thus, the presence of an eight-nt bubble partially
restored helix destabilization activity observed for RPA·32Asp8 and
RPA·70N168 with dsDNA. This indicates that these forms of RPA
have defects in both nucleation and strand separations steps of unwinding.
Kinetics of Helix Destabilization—To investigate the kinetics of the nucleation event, a time course of helix destabilization was examined. Saturating amounts of RPA were incubated under helix destabilization conditions, and 15-µl samples were removed at the indicated time points. The amount of ssDNA present at each time point was quantitated and plotted as a function of time.
Wild-type RPA, RPA·32Ala10, and RPA·3233 all completed
destabilization of both DNA substrates within 5 min
(Fig. 4, A and
B). The RPA·32Asp8 mutant took 15 min longer to
complete destabilization of the dsDNA substrate than wild-type RPA
(Fig. 4A) but was
still able to reach the same final level of destabilized DNA. In contrast,
RPA·32Asp8 required only 1 min for destabilization of the eight-nt
bubble substrate. This indicates that RPA·32Asp8 has a kinetic defect
that affects the generation or recognition of a small region of ssDNA.
RPA·70N168 shows similar activity to RPA·32Asp8 helix
destabilization is slower than wild-type RPA, and the time required to
destabilize the eight-nt bubble substrate (10 min) is much faster than with
the dsDNA substrate (60 min) (data not shown). We conclude that
RPA·32Asp8 and RPA·70N168 mutants have defects in both
the nucleation step and in the helix destabilization steps of unwinding. These
studies cannot resolve fast kinetics in the <10 s time scale.
|
A striking result of these studies is that removing DBD F
(RPA·70N168) or the addition of negative charges to the
phosphorylation domain of RPA32 (RPA·32Asp8) caused similar effects on
RPA interactions with complex DNA substrates. Furthermore, the absence of an
altered affinity to oligonucleotides and altered kinetics of helix
destabilization in these mutants indicated that these mutations are acting
through a mechanism distinct from those that affect ssDNA binding. These data
suggest a link between DBD F and the N-terminal phosphorylation domain of
RPA32. We hypothesized that an interaction between the negatively charged
phosphorylation domain and the basic cleft in DBD F may exist that prevents
the N-terminal region of RPA70 from contacting DNA and contributing to
destabilization (Fig. 5).
|
Residue-specific Interactions Monitored by HSQC NMR—To test
the intersubunit interaction hypothesis, peptides corresponding to the
non-phosphorylated (RPA32-WT) and the negatively charged (RPA32-Asp)
phosphorylation domain (Fig. 1)
were individually incubated with 15N-labeled
RPA70169–616 (DBD F; the first 168 residues of RPA70). HSQC NMR
was used to monitor changes in the backbone of RPA70169–616 at
five different concentrations of peptide. The final difference in
1H and 15N chemical shift upon addition of RPA32-WT or
RPA32-Asp peptide was determined and plotted against residue number
(Fig. 6, A and
B). Only the addition of RPA32-Asp peptide caused a
significant shift in either the backbone nitrogen or the proton spectra of
RPA70169–616. A summary of the residues perturbed by the
RPA32-Asp peptide is shown in Fig.
6C. The residues perturbed by the RPA32-Asp peptide
correlated well with the area of the basic cleft and residues that interact
with ssDNA (15). We conclude
that the addition of negative charges to the phosphorylation domain
(RPA·32Asp8) can cause it to interact with DBD F. This suggests that
phosphorylation can cause a conformational change in RPA in which there are
increased interactions between the N-terminal domains of RPA70 and RPA32.
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
Several lines of evidence now indicate that the phosphorylation domain of RPA32 modulates RPA activity. (i) RPA is phosphorylated in a cell cycle-dependent manner (26, 27). (ii) Cell extracts exposed to UV radiation that contain phosphorylated RPA are deficient in SV40 replication. Replication activity is restored with the addition of purified non-phosphorylated RPA (30). (iii) RPA phosphorylation modulates RPA-protein interactions. The addition of negative charges either by mutation or phosphorylation of the phosphorylation domain causes decreased interactions with T antigen and DNA polymerase
. The negatively charged phosphorylation domain also enhances RPA-p53
interactions (37).2
(iv) A form of RPA lacking the phosphorylation domain, RPA·3233,
was not able to support SV40 DNA replication after pretreatment with DNA-PK.
Without pretreatment of DNA-PK, RPA·3233 is able to support SV40
replication. This suggests that under some conditions the phosphorylation
domain may be required to overcome inhibitory effects of DNA-PK on components
of the replication machinery
(34). (v) Saccharomyces
cerevisiae with the serine/threonine to aspartic acid mutations in the
phosphorylation domain as the only functional copy of RPA32 is hypersensitive
to methyl methane sulfonate and hydroxyurea treatment indicating that DNA
damage is not being repaired.2 (vi) Here we show that a negatively
charged phosphorylation domain causes decreased helix destabilization,
suggesting that RPA phosphorylation modulates RPA-DNA interactions. Recently
Oakley et al. (37)
have shown similar changes with dsDNA interactions using RPA phosphorylated
in vivo. There is one report in the literature that cdc2-phosphorylated calf thymus RPA stimulates helix destabilization (38). These findings are not consistent with those presented here or recent studies by Oakley et al. (37). They are also not consistent with a decrease in helix destabilization activity observed with RPA phosphorylated by DNA-PK (47). The disparity remains unresolved but could be because of differences in phosphorylation sites or differences in assay conditions.
Helix Destabilization—RPA efficiently destabilizes dsDNA
(20,
21). The destabilizing
reaction has been functionally separated into two steps. The first step,
termed nucleation, is the generation of a small region of ssDNA, either by DNA
breathing or RPA association, culminating with the stable binding of RPA. The
second step is efficient strand separation. As the first step of helix
destabilization includes the stable binding of RPA to a small region of ssDNA,
RPA mutants that effect ssDNA binding also effect destabilization
(Table I)
(22); there is a direct
correlation between ssDNA binding and helix destabilization. There are two
exceptions to this generalization. RPA·70N168 has wild-type
ssDNA binding activity but decreased dsDNA binding and helix destabilization
(22). In addition, the
phosphorylation domain mutant, RPA·32Asp8, also showed wild-type ssDNA
binding but decreased helix destabilization activity
(Table I). The finding that
separate mutations affecting either the basic cleft of DBD F
(RPA·70N168) or the RPA32 phosphorylation domain
(RPA·32Asp8) do not affect ssDNA binding but do have similar effects on
helix destabilization suggests a link between these two domains. We
hypothesized a direct interaction between the phosphorylation domain and the
basic cleft. This interaction may prevent the N-terminal region of RPA70 from
contacting the DNA and contributing to destabilization.
A direct interaction between the basic cleft of RPA70 and the negatively charged phosphorylation domain was observed by HSQC NMR experiments. A peptide corresponding to the negatively charged (RPA32-Asp) phosphorylation domain was found to interact with DBD F whereas a peptide corresponding to the native, unphosphorylated sequence did not (Fig. 6). Furthermore, the residues perturbed by the RPA32-Asp peptide were predominantly in the basic cleft and correlated well with residues that interact with ssDNA (15). These data support the hypothesis that the negatively charged phosphorylation domain of RPA32 interacts specifically with DBD F in RPA70.
Model for RPA Intersubunit Interaction—We have presented
evidence for a specific intersubunit interaction between a negatively charged
phosphorylation domain of RPA32 and the basic cleft in DBD F. The negative
charges can be a result of phosphorylation or aspartic acid mutations as in
RPA·32Asp8. Increased interactions between the phosphorylation domain
and DBD F are likely to cause a global conformational change in RPA. These
intersubunit interactions also probably keep DBD F from interacting with DNA
and contributing to dsDNA destabilization
(Fig. 4). This model provides a
mechanism by which the phosphorylation domain can modulate RPA activity. This
conformational switch is consistent with the finding that the phosphorylation
domain affects RPA-protein interactions indirectly by regulating the contacts
of DBD F with T antigen and DNA polymerase
(37).2 The proposed
interactions are also consistent with recent evidence that DBD F rotates
independently of DBD allowing domain F to switch between conformations
(15).
The Basic Cleft and Phosphorylation Domain—There are six residues, five arginines and one lysine, that form the basic cleft in DBD F of human RPA (Fig. 2). The charge of the basic cleft in human DBD F has been found to be conserved in other RPA homologues, with the exception of two eukaryotes that are missing the entire DBD F (Crithidia fasciculata (trypanosome), 23.9% identity; Pyrococcus furiosus (archeae bacteria), 11.8% identity). The basic cleft is also not conserved in Cryptospordium parvum, a protozoal parasite. We conclude that the basic cleft is evolutionarily conserved and present in all metazoans and yeast that have been examined.
Several mutations within DBD F have been characterized in yeast. Many of those with UV and methyl methane sulfonate hypersensitivity phenotypes map to regions within the basic cleft (48). This sensitivity to methyl methane sulfonate and UV is similar to the phenotype observed in RPA32 mutants with negatively charged N-terminal phosphorylation domains of RPA32.2 Decreased rates of recombination, double-strand break repair and HO-gene conversion are also associated with mutations near the basic cleft (48–51). In addition, some DBD F mutations are temperature-sensitive (49) and have defective DNA damage checkpoints (52, 53). Recently, S. cerevisiae RPA trimer containing the rfa1 mutant t11 (K45Q) has been expressed and found to have similar ssDNA binding affinity to wild-type S. cerevisiae RPA but impaired Rad51 displacement from ssDNA (54). The genetic data demonstrates that DBD F is playing a role in DNA metabolism, particularly DNA repair. The studies presented here show that mutations that alter the structure and charge distribution of the basic cleft or the addition of negative charges to the phosphorylation domain have similar effects on RPA interactions with dsDNA. These mutations may prevent the basic cleft from participating in RPA-protein or RPA-DNA interactions that are needed for an appropriate cellular response to DNA damage.
The serines and threonines in the phosphorylation domain of RPA32 are conserved to varying degrees in eukaryotes. The total number ranges from eight to 12 for most eukaryotes examined (data not shown). The two RPA homologues that have less than eight serine and threonine residues have the remaining serines and threonines in conserved sites. The major exception to the high level of conservation is Schizosaccharomyces pombe, which has only three serine/threonines at non-conserved locations in its N-terminal phosphorylation domain. This suggests that S. pombe uses a different mechanism of regulation not involving a basic cleft. A homolog of RPA32 called RPA4 was found to be expressed in quiescent cells (55). RPA4 lacks four serines (S12I, S23L, S29T, and S33K) and one threonine (T21D) found in RPA32. The absence of the serines in RPA4 may reflect the proliferation status of the cell types it is expressed in or that it is part of complexes that are regulated differently.
Implications for RPA32 Phosphorylation—The effect RPA phosphorylation has on DNA repair pathways is not known. Phosphorylated and non-phosphorylated forms of RPA are equally active in in vitro SV40 DNA replication and nucleotide excision repair assays (56).2 However, deletion of the phosphorylation domain and part or all of DBD F decreases the ability of RPA to support nucleotide excision repair (57). The phosphorylation state of RPA has no effect on RPA-xeroderma pigmentosum complementation group A (XPA) interactions (57).5 XPA has been shown to inhibit RPA-dependent separation of a 24-mer from M13 (58). This may be an example of a coordinated effort of phosphorylated RPA and XPA to limit the opening of DNA around the lesion site. It remains possible that phosphorylation of RPA may modulate RPA function in nucleotide excision repair in vivo. In addition, the effect of RPA phosphorylation on recombination and base excision repair remains to be determined. We have shown that negative charges in the N-terminal phosphorylation domain can modulate RPA-DNA interactions and present data that these negative charges promote an intersubunit interaction with the basic cleft of the N-terminal region of RPA70. This conformational change can modulate RPA70-mediated interactions with DNA and proteins and cause modulation of RPA activity.
|
FOOTNOTES |
|---|
* This work was supported in part by NIGMS, National Institutes of Health Grant GM4471 (to M. S. W.) and in part by the Office of Science (Biological and Environmental Research), United States Department of Energy Contract DE-AC06-76RL01830 (to D. F. L.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Present address: Lexicon, Inc., Houston, TX.
|| To whom correspondence should be addressed: Dept. of Biochemistry, University of Iowa College of Medicine, 51 Newton Rd., Iowa City, IA 52242-1109. Tel.: 319-335-6784; Fax: 319-384-4770; E-mail: marc-wold{at}uiowa.edu.
1 The abbreviations used are: RPA, replication protein A; ss,
single-stranded; ds, double-stranded; HSQC, heteronuclear single quantum
correlation; nt, nucleotide; DBD, DNA binding domain; OB-fold, oligonucleotide
binding fold; WT, wild-type; TBE, Tris borate/EDTA; DNA-PK, DNA-dependent
protein kinase.
2 K. A. Braun, A. P. Walther, Y. Lao, L. A. Henricksen, S. K. Binz, C. G.
Lee, T. Carter, S. P. Lees-Miller, and M. S. Wold, manuscript in
preparation.
3 Y. Lao and M. S. Wold, unpublished data.
4 M. S. Wold, unpublished data.
5 Y. Lao and M. S. Wold, unpublished data.
|
ACKNOWLEDGMENTS |
|---|
We thank the members of the Wold laboratory for scientific discussions and critical reading of this manuscript. We thank Kitty Dixon for communication of data prior to publication. We also thank the University of Iowa DNA Core Facility for oligonucleotide synthesis and DNA sequencing.
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
- Wold, M. S. (1997) Annu. Rev. Biochem. 66, 61–92[CrossRef][Medline] [Order article via Infotrieve]
- Iftode, C., Daniely, Y., and Borowiec, J. A. (1999) CRC Crit. Rev. Biochem. 34, 141–180[Medline] [Order article via Infotrieve]
- Wobbe, C. R., Weissbach, L., Borowiec, J. A., Dean, F. B.,
Murakami, Y., Bullock, P., and Hurwitz, J. (1987)
Proc. Natl. Acad. Sci. U. S. A.
84,
1834–1838
[Abstract/Free Full Text] - Wold, M. S., and Kelly, T. (1988) Proc.
Natl. Acad. Sci. U. S. A. 85,
2523–2527
[Abstract/Free Full Text] - Fairman, M. P., and Stillman, B. (1988) EMBO J. 7, 1211–1218[Medline] [Order article via Infotrieve]
- Ishibashi, T., Kimura, S., Furukawa, T., Hatanaka, M., Hashimoto, J., and Sakaguchia, K. (2001) Gene 272, 335–343[CrossRef][Medline] [Order article via Infotrieve]
- Wold, M. S., Weinberg, D. H., Virshup, D. M., Li, J. J., and Kelly,
T. J. (1989) J. Biol. Chem.
264,
2801–2809
[Abstract/Free Full Text] - Kim, C., Paulus, B. F., and Wold, M. S. (1994) Biochemistry 33, 14197–14206[CrossRef][Medline] [Order article via Infotrieve]
- Kim, C., and Wold, M. S. (1995) Biochemistry 34, 2058–2064[CrossRef][Medline] [Order article via Infotrieve]
- Walther, A. P., Gomes, X. V., Lee, C. G., and Wold, M. S. (1999) Biochemistry 38, 3963–3973[CrossRef][Medline] [Order article via Infotrieve]
- Philipova, D., Mullen, J. R., Maniar, H. S., Gu, C., and Brill, S.
J. (1996) Genes Dev.
10,
2222–2233
[Abstract/Free Full Text] - Bochkarev, A., Pfuetzner, R. A., Edwards, A. M., and Frappier, L. (1997) Nature 385, 176–181[CrossRef][Medline] [Order article via Infotrieve]
- Murzin, A. G. (1993) EMBO J. 12, 861–867[Medline] [Order article via Infotrieve]
- Jacobs, D. M., Lipton, A. S., Isern, N. G., Daughdrill, G. W., Lowry, D. F., Gomes, X., and Wold, M. S. (1999) J. Biomol. NMR 14, 321–331[CrossRef][Medline] [Order article via Infotrieve]
- Daughdrill, G. W., Ackerman, J., Isern, N. G., Botuyan, M. V.,
Arrowsmith, C., Wold, M. S., and Lowry, D. F. (2001)
Nucleic Acids Res. 29,
3270–3276
[Abstract/Free Full Text] - Lao, Y., Gomes, X. V., Ren, Y. J., Taylor, J. S., and Wold, M. S. (2000) Biochemistry 39, 850–859[CrossRef][Medline] [Order article via Infotrieve]
- Gomes, X. V., and Wold, M. S. (1996) Biochemistry 35, 10558–10568[CrossRef][Medline] [Order article via Infotrieve]
- Bochkareva, E., Korolev, S., Lees-Miller, S. P., and Bochkarev, A. (2002) EMBO J. 21, 1855–1863[CrossRef][Medline] [Order article via Infotrieve]
- Mer, G., Bochkarev, A., Gupta, R., Bochkareva, E., Frappier, L., Ingles, C. J., Edwards, A. M., and Chazin, W. J. (2000) Cell 103, 449–456[CrossRef][Medline] [Order article via Infotrieve]
- Georgaki, A., Strack, B., Podust, V., and Hübscher, U. (1992) FEBS Lett. 308, 240–244[CrossRef][Medline] [Order article via Infotrieve]
- Treuner, K., Ramsperger, U., and Knippers, R. (1996) J. Mol. Biol. 259, 104–112[CrossRef][Medline] [Order article via Infotrieve]
- Lao, Y., Lee, C. G., and Wold, M. S. (1999) Biochemistry 38, 3974–3984[CrossRef][Medline] [Order article via Infotrieve]
- Iftode, C., and Borowiec, J. A. (1998)
Nucleic Acids Res. 26,
5636–5643
[Abstract/Free Full Text] - He, Z., Brinton, B. T., Greenblatt, J., Hassell, J. A., and Ingles, C. J. (1993) Cell 73, 1223–1232[CrossRef][Medline] [Order article via Infotrieve]
- Li, R., and Botchan, M. R. (1993) Cell 73, 1207–1221[CrossRef][Medline] [Order article via Infotrieve]
- Din, S., Brill, S. J., Fairman, M. P., and Stillman, B.
(1990) Genes Dev.
4,
968–977
[Abstract/Free Full Text] - Dutta, A., and Stillman, B. (1992) EMBO J. 11, 2189–2199[Medline] [Order article via Infotrieve]
- Lee, S.-H., and Kim, D. K. (1995) J. Biol.
Chem. 270,
12801–12807
[Abstract/Free Full Text] - Zernik-Kobak, M., Vasunia, K., Connelly, M., Anderson, C. W., and
Dixon, K. (1997) J. Biol. Chem.
272,
23896–23904
[Abstract/Free Full Text] - Carty, M. P., Zernik-Kobak, M., McGrath, S., and Dixon, K. (1994) EMBO J. 13, 2114–2123[Medline] [Order article via Infotrieve]
- Liu, V. F., and Weaver, D. T. (1993) Mol.
Cell Biol. 13,
7222–7231
[Abstract/Free Full Text] - Shao, R. G., Cao, C. X., Zhang, H. L., Kohn, K. W., Wold, M. S., and Pommier, Y. (1999) EMBO J. 18, 1397–1406[CrossRef][Medline] [Order article via Infotrieve]
- Treuner, K., Okuyama, A., Knippers, R., and Fackelmayer, F. O.
(1999) Nucleic Acids Res.
27,
1499–1504
[Abstract/Free Full Text] - Henricksen, L. A., Carter, T., Dutta, A., and Wold, M. S.
(1996) Nucleic Acids Res.
24,
3107–3112
[Abstract/Free Full Text] - Gomes, X. V., Henricksen, L. A., and Wold, M. S. (1996) Biochemistry 35, 5586–5595[CrossRef][Medline] [Order article via Infotrieve]
- Kraulis, P. J. (1991) J. Appl. Crystallogr. 24, 946–950[CrossRef]
- Oakley, G. G., Patrick, S. M., Yao, J., Carty, M. P., Turchi, J. J., and Dixon, K. (2003) Biochemistry 42, 3255–3264[CrossRef][Medline] [Order article via Infotrieve]
- Georgaki, A., and Hübscher, U. (1993)
Nucleic Acids Res. 21,
3659–3665
[Abstract/Free Full Text] - Fried, L. M., Koumenis, C., Peterson, S. R., Green, S. L., Van
Zijl, P., Allalunis-Turner, J., Chen, D. J., Fishel, R., Giaccia, A. J.,
Brown, J. M., and Kirchgessner, C. U. (1996) Proc.
Natl. Acad. Sci. U. S. A. 93,
13825–13830
[Abstract/Free Full Text] - Treuner, K., Findeisen, M., Strausfeld, U., and Knippers, R.
(1999) J. Biol. Chem.
274,
15556–15561
[Abstract/Free Full Text] - Loo, Y. M., and Melendy, T. (2000) Nucleic
Acids Res. 28,
3354–3360
[Abstract/Free Full Text] - Dimitrova, D. S., and Gilbert, D. M. (2000) Exp. Cell Res. 254, 321–327[CrossRef][Medline] [Order article via Infotrieve]
- Henricksen, L. A., Umbricht, C. B., and Wold, M. S.
(1994) J. Biol. Chem.
269,
11121–11132
[Abstract/Free Full Text] - Gomes, X. V., and Wold, M. S. (1995) J.
Biol. Chem. 270,
4534–4543
[Abstract/Free Full Text] - Johnson, M. L., and Frasier, S. G. (1985) Methods Enzymol. 117, 301–342
- Iftode, C., and Borowiec, J. A. (1997) Mol. Cell Biol. 17, 3876–3883[Abstract]
- Lao, Y. (2000) Multiple Interactions Between DNA and Human Replication Protein A (RPA), Ph.D. Dissertation, University of Iowa
- Umezu, K., Sugawara, N., Chen, C., Haber, J. E., and Kolodner, R.
D. (1998) Genetics
148,
989–1005
[Abstract/Free Full Text] - Longhese, M. P., Plevani, P., and Lucchini, G. (1994)
Mol. Cell Biol. 14,
7884–7890
[Abstract/Free Full Text] - Soustelle, C., Vedel, M., Kolodner, R., and Nicolas, A.
(2002) Genetics
161,
535–547
[Abstract/Free Full Text] - Firmenich, A. A., Elias-Arnanz, M., and Berg, P. (1995) Mol. Cell Biol. 15, 1620–1631[Abstract]
- Longhese, M. P., Neecke, H., Paciotti, V., Lucchini, G., and
Plevani, P. (1996) Nucleic Acids Res.
24,
3533–3537
[Abstract/Free Full Text] - Kim, H. S., and Brill, S. J. (2001) Mol.
Cell Biol. 21,
3725–3737
[Abstract/Free Full Text] - Kantake, N., Sugiyama, T., Kolodner, R. D., and Kowalczykowski, S. C. (2003) J. Biol. Chem.
- Keshav, K. F., Chen, C., and Dutta, A. (1995) Mol. Cell Biol. 15, 3119–3128[Abstract]
- Pan, Z.-Q., Park, C.-H., Amin, A. A., Hurwitz, J., and Sancar, A.
(1995) Proc. Natl. Acad. Sci.
92,
4636–4640
[Abstract/Free Full Text] - Stigger, E., Drissi, R., and Lee, S. H. (1998)
J. Biol. Chem. 273,
9337–9343
[Abstract/Free Full Text] - Missura, M., Buterin, T., Hindges, R., Hubscher, U., Kasparkova, J., Brabec, V., and Naegeli, H. (2001) EMBO J. 20, 3554–3564[CrossRef][Medline] [Order article via Infotrieve]
- Henricksen, L. A., and Wold, M. S. (1994)
J. Biol. Chem. 269,
24203–24208
[Abstract/Free Full Text]
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Reddit 
Technorati What's this?
This article has been cited by other articles:
|
|
 |
|
  A. M. Dickson, Y. Krasikova, P. Pestryakov, O. Lavrik, and M. S. Wold Essential functions of the 32 kDa subunit of yeast replication protein A Nucleic Acids Res., April 1, 2009; 37(7): 2313 - 2326. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 S. K. Binz and M. S. Wold Regulatory Functions of the N-terminal Domain of the 70-kDa Subunit of Replication Protein A (RPA) J. Biol. Chem., August 1, 2008; 283(31): 21559 - 21570. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. J. Haring, A. C. Mason, S. K. Binz, and M. S. Wold Cellular Functions of Human RPA1: MULTIPLE ROLES OF DOMAINS IN REPLICATION, REPAIR, AND CHECKPOINTS J. Biol. Chem., July 4, 2008; 283(27): 19095 - 19111. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 K. C. Manthey, S. Opiyo, J. G. Glanzer, D. Dimitrova, J. Elliott, and G. G. Oakley NBS1 mediates ATR-dependent RPA hyperphosphorylation following replication-fork stall and collapse J. Cell Sci., December 1, 2007; 120(23): 4221 - 4229. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 H. L. Ball, M. R. Ehrhardt, D. A. Mordes, G. G. Glick, W. J. Chazin, and D. Cortez Function of a Conserved Checkpoint Recruitment Domain in ATRIP Proteins Mol. Cell. Biol., May 1, 2007; 27(9): 3367 - 3377. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K. Herzberg, V. I. Bashkirov, M. Rolfsmeier, E. Haghnazari, W. H. McDonald, S. Anderson, E. V. Bashkirova, J. R. Yates III, and W.-D. Heyer Phosphorylation of Rad55 on Serines 2, 8, and 14 Is Required for Efficient Homologous Recombination in the Recovery of Stalled Replication Forks Mol. Cell. Biol., November 15, 2006; 26(22): 8396 - 8409. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. R. Salas, I. Petruseva, O. Lavrik, A. Bourdoncle, J.-L. Mergny, A. Favre, and C. Saintome Human replication protein A unfolds telomeric G-quadruplexes Nucleic Acids Res., October 18, 2006; 34(17): 4857 - 4865. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
E. Fanning, V. Klimovich, and A. R. Nager A dynamic model for replication protein A (RPA) function in DNA processing pathways Nucleic Acids Res., September 10, 2006; 34(15): 4126 - 4137. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. Guo, Y. Zhang, F. Yuan, Y. Gao, L. Gu, I. Wong, and G.-M. Li Regulation of Replication Protein A Functions in DNA Mismatch Repair by Phosphorylation J. Biol. Chem., August 4, 2006; 281(31): 21607 - 21616. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 R. Sakasai, K. Shinohe, Y. Ichijima, N. Okita, A. Shibata, K. Asahina, and H. Teraoka Differential involvement of phosphatidylinositol 3-kinase-related protein kinases in hyperphosphorylation of replication protein A2 in response to replication-mediated DNA double-strand breaks Genes Cells, March 1, 2006; 11(3): 237 - 246. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 T. Ishibashi, S. Kimura, and K. Sakaguchi A Higher Plant Has Three Different Types of RPA Heterotrimeric Complex J. Biochem., January 1, 2006; 139(1): 99 - 104. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A. J. Bartrand, D. Iyasu, S. M. Marinco, and G. S. Brush Evidence of Meiotic Crossover Control in Saccharomyces cerevisiae Through Mec1-Mediated Phosphorylation of Replication Protein A Genetics, January 1, 2006; 172(1): 27 - 39. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 D. Nakada, Y. Hirano, Y. Tanaka, and K. Sugimoto Role of the C Terminus of Mec1 Checkpoint Kinase in Its Localization to Sites of DNA Damage Mol. Biol. Cell, November 1, 2005; 16(11): 5227 - 5235. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 E. Bochkareva, L. Kaustov, A. Ayed, G.-S. Yi, Y. Lu, A. Pineda-Lucena, J. C. C. Liao, A. L. Okorokov, J. Milner, C. H. Arrowsmith, et al. Single-stranded DNA mimicry in the p53 transactivation domain interaction with replication protein A PNAS, October 25, 2005; 102(43): 15412 - 15417. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Y. Liu, M. Kvaratskhelia, S. Hess, Y. Qu, and Y. Zou Modulation of Replication Protein A Function by Its Hyperphosphorylation-induced Conformational Change Involving DNA Binding Domain B J. Biol. Chem., September 23, 2005; 280(38): 32775 - 32783. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
P. D. Vise, B. Baral, A. J. Latos, and G. W. Daughdrill NMR chemical shift and relaxation measurements provide evidence for the coupled folding and binding of the p53 transactivation domain Nucleic Acids Res., April 11, 2005; 33(7): 2061 - 2077. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. G. Robison, L. Lu, K. Dixon, and J. J. Bissler DNA Lesion-specific Co-localization of the Mre11/Rad50/Nbs1 (MRN) Complex and Replication Protein A (RPA) to Repair Foci J. Biol. Chem., April 1, 2005; 280(13): 12927 - 12934. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K. Weisshart, P. Pestryakov, R. W. P. Smith, H. Hartmann, E. Kremmer, O. Lavrik, and H.-P. Nasheuer Coordinated Regulation of Replication Protein A Activities by Its Subunits p14 and p32 J. Biol. Chem., August 20, 2004; 279(34): 35368 - 35376. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. G. Robison, J. Elliott, K. Dixon, and G. G. Oakley Replication Protein A and the Mre11{middle dot}Rad50{middle dot}Nbs1 Complex Co-localize and Interact at Sites of Stalled Replication Forks J. Biol. Chem., August 13, 2004; 279(33): 34802 - 34810. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. J. Bartrand, D. Iyasu, and G. S. Brush DNA Stimulates Mec1-mediated Phosphorylation of Replication Protein A J. Biol. Chem., June 18, 2004; 279(25): 26762 - 26767. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
P. E. Pestryakov, D. Y. Khlimankov, E. Bochkareva, A. Bochkarev, and O. I. Lavrik Human replication protein A (RPA) binds a primer-template junction in the absence of its major ssDNA-binding domains Nucleic Acids Res., March 26, 2004; 32(6): 1894 - 1903. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
V. M. Vassin, M. S. Wold, and J. A. Borowiec Replication Protein A (RPA) Phosphorylation Prevents RPA Association with Replication Centers Mol. Cell. Biol., March 1, 2004; 24(5): 1930 - 1943. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
D. M. Clifford, S. M. Marinco, and G. S. Brush The Meiosis-specific Protein Kinase Ime2 Directs Phosphorylation of Replication Protein A J. Biol. Chem., February 13, 2004; 279(7): 6163 - 6170. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
W. D. Block, Y. Yu, and S. P. Lees-Miller Phosphatidyl inositol 3-kinase-like serine/threonine protein kinases (PIKKs) are required for DNA damage-induced phosphorylation of the 32 kDa subunit of replication protein A at threonine 21 Nucleic Acids Res., February 10, 2004; 32(3): 997 - 1005. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K. Unsal-Kacmaz and A. Sancar Quaternary Structure of ATR and Effects of ATRIP and Replication Protein A on Its DNA Binding and Kinase Activities Mol. Cell. Biol., February 1, 2004; 24(3): 1292 - 1300. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| All ASBMB Journals | Molecular and Cellular Proteomics |
| Journal of Lipid Research | ASBMB Today |