Replication Protein A and the Mre11 (cid:1) Rad50 (cid:1) Nbs1 Complex Co-localize and Interact at Sites of Stalled Replication Forks*

In response to replicative stress, cells relocate and activate DNA repair and cell cycle arrest proteins such as replication protein A (RPA, a three subunit protein complex required for DNA replication and DNA repair) and the MRN complex (consisting of Mre11, Rad50, and Nbs1; involved in DNA double-strand break repair). There is increasing evidence that both of these complexes play a central role in DNA damage recognition, activation of cell cycle checkpoints, and DNA repair pathways. Here we demonstrate that RPA and the MRN complex co-localize to discrete foci and interact in response to DNA replication fork blockage induced by hydroxyurea (HU) or ultraviolet light (UV). Members of both RPA and the MRN complexes become phosphorylated during S-phase and in response to replication fork blockage. Analysis of RPA and Mre11 in fractionated lysates (cytoplasmic/nucleoplasmic, chromatin-bound, and nuclear matrix fractions) showed increased hyper-phosphorylated-RPA and phosphorylated-Mre11 in the chromatin-bound fractions. HU and UV treatment also led to co-localization of hyperphosphorylated RPA and Mre11 to discrete detergent-resistant nuclear foci. An interaction between RPA and Mre11 was demonstrated by co-immunoprecipitation of both protein complexes with anti-Mre11, anti-Rad50, anti-NBS1, or anti-RPA antibodies. Phosphatase treatment with calf intestinal phosphatase

Stalled replication forks threaten DNA replication fidelity and genomic integrity. Stalled forks occur after deficiencies in replication substrates, inhibition of replication machinery, or blocking of the replication machinery by DNA damage or DNAprotein complexes (1,2). Replication fork stalling with subsequent replication stress is also thought to occur during normal DNA replication, particularly at DNA sequences that are prone to form secondary structures (e.g. tRNA genes) or in regions where transcription complexes may collide (2). Failure to resolve this replication stress may result in the collapse of stalled forks and genomic instability (3). To prevent such instability, replication stress triggers the activation of a DNA damage response. This response involves the recruitment and activation of proteins involved in DNA repair and cell cycle regulation. This DNA damage response uses proteins to detect damage, signal the site of damage, and transduce and amplify the signal to activate needed effector proteins. Many proteins involved in these three aspects accumulate and form large nuclear foci after DNA damage. Examples of such proteins include ␥H2AX, 53BP1, ATM, ATR, BRCA1, Werner's protein, the MRN 1 complex, and RPA1 (4,5). The functions of these foci are not fully understood, but they may represent sites of fork reactivation, protein activation, DNA repair, and/or non-repairable damage (6 -9).
Replication protein A (RPA), the major single-stranded DNA (ssDNA)-binding protein in eukaryotic cells, accumulates along stretches of ssDNA generated by stalled replication forks and/or DNA damage (10,11). It has been suggested that the RPA⅐DNA complex created by the accumulation of RPA at these sites may signal the presence of damage and activate the DNA damage response. In support of this, Dart et al. (12) show that recruitment of ATR to nuclear foci after replication fork stalling is dependent on RPA (12). Additionally, it has been shown that RPA promotes DNA binding and activation of ATR/ ATRIP in vitro (13). RPA is also required to recruit and activate Rad17 complexes for checkpoint signaling in vivo (14). Further evidence that RPA acts as a common intermediate for signaling stalled replication forks/DNA damage is demonstrated by the RPA-dependent binding of Cut5 to chromatin after DNA damage and its subsequent recruitment of ATR and DNA polymerase ␣ to chromatin (15). In that report, Parrilla-Castellar and Karnitz (15) put forth a model suggesting that ssDNA generated at a stalled replication fork is coated with RPA, leading to Cut5 chromatin association. Cut5 then facilitates the chromatin association of ATR and DNA Pol␣, which in turn leads to the loading of the 9-1-1 complex (15). These data have given credence to the hypothesis that RPA-coated ssDNA acts as a common intermediate for signaling-stalled replication forks and/or DNA damage and subsequent recruitment and activation of DNA damage response proteins. It is likely that RPA plays a dual role in the damage response network, that of a sensor of damage and also as an effector. Replication protein A is an essential component of most, if not all, DNA repair processes.
Petrini and Stracker (16) postulate that, similar to RPA, the MRN complex acts as a sensor of DNA damage needed to activate the DNA damage response. Recent data suggest that the MRN complex functions as a damage sensor upstream of ATM/ATR activation in addition to acting as an effector downstream of ATM/ATR in double strand break repair and cell cycle checkpoints (17)(18)(19)(20). Although it is known that the MRN complex binds to DNA ends, there is no clearly defined mechanism by which the MRN complex recognizes other types of DNA damage. Because both RPA and the MRN complexes are believed to be involved in the sensing and signaling of DNA damage, we postulated that these two complexes might interact at sites of stalled replication forks and DNA damage. To test this hypothesis, we investigated the interaction of these proteins in response to hydroxyurea (HU)-and ultraviolet light (UV)-induced replication stress and DNA damage. After treatment with these agents, RPA and Mre11 became phosphorylated and co-localized to discrete chromatin-bound nuclear foci. RPA and the MRN complex also co-immunoprecipitated together, suggesting that these proteins interact either directly or indirectly. The interaction between RPA and the MRN complex was abrogated by phosphatase treatment. Taken together, our data suggest that after replication stress induced by HU or UV, RPA and the MRN complex co-localized and interacted at sites of stalled replication forks and that the protein interaction may be mediated at least in part by protein phosphorylation.

EXPERIMENTAL PROCEDURES
Cell Lines and Treatments-HeLa cells were obtained from American Type Culture Collection (ATCC; Manassas, VA) and maintained at 37°C and 5% CO 2 in Dulbecco's modified Eagle's medium (Invitrogen) supplemented with 10% fetal bovine serum (Hyclone, Logan, UT) and 1% penicillin-streptomycin (Invitrogen). For UV exposure, the growth medium was removed (and held at 37°C), and cells were washed with phosphate-buffered saline (PBS). The PBS was replaced with minimum essential medium (Invitrogen) without phenol red, and cells were treated with 30 J/m 2 UVC (for asynchronous cells) or 20 J/m 2 (for cells synchronized in S-phase) using a low pressure mercury lamp (Mineralight lamp; model UVG-11; UVP, Inc., San Gabriel, CA) with a maximal output at 254 nm. After UV exposure, the minimum essential medium was removed and replaced with the original growth medium, and cells were allowed to recover for 8 h at 37°C before harvesting. For HU (Sigma-Aldrich) treatment, cells were incubated in growth medium containing HU (2 mM) for 3 h before harvesting.
Cell Synchronization-Cells were synchronized in S-phase and G 1phase of the cell cycle as previously described (21). To synchronize in S-phase, cells were incubated in growth medium containing aphidocolin (final concentration of 1 M; Sigma-Aldrich) for 15 h. After incubation, the aphidocolin-containing medium was removed, and cells were washed with PBS and then incubated in fresh medium without aphidocolin for an additional 2 h at 37°C. For nocodazole synchronization in G 1 -or S-phase of the cell cycle, cells were incubated in medium containing nocodazole (0.3 M final concentration; Sigma-Aldrich) for 17 h. The mitotic cells, which become detached from the culture dish as they entered M-phase, were collected and pelleted at 500 ϫ g. Mitotic cells were released from nocodazole treatment by incubating in fresh medium. We have shown previously that HeLa cells enter G 1 -phase about 5 h after release and enter S-phase about 12 h after release (21). For the experiments reported here cells were treated with HU from 4.5 to 7.5 h after release (for G 1 -phase) or from 12 to 15 h after release (for S-phase) and then harvested. Subcellular Fractionation-The cellular protein fractionation protocol was performed as previously described with slight modifications (22). Briefly, S-phase-synchronized HeLa cells were treated with either HU, UV, or mock-treated. The free cytoplasmic/nucleoplasmic fraction was prepared by allowing cells to lyse for 10 min on ice in 0.5% Triton X-100 in cell lysis buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0.1% Nonidet P-40, 5 l/ml pepstatin, 5 g/ml leupeptin, 5 g/ml aprotinin, 10 mM NaF, 10 mM ␤-glycerophosphate, 1 mM Na 3 VO 4 , 1 mM phenylmethylsulfonyl fluoride). The insoluble fraction was pelleted by centrifugation at 13,000 ϫ g for 10 min at 4°C, and the supernatant (free cytoplasmic/nucleoplasmic; fraction FCN) was collected. The pellet was washed with PBS and treated with 100 g/ml DNase I in cell lysis buffer at 37°C for 15 min followed by the addition of ammonium sulfate (250 mM final concentration) and further incubation at room temperature for 10 min. The insoluble fraction was pelleted by centrifugation at 13,000 ϫ g for 10 min at 4°C, and the supernatant was collected and designated as the chromatin-bound fraction (fraction CB). The remaining insoluble material was washed with PBS, suspended in SDS buffer (2% SDS, 20 mM Tris-HCl, pH 7.5, 1 M ␤-mercaptoethanol, 10% glycerol), and designated the nuclear matrix fraction (fraction NM). The presence of glucose-3-phosphate dehydrogenase, histone H1, and lamin A/C within the free nucleoplasmic/cytoplasmic fraction, chromatinbound fraction, and nuclear matrix fraction, respectively, verified the validity of this fractionation method.
Co-immunoprecipitation Assays-For co-immunoprecipitation reactions, 50 l of protein G-agarose beads (Invitrogen) were incubated with 3.0 -5.0 g of anti-Mre11 (Novus Biological), anti-RPA-p70 (Bethyl Laboratories, Inc.), or normal rabbit IgG (Oncogene, San Diego, CA) antibodies in PBS for 1 h at room temperature with end-over-end mixing. After the addition of ϳ1000 g of cell lysate, the immunoprecipitation reactions were incubated for 20 -24 h at 4°C with end-over-end mixing. The immunoprecipitates were separated from the supernatant by centrifugation and washed with PBS containing 0.05% Nonidet P-40. Proteins were extracted from the agarose beads by resuspending in 1ϫ Laemmli gel-loading buffer and separated on 12% SDS-polyacrylamide gels.
When antibodies cross-linked to the agarose beads were used, 50 l of protein G-agarose beads (Invitrogen) were incubated with 3.0 -5.0 g of anti-Mre11 (Novus Biological) for 1 h at room temperature with end-over-end mixing. The beads were washed twice with 200 mM triethanolamine (Sigma-Aldrich) and then incubated in 20 mM dimethyl pimelimidate (Sigma-Aldrich) in 200 mM triethanolamine for 30 min at room temperature. The dimethyl pimelimidate solution was replaced with 50 mM Tris-HCl, pH 7.5, for an additional 15 min. The Tris-HCl was removed, and the beads were suspended in PBS and stored at 4°C until the addition of cell lysate.
Phosphatase Treatment-For phosphatase treatments of cellular lysates, 10ϫ NEBuffer 3 (New England Biolabs, Beverly, MA; 100 mM NaCl, 50 mM Tris-HCl, 10 mM MgCl 2 , 1 mM dithiothreitol) was added to ϳ1000 g of total protein to give a final concentration of 1ϫ NEBuffer. 200 units of calf intestinal alkaline phosphatase (CIP) was added to the mixture and then incubated for 2 h at 37°C. For phosphatase treatment of immunoprecipitates, samples were prepared by washing immunoprecipitates with PBS to remove non-specifically bound proteins and the remaining medium. For CIP treatment, the pellets were resuspended in 1ϫ NEBuffer 3 with the addition of 50 units of CIP. The reaction mixtures were incubated at 37°C for 2 h before collection of the Mre11associated pellet and supernatant. For -phosphatase (PPase) treatment, the pellets were resuspended in 1ϫ PPase buffer (New England Biolabs) with 2 mM MnCl 2 or 1ϫ PPase buffer with 1 mM Na 2 VO 4 and 10 mM NaF (known inhibitors of PPase). 400 units of PPase was added to each reaction tube and then incubated at 30°C for 30 min.

Replication Stress Induces Hyperphosphorylation of RPA
and Phosphorylation of Mre11-Both RPA-p34 and Mre11 are phosphorylated in a DNA damage-dependent and a DNA replication-dependent manner (23,24). To verify that phosphorylation of RPA-p34 and Mre11 occurred after exposure of HeLa cells to 2 mM HU for 3 h and 20 J/m 2 or 30 J/m 2 UV, whole cell lysates from treated cells were visualized for RPA-p34 and Mre11 via Western blotting. Similar to previous reports, these treatments resulted in the hyperphosphorylation of the RPA-p34 subunit and phosphorylation of Mre11 (Fig. 1, A (lanes 2-3  and 5-6) and B (lanes 2-3 and 8 -9)). When cells were synchronized in S-phase before treatment with HU or UV, the proportion of hyperphosphorylated RPA and phosphorylated Mre11 increased (Fig. 1, A (lanes 5-6) and B (lanes 8 -9)). This increase in RPA and Mre11 phosphorylation in treated S-phase cells is most likely due to the replication stress induced by these agents. An antibody that recognizes phosphorylated serine 4 and serine 8 of RPA-p34 specifically recognized only the DNA damage-dependent hyperphosphorylated form of RPA-p34 ( Fig. 1, B (lanes 4 -6 and 10 -12)). The phosphorylation of RPA and Mre11 suggests that these proteins are involved in the cellular response to replication stress.
Phosphorylated RPA and Mre11 Are More Tightly Bound to the Chromatin-It has been previously reported that after DNA damage, RPA becomes re-distributed within the nucleus and is found predominantly bound to the chromatin (22). Mre11, which has increased chromatin binding in S-phase, does not show a re-distribution within the nucleus after DNA damage in any stage of the cell cycle (6,25). To investigate the nuclear re-distribution of the phosphorylated isoforms of RPA and Mre11 after replication fork stalling and DNA damage, cellular proteins of S-phase-synchronized cells were separated into three fractions: 1) free cytoplasmic/nucleoplasmic, 2) chromatin-bound, and 3) nuclear matrix fractions. To confirm the separation of the three fractions, we verified the presence of glucose-3-phosphate dehydrogenase (a cytosolic protein), histone H1, and lamin A/C (a nuclear matrix protein) within the free nucleoplasmic/cytoplasmic fractions, chromatin-bound fractions, and nuclear matrix fractions, respectively (data not shown). In mock-treated cells, ϳ60% of the total RPA was in the free cytoplasmic/nucleoplasmic fraction, whereas Mre11 was present in all three fractions, with the chromatin-bound fraction containing the largest percent (Fig. 2, A (lanes 2-3) and B). After HU or UV treatment, there was a redistribution of RPA to the chromatin-bound fraction (Fig. 2, A (lanes 6 -8 and  10 -12) and B), with the percent of total RPA changing from 40% chromatin-bound in mock-treated cells to ϳ85% in HU-or UV-treated cells. As previously reported (22), the damage-induced hyperphosphorylated isoform of RPA was found predominantly in the chromatin-bound fraction. Although the distribution of Mre11 between the different fractions did not change after damage, the majority of the damage-induced phosphorylated form of Mre11 was found in the chromatin-bound fraction (Fig. 2, A (lanes 7 and 11) and B).
RPA and Mre11 Co-localize to Discrete Nuclear Foci after HU or UV Treatment-After the induction of DNA damage, many proteins associated with DNA damage signaling, DNA repair, and cell cycle checkpoints become localized to sites of damage and form nuclear "foci" (4). To visualize these foci and to determine whether Mre11 and RPA were present in these "repair foci," we used immunofluorescent staining. Pre-extraction of cells with a detergent-containing buffer removes the nucleoplasmic and cytoplasmic proteins, leaving behind the chromatin-bound and matrix-associated proteins (6). In mock-treated cells stained with antibodies against RPA and Mre11, there was a diffuse nuclear staining. After HU or UV treatment, there was a redistribution of RPA and Mre11 to discrete foci that co-localized (Fig. 3A, panels D, H, and L). Although studies have shown that both RPA and the MRN complex are able to form nuclear foci after DNA damage by various genotoxic agents (6, 7, 11, 26 -29), no previous reports have indicated that RPA and Mre11 foci co-localize. We also looked for colocalization of Mre11 with hyperphosphorylated RPA using the RPA-p34-S P 4-S P 8 phospho-specific antibody. Under mocktreatment conditions, there is no staining with the phosphospecific RPA antibody as expected (Fig. 3B, panel B). After treatment with HU or UV, similar to staining with the antibody that recognizes all isoforms of RPA, phosphorylated-RPA aggregated into nuclear foci that co-localized with Mre11 (Fig.  3B, panels D, H, and L).
In addition to co-localization of RPA and hyperphosphorylated-RPA with Mre11 and similar to previous reports, these foci also showed limited co-localization with phosphorylated histone H2AX (␥H2AX) (6, 29 -31) and with Werner's protein (Wrn) (data not shown) (5,26). ␥H2AX foci formation is a marker of double-strand breaks (DSBs) and has also been reported to form after replication stress and at replication forks (6,8,32). Wrn protein is a RecQ-class DNA helicase that localizes to sites of stalled replication where it is involved in the resolution and prevention of aberrant recombination events (5) and is able to directly interact with RPA (5, 33) and the Mre11 complex (34). Co-localization of hyperphosphorylated RPA and the MRN complex with ␥H2AX and Wrn along with previous reports (5,6) indicates that these foci are at sites of stalled replication forks, which at the time points investigated may have progressed to the point of collapse and generation of DSBs.
RPA and the MRN Complex Interact-The observation of co-localization of RPA and the MRN complex raised the possibility that these proteins may directly interact. We performed co-immunoprecipitation experiments using whole cell lysates of S-phase-synchronized HeLa cells from HU, UV, or mocktreated cells to probe for such RPA⅐MRN complex interactions. Rabbit anti-Mre11, anti-Rad50, or anti-Nbs1 antibodies was able to immunoprecipitate RPA, and rabbit anti-RPA-p70 antibodies were able to immunoprecipitate Mre11 (Fig. 4, A (lanes  7-12), B (lanes 3-5), and C). Normal rabbit IgG did not immunoprecipitate RPA or Mre11 (Fig. 4, A (lanes 4 -6) and B (lane 6)), demonstrating the co-immunoprecipitation of RPA and Mre11 was not due to nonspecific antibody binding.
FIG. 3. RPA and Mre11 co-localize to discrete nuclear foci following HU and UV treatment. Asynchronous HeLa cells were treated with 2 mM HU for 3 h, 30 J/m 2 UV and allowed recover for 8 h, or mock-treated. After extraction of cytoplasmic and nucleoplasmic proteins with PBS containing 0.5% Triton X-100, cells were fixed in paraformaldehyde, incubated in primary and secondary antibodies, and visualized by fluorescent microscopy. A, RPA-p34 and Mre11 co-localize to HU-and UV-induced nuclear foci. Cells were stained with anti-Mre11 antibodies (green, panels B, F, and J) and anti-RPA-p34 antibodies (red, panels C, G, and K). Panels A, E, and I are the 4,6diamidino-2-phenylindole (DAPI)-stained nuclei, and panels D, H, and L are the merged images of the anti-Mre11 and anti-RPA stained cells. B, hyperphosphorylated-RPA-p34 and Mre11 co-localize to HU-and UV-induced nuclear foci. Cells were stained with damage-induced hyperphosphorylation-specific anti-RPA-p34-S P 4-S P 8 antibodies (green, panels B, F, and I) and anti-Mre11 antibodies (red, panels C, G, and K). Panels A, E, and I are the 4,6-diamidino-2-phenylindole-stained nuclei, and panels D, H, and L are the merged images of the anti-RPA-p34-S P 4-S P 8-and anti-Mre11-stained cells.
RPA is a very abundant protein in the cell. Therefore, we considered the possibility that the apparent co-immunoprecipitation of RPA with Mre11 using anti-Mre11 antibodies could be nonspecific and simply due to RPA abundance. In addition, the co-immunoprecipitation of Mre11 with RPA using anti-RPA-p34 antibodies was barely detectable, and co-immunoprecipitations using anti-RPA-p70 antibodies was present but at very low levels. To demonstrate that the co-immunoprecipitation of RPA and MRN was specific, we carried out the following experiment. Cell lysates were incubated with anti-Mre11 antibodies cross-linked to protein-G-agarose beads, and bound protein was eluted with sodium citrate, pH 3.0, neutralized, and then immunoprecipitated again with anti-RPA-p70, anti-Mre11, or anti-IgG antibody-coated agarose beads. The sequential immunoprecipitations removed the excess RPA from the lysate, allowing for a more stoichiometrically equal amount of RPA and MRN complex to interact in the second round. The ability of anti-RPA-p70 antibodies to immunoprecipitate Mre11 was increased under these conditions (Fig. 4B, lanes 3  and 4), confirming the specificity of the interaction.
In addition to co-immunoprecipitation of RPA and MRN in lysates from damaged cells, co-immunoprecipitation was also observed in lysates from mock-treated cells (Fig. 4, A (lane 7)  and B (lane 3)). This suggested that the interaction might not  1-3). Proteins from the immunoprecipitates were detected by Western blotting using anti-RPA-p34 and anti-Mre11antibodies. B, S-phase-synchronized HeLa cells were either treated with 2 mM HU for 3 h (lanes 2 and 4 -6) or mock-treated (lanes 1 and 3). Whole cell lysates were incubated with agarose beads cross-linked with anti-Mre11 antibodies as the primary immunoprecipitate. The proteins were eluted from the beads using sodium citrate and then incubated with anti-RPA-p70 (lanes 3 and 4), anti-Mre11 (lane 5), or anti-IgG (lane 6) antibody-coated agarose beads for the secondary immunoprecipitation. The proteins were eluted from the second set of antibody-coated beads with Laemmli loading buffer and separated on a 12% SDS-PAGE gel. Proteins were visualized by Western blotting using anti-RPA-p34 and anti-Mre11 antibodies. C, S-phase-synchronized HeLa cells were treated with 2 mM HU for 3 h. Whole cell lysates were incubated with agarose beads coated with anti-Mre11, anti-Rad50, or anti-Nbs1 antibodies. The immunoprecipitated proteins were eluted with Laemmli loading buffer and subjected to Western blotting using anti-RPA-p34 antibodies. D, HeLa cells were synchronized in M-phase of the cell cycle with nocodazole and allowed time to enter G 1 -phase (lanes 1 and 2) or S-phase of the cell cycle with aphidicolin (lanes 3 and 4) as described under "Experimental Procedures." Cells were either mock-treated or treated with 2 mM HU for 3 h before harvesting and whole cell lysate formation. Lysates were incubated with anti-Mre11 antibody-coated agarose beads and eluted off using Laemmli gel loading buffer. The immunoprecipitated proteins were subjected to Western blotting using anti-RPA-p34 and anti-Mre11 antibodies. E, S-phase-synchronized HeLa cells were treated with 2 mM HU for 3 h. Whole cell lysates were either treated with 100 g/ml DNase I for 20 min at 37°C (lane 3) or 50 g/ml EtBr on ice for 30 min (lane 9). Pretreated lysates, lysates without DNase, or EtBr-incubated under similar conditions (lanes 2 and 8) as well as lysates with no previous incubation were immunoprecipitated with anti-Mre11 antibody-coated agarose beads. The two immunoprecipitation reactions with non-treated lysates were washed with PBS and incubated for 20 min at 37°C with or without DNase I in cell lysis buffer (100 g/ml, lanes 5 and 6, respectively). Proteins from all the immunoprecipitation reactions were eluted from the beads with Laemmli gel-loading buffer and separated on 12% SDS gels. The immunoprecipitated proteins were subjected to Western blotting using anti-RPA-p34 antibodies.
be dependent on DNA damage. It has been shown that the MRN complex is required for the resolution of breaks that occur spontaneously during DNA replication (24), so we considered the possibility that the co-immunoprecipitation in the mocktreated samples might be an S-phase phenomenon due to normal replication. To investigate this possibility, immunoprecipitation reactions were done using whole cell lysates from mocktreated and HU-treated cells synchronized in G 1 -phase or S-phase of the cell cycle. The amount of RPA that co-immunoprecipitated with Mre11 in mock-treated cells in S-phase of the cell cycle was six times more than mock-treated G 1 -phase cells as measured by densitometry (Fig. 4D, lanes 1 and 3). After HU treatment, the amount of immunoprecipitated RPA increased 8-fold when comparing S-phase to G 1 -phase. (Fig. 4D, lanes 2  and 4). This suggests that the interaction between RPA and the MRN complex, although present in both G 1 -and S-phase of the cell cycle, is increased when cells are in S-phase and increased to a greater extent after damage in S-phase. The increase in interaction in mock-treated cells in S-phase may represent interactions that occur at spontaneously stalled replication forks during normal DNA replication.
We also considered the possibility of an indirect interaction between RPA and the MRN complex mediated by independent binding of both protein complexes to DNA. To test this possibility, we pretreated lysates with ethidium bromide (EtBr) or DNase I or treated immunoprecipitates with DNase I. EtBr is a DNA intercalator known to disrupt DNA-protein interactions (35,36). Pretreatment of lysates with 50 g/ml EtBr did not alter the ability of anti-Mre11 antibodies to immunoprecipitate RPA (Fig. 4E, lane 9). DNase I treatment of lysates or immunoprecipitates using the same conditions as the subcellular fractionation protocol did not affect the RPA⅐MRN complex interaction as well (Fig. 4E, lanes 3 and 6, respectively), suggesting this interaction is not indirect via DNA-protein interactions.
Phosphatase Treatment Abrogates the RPA⅐MRN Complex Interaction-The Nbs1 protein of the MRN complex contains two domains associated with protein-protein interactions; the forkhead-associated domain and the BRCA1 carboxyl-terminal domain (37,38). The forkhead-associated domain and BRCA1 carboxyl-terminal domains are both known to mediate proteinprotein interactions in a phosphorylation-dependent manner (39,40), which is demonstrated in the interaction of Nbs1 with ␥H2AX and BRCA1 with BACH1 (30,40). Because increased RPA⅐MRN complex interactions occur under conditions of increased RPA-p34 phosphorylation, specifically RPA hyperphosphorylation, we wanted to investigate if protein phosphorylation played a part in the RPA⅐MRN complex interaction. To address this question, whole cell lysates from HU-treated cells were incubated for 2 h at 37°C with or without CIP before incubation with anti-Mre11 antibody-coated agarose beads. Pretreatment with CIP abrogated the ability of anti-Mre11 antibodies to immunoprecipitate RPA (Fig. 5A, lane 3), whereas the 2-h incubation at 37°C had no effect (Fig. 5B, lane  2). To verify that the CIP treatment resulted in protein dephosphorylation, the supernatant from the CIP-treated sample  1 and 2) or specific phosphatase inhibitors (lanes 5 and 6) as indicated. Samples were incubated at 30°C for 30 min, and the immunoprecipitate pellets (P) and supernatants (S) were separated by centrifugation and subjected to Western blotting using anti-RPA-p34 antibodies. Whole cell lysate (10% of total protein used in immunoprecipitation reaction) was included as a loading control (Input, lane 1).
was analyzed for RPA-p34. Loss of the hyperphosphorylated form of RPA-p34 and retention of non-phosphorylated RPA (Fig. 5B, lane 4) indicated that protein de-phosphorylation had occurred.
To continue investigating the effect of phosphatase activity on the RPA⅐MRN complex interaction, we immunoprecipitated proteins from whole cell lysates of HU-treated cells with anti-Mre11 antibody-coated beads. The resulting immunoprecipitate treated with CIP demonstrated a loss of RPA in the Mre11associated pellet and appearance of non-phosphorylated RPA in the supernatant (Fig. 5B, lanes 2 and 3).
To verify that the abrogation of the RPA⅐MRN interaction was dependent upon phosphatase activity and not just the physical presence of the phosphatase enzymes, we used another phosphatase, PPase, and specific phosphatase inhibitors. Immunoprecipitates from HU-treated lysates immunoprecipitated with anti-Mre11 antibodies were treated with PPase with (Fig. 5C, lanes 5 and 6) and without the presence of specific PPase inhibitors (sodium orthovanadate and sodium fluoride; Fig. 5C, lanes 3 and 4). The addition of the PPase inhibitors preserved the RPA⅐MRN complex interaction (and the presence of phosphorylated RPA) with the anti-Mre11 antibody-coated beads (Fig. 5C, lanes 5 and 6), suggesting that the abrogation of the RPA⅐MRN complex interaction was indeed dependent upon phosphatase activity. Together, these data suggest that the RPA⅐MRN complex interaction may be mediated by protein phosphorylation. These data, however, do not address whether the abrogation of the RPA⅐MRN complex interaction is due to de-phosphorylation of RPA, the MRN complex, or some other protein. DISCUSSION The results presented here offer insight into the role of RPA and the MRN complex in the replicative stress-induced DNA damage response. Upon treatment with HU or UV, RPA becomes hyperphosphorylated, and Mre11 is phosphorylated. The relative proportion of the phosphorylated isoforms of RPA and Mre11 were increased when cells were synchronized in the S-phase of the cell cycle before treatment. The phosphorylated isoforms of RPA and Mre11 were chromatin-bound, most likely at sites of stalled and/or collapsed replication forks (6). Although redistribution of phosphorylated RPA to chromatinbound fractions has been previously reported (22), this may seem to be in contradiction to reports that show phosphorylated RPA has a decreased affinity for double-stranded DNA (41,42). Two possible explanations may account for this apparent discrepancy. First, stalled replication forks lead to the generation of large regions of ssDNA. Although phosphorylated RPA has decreased affinity for double-stranded DNA, it shows no difference in ssDNA binding activity (41). Alternatively, after replication fork stalling and/or DNA damage, RPA may have increased interactions with a protein that is itself associated with the chromatin (possibly the MRN complex or some other as yet unidentified protein). Either scenario could explain the increased chromatin association of phosphorylated RPA we observed after HU or UV treatment.
Consistent with a previous report showing constitutive chromatin association of the MRN complex (25), we saw similar amounts of Mre11 in the chromatin-bound fraction in mocktreated and HU-and UV-treated samples. This is also consistent with a report that shows increased chromatin association of the MRN complex in S-phase as compared with G 1 or G 2 /M but no change in S-phase chromatin-association with or without damage (6). Interestingly, the damage-dependent phosphorylated Mre11 is predominantly contained within the chromatinbound Mre11 pool. These data suggest that phosphorylated Mre11 may have increased DNA binding affinity or that Mre11 may have to be chromatin-bound to be phosphorylated.
Although studies have shown that both RPA and MRN form nuclear foci after treatment with a variety of genotoxic agents (6, 7, 11, 26 -29), no one has previously reported that these foci co-localize. Mre11 foci formation after UV-induced damage has been reported to occur only in xeroderma pigmentosum variant cells and not in "normal" cells (7,29,43). We observed UVinduced Mre11 foci in both HeLa and a normal telomerasetransformed cell line (data not shown) 8 h after UV irradiation. It has recently been reported that the large T-antigen in SV40transformed cell lines disturbs the formation of Mre11 nuclear foci (44). Although previous studies that reported Mre11 foci only in xeroderma pigmentosum variant cell lines investigated similar time points and doses of UV, their use of SV40-transformed cells may explain the discrepancies with our results.
Both RPA and the MRN complex are known to bind DNA and interact with numerous other proteins. However, our data demonstrate that DNA binding is not required for the RPA⅐MRN complex interaction. In addition, we observed that protein phosphorylation enhanced RPA⅐MRN complex interaction, similar to the reported phosphorylation-enhanced BRAC1/BACH1 interaction (40).
Our data as well as previous reports (12,13,45) have led to our proposed model of events associated with stalled replication forks (Fig. 6). RPA and the MRN complex are normally associated with replication forks during S-phase (6,10,46). Stalled replication forks generate stretches of ssDNA that become coated with RPA, leading to the recruitment and activation of ATR/ATRIP (12,13) and possibly ATM (12). The close proximity of these proteins at stalled forks would allow for ATR/ATM phosphorylation of RPA and the MRN complex. This enhances recruitment of additional MRN complex and phosphodependent RPA⅐MRN interaction. Phosphorylation most likely has functional consequences in addition to altering protein interactions. For example, phosphorylation of Mre11 is thought to increase its nuclease activity (24), which could increase DNA processing necessary for resolution of the stalled fork. Phos-phorylation of RPA is known to decrease its unwinding or destabilization activity of double-stranded DNA but does not effect ssDNA binding ability (41,42). This may help prevent further migration of the replication fork and provide the opportunity for further protein recruitment. The recruited proteins, RPA, ATR, ATRIP, Mre11⅐Rad50⅐Nbs1 (and possibly ATM), may then initiate DNA repair and act as a scaffold to activate other proteins such as Rad17, Rad9⅐Rad1⅐Hus1, or Wrn (5,14). Possible outcomes may include cell cycle arrest, resolution of Holliday junctions, and continuation of replication, non-homologous end joining (NHEJ)-mediated repair of subsequent DSBs or recombination-like repair of subsequent DSBs. This model suggests that RPA and the MRN complex work together at sites of stalled or collapsed replication forks and that this cooperative interaction occurs via the MRN ability to interact with RPA in a phosphorylation-dependent manner.
In this study a single time point for each DNA-damaging agent was selected based on peak incidence of RPA-p34 hyperphosphorylation (data not shown). Future work needs to investigate additional time points to determine the exact role of RPA and MRN in the replication stress-induced damage response and if additional proteins mediate the RPA⅐MRN complex interaction.