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Previous Article | Next Article 
Molecular and Cellular Biology, February 2004, p. 1292-1300, Vol. 24, No. 3
0270-7306/04/$08.00+0 DOI: 10.1128/MCB.24.3.1292-1300.2003
Copyright © 2004, American Society for Microbiology. All Rights Reserved.
Department of Biochemistry and Biophysics, University of North Carolina School of Medicine, Chapel Hill, North Carolina 27599
Received 5 October 2003/ Returned for modification 5 November 2003/ Accepted 6 November 2003
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ABSTRACT |
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INTRODUCTION |
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The PIKK family encompasses three proteins, DNA-dependent protein kinase (DNA-PK), ATM, and ATR (1, 6, 30). These are large proteins with molecular masses in the range of 301 to 469 kDa. Although PIKKs exhibit sequence similarities in their carboxy-terminal domains to phosphoinositide 3-kinsases, they have no lipid kinase activities. Instead, all three PIKK members are Ser/Thr kinases with preference for amino acids within the SQ/TQ context (11). The prototype of the family, DNA-PK, is a heterotrimeric complex of a 469-kDa catalytic subunit, DNA-PKcs, and a dimer of Ku70 and Ku86 (8). Ku70-Ku86 is a rather stable dimer which binds to DNA ends and recruits DNA-PKcs to the sites of double-strand breaks. DNA-PK plays an essential role in double-strand break repair necessary for V(D)J recombination and in nonhomologous end joining of double-strand breaks induced by ionizing radiation and radiomimetic drugs (32). Whether DNA-PK signals double-strand breaks to the checkpoint machinery still remains controversial, but DNA-PK mutants do not have an overt DNA damage checkpoint defect (24). In contrast, ATM and ATR seem to function almost exclusively in the DNA damage checkpoint response, although it has been found that ATM does contribute to double-strand break repair during V(D)J recombination (22). Both ATM and ATR are DNA binding proteins. ATM binds preferentially to DNA termini, apparently in a monomeric form (33), whereas ATR binds to the DNA backbone with no preference for ends but with some preference for UV-induced (6-4) photoproducts (34). Consistent with these properties, ATM is the main kinase that initiates the immediate checkpoint response to ionizing radiation, and ATR is the predominant initiator of the checkpoint response to UV-induced DNA damage (1, 29, 30).
Because DNA-PKcs is recruited to DNA by the Ku complex, it has been suggested that ATM and ATR may be recruited to DNA in a similar manner, that is, through the intermediacy of smaller subunits (6). Indeed, in Schizosaccharomyces pombe spRad3 (ATR/ATM ortholog) forms a complex with spRad26 (7), and in Saccharomyces cerevisiae, scMec1 (ATR ortholog) forms a complex with scDdc2 (20, 25, 35); some evidence was obtained that these smaller proteins play a role in regulating the kinase and DNA binding activities, respectively, of the corresponding PIKK members (27, 37). Search for similar proteins in mammalian cells led to the discovery of an ATR-interacting protein, ATRIP (5). However, the role of ATRIP in regulating ATR, especially recruiting it to DNA, is uncertain because ATRIP-free ATR binds to DNA relatively efficiently (34). So far, no small subunit for ATM has been found, but it is thought that the ATM activity is regulated by the change in the oligomeric state of the protein (2).
Using immunoprecipitation assays coupled with site-specific mutagenesis, it was found that in undamaged human cells ATM was in the form of a homodimer or a higher-order oligomer that upon DNA damage became phosphorylated by an intermolecular reaction and dissociated to monomers, which appear to be the active form of the enzyme (2). In support of oligomerization as a general regulatory mechanism for PIKK family proteins, it was found that these proteins contain many HEAT repeats, which are known to be involved in protein-protein interactions (23). Because ATR, like ATM, catalyzes autophosphorylation, as we show below, we reasoned that ATR might similarly be an oligomeric protein whose activity is regulated by a change in its oligomerization state induced by DNA damage. Indeed, a preliminary study using gel filtration chromatography concluded that ATR was found predominantly in complexes with an average molecular mass of 1,000 kDa (40). We wished to expand upon this finding and to determine if the ATR activity may be modulated by a change in its quaternary structure, as has been found for ATM.
In this study we used hydrodynamic methods to determine the molecular weight of native ATR protein and the other members of the PIKK family. In agreement with earlier reports, we find that DNA-PKcs is a monomer (13). We also found that ATM is predominantly a dimer that is in equilibrium with monomeric and larger oligomeric forms. Importantly, in contrast to an earlier report (40), ATR behaves as a monomer, as analyzed both by gel filtration chromatography and by glycerol gradient sedimentation velocity centrifugation. Moreover, ATRIP, which has been found in ATR immunoprecipitates and is considered to be essential for the stability of ATR in vivo (5), readily dissociates from ATR during gel filtration or sedimentation under relatively mild conditions. Analyses of DNA binding and kinase activities of ATR and the ATR-ATRIP complex revealed that both forms bound to naked and replication protein A (RPA)-covered DNA with comparable affinities. The ATR-ATRIP complex also binds to RPA and phosphorylates the RPA32 subunit in a DNA-dependent manner. The binding to RPA is mediated by ATRIP, which also stimulates the kinase activity of ATR. Taken together, our data indicate that ATR has a distinct quaternary structure, unrelated to the other members of the PIKK family, and that it can directly bind to DNA without the intermediacy of another protein.
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MATERIALS AND METHODS |
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Mouse monoclonal anti-Flag antibody was obtained from Sigma. Rabbit polyclonal ATRIP-N antibody was a gift from Stephen J. Elledge (5). Mouse monoclonal antibodies against RPA32 and RPA70 were obtained from Oncogene Research Products. Rabbit polyclonal antibody against ATR was obtained from Affinity Bioreagents. Mouse monoclonal antibody against DNA-PKcs was a kind gift from Dale A. Ramsden (Department of Biochemistry and Biophysics, University of North Carolina). Rabbit polyclonal antibody against ATM was a kind gift from Richard S. Paules (National Institute of Environmental Health Sciences).
Expression and purification of recombinant proteins. HEK293T cells were grown in Dulbecco's modified Eagle medium (GIBCO/BRL) supplemented with 10% fetal bovine serum and 100 U of penicillin and streptomycin/ml. Cells (8 x 106/225-cm2 flask) were transfected either with 25 µg of pcDNA3-Flag-ATR plasmid alone or cotransfected with 15 µg of Flag-ATR and 10 µg of Myc(3x)-tagged ATRIP plasmids by using a calcium phosphate transfection method (10). Forty-eight hours after transfection cells were washed with phosphate-buffered saline and lysed in 5 ml of lysis buffer (50 mM Tris-HCl [pH 7.5], 150 mM NaCl, 10 mM ß-glycerophosphate, 10% glycerol, 1% Tween 20, 0.1% NP-40, 1 mM Na3VO4, 1 mM NaF, and protease inhibitors [Roche Molecular Biochemicals]) for 30 min on ice. After centrifugation at 32,000 x g for 30 min, the supernatants were incubated with 40 µl of anti-Flag M2 affinity gel (Sigma) at 4°C overnight. For Flag-ATR-transfected cells, the Flag-ATR-bound beads were washed three times with 1.5 ml of washing buffer that contained 50 mM Tris-HCl (pH 7.5) and either 150 mM or 1 M NaCl and the proteins were eluted in 200 µl of elution buffer that contained 50 mM Tris-HCl (pH 7.5), 150 mM NaCl, 10% glycerol, 1 mM dithiothreitol [DTT], and 200 µg of Flag-peptide/ml. For Flag-ATR- and Myc(3x)-ATRIP-cotransfected cells, washing was performed only under low salt conditions (150 mM NaCl).
Baculovirus for expression of His6-Flag-ATRIP was generated with the BAC-TO-BAC baculovirus expression system (GIBCO/BRL) and protocols suggested by the manufacturer. The optimal multiplicity of infection for recombinant virus was empirically determined. A monolayer of Sf21 insect cells (Invitrogen) grown in Grace's insect medium (GIBCO/BRL) supplemented with 10% fetal bovine serum and 100 U of penicillin and streptomycin/ml was infected with virus and harvested 48 h later. The cells were lysed and centrifuged as described for HEK293T cells, and protein-bound Flag-beads were washed three times in a buffer containing 50 mM Tris-HCl (pH 7.5) and 300 mM NaCl, and the proteins were eluted in elution buffer with 200 µg of Flag-peptide/ml. The 9-1-1 complex containing Flag-Hus1 was purified from High Five insect cells as previously described (15). The recombinant RPA was expressed and purified from BL21(DE3) cells as described previously (9).
Gel filtration analysis. HEK293T whole-cell extract was prepared in hypotonic buffer (20 mM HEPES [pH 7.9], 5 mM KCl, 1.5 mM MgCl2, 1 mM DTT) as described previously (14). A Superdex-200 PC 3.2/30 (2.4-ml) gel filtration column was equilibrated at 4°C in a buffer with 20 mM HEPES (pH 7.9), 250 mM NaCl, 1.5 mM MgCl2, and 2 mM DTT. Forty microliters of whole-cell extracts (25 mg/ml) which contain approximately 1.5 to 2 pmol of ATR and 0.35 to 0.5 pmol of ATRIP (as determined by Western blotting using recombinant proteins as standards) was loaded onto the column, and the column was developed with the same buffer. After determining the excluded volume (Vo = 0.87 ml) and included volume (Vi = 1.708) of the column, the elution volume (Ve) was measured for standard proteins with a known Stokes radius. For each protein, Kav was calculated from the equation that follows, and a linear relationship was obtained by plotting (-logKav)1/2 against the Stokes radius: Kav = (Ve - Vo)/(Vi - Vo). The standard proteins used were bovine serum albumin (67 kDa, 1.46 ml), rabbit muscle aldalose (158 kDa, 1.33 ml), bovine liver catalase (232 kDa, 1.28 ml), horse spleen ferritin (440 kDa, 1.12 ml), and bovine thyroid thyroglobulin (669 kDa, 0.98 ml). Elution profiles of the proteins of interest were determined by immunoblot analysis.
Glycerol gradient centrifugation. Whole-cell extracts (500 µg) obtained from nonirradiated or UV-irradiated (50 J/m2, extract made 1 h after irradiation) HEK293T cells in a total volume of 20 µl (25 mg/ml) were layered on top of 2.4-ml 15 to 35% glycerol gradients in buffer containing 20 mM HEPES (pH 7.9), 50 mM KCl, 7 mM MgCl2, and 0.5 mM DTT. The gradients were spun for 6 h at 55,000 rpm in an SW60 rotor at 4°C. Fractions of 200 µl were collected from the bottom of the tube, and 75-µl aliquots were analyzed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). To determine the sedimentation coefficient of ATRIP in the absence of ATR, 4 µg of His6-Flag-tagged ATRIP (purified from insect cells) in a total volume of 40 µl was loaded onto the glycerol gradient and processed as for the whole-cell extract. The elution positions of the proteins of interest were determined by Western blot analysis. Reference proteins with known sedimentation coefficients were used to calibrate the gradients: chicken ovalbumin (3.55 S), rabbit muscle aldalose (7.3 S), bovine liver catalase (11.3 S), bovine thyroid thyroglobulin (19 S).
Biotinylated DNA binding assay. Streptavidin-conjugated Dynabeads M280 (Dynal) were washed three times with Tris-EDTA (TE) buffer plus 1 M NaCl. A 5'-biotinylated 80-nucleotide (nt) oligomer was added to the beads in the same buffer and incubated at room temperature for 1 h. The sequence of the 5'-biotinylated oligonucleotide used in this study is 5'TTTTTTTTTTTTTTTTTTTTTTTTTTTTTTCTCCCTTCTTCTCCTCCCTCTCCCTTCCCTTTTTTTTTTTTTTTTTTTTT 3'.
For each reaction, 7 pmol of DNA and 5 µl of Dynabeads were used. The beads and DNA mixture were washed three times with TE buffer plus 1 M NaCl followed by two washes with TE buffer. The DNA-bound beads were incubated with purified proteins at 30°C for 30 min in a reaction volume of 400 µl with a binding buffer (10 mM MgCl2, 2 mM MnCl2, 10 mM Tris-Cl [pH 7.5], 50 mM NaCl, 0.2 mM EDTA, 10% glycerol, 1 mM DTT, 10 µg of bovine serum albumin/ml). For each binding reaction, 7 pmol of RPA, 4 pmol of ATRIP, and 4 pmol of ATR were used. These amounts of DNA and protein were chosen for the binding assays so as to have concentrations of reactants comparable to those used in a recent study (43). After the beads were washed three times with washing buffer (binding buffer with 100 mM NaCl and 0.01% NP-40), the bound proteins were analyzed by Western blotting. In reaction mixtures containing RPA-covered DNA, the 7 pmol of 5'-biotinylated 80-nt DNA was first incubated with RPA at 30°C for 30 min and then either ATR, ATRIP, or ATR-ATRIP complex reconstituted by preincubating the purified ATR and ATRIP on ice for 30 min was added to RPA-coated DNA and further incubated for 30 min at 30°C. After the beads were washed to remove the unbound proteins, the bound proteins were eluted by boiling the beads in SDS buffer and analyzed by Western blotting.
Kinase assay.
Kinase assays were performed in 50-µl reaction mixtures by using 50 ng of ATR, 200 ng of RPA, 200 ng of ATRIP, and 25 µg of single-stranded DNA (ssDNA) in a kinase buffer containing 25 mM HEPES (pH 7.9), 50 mM KCl, 10 mM MgCl2, 2 mM MnCl2, 20% glycerol, 0.1% NP-40, 1 mM DTT, 1 mM Na3VO4, and 1 mM NaF. The reaction mixtures were incubated on ice for 5 min before ATP addition. ATP was added to a final concentration of 50 µM along with 5 µCi of [
-32P]ATP. Reaction mixtures were incubated at 30°C for the indicated times. Phosphorylated proteins were separated on SDS-10% polyacrylamide gels, visualized by silver staining, and exposed to a Molecular Dynamics PhosphorImager screen for quantitation by IMAGEQUANT software.
Immunoprecipitation assays. HEK293T cells were either singly transfected or cotransfected with the indicated plasmids by a calcium phosphate method. After 16 h of incubation at 37°C, the cells were washed twice in serum-free Dulbecco's modified Eagle medium, fresh medium was added, and the cells were incubated for another 48 h. The cells were lysed in ice-cold lysis buffer for 30 min. The cell lysates were centrifuged for 30 min at 30,000 x g, and the supernatants were incubated with anti-Flag M2 affinity gel (Sigma) at 4°C for 4 h. The beads were then washed three times with TBS buffer (50 mM Tris-HCl [pH 7.5], 150 mM NaCl), and bound proteins were eluted with TBS buffer containing 200 µg of Flag-peptide (Sigma)/ml.
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RESULTS |
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oNaS/(1 - 
), where S is the sedimentation coefficient and a is the Stokes radius of the protein (see Table 1, footnote a, for other definitions).
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Effect of UV irradiation on ATR.
It was shown that ionizing radiation drastically alters ATM quaternary structure by causing transphosphorylation followed by dissociation to the monomeric form (2). We wished to find out if DNA damage affected ATR by changing its interactions with other cellular proteins, in particular with ATRIP. Since ATR is the principal sensor in the UV-induced checkpoint response, we used UV irradiation as the genotoxicant and compared the behavior of the protein on a glycerol gradient before and after UV irradiation. In particular, we were interested in finding out if there might be a change in ATR-ATRIP interaction following DNA-damage. Figure 3 shows that the sedimentation of ATR and ATRIP in the glycerol gradient before and after UV irradiation is essentially the same. Importantly, there does not appear to be any change in the strength of the ATR-ATRIP interaction. ATRIP sediments faster than would be expected from its native molecular weight under both conditions, indicative of ATR-ATRIP 
ATR + ATRIP equilibrium that is not affected by DNA damage. It should be noted that the faster sedimentation of ATRIP is not caused by an unusual shape, as purified ATRIP, free of ATR, has an S value consistent with its monomeric molecular weight (Fig. 3 and Table 1). It must be also noted that under the irradiation conditions we used for Fig. 3 there is an ATR-mediated checkpoint activation, as evidenced by Chk1-S345 phosphorylation (data not shown). Taken together, these data lead us to conclude that after UV damage there is no long-lasting measurable change in the interaction of ATR with its smaller partner, ATRIP, or any other cellular protein.
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DISCUSSION |
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Quaternary structure of ATR. Hydrodynamic analysis of ATR in human whole-cell extract by gel filtration chromatography and glycerol gradient velocity sedimentation revealed that ATR behaved as a protein with a Stokes radius of 72.8 Å and a sedimentation coefficient of 12.6, from which an apparent molecular weight of 385,500 is calculated. This is in reasonable agreement with the theoretical molecular weight of 391,382 for the ATR-ATRIP complex. Thus, it appears that ATR is in the form of an ATR-ATRIP heterodimer in equilibrium with the individual components. Even though ATR is known to interact with many other proteins involved in the cell cycle checkpoint or DNA repair those interactions are probably transient and of not sufficient avidity for the proteins to be considered part of an ATR-containing multiprotein complex. This finding is in disagreement with an earlier report that concluded that ATR was predominantly in complexes with an average molecular mass of 1,000 kDa based on its elution profile in a gel filtration (40). The lysis procedure and the ionic strength of the gel filtration column used in the previous work were quite similar to ours, which excludes nonspecific aggregation as the cause of the abnormally high molecular mass found in the previous study. We have no satisfactory explanation for the discrepancy. However, as we obtain similar molecular weights both by gel filtration and sedimentation and by using the Siegel-Monty (31) formula, which is more rigorous than those obtained by either method alone, we consider ATR to be mostly in the form of an ATR-ATRIP heterodimer in vivo. We also find that ATM is in equilibrium among various oligomerization states, with the dimer form being the predominant species. This is the first hydrodynamic evidence for the recent report (2) that the native form of ATM in undamaged cells is a dimer or a higher-order oligomer.
It has been reported that DNA damage leads to transphosphorylation of ATM and its consequent conversion into a monomer, which is the active form that initiates the checkpoint cascade (2). We considered the possibility that ATR may undergo a similar change in its quaternary structure by a change in the interaction state of ATR and ATRIP. However, we did not detect any change in the ATR-ATRIP ATR + ATRIP equilibrium following UV damage, which activates the DNA damage checkpoint pathway mainly through ATR. Even though our experimental design would have not detected a transient change in this equilibrium we believe that is unlikely and that ATR, in contrast to ATM, is not regulated by a change in quaternary structure caused by autophosphorylation. The different behavior of these two PIKK family members is not unexpected. Despite being members of the PIKK family and playing important roles in DNA damage checkpoint signaling, ATM and ATR do have some fundamental differences (1, 29, 30): ATM is most effectively activated by double-strand breaks, whereas ATR is the initiator of the checkpoint response for UV damage and replication stress. ATM is not essential for cell viability, but ATR is because of an apparently indispensable role during normal DNA replication. Finally, ATR has a smaller molecular weight partner called ATRIP, but ATM does not. Assuming that ATR is a monomeric protein bound to ATRIP only with moderate affinity, it might be asked why transfecting cells with the ATR-kinase-inactive mutant causes a dominant negative phenotype. The most common mechanism for the dominant negative phenotype is the formation of complexes between mutant and wild-type protomers of an oligomeric protein, as is the case for ATM (2). However, an equally plausible mechanism is the sequestration of either the upstream or the downstream proteins in a signal transduction pathway by the mutant protein. It is likely that kinase-inactive ATR exerts its dominant negative effect by both mechanisms. It may heterodimerize with ATRIP in competition with wild-type ATR and cause the degradation of the wild-type protein (5), and it may bind to downstream targets such as Claspin and Chk1 and sequester them from the wild-type ATR-initiated signaling pathway.
Finally, in comparing ATR with ATM some comment must be made on the role of autophosphorylation. DNA damage induces autophosphorylation of ATM (2). It has been shown that autophosphorylation in trans converts ATM to the active monomeric state. We find no change in hydrodynamic behavior of ATR upon DNA damage even though ATR is capable of autophosphorylation. Preliminary data with site-specific mutants of ATR suggest that elimination of major phosphorylation sites do not affect its known activities, including ATRIP binding, DNA binding, and kinase activity on other substrates (data not shown). Clearly, the significance of ATR autophosphorylation remains to be determined.
Role of ATRIP in ATR function. Downregulation of ATRIP by siRNA in vivo causes a drastic decrease in ATR level, suggesting that ATRIP is necessary for the stability of ATR (5). This phenomenon is commonly observed with proteins that make very stable complexes such as the XPF and ERCC1 excision repair proteins (28, 39). In a previous study we found that ATRIP was readily separated from ATR by washing ATR immunoprecipitates with 1 M NaCl (34). Dissociating a heterodimer with such relatively high ionic strength was not unexpected. However, we were surprised to see that ATRIP partly separated from ATR under nearly physiological ionic strength in both a gel filtration column and during sedimentation in a glycerol gradient (Fig. 1 and 3). Lacking a concentration-dependent analysis of monomer-dimer equilibrium we are unable to calculate the equilibrium dissociation constant of the heterodimer. However, an approximate consideration of the available data would indicate that the equilibrium dissociation constant is in the micromolar range. These data strongly indicate that in vivo at any given time a significant fraction of ATR is not in complex with ATRIP. In this regard, the ATR-ATRIP interaction is more similar to DNA-PKcs-Ku interaction rather than those of other more stable heterodimers such as the XPF-ERCC1 repair/recombination endonuclease, which is quite stable in ionic strength up to 1 M and purifies through several columns as a complex (28, 39).
The role of ATRIP in ATR functions is of particular interest because it was presumed that ATRIP was the functional analogue of Rad26 and Ddc2 in S. pombe and S. cerevisiae, respectively. However, there is some evidence that the roles of these Rad26-related proteins may differ among different organisms. It has been shown that spRad26 is required for spRad3 kinase activity in fission yeast (37), but scDdc2 is not required for scMec1 kinase activity in budding yeast (35) while ATRIP is required for stability of ATR in humans (5). Likewise, it was shown that scMec1 and scDdc2 colocalize to sites of DNA damage (12, 17) and that localization of scDdc2 in vivo requires scMec1 (17); however, fission yeast spRad26 seems to respond to DNA damage independently of spRad3 (38), as the localization pattern of spRad26 before and after DNA damage was unaffected by a null mutation in spRad3. Finally, it has been reported that although scDdc2 binds DNA independently of scMec1, both in vivo and in vitro binding of scMec1 to DNA requires scDdc2 (27). It was previously reported that, in contrast to scMec1, ATR was capable of binding to DNA without the aid of ATRIP (34). This finding has recently been confirmed and extended in a study that demonstrated that ATR free of ATRIP can bind to single- and double-stranded DNAs with comparable affinities (43). However, it was also reported that ATRIP was incapable of binding to ssDNA unless the DNA was covered with RPA and moreover that as a consequence of this property of ATRIP the ATR-ATRIP complex failed to bind ssDNA but bound to RPA-covered ssDNA (43). Therefore, it was suggested that binding of ATRIP to RPA-coated DNA enables the ATR-ATRIP complex to associate with RPA-covered ssDNA, which is a feature of a stalled replication fork (43). In the present study, we observed comparable bindings of ATR, ATRIP, and the ATR-ATRIP complex to both naked and RPA-covered DNA under our experimental conditions. However, we did observe a modest stimulation of ATR binding to RPA-covered DNA by ATRIP. Although there are some subtle differences between the reaction conditions used in the two studies, we believe our data strongly suggest that under physiological conditions ATRIP is not necessary for recruitment of ATR to a stalled replication fork. Considering our hydrodynamic analysis of ATR, which shows that a significant fraction of ATR in cells might be free of ATRIP, together with our DNA binding studies, we think it is possible that ATR and ATRIP in monomer forms may orchestrate different steps of the DNA damage checkpoint pathway. Clearly, more work is needed to deconvolute the ATRIP-dependent and independent functions of ATR.
Effect of ATR on RPA. When ATR is immunopurified under relatively mild conditions such that ATRIP is retained, we routinely observe significant amounts of RPA in the immunoprecipitate as well. Therefore, we investigated the effects of ATR and RPA on one another's activity. First, we find that the RPA-ATR interaction is dependent on the presence of ATRIP and is most likely mediated by ATRIP. Second, ATR weakly phosphorylates RPA32 in the absence of DNA, but this is greatly stimulated by ssDNA, in agreement with a recent report (3). Finally, when the effect of ATRIP on RPA32 phosphorylation was tested it was found that ATR does phosphorylate RPA32 even in the absence of ATRIP but that ATRIP stimulated the kinase activity of ATR significantly both with regard to autophosphorylation of ATR and the phosphorylation of RPA32. The phosphorylation of RPA32 in a cell-cycle-dependent manner has been known for some time; however, its significance is unknown (36). Phosphorylation does modulate RPA-DNA interactions (4, 19). However, in vitro, phosphorylation of RPA32 does not affect the RPA activity in replication or in excision repair (21). The recent finding that the kinase activity of ATR is essential for irradiation-induced translocation of RPA to sites of DNA damage (3) seemed to be an important clue to the significance of RPA32 phosphorylation during S phase and DNA damage. Paradoxically, however, it was shown that when the phosphorylation sites of RPA32 were mutated, the nonphosphorylatable form of RPA32 did not affect the DNA damage checkpoint response (3). Thus, this issue remains unresolved. Nevertheless, the interactions of among ATR, ATRIP, RPA, and DNA that have been uncovered in this and other recent studies are likely to contribute to a better understanding of the damage-sensing step of the DNA damage checkpoint response.
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ACKNOWLEDGMENTS |
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We thank Stephen J. Elledge (Baylor College of Medicine) and David Cortez (Vanderbilt University) for the ATRIP plasmid and ATRIP antibodies, Mark C. Hall (University of North Carolina) for mass spectroscopy, Dale A. Ramsden (University of North Carolina) for help with gel filtration chromatography, and Jerard Hurwitz (Sloan-Kettering Institute) and Laura Lindsey-Boltz (University of North Carolina) for critical comments on the manuscript.
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FOOTNOTES |
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-Flag)-agarose under low-salt (150 mM NaCl) conditions (lane 1). The positions of ATR as well as the ectopically expressed and endogenous (Endo) ATRIP are indicated. Flag-tagged ATR purified from transiently transfected HEK293T cells with anti-Flag-agarose by a low-salt (150 mM NaCl) wash contains endogenous ATRIP (lane 2). Flag-ATR prepared by high-salt (1 M NaCl) wash has no detectable ATRIP (lane 3). (B) The proteins purified in panel A were used in a DNA binding assay with streptavidin-biotin-immobilized DNA. For each binding reaction, 7 pmol of 5'-biotinylated 80-nt oligomer was attached to 5 µl of streptavidin-coated beads and the ssDNA-coated beads were incubated with 5 pmol of the indicated protein samples at 30°C for 30 min. The protein samples from panel A in lanes 1, 2, and 3 were used for lanes 1 to 3, 4 to 6, and 7 to 9, respectively. After the unbound proteins were washed away from the beads, ATR and ATRIP associated with ssDNA were separated by SDS-PAGE and the proteins were detected by immunoblotting with corresponding antibodies. Input represents one-fourth of the total protein used in the experiment. At the bottom, the 9-1-1 complex was applied to the DNA-biotin-streptavidin-beads as a negative control for nonspecific adsorption to the beads. The Hus-1 subunit of the complex was Flag tagged and was used to locate the complex. Input is the total protein used in the experiment.
4 µg) Flag-agarose beads. The protein mixture was incubated for 30 min on ice, and then the Flag-agarose beads were extensively washed three times. The bound and unbound proteins were identified by Western blotting with the indicated antibodies.
