4070 Research Article

IntroductionIn response to genotoxic stress, eukaryotic cells induce signalingpathways that result in the regulation of cell-cycle progression andmobilization of repair proteins. An important, although poorlyunderstood, pathway involves the response to replication stress(Ben-Yehoyada et al., 2007). During S phase, conditions that causeuncoupling of the replicative DNA helicase from the DNA-polymerase machinery (e.g. by DNA-polymerase stalling) lead tothe generation of significant lengths of single-stranded DNA(ssDNA) (Byun et al., 2005). The ssDNA becomes bound byreplication protein A (RPA) and the resulting RPA-ssDNA entityserves to recruit the ATR checkpoint kinase and the associated ATR-interacting protein (ATRIP) (Namiki and Zou, 2006; Zou andElledge, 2003). Binding leads to kinase activation andphosphorylation of downstream effector molecules such as the tumorsuppressors BRCA1 (Yarden et al., 2002) and p53 (Tibbetts et al.,1999), the histone variant H2AX (yielding H2AX) (Ward andChen, 2001), and the Chk1 checkpoint kinase (Guo et al., 2000;Liu et al., 2000; Zhao and Piwnica-Worms, 2001). Study of thefunction of ATR and its homologs (e.g. Mec1 in Saccharomycescerevisiae) indicate that it stabilizes stalled or damaged DNA-replication forks (Lopes et al., 2001; Tercero and Diffley, 2001;Sogo et al., 2002) and arrests the progression of non-damaged forks(Seiler et al., 2007; Unsal-Kacmaz et al., 2007).

RPA is the primary eukaryotic ssDNA-binding protein and isessential for chromosomal DNA replication and certain DNA repairreactions (Binz et al., 2004; Iftode et al., 1999; San Filippo et al.,2008). In addition to being crucial for recruitment and activationof ATR, RPA is also an ATR target (Block et al., 2004; Olson et

al., 2006). Human RPA is a heterotrimer composed of the RPA1,RPA2 and RPA3 subunits (also termed RPA70, RPA32 and RPA14,respectively, for their molecular masses). The protein is primarilyphosphorylated within the N-terminal 33 residues of RPA2, in whichapproximately nine sites have been identified (Binz et al., 2004).Two of the sites (T21 and S33) are consensus forphosphatidylinositol 3-kinase-like kinase (PIKK)-family members(ATM, ATR and DNA-PK), whereas others are targets of the cyclin-Cdk complex (S23 and S29) or DNA-PK (S4, S8, S11, S12 andS13; Fig. 1A) (Anantha et al., 2007; Block et al., 2004; Dutta andStillman, 1992; Fang and Newport, 1993; Liu et al., 2006; Mantheyet al., 2007; Niu et al., 1997; Oakley et al., 2003; Olson et al., 2006;Pan et al., 1994; Zernik-Kobak et al., 1997). Treatment of cells withthe DNA-damaging agent camptothecin (CPT) causes initial RPAphosphorylation on S33 by ATR; this phosphorylation subsequentlystimulates phosphorylation by cyclin-Cdk and DNA-PK to yieldhyperphosphorylated RPA (Anantha et al., 2007). A variety of otherdamage conditions, including UV irradiation or ionizing irradiation(IR), treatment with DNA-damaging chemicals and use of DNA-polymerase inhibitors [hydroxyurea (HU), aphidicolin], stimulateRPA phosphorylation by ATM, ATR, DNA-PK and cyclin-Cdk(Anantha et al., 2007; Gibbs et al., 1996; Liu et al., 2000; Pan etal., 1994; Wang et al., 2001; Zernik-Kobak et al., 1997) [also seereview by Binz et al. (Binz et al., 2004) and references therein].

Although significant understanding exists regarding the kinasesthat are responsible for RPA phosphorylation, the role ofphosphorylation in regulating RPA activity remains unclear. RPAphosphorylation does not have profound effects on ssDNA bindingor support of SV40 DNA replication (Brush et al., 1994; Henricksen

ATR is an essential kinase activated in response to DNA-replication stress, with a known target being the RPA2 subunitof human replication protein A (RPA). We find that S33-RPA2phosphorylation by ATR occurs primarily in the late-S and G2phases, probably at sites of residual stalled DNA-replicationforks, with S33-P-RPA2 contained within nuclear repair centers.Although cells in which endogenous RPA2 was ‘replaced’ withan RPA2 protein with mutations T21A and S33A (T21A/S33A-RPA) had normal levels of DNA replication under non-stressconditions, the mutant cells were severely deficient in theamount of DNA synthesis occurring during replication stress.These cells also had abnormally high levels of chromatin-boundRPA, indicative of increased amounts of single-stranded DNA(ssDNA) and showed defective recovery from stress. Cells

replaced with the mutant RPA2 also generated G1 cells with abroader DNA distribution and high levels of apoptosis followingstress, compared with cells expressing wild-type RPA2.Surprisingly, cells expressing the wild-type RPA2 subunit hadincreased levels of stress-dependent DNA breaks. Our datademonstrate that RPA phosphorylation at the T21 and S33 sitesfacilitates adaptation of a DNA-replication fork to replicationstress.

Supplementary material available online athttp://jcs.biologists.org/cgi/content/full/122/22/4070/DC1

Key words: ATR, DNA damage, DNA replication, RPA, Replicationstress

Summary

Human RPA phosphorylation by ATR stimulates DNAsynthesis and prevents ssDNA accumulation duringDNA-replication stressVitaly M. Vassin, Rachel William Anantha, Elena Sokolova, Shlomo Kanner and James A. Borowiec*Department of Biochemistry and New York University Cancer Institute, New York University School of Medicine, New York, NY 10016, USA*Author for correspondence (james.borowiec@med.nyu.edu)

Accepted 30 August 2009Journal of Cell Science 122, 4070-4080 Published by The Company of Biologists 2009doi:10.1242/jcs.053702

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et al., 1996; Oakley et al., 2003; Pan et al., 1995; Vassin et al.,2004). Phosphorylation of RPA also has not been found to directlyinfluence checkpoint activation in response to interphase DNAdamage (Morgan and Kastan, 1997; Olson et al., 2006). By contrast,tests of RPA2 mutants that mimic hyperphosphorylation indicatethat phosphorylation inhibits RPA from participating inchromosomal DNA replication (Vassin et al., 2004; Olson et al.,2006). Because these mutant factors are competent to bind to sitesof DNA damage, it has been argued that hyperphosphorylationinhibits RPA loading onto ssDNA by the replication machinery(Vassin et al., 2004). Mitotic RPA2 hyperphosphorylation inresponse to mitotic DNA damage facilitates release of cells into G1and increases cell viability, apparently by a signaling mechanism(Anantha and Borowiec, 2009; Anantha et al., 2008). In responseto genotoxic stress, it was shown that cells expressing an RPA2variant that was mutated at both Cdk2 sites (S23 and S29) hadaberrant RPA foci and increased persistence of the DNA-damagemarker H2AX following genotoxic stress (Anantha et al., 2007).Such data indicate that RPA phosphorylation stimulateschromosomal DNA repair, although the molecular basis for theseeffects are not understood.

To elucidate the functional role of RPA phosphorylation, weexamined the effect of mutation of the two PIKK sites (T21 andS33). Testing cells in which the endogenous RPA2 was ‘replaced’with an ectopic version, we found that cells expressing the mutantRPA2 were severely defective in DNA synthesis under conditions

of DNA-replication stress, causing high levels of apoptosis. Thisdecrease in synthesis occurred concomitantly with an increase inthe amount of RPA bound to DNA, an event also seen in cellsdeficient for ATR. Our data demonstrate that RPA phosphorylationminimizes ssDNA generation under replication-stress conditions inpart by stimulating repair DNA synthesis.

ResultsRPA2 S33 is phosphorylated in response to replication stressRPA is phosphorylated in response to a variety of genotoxicstresses. In this study, we focused on mild DNA-replication stressto understand the cellular response to impeded replication-forkprogression through particular chromosomal regions (e.g. fragileloci) in non-stressed cells or during the early response to DNAlesions.

Human U2-OS cells were first treated with HU, and the effecton RPA2 phosphorylation was examined at various time points usingantibodies specific to RPA2 phosphorylated at S33 (S33-P), T21(T21-P), S29 (S29-P), and doubly modified S4 and S8 (S4-P-S8-P).Previous characterization of these antibodies found no significantrecognition of non-phosphorylated RPA2 or of bands that co-migratewith RPA2 (e.g. Anantha et al., 2007). In the absence of treatment,only two weak S33-P-positive bands were seen, and termed thelow (L) and intermediate (I) forms (Fig. 1B, lane 1). The I formalso cross-reacted with the S29-P antibody. Test of a ‘low’concentration of HU that has partial inhibitory properties on

Fig. 1. RPA2 S33 is phosphorylated in response to replication stress. (A)Human RPA2 phosphorylation sites. Phosphorylation sites are indicated by underlines,with known or putative kinases shown. (B)Time course of RPA modification in response to HU treatment. U2-OS cells were treated with 1 or 5 mM HU forvarious times, as indicated. Lysates were then prepared and subjected to western analysis for the presence of S33-P-, T21-P-, S4-P-S8-P- and S29-P-RPA2, andtotal RPA2. The relative intensities of the S33-P-positive ‘L’ bands in each lane, normalized to total RPA2 levels, are indicated beneath the S33-P panel. (C)RPAmodification of S33 in response to aphidicolin treatment. U2-OS cells were mock-treated or treated with 30M aphidicolin for various times, as indicated. Lysateswere then probed by western blotting for S33-P-RPA2 and total RPA2. (D)U2-OS cells were subjected to various treatments as follows: mock (‘M’), 5 mMcaffeine for 5 hours (‘C’), 5M aphidicolin for 5 hours (‘A’), 5 mM caffeine and 5M aphidicolin for 5 hours (‘A/C’), 2.5 mM HU for 5 hours (‘H’), 2.5 mM HUand 5 mM caffeine for 5 hours (‘H/C’), and 2M camptothecin for 1 hour (‘Cpt’). Lysates were subjected to western analysis using antibodies specific for S33-P-RPA2, S4-P-S8-P-RPA2 and total RPA2. (B-D)The migration of non-phosphorylated RPA2 (‘B’ for basal), RPA2 phosphorylated to yield species with low (‘L’) orintermediate (‘I’) mobilities, and hyperphosphorylated RPA2 (‘H’) are indicated.

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nucleotide incorporation (1 mM; see below) demonstrated that theL form of S33-P-RPA2 was induced at early times post-treatment(30 minutes), peaked at 2 hours and decreased thereafter (Fig. 1B,lanes 2-6). Other than a slight increase in S29-P formation (Fig.1B, compare lanes 1 and 3) that diminished at late times of treatment,no other modification was detected. A similar biphasic profile wasnoted using aphidicolin to cause DNA-replication stress (Fig. 1C).Previous studies testing various RPA2 phosphorylation mutantsindicated that the L species corresponds to RPA2 phosphorylatedsolely on S33, whereas the S33-P-positive species migrating withintermediate mobility is also modified at the S23 and/or S29 cyclin-Cdk sites (Anantha et al., 2007). Thus, conditions of DNA-replication stress cause an increase in RPA2 phosphorylation at theS33, S29 and S23 sites. In addition, these data indicate thatactivation of the replication-stress checkpoint under these conditionsis transient in that the cell can respond in a manner that eventuallycauses a reduction in the checkpoint signal.

In response to replication stress caused by treatment with 5 mMHU, the S33-P signal initially gained intensity in the first 2 hoursof treatment and then partially diminished (Fig. 1B, lanes 7-11). Atthe two later time points (i.e. 6 and 11 hours; Fig. 1B, lanes 10 and11, respectively), an additional band migrating even more slowlywas detected (termed ‘H’). This RPA2 species also reacted withthe T21-P, S29-P and S4-P-S8-P antibodies, and therefore ishyperphosphorylated RPA2. The H species has been previouslyfound to be formed in response to agents that cause double-strandDNA breaks (DSBs), including CPT and bleomycin, and that alsolead to the formation of H2AX (Vassin et al., 2004; Anantha etal., 2007; Anantha et al., 2008). Notably, evidence from cell-freeand cell-culture studies indicate that phosphorylation of S4 and S8is primarily catalyzed by the DSB-responsive DNA-PK (Zernik-Kobak et al., 1997; Anantha et al., 2007). We interpret these datato indicate that extended incubation in high HU concentrations causea fraction of DNA-replication forks to collapse, generating DSBsand RPA2 hyperphosphorylation (see also below).

We treated cells with aphidicolin or HU in the presence of caffeine.Our previous work has demonstrated that the use of caffeine underDNA-damage conditions blocks the checkpoint response, inducingreplication-fork collapse and hence more-extensive damage (Vassinet al., 2004). Note that the effect of caffeine treatment is functionallysimilar to that caused by deficiencies in Mec1p and Rad53p (thebudding yeast homologs of ATR and Chk1, respectively), followingexposure to DNA-replication stress (Lopes et al., 2001; Tercero andDiffley, 2001). Treatment with either aphidicolin (Fig. 1D, lane 4)or HU (Fig. 1D, lane 6) in combination with caffeine gave rise to ahyperphosphorylated RPA2 species that was reactive to both the S33-P and S4-P-S8-P antibodies. Incubation of cells with CPT to causeDSBs similarly caused the formation of a S33-P and S4-P-S8-P-positive hyperphosphorylated RPA2 (Fig. 1D, lane 7). These dataindicate that replication stress (i.e. treatment with aphidicolinor HU) induces limited RPA2 modification including formationof S33-P. By contrast, more-extensive damage causes RPA2hyperphosphorylation, which includes S4 and S8 modification (aswell as formation of S29-P and T21-P; see Fig. 3B) (Anantha et al.,2007).

S33-P-RPA2 is formed in late-S and G2 phasesWe examined the cell-cycle dependence of S33 phosphorylation inthe absence of induced genotoxic stress. Cells were arrested inmitosis and then released into the cell cycle for various times (Fig.2A). At each time point, both the cell-cycle distribution and the

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phosphorylation status of S33 were quantitated (Fig. 2B and 2C,respectively). Cells in G1 showed low levels of S33-P (4 and 8hours), which increased as cells entered S phase (16 hours). Theamount of S33-P-RPA2 continued to increase and reached a peakat 23 hours when cells were predominantly in late S and in G2.Throughout the cell cycle, the predominant S33-P (L) band migratednear the basal RPA2 form. This S33-P-RPA2 species was alsodetected using the general anti-RPA2 antibodies as a weak bandmigrating just above basal RPA2 (Fig. 2C, marked with ‘*’) andwith similar kinetics of appearance. Weaker S33-P-positive RPA2species migrating in the I and H forms were also detected. Thesedata indicate that a fraction of the RPA pool becomes phosphorylatedon S33-RPA2 in a normal cell cycle, as cells proceed through Sphase into G2. Use of epifluorescence microscopy similarlydemonstrated the presence of a small number of S33-P-RPA2-positive foci (usually in the range of two to five foci), which were

Fig. 2. S33-P-RPA2 foci are formed in the late-S and G2 phases, andcolocalize with BLM foci. (A)Schema of treatment. (B,C)U2-OS cells weretreated with nocodazole for 8 hours and the mitotic cells then shaken off andresuspended in media lacking nocodazole. Cells were allowed to grow forvarious times, and aliquots taken and either subjected to FACS analysis forcell-cycle position (B) or lysed and analyzed by western blotting for totalRPA2 and S33-P-RPA2 (C). (C)The positions of the various RPA2 species areindicated as described in the Fig. 1 legend. The ‘*’ band indicates the putativeRPA2 species phosphorylated only on S33, and migrating slightly above basalRPA2. (D)Nocodazole-arrested U2-OS cells were released for 23 hours,yielding a population of cells mostly in either the late-S or G2 phases. Justprior to fixation, an aliquot of cells were incubated with biotin-11-dUTP(biodU) to label sites of DNA synthesis. Cells were extracted to remove mostsoluble RPA, such that only DNA-bound RPA (at sites of replication or repair)remained. Images show representative cells stained for: (top) incorporatedbiodU, total RPA2 and S33-P-RPA2; and (bottom) S33-P-RPA2, total RPA2and BLM. The signals are shown in grey tones for the left three images. Theright image for each row shows the combined staining for all three markers.The colors of the three markers in the merged images are indicated by thecolor of letters for each stain.

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present selectively in the nuclei of late-S and G2 cells(supplementary material Fig. S1). From the results described above(see Fig. 1), we argue that this phosphorylation reflects the presenceof residual stalled DNA-replication forks in a normal S phase.

The significance of S33 phosphorylation was furthercharacterized by immunofluorescence microscopy. To determinewhether S33-P-RPA2 is found at sites of DNA synthesis, cells werepermeabilized and incubated with biotin-11-dUTP (biodU) to labelsites of incorporation. Following fixation, cells were also stainedfor general RPA2 and S33-P-RPA2. Note that biodU is used hererather than BrdU because the procedure for detecting incorporatedBrdU generally involves harsh conditions (e.g. 2 M HCl) that caninterfere with subsequent detection of proteins. As shown in animage of a typical late-S–G2 cell (Fig. 2D, top panel), a smallnumber of S33-P-positive foci were detected that colocalized withtotal RPA2 and sites of biodU incorporation. Additional images ofnon-stressed cells demonstrate similar patterns of colocalizing S33-P-RPA2 and biodU foci (supplementary material Fig. S2), with suchpatterns observed in a majority of cells. We conclude that S33-P-RPA2 formation is associated with sites of DNA synthesis, probablystalled (or slowly advancing) DNA-replication forks.

BLM is a recQ-like DNA helicase whose gene is mutated inBloom syndrome, a disease resulting in cancer predisposition anddevelopmental defects (Ouyang et al., 2008). Under genotoxic-stressconditions, BLM moves to DNA-damage foci (Davalos andCampisi, 2003). Testing the colocalization of RPA and BLM, weobserved that nearly all BLM-positive foci also showed S33-Pstaining (Fig. 2D, bottom panel). S33-P-RPA2 was also found tocolocalize with H2AX (see below). These data indicate that S33-P forms at sites of DNA damage.

ATR phosphorylates S33 in response to replication stressWe determined the kinase(s) responsible for S33 phosphorylationunder replication-stress conditions. Because S33 is a PIKK site, werestricted our analysis to DNA-PK, ATM and ATR. The role ofDNA-PK in modifying S33 was tested using the M059J (DNA-PK–/–) or M059K (DNA-PK+/+) paired cell lines (Allalunis-Turneret al., 1993). Aliquots of each cell type were mock-treated or treatedwith HU to cause replication stress, and the lysates then subjectedto western blot analysis for S33-P-RPA2. The absence of DNA-PK did not have any notable effects on the L and I forms ofphosphorylated RPA2 (Fig. 3A). A similar test of ATM using cellsdefective for ATM kinase activity (ATM–/–) and a paired wild-typeATM control cell line (ATM+/+) showed no significant differencebetween the control and mutant cell lines on S33-P formation.Finally, the influence of ATR on S33 phosphorylation was examinedusing RNA interference to knock down ATR levels (Fig. 3C).Transfection of a small interfering RNA (siRNA) specific for ATRwas observed to reduce ATR protein levels by >90%, comparedwith control cells transfected with a luciferase siRNA (Luc).Treatment with HU demonstrated that the appearance of both theS33-P-positive L and I species of RPA2 was selectively inhibitedin the cells lacking ATR (Fig. 3C, lane 4). These data indicate thatATR, but not ATM or DNA-PK, is the primary S33 kinase inresponse to replication stress. A similar conclusion was recentlymade by Olson et al. (Olson et al., 2006).

Chk1 is a key effector kinase activated by ATR phosphorylation(Paulsen and Cimprich, 2007). Chk1 modification by ATR is alsoindirectly dependent upon the Rad17-RFC complex. Rad17-RFCloads the Rad9-Hus1-Rad1 (9-1-1) complex onto DNA at sites ofdamage (Kondo et al., 2001; Melo et al., 2001; Zou et al., 2002),

with the bound 9-1-1 supporting recruitment and activation of Chk1(Zou et al., 2002). Because RPA is a direct ATR target, weexamined whether S33 phosphorylation is also dependent on theRad17-RFC complex. The human Rad17 protein was down-modulated by use of a specific siRNA (Fig. 3D, lane 2), incomparison with cells transfected with a luciferase (control) siRNA(Fig. 3D, lane 1). HU treatment of control cells showed robustphosphorylation of S317-Chk1 and S33-RPA2 (Fig. 3D, lane 3).Rad17-deficient cells were found to have defective S317-Chk1, butnot S33-RPA2, phosphorylation (Fig. 3D, lane 4). These datademonstrate that S33-RPA2 phosphorylation is independent ofRad17-RFC- and presumably 9-1-1-complex loading at sites ofDNA-replication stress.

Response of DNA replication to HU treatmentWe examined in more detail the effect of HU treatment onchromosomal DNA synthesis and chromatin association of RPA inU2-OS cells. At various times post-treatment, cells were incubatedwith BrdU and then fixed. Cells were then stained with anti-BrdUand -RPA2 antibodies and imaged. Under our fixation conditions,soluble RPA is extracted, so that the retained RPA fraction isprimarily chromatin-bound (Dimitrova and Gilbert, 2000). Wequantitated both the fraction of BrdU- and RPA-positive cells, andthe average BrdU and RPA signals in these cells. A representativeexperiment demonstrating the appreciable BrdU staining signal incells exposed to 1.5 mM HU is included (supplementary materialFig. S3). Following treatment with a low HU concentration (1 mM),the fraction of BrdU-positive cells initially decreased from >55%to ~15%, before recovering to near control levels (Fig. 4A). Thelevel of BrdU incorporation in these cells showed an even more

Fig. 3. S33-RPA2 phosphorylated by ATR in response to replication stress.(A,B)The M059J (DNA-PK–/–) or M059K (DNA-PK+/+) paired cell lines (A),and GM637 (ATM+/+) or GM5849 (ATM–/–) lines (B) were either mock-treatedor treated with 2.5 mM HU for 3 hours to cause replication stress. (C)U2-OScells were transfected with an siRNA directed against ATR or a control siRNAagainst luciferase (Luc). After 48 hours, cells were either mock-treated ortreated with 2.5 mM HU for 3 hours. (D)U2-OS cells were transfected with ansiRNA directed against Rad17 or luciferase. For all panels, cells were lysedfollowing treatment and the lysates subjected to SDS-PAGE and westernblotting using antibodies specific to total RPA2, S33-P-RPA2, ATR, Rad17 orS317-P-Chk1, as indicated. The total RPA2 signal was used as a loadingcontrol. The positions of the various RPA2 species are indicated as in the Fig.1 legend.

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dramatic drop at initial times of treatment, followed by a weakrecovery in which cells were incorporating BrdU at ~15% of controllevels (Fig. 4B). The formation of S4-P S8-P was low under theseconditions (see Fig. 1B, lanes 2 to 6), indicating that replication-fork collapse does not appreciably occur under these conditions.By contrast, when a high HU concentration (5 mM) was used, anearly complete abolition of DNA synthesis occurred, and only amarginal recovery phase was seen. Nearly all cells stained positivefor RPA over the course of the experiment, regardless of HUconcentration, although those treated with high HU also showed acorresponding increase in formation of S4-P-S8-P-RPA2, indicativeof DSB formation following replication-fork collapse (Fig. 4C).

Of note, the average RPA2 signal per cell increased followingHU treatment, with the use of high HU levels found to cause an~sevenfold increase in chromatin-bound RPA at late times post-treatment (Fig. 4D). Because the extraction procedure retains onlychromatin-bound RPA, the level of non-extracted RPA provides ageneral indicator of the amount of ssDNA present in cells.Chromatin-associated RPA has been validated as an indicator ofssDNA by others (e.g. Raderschall et al., 1999). This increase inssDNA probably occurs as a consequence of DNA-polymerasestalling even while the replicative DNA helicase continues to unwindthe duplex DNA at the fork.

The cell-cycle profiles of U2-OS cells were determined aftera 48-hour exposure to HU. Following treatment with 1 mM HU,we saw a decline in the fractions of both S (from 35.8 to 19.5%)-and G2-M (from 19.2 to 8.1%)-phase cells, whereas G1-phasecells increased (from 45.0 to 72.4%) (Fig. 4E). Although cellstreated with 5 mM HU had a similar decrease in G2-M cells (to8.6%), the loss in S-phase cells was somewhat reduced (to25.0%). These data indicate that a significant fraction of replicatingcells exposed to 1 mM HU are eventually able to exit S phase

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and reach the subsequent G1 phase. Cells incubated with the higher(5 mM) HU concentration evinced a smaller fraction of cells ableto exit S phase.

The S33A-RPA2 mutation reduces DNA synthesis catalyzedduring DNA-replication stressTo examine the functional role of RPA phosphorylation, wegenerated stable cell lines that allow inducible expression of anuntagged mutant or wild-type RPA2 subunit. Clones were chosenthat express RPA2 at a level similar to that of the endogenous factor(e.g. see supplementary material Fig. S4). Expression is coupledwith knock down of endogenous RPA2 using an siRNA duplex thattargets a 3� UTR sequence specific to the endogenous RPA2mRNA. To minimize stress caused by replacement, the ectopicRPA2 is first induced, followed by knock down of the endogenousprotein (Fig. 5A). The ‘replaced’ cells are then assayed 72 hourspost-siRNA transfection. The efficiency of the approach and keycontrols were previously presented (Anantha et al., 2007; Ananthaet al., 2008) (see also below).

Using this replacement strategy, we examined the biologicaleffects of RPA2 subunits mutated at both PIKK-phosphorylationsites (i.e. S33 and T21) to alanines. Replaced cells were treatedwith HU and then incubated with BrdU to allow quantitation ofchromosomal DNA synthesis and fixed. Cells were stained withanti-BrdU antibodies and imaged, quantitating the average BrdUsignal from all cells. In the absence of exogenous stress, the mutantclones (expressing RPA2 with mutations T21A and S33A;T21A/S33A-RPA) were seen to have a similar overall BrdU signalas the two wild-type RPA2 (wt-RPA2) clones (Fig. 5B). These datademonstrate that the T21A/S33A mutation does not have significanteffects on the overall level of DNA synthesis during chromosomalDNA replication.

Fig. 4. Effects of HU on chromosomal DNA replication and RPAchromatin loading. U2-OS cells on coverslips were incubated with 1or 5 mM HU for various times. (A,B)For measurement of DNAsynthesis, cells were pulsed with BrdU and then processed asindicated in Materials and Methods to quantitate both the percentageof BrdU-positive cells (A) and the average BrdU signal in BrdU-positive cells (B). For the latter determination, the signal in cells nottreated with HU (i.e. 0-hour time point) was normalized to 100%.See supplementary material Fig. S3 for representative images ofcells stained for BrdU. (C,D)Following HU treatment, cells werefixed and then stained with anti-RPA2 antibodies and imaged byepifluorescence microscopy. Both the fraction of RPA2-positive cells(in percent; C) and the relative RPA2 signal in RPA2-positive cells(D) were determined. (D)The average RPA2 signal in mock-treatedcells was normalized to a value of 1.0. (E)Effect of HU on cell-cycledistribution. U2-OS cells were either mock-treated or subjected totreatment with 1 or 5 mM HU for 48 hours. Cells were then analyzedby FACS for DNA content. The relative percentages of cells in G1, Sand G2-M phases were quantitated and are shown on the right.

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Next, the same clones were treated with a low concentration ofHU to cause replication stress, and similarly analyzed. The levelof BrdU incorporation for the HU-treated cells replaced with wt-RPA2 was ~10% of that found in the same cells not exposed toHU. Significantly, under HU-treatment conditions, the S33Amutation caused a further >tenfold reduction in the level of DNAsynthesis compared with wt-RPA2 cells. The T21A/S33A-RPA2clones had both a reduced fraction of BrdU-positive cells and alower BrdU signal in these cells. Thus, expression of theT21A/S33A-RPA2 mutant results in the inhibition of DNA synthesisselectively following HU treatment. Because of the reproducibilityof the approach, hereafter we generally show the results from onlya single representative clone, even though three clones wereinvariably analyzed for each RPA2 variant.

Lack of T21/S33-RPA2 phosphorylation increases ssDNAformation during replication stressWe examined the consequences of the T21A/S33A-RPA2 mutationon the generation of ssDNA during DNA-replication stress. We firsttested the effect of ATR down-modulation on RPA-chromatinassociation. U2-OS cells were either treated with an siRNA specificfor ATR, or a luciferase control. Following mock or HU treatment,cells were extracted and fixed, and the level of retained RPAquantitated after immunofluorescence microscopy (Fig. 5C). In theabsence of genotoxic stress, there was only a minor effect of ATRdown-modulation on the level of chromatin-associated RPA.Following exposure to HU, whereas control knockdown cells hada 1.5-fold increase in RPA staining, ATR-knockdown cells had a>sixfold increase in RPA staining. These data indicate that ATRactivity serves to reduce the amount of ssDNA generated in cellsundergoing replication stress. A similar phenomenon has beenobserved in yeast defective in the Mec1 pathway (Lopes et al., 2001;Sogo et al., 2002).

We next tested the effects of mutation of the two consensus PIKKsites on RPA-chromatin association (Fig. 5D). Under mock

treatment conditions, no significant differences were observed inthe level of RPA-chromatin association, indicative of similar levelsof ssDNA being present in each cell line on average. Following a1.5-hour treatment with HU, cells replaced with S33A-RPA2 orT21A/S33A-RPA2 each had a ~3.3-fold increase in RPA-chromatinassociation, which is modestly higher than the 2.5-fold increase seenin cells replaced with wt-RPA2. After a longer treatment with HU(4.5 hours), whereas the wt-RPA2 cells showed a threefold increasein chromatin-associated RPA, the S33A-RPA2 and T21A/S33A-RPA2 cells had 6.3-fold and 8.4-fold increases in chromatin-boundRPA, respectively. These higher levels of retained RPA indicate thatcells replaced with S33A-RPA2 and T21A/S33A-RPA2 have moressDNA present under replication-stress conditions compared withcells replaced with wt-RPA2.

Mutation of the T21 and S33 sites inhibits recovery fromgenotoxic stressWe examined the role of the T21 and S33 sites in recovery fromgenotoxic stress, first testing HU. Replaced cells were firstsynchronized in G1 with mimosine and then released. Afterallowing cells to proceed into S phase, cells were treated with 5mM HU for 4 hours, released from the HU block for 40 minutesand the relative BrdU incorporation then determined (Fig. 6A).Whereas the level of DNA synthesis in wt-RPA2 cells recoveredto ~60% of the level found in non-stressed cells, only a ~22%recovery was detected in cells replaced with the T21A/S33A-RPA2mutant. The recovery of the S33A-RPA2 mutant was intermediatebetween the wt-RPA2 and T21A/S33A-RPA2 cells. Similarly, 30minutes after release from a 7-hour treatment with aphidicolin, cellsreplaced with S33A-RPA2 showed a significantly reduced level ofDNA synthesis compared with wt-RPA2 cells (Fig. 6B). The S33A-RPA2 cells were also defective in recovery from UV irradiation(Fig. 6C). These data demonstrate that cells expressing RPA2phosphorylation mutants are defective in recovery from replicationstress.

Fig. 5. The T21A-S33A-RPA2 mutation inhibits DNA repair synthesis and increases ssDNA accumulation during replication stress. (A)Schematic indicating thesteps involved in cellular RPA2 replacement. ‘–Dox’ indicates the removal of doxycycline to cause the induction of ectopic RPA2 expression. Three U2-OS cloneseach that express either wt-RPA2 or a mutant RPA2 variant were invariably tested. Ectopic RPA2 was induced and, 24 hours later, the endogenous RPA2 wassilenced. 3 days later, cells were either mock-treated (no stress) or treated with HU for various times and processed. (B)U2-OS clones were replaced with either wt-RPA2 or T21A-S33A-RPA2, and then either mock- or HU-treated (1.5 mM) for 4.5 hours. Cells were then incubated with BrdU and processed as described inMaterials and Methods for analysis of BrdU incorporation. Approximately 500 cells were analyzed for each clone, quantitating the average BrdU signal from allcells. (C,D)U2-OS cells were transfected with siRNA molecules against luciferase (Luc; control) or ATR (C), or cloned cells replaced with either wt-, S33- orT21A-S33A-RPA2 (D). Cells were then either mock- or HU-treated (5 mM) for 1.5 or 4.5 hours (as indicated). Following extraction and fixation, cells werestained for RPA2 and imaged by epifluorescence microscopy. The average signals of retained, and hence chromatin-bound, RPA2 were determined.

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Correlation between S33-P and H2AX signalsWe examined the levels of the DNA-damage marker H2AX in celllines replaced with wt- or S33A-RPA2 (Fig. 7A). Cells were treatedwith a low concentration of HU (1 mM for 20 hours), which allowsresidual DNA replication yet causes replication stress (e.g. see Fig.4B). Lysates were prepared and subjected to western analysis againstvarious targets (Fig. 7B). Although mock-treated wt-RPA2 cells hadonly background levels of S33-P, this signal increased after HU

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treatment. Whereas the signal was mostly found at the L and Ipositions, indicative of replication-fork stalling, a clear S33-P signalmigrating at the hyperphosphorylated (H) position was also detected(Fig. 7B, lanes 7-9). Importantly, cells expressing wt-RPA2 alsoshowed significant signals of both S4-P-S8-P-RPA2 and H2AX.

As expected, cells replaced with S33A-RPA2 showed a great lossof the S33-P signal, indicating the efficiency of the replacementstrategy. These cells also showed a reduced S4-P-S8-P signal, also

Fig. 6. Effect of mutation of RPA2 PIKK sites on recovery from DNA-replication stress. U2-OS clones were replaced with either wt-, S33- or T21A/S33A-RPA2.(A)Cells were synchronized in G1 with 0.5 mM mimosine for 24 hours and then released from the block for 7 hours. Cells were then either mock- or HU-treated(5 mM) for 4 hours and released into fresh media for 40 minutes. (B)Replaced cells were mock or aphidicolin-treated (30M) for 7 hours and then released intofresh media for 40 minutes. (C)Replaced cells were subjected to UV irradiation (70 J/m2 of UVB) and further incubated for 4 or 7 hours. In all cases, cells werethen incubated with 20M BrdU for 30 minutes, fixed and processed as described in Materials and Methods for analysis of BrdU incorporation.

Fig. 7. Cells replaced with the T21A/S33A-RPA2 mutant have a reduced H2AX signalduring replication stress. (A)Schematicindicating the experimental protocol.(B)Three U2-OS clones each were replacedwith wt- or S33A-RPA2. At 3 days post-silencing, cells were either mock-treated orincubated with 1 mM HU for 20 hours.Lysates were then prepared and subjected towestern analysis against H2AX, RPA2(total), S33-P-RPA2, S4-P-S8-P-RPA2 and-actin (loading control). Also included as acontrol is a lysate from U2-OS cells thatwere silenced but not replaced (lane 13).Numbers below H2AX panel indicate theintensity of H2AX staining, normalized to-actin levels. (C)Identical to panel Bexcept that two clones each of cells replacedwith wt- or T21A/S33A-RPA2 were treatedwith 5 mM HU for 5 hours. (D)U2-OS cellswere treated with 1 mM HU for 10 hoursand then fixed. Cells were stained withantibodies against S33-P-RPA2 andH2AX. Approximately 300 cells wereanalyzed by immunofluorescencemicroscopy for the intensity of two stains,and these signals plotted against each other.(E)Similar to panel D, except that cellswere treated with 5 mM HU for 5 hours andassayed for S4-P-S8-P-RPA2 and H2AX.(F,G)Aliquots of cells described in D werestained for S33-P-RPA2 and either H2AX(F) or a modified TUNEL assay to directlydetect DNA ends (G). Cells were thenimaged by immunofluorescencemicroscopy. The right image for each panelshows the combined staining for the twomarkers, with the colors of each markerindicated by the color of letters for eachstain.

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not unexpected because we found previously that S4 and S8phosphorylation is stimulated by S33-P formation (Anantha et al.,2007). Surprisingly, clones replaced with S33A-RPA2 showed a~2.5-fold decrease in the H2AX signal, compared with the H2AXlevels in cells replaced with wt-RPA2 (Fig. 7B, lanes 10-12).Similarly, cells expressing T21A/S33A-RPA2 (Fig. 7C, lanes 7 and8) had an ~fourfold lower H2AX signal compared with wt-RPA2cells (Fig. 7C, lanes 5 and 6), selectively following HU (5 mM)treatment.

As mentioned above, past studies have failed to detect any notableeffects of RPA phosphorylation on checkpoint kinase activationfollowing interphase DNA damage (Morgan and Kastan, 1997;Olson et al., 2006). The greater levels of H2AX are thereforeindicative of more DNA breaks, rather than any defects incheckpoint signaling. We interpret these results to indicate that RPAphosphorylation can stimulate DNA synthesis during replicationstress (see Fig. 5B), but this incorporation heightens the possibilityof DNA breakage. Nevertheless, we demonstrate below that RPAphosphorylation enhances overall cell viability during stress.

To examine the link between the S33-P and H2AX signals ingreater detail, U2-OS cells were subjected to HU treatment (1 mMfor 10 hours), dually stained with antibodies against S33-P andH2AX, and then analyzed by immunofluorescence microscopy.Nearly 300 cells were analyzed for the levels of both the S33-Pand H2AX signals, and these were then plotted against each other(Fig. 7D). As expected, the two signals demonstrated significantcorrelation. Note that the low concentration of HU in this experimentfavor RPA2 phosphorylation up to the L and I levels, rather thanformation of hyperphosphorylated RPA (see Fig. 1A). The S4-P-S8-P signal for hyperphosphorylated RPA was also examinedfollowing treatment with 5 mM HU (Fig. 7E). A similar relationshipwas found between the cellular levels of S4-P-S8-P-RPA2 andH2AX, even though the actual levels of staining showed a weakercorrespondence as compared with the S33-P and H2AX signals.

The correlation between S33-P and H2AX signals also occurredat the level of nuclear foci, because immunofluorescence imagingof HU-treated cells showed that a significant fraction of S33-P-positive foci colocalized with H2AX (Fig. 7F). Because H2AXis only an indirect indicator of DNA damage, we verified thisassociation by an independent assay, using immunofluorescencemicroscopy in combination with a modified TUNEL assay todirectly detect DNA nicks and/or ends. Using this assay, S33-P-positive foci were found also to contain DNA ends, and hence weresites of DSBs or nicks (Fig. 7G). In summary, we conclude thatcells expressing wt-RPA2 support increased DNA synthesis underDNA-replication-stress conditions, yet this synthesis correlates withan increased level of DNA breaks.

RPA phosphorylation-site mutations alter cell-cycle profile andreduce viabilityWe examined whether the loss of RPA2 phosphorylation at the twoconsensus PIKK sites had additional functional consequences.Strong differences were noted when cell proliferation was tested inU2-OS cells replaced with wt-RPA2 or T21A/S33A-RPA2. Whereasclones expressing wt-RPA2 demonstrated a ~threefold expansionwhen assayed 48 hours after plating, the T21A/S33A-RPA2 clonesshowed little change in growth (Fig. 8A). Under replication-stressconditions (5 mM HU for 48 hours), the wt-RPA2 clones hadminimal changes in cell density. By contrast, only 30% of cellsexpressing T21A/S33A-RPA2 survived, with visible accumulationof cell debris (data not shown). These data indicated that the

T21A/S33A mutation had deleterious effects on cell viability. Totest this directly, U2-OS cells replaced with the wt-RPA2 orT21A/S33A-RPA2 variants were either mock-treated or exposed to5 mM HU for 24 hours, and then assayed for apoptosis (Fig. 8B).The level of apoptosis was low in both cell lines under controlconditions, even though an ~sevenfold higher fraction ofT21A/S33A-RPA2 cells were TUNEL-positive (apoptotic),compared with cells replaced with wt-RPA2. Following HUtreatment, a massive fraction of the T21A/S33A-RPA2 cellsunderwent apoptosis (>27%), compared with ~6% of the wt-RPA2cells. Thus, the lack of proper RPA phosphorylation significantlyreduces cell viability, both under control and replication-stressconditions.

The cell-cycle distribution of the different lines was analyzed byFACS, using similar conditions and gating the populations toeliminate cellular debris. In the absence of HU, cells expressingwt-RPA2 or T21A/S33A-RPA2 had similar distributions, althoughwe did reproducibly observe a slightly higher fraction of cells

Fig. 8. Mutation of RPA2 PIKK sites impairs cell growth and alters the cell-cycle profile. (A)Two U2-OS clones each were replaced with either wt- orT21A/S33A-RPA2 (DM, double mutant). At 72 hours post-siRNAtransfection, cells were plated in triplicate to an equivalent relative density(0.35; indicated by dashed line) and then either mock-treated or incubated with5 mM HU. 48 hours later, cells were released from the plate by trypsin-EDTAtreatment and cell count in each plate determined. (B)U2-OS clones replacedwith either wt- or T21A/S33A-RPA2 (DM) were, at 72 hours post-siRNAtransfection, mock-treated or incubated with 5 mM HU for 24 hours. Cellswere then processed to quantify the percentage of cells undergoing apoptosis,using a TUNEL assay. (C,D)Similar aliquots of cells described in A wereeither mock- or HU-treated (5 mM for 48 hours) and then subjected to FACSanalysis for DNA content. Cell-cycle profiles and relative distributions (inpercent of total) in the G1, S and G2-M phases are shown. (E)Overlay ofrepresentative cell distributions from clones replaced with wt-RPA2 (green) orT21A/S33A-RPA2 (red), prepared as described in B and C. Regions of overlapare shown by yellow coloring.

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replaced with the doubly-mutated RPA2 in the S and G2-M phases,as compared with wt-RPA2 cells (Fig. 8C). A similar finding waspreviously made in our studies of cells expressing RPA2 mutatedat the two cyclin-Cdk sites (Anantha et al., 2007). Following a 48-hour treatment with HU, similar fractions of wt-RPA2 andT21A/S33A-RPA2 cells exited S phase and became synchronizedat the G1-S boundary of the subsequent cell cycle (Fig. 8D), similarto the effect of HU on the parental U2-OS cells (Fig. 4E). Twosignificant differences were noted between HU-treated wt-RPA2and T21A/S33A-RPA2 cells. First, the residual G2-M-phase cellsreplaced with T21A/S33A-RPA2 were nearly completely depletedin comparison with the wt-RPA2 cells (i.e. 2 vs 7%, respectively).Second, we found that the distribution of G1-phase cells expressingT21A/S33A-RPA2 was significantly wider than the wt-RPA2 cells,with a clear shift of mutant cells towards the <2N side of the G1peak. This effect is more clearly visible when the two distributionswere overlaid (Fig. 8E). From these data, we suggest that themutation of RPA2 phosphorylation sites causes a defect in the propertransmission of chromosomal DNA to the subsequent cell cycle,following severe replication stress.

DiscussionConditions of DNA-replication stress cause the stalling of one ormore replicative DNA polymerases, resulting in uncoupling of theadvancing DNA helicase from the polymerase machinery. Theseevents increase the steady-state amount of ssDNA, as seen both inthis study and previously by others (Byun et al., 2005; Lopes et al.,2001; Sogo et al., 2002), and lead to activation of the ATR kinase(Zou and Elledge, 2003; Namiki and Zou, 2006). The resultingphosphorylation of RPA by ATR facilitates additionalphosphorylation events on RPA by other kinases, dependent on thedegree of stress and cell-cycle position (Anantha et al., 2007;Anantha et al., 2008; Anantha and Borowiec, 2009) (and this work).We demonstrate that the phosphorylated RPA formed underreplication-stress conditions stimulates DNA synthesis and reducesthe amount of ssDNA generated as assayed by chromatin-boundRPA. In the absence of stress-dependent RPA phosphorylation, cellsundergo high levels of apoptosis and a reduction in the rate of cellproliferation under both normal and stress conditions.

Increased ssDNA accumulation following replication stress wasobserved both in cells expressing RPA2 phosphorylation mutantsand in cells that were down-modulated for ATR. Because the mutantcells also show reduced DNA synthesis during stress, a simpleinterpretation is that these two events are directly coupled. That is,phosphorylated RPA stimulates stress-dependent DNA synthesis byan as-yet-unidentified DNA polymerase on the persistent ssDNA,thereby minimizing ssDNA accumulation (Fig. 9). This model isconsistent with our finding that, in response to a low concentrationof HU, cells can partially recover DNA synthesis from levelsobserved soon after stress onset (see Fig. 4B). Thus, an importantconclusion from this study is that RPA not only signals the presenceof stress and stimulates the activation of ATR, but the accompanyingphosphorylation of RPA facilitates events at the site of the stalledfork that alleviate the consequences of stress. In other words, theinitial modification of RPA in response to stress stimulates changesto the replication and/or repair machineries that eventually lead toa reduction in stress signaling and loss of RPA phosphorylation.

We found that the down-modulation of ATR caused more RPA-chromatin binding, and hence ssDNA accumulation, compared witheither of the two RPA2 mutants tested (see Fig. 5). These dataindicate that, although RPA2 is an important ATR target for

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counteracting the effects of replication stress, it is clearly not theonly target. Recent analysis of ATM and ATR targets that aremodified during genotoxic stress (i.e. IR) found numerous substratesinvolved in DNA replication and repair, including subunits of theMCM (the putative replicative helicase) and RFC (PCNA clamploader) complexes, and the DNA polymerase- catalytic subunit(Matsuoka et al., 2007). It is apparent that alteration of the activitiesof such factors could have complementary effects to RPAphosphorylation in the response to replication stress and theminimization of ssDNA accumulation.

One of the more intriguing findings of our study is that cellsreplaced with wt-RPA2 show higher levels of both H2AX and DNAends during replication stress compared with cells replaced withthe mutant RPA2 variants. Note that RPA phosphorylation does nothave significant effects on checkpoint activation in interphase cells(Morgan and Kastan, 1997; Olson et al., 2006), indicating that thisresult is not a result of defective cell-cycle checkpoints. In ourexperiments, replication stress was induced by treatment with theribonucleotide-reductase inhibitor HU, which reduces cellulardNTP pools (Yarbro, 1992). It is possible that the replicationcomplex catalyzing DNA synthesis under such conditions is proneto undergo collapse and further damage, including DSBs.Alternatively, the observed increase in DNA breaks might not bereflective of true damage, but rather a normal event in the processingof stalled DNA-replication forks. Regardless of the cause, it is clearthat the events stimulated by RPA phosphorylation significantlyincrease cell viability both under stress and non-stress conditions.

Although we interrogated the role of RPA phosphorylation in thepresence of HU, formation of S33-P-RPA2 was also detected innon-stressed cells, particularly at the S-G2-phase transition of thecell cycle. These data presumably reflect the gradual accumulation

Fig. 9. Model indicating the roles of RPA phosphorylation in response toDNA-replication stress. See text for details.

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of stalled DNA-replication forks as cells progress through S phase.S33-P-RPA2 colocalized with the BLM protein, as well as withsites of DNA synthesis, indicating that the molecular eventsregulated by phosphorylated RPA probably occur within nuclearrepair foci. That said, mutation of the RPA2 phosphorylation siteswas not seen to have any dramatic effects on BLM-foci formation(data not shown), indicating that BML localization to sites of DNA-replication stress is independent of modification of these RPA2residues.

Lastly, our data suggest that abnormal RPA phosphorylationincreases genomic instability. We reproducibly observed thatchronically stressed cells replaced with the T21A/S33A-RPA2mutant yielded a distribution of G1 cells with a broader DNAdistribution than that seen with wt-RPA2 cells. The mutantdistribution was skewed to the <2N side of the G1 peak, potentiallyindicative of a significant loss of chromosomal DNA. Althoughidentification of the molecular basis for the effect is ongoing, twopossibilities come to mind. First, replication forks stalled by HUtreatment are not properly resuscitated in cells expressing theT21A/S33A-RPA2 mutant. These stalled replication structures aresubsequently repaired by error-prone pathways, leading to loss andgain of DNA from chromosomal DNA regions and therebygenerating the distribution of G1 cells we observe by FACS.Alternatively, recent work by our laboratory found a role for RPAphosphorylation in stimulating cellular transit through a damagedmitosis (Anantha and Borowiec, 2009; Anantha et al., 2008). Wefound that mitotic DNA damage in cells replaced with an RPA2phosphorylation mutant (i.e. in which the two cyclin-Cdk sites atS23 and S29 were changed to alanines) had an increased failure ofcytokinesis compared with cells replaced with wt-RPA2. These dataraise the possibility that defective RPA2 phosphorylation reducesthe fidelity of mitotic chromosomal segregation under conditionsof severe DNA-replication stress, thereby increasing cellularaneuploidy. Further investigation will be required to analyze thespectrum of mutagenic events that our data suggest is induced bydefective RPA phosphorylation under conditions of DNA-replicationstress.

Materials and MethodsCell cultureU2-OS cell lines were cultured in McCoy’s medium supplemented with 10% FBS.Cell lines positive (MO59K) or null (MO59J) for the DNA-PK catalytic subunit werecultured in DME/F12 (1:1; v:v) containing 10% FBS. Human fibroblast cell linespositive (GM637) or null (GM5849) for ATM (Coriel Institute, Camden, NJ) werecultured in McCoy’s medium with 15% FBS. For synchronization experiments, cellswere arrested in prometaphase by treatment with 100 ng/ml nocodazole (Sigma-Aldrich) for 8 hours and the mitotic cells isolated by shake-off.

All experiments involved the use of untagged RPA2. To generate the S33A-RPA2and T21A/S33A-RPA2 variants, the parental pRetro-Off retroviral vector (Clontech)containing the wt-RPA2 coding sequence (Anantha et al., 2007) was subjected tosite-directed mutagenesis using the Stratagene QuikChange Kit. U2-OS clonescontaining the integrated pRetro-Off RPA2 expression vector were prepared andisolated as described by Anantha et al. (Anantha et al., 2007). For RPA2 replacement,the appropriate U2-OS clone was first grown for 24-48 hours in medium lackingdoxycycline to induce expression of the ectopic RPA2. Cells were then transfected(Hiperfect; Qiagen) with an siRNA molecule that selectively targets the endogenousRPA2 mRNA (Anantha et al., 2007), and the cells then tested 72 hours post-transfection. A representative western analysis of RPA2 protein levels followingendogenous RPA2 silencing and ectopic RPA2 induction is shown (supplementarymaterial Fig. S4).

Immunoblotting and antibodiesFor western analysis, cells were directly lysed in SDS-PAGE sample buffer and thelysate proteins were separated by SDS-PAGE. Proteins were immobilized onto Protrannitrocellulose membranes (0.2-m pore size). The antibodies used in this study wereagainst general RPA2 (NeoMarkers), H2AX (Upstate), S4-P-S8-P-RPA2 and S33-P-RPA2 (Bethyl Laboratories), T21-P-RPA2 (Abcam), ATR (Novus Biologicals),

PML (Abnova), BLM (Santa Cruz), Rad17 (Santa Cruz), and S317-P-Chk1 (BethylLaboratories). The S29-P-RPA2 antibody was custom-synthesized as described(Anantha et al., 2007).

Immunofluorescence microscopyFor detection of chromatin-bound RPA, S33-P-RPA2 or BLM, cells were firstextracted with CSK buffer (10 mM HEPES-KOH, pH 7.4, 300 mM sucrose, 100mM NaCl, 3 mM MgCl2) containing 0.5% (v/v) Triton X-100 for 5 minutes on ice.Cells were then washed with CSK buffer lacking Triton X-100 and fixed with 4%(w/v) paraformaldehyde for 30 minutes. To quantitate chromatin-bound RPA, cellswere stained with a mouse anti-RPA2 primary antibody and an anti-mouse Cy3-conjugated secondary antibody. A total of seven fields containing 300-400 cells wereanalyzed for each experiment, using IPLab software (BD Biosciences) to quantitatethe Cy3 (i.e. RPA2) signal within the DAPI-stained nuclear region. In all cases, theaverage background signal in cells considered to be RPA2-negative was subtractedto determine the ‘true’ RPA2 signal. When ‘RPA2-positive’ cells were assayed, onlythose cells with RPA2 signals above background were analyzed.

BrdU incorporationCells were incubated in media containing 20 M BrdU for 20 minutes and 1 hourfor untreated and HU-treated cells, respectively, to yield a BrdU signal in the dynamicrange. Cells were briefly extracted with CSK buffer containing 0.5% (v/v) Triton X-100 and fixed as above. Cells were then treated with 2 M HCl for 50 minutes andsubjected to immunofluorescence microscopy using rat anti-BrdU primary antibodies(Harlan Sera-Lab) and Alexa-Fluor-499-conjugated secondary antibodies (Invitrogen).Photographs were taken on Zeiss AxioCam camera with a 40� objective and analyzedas described above for RPA2-signal quantitation. See supplementary material Fig.S3 for a representative experiment.

BiodU incorporationU2-OS cells were arrested with nocodazole and the mitotic cells then released intonocodazole-free medium for 18 hours. Cells (on 18-mm coverslips) were washed at37°C with ABC buffer (50 mM KCl, 20 mM creatine phosphate, pH 7.4, 3 mMMgCl2, 1 mM ATP and 11 mM K2HPO4) and then incubated at 37°C for 5-10 minuteswith ABC buffer containing 250 M each of dNTPs (dATP, dCTP and dGTP), 100M each of rNTPS (UTP, CTP and GTP), 50 M biotin-11-dUTP, 0.5 U/lstreptolysin-O (Sigma-Aldrich) and 20 g/ml creatine phosphokinase. Cells werewashed with cold CSK buffer, incubated on ice for 2 minutes with CSK buffercontaining 0.5% (v/v) Triton X-100 and then washed again with cold CSK buffer.Cells were fixed with 4% (w/v) paraformaldehyde for 30 minutes at room temperature,washed three times with PBS and then stained with a goat anti-biotin primary antibody(Pierce).

To measure apoptosis, two U2-OS clones were each treated to replace endogenousRPA2 with wt-RPA2 or T21A/S33A-RPA2. At 72 hours post-siRNA transfection,cells were mock-treated or incubated with 5 mM HU for 24 hours. Apoptosis wasthen assayed using the In Situ Cell Death Detection Kit (Roche), as per themanufacturer’s instructions.

Flow cytometryFACS analysis for DNA content was performed as described by Anantha et al.(Anantha et al., 2007). FACS was performed on a Becton Dickinson flow cytometerusing the Cell Quest software. Cell-cycle analysis was performed using the Mod-Fitsoftware.

We thank Priscilla Maldonado for critical reading of the manuscript.This work was supported by an Exceptional Project Award Grant fromthe Breast Cancer Alliance, NIH grant R01 GM083185, the New YorkUniversity Cancer Institute, the Rita J. and Stanley KaplanComprehensive Cancer Center, and National Cancer Institute GrantP30CA16087. Deposited in PMC for release after 12 months.

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