Ddc2ATRIP promotes Mec1ATR activation at RPA-ssDNA tracts

The DNA damage checkpoint response is controlled by the phosphatidylinositol 3-kinase-related kinases (PIKK), including ataxia telangiectasia-mutated (ATM) and ATM and Rad3-related (ATR). ATR forms a complex with its partner ATRIP. In budding yeast, ATR and ATRIP correspond to Mec1 and Ddc2, respectively. ATRIP/Ddc2 interacts with replication protein A-bound single-stranded DNA (RPA-ssDNA) and recruits ATR/Mec1 to sites of DNA damage. Mec1 is stimulated by the canonical activators including Ddc1, Dpb11 and Dna2. We have characterized the ddc2-S4 mutation and shown that Ddc2 not only recruits Mec1 to sites of DNA damage but also stimulates Mec1 kinase activity. However, the underlying mechanism of Ddc2-dependent Mec1 activation remains to be elucidated. Here we show that Ddc2 promotes Mec1 activation independently of Ddc1/Dpb11/Dna2 function in vivo and through ssDNA recognition in vitro. The ddc2-S4 mutation diminishes damage-induced phosphorylation of the checkpoint mediators, Rad9 and Mrc1. Rad9 controls checkpoint throughout the cell-cycle whereas Mrc1 is specifically required for the S-phase checkpoint. Notably, S-phase checkpoint signaling is more defective in ddc2-S4 mutants than in cells where the Mec1 activators (Ddc1/Dpb11 and Dna2) are dysfunctional. To understand a role of Ddc2 in Mec1 activation, we reconstituted an in vitro assay using purified Mec1-Ddc2 complex, RPA and ssDNA. Whereas ssDNA stimulates kinase activity of the Mec1-Ddc2 complex, RPA does not. However, RPA can promote ssDNA-dependent Mec1 activation. Neither ssDNA nor RPA-ssDNA efficiently stimulates the Mec1-Ddc2 complex containing Ddc2-S4 mutant. Together, our data support a model in which Ddc2 promotes Mec1 activation at RPA-ssDNA tracts.


Introduction
Chromosomes are constantly damaged by exogenous and endogenous threats [1]. The repair of damaged chromosomes is therefore critical for maintaining genome stability. The DNA damage response consists of multiple pathways controlled by the phosphatidylinositol 3-kinase-related kinases (PIKK) [2,3]. These kinases include DNA-dependent protein kinase catalytic subunit (DNA-PKcs), ataxia telangiectasia-mutated (ATM), and ATM and Rad3-related (ATR). Although all these PIKKs respond to DNA damage, their DNA damage specificities are different. ATM and DNA-PKcs are activated by double-stranded DNA breaks (DSBs), whereas ATR responds to various types of DNA lesions with single-stranded DNA (ssDNA) [4,5].
We have identified a separation-of-function ddc2 mutation (ddc2-S4) that causes defects in Mec1 activation but does not affect Mec1 recruitment [49]. However, it is not known how Ddc2-dependent Mec1 activation triggers checkpoint signaling. Moreover, the underlying mechanism of Ddc2-dependent Mec1 activation remains to be determined. To understand the significance of Ddc2-dependent Mec1 activation, we further characterized the ddc2-S4 mutation by carrying out genetic experiments. We found that the ddc2-S4 mutation impaired Rad9 and Mrc1 phosphorylation after DNA damage, consistent with the previous observation that the ddc2-S4 mutation is defective in checkpoint activation throughout the cell cycle [49]. The Ddc1-Dpb11 axis and Dna2 contribute to Mec1 activation in S phase [26]. We also found that S-phase checkpoint signaling is more significantly defective in ddc2-S4 mutants than in ddc1and dna2-deficient mutants. Thus, Ddc2 appears to promote Mec1 activation independently of Ddc1/Dpb11 and Dna2 function. We further examined the effect of ddc2-S4 mutation on kinase activity of Mec1-Ddc2 using an in vitro reconstitution system. Whereas ssDNA stimulated Mec1 activity, RPA did not. However, RPA was found to promote ssDNA-dependent Mec1 activation. The Mec1-Ddc2 complex containing Ddc2-S4 mutant (Mec1-Ddc2-S4) exhibited a basal kinase activity in vitro. However, neither ssDNA nor RPA-ssDNA efficiently stimulated Mec1-Ddc2-S4. Our results support a model in which Ddc2 promotes Mec1 activation through ssDNA recognition.

Results
The ddc2-S4 mutation impairs Rad9 phosphorylation and recruitment after DNA damage Mec1 phosphorylates the Rad9 checkpoint mediator to promotes Rad9-Rad53 interaction, which is essential for Rad53 activation [30,31]. The ddc2-S4 mutation confers defects in DNA damage checkpoint activation and damage-induced Rad53 phosphorylation at G2/M [49]. To understand the significance of Ddc2-dependent Mec1 activation, we first examined the effect of ddc2-S4 mutation on Rad9 phosphorylation after DNA damage at G2/M (Fig 1A). Cells expressing HA-tagged Rad9 protein (Rad9-HA) were arrested with nocodazole and exposed to methyl methanesulfonate (MMS). Cells were then subjected to immunoblotting analysis with anti-HA antibodies to monitor Rad9 phosphorylation (Fig 1A). Rad9 underwent phosphorylation in wild-type cells after MMS treatment. Phosphorylation was decreased in ddc2-S4 mutants but less significantly compared with in ddc2Δ mutants. Thus, the ddc2-S4 mutation impairs Rad9 phosphorylation after DNA damage.
One explanation for Rad9 phosphorylation defect in ddc2-S4 mutants is that Rad9 does not efficiently localize to sites of DNA damage. We next examined the effect of ddc2-S4 mutation on Rad9 accumulation at sites of DNA damage by chromatin immunoprecipitation (ChIP) assay ( Fig 1B). In budding yeast, HO endonuclease introduces a sequence-specific DSB. We used an experimental system in which cells carry the GAL-HO plasmid and contain a single HO cleavage site at the ADH4 locus [49]. In this system, HO endonuclease, expressed after incubation with galactose, generates a single DSB at the ADH4 locus. Cells expressing HAtagged Rad9 (Rad9-HA) protein were transformed with the GAL-HO plasmid. Transformants were grown initially in sucrose to repress HO expression, and then transferred to medium containing nocodazole to arrest at G2/M. After arrest, galactose was added to induce HO expression. Cells were then analyzed by the ChIP assay using anti-HA antibodies. Rad9 , ddc2-S4 (HB10) or ddc2Δ (KSC1536) cells expressing Rad9-HA were treated with nocodazole to arrest at G2/M. Cells were then exposed to 0.05% MMS. Cells were collected at the indicated time points, and extracts were subjected to immunoblotting analysis with anti-HA antibodies or anti-tubulin antibodies. (B) Effect of ddc2-S4 or ddc2Δ on Rad9 localization to a HO-induced DSB. Wild-type (HB09), ddc2-S4 (HB10) or ddc2Δ (KSC1536) cells expressing Rad9-HA were transformed with the YCpA-GAL-HO plasmid. Transformed cells were grown in sucrose and treated with nocodazole. After arrest at G2/M, the culture was incubated with galactose for 3 hr to induce HO expression, while half of the culture was maintained in sucrose to repress HO expression. Cells were subjected to chromatin immunoprecipitation with anti-HA antibodies. Association of Rad9 with a HO-induced DSB was analyzed by realtime PCR. Relative enrichment was determined from three independent experiments. The error bars indicate standard deviation.
Rad9 limits the Sae2-and Sgs1-dependent pathway and interferes with DNA end resection [50]. We next addressed whether the ddc2-S4 mutation affects the kinetics of DNA end resection. To this end we monitored ssDNA generation at two EcoRI restriction sites near the HO cleavage site (at 0.8 kb or 5.8 kb from the site) by a quantitative PCR-based method [51] (Fig  2A). PCR amplifies only resected DNA because the EcoRI restriction enzyme can cleave unprocessed DNA (S1 Fig). The PCR amplification, normalized to the efficiency of HO cleavage, corresponds to the rate of DNA end resection [51]. The ddc2-S4 mutation did not significantly affect DNA end resection ( Fig 2B). RPA, consisting of Rfa1, Rfa2 and Rfa3, binds to ssDNA tracts [8]. We also examined whether the ddc2-S4 mutation affects RPA accumulation near the DSB (Fig 2C). No apparent defect in Rfa2 association was observed in ddc2-S4 mutants. These result support the previous finding that the ddc2-S4 mutation does not affect Mec1 localization to sites of DNA damage [49]. Our results are also consistent with the current view that Mec1 positively controls DNA end resection although it promotes Rad9 accumulation at sites of DNA damage [52].
Mec1 phosphorylates Ddc1 to recruit Rad9 near sites of DNA damage through the Dpb11 scaffold [39,44,48]. We next examined the effect of ddc2-S4 mutation on Ddc1 phosphorylation after DNA damage ( Fig 3A). Cells expressing HA-tagged Ddc1 protein were treated as above and subjected to immunoblotting analysis to monitor Ddc1 phosphorylation. Ddc1 phosphorylation was decreased in ddc2-S4 mutants compared with wild-type cells. Mec1 and Ddc1 are independently recruited to sites of DNA damage [53,54]. We confirmed that the ddc2-S4 mutation has no impact on Ddc1 accumulation at sites of DNA damage ( Fig 3B). Thus, the ddc2-S4 mutation impairs Ddc1 phosphorylation that promotes Ddc1-Dpb11-Rad9 assembly at sites of DNA damage [39,44,48].

The ddc2-S4 mutation is defective in S phase checkpoint signaling
Mec1 phosphorylates the Mrc1 checkpoint mediator that is essential for Rad53 activation during S phase [32,35]. Notably, Mrc1-dependent Rad53 activation does not require Rad9 or Ddc1/Dpb11 function [55]. We further examined the effect of ddc2-S4 mutation on Mrc1 phosphorylation after DNA damage ( Fig 4A). Wild-type, ddc2-S4 or ddc2Δ cells expressing HA-tagged Mrc1 protein were arrested in G1 with α-factor and released into medium containing MMS. Cells were harvested at the indicated times after release. We confirmed that cells remained within S phase at the time point after release (Fig 4A). Mrc1 phosphorylation was decreased in ddc2-S4 mutants but less significantly than in ddc2Δ mutants. Thus, Ddc2-dependent Mec1 activation also controls the Mrc1-dependent checkpoint pathway during S phase.
The Ddc1-Dpb11 axis and Dna2 control DNA damage checkpoints in S phase [26]. Two substitution mutations at the N-terminus of Dna2 (dna2-W128A, Y130A; hereafter called dna2-AA) abrogate its checkpoint function [26]. We next compared Rad53 phosphorylation in ddc2-S4, ddc2Δ and ddc1Δ dna2-AA mutants in S phase ( Fig 4B). Cells expressing HA-tagged Rad53 protein were arrested at G1 and released into MMS or hydroxyurea (HU) as above. Rad53 phosphorylation was decreased in ddc1Δ dna2-AA mutants. However, a more significant defect was observed in ddc2-S4 or ddc2Δ mutants ( Fig 4B). Our results agree with the previous observation that ddc1Δ dna2-AA cells are less defective in Rad53 activation than mec1Δ cells [56]. We next compared DNA damage sensitivity of ddc2-S4 and ddc1Δ dna2-AA mutants. While the ddc2-S4 and the ddc1Δ dna2-AA mutation caused similar sensitivities to MMS, ddc2-S4 mutants were more sensitive to HU than ddc1Δ dna2-AA mutants ( Fig 4C).
We further addressed whether the residual checkpoint activation in ddc1Δ dna2-AA mutants depends on Mec1 function in S phase ( Fig 4D). The introduction of a mec1Δ mutation decreased damage-induced Rad53 phosphorylation in ddc1Δ dna2-AA mutants. Together, these results suggest that Ddc2 promotes Mec1 activation through a Ddc1/Dpb11/Dna2-independent mechanism. The ddc2-S4 mutation is defective in RPA phosphorylation in vivo and in vitro Mec1 phosphorylates two subunits of RPA, Rfa1 and Rfa2, in response to DNA damage [57,58] although the significance of RPA phosphorylation in checkpoint signaling is not fully understood [59]. We examined the effect of ddc2-S4 mutation on Rfa2 phosphorylation after DNA damage (Fig 5A). Wild-type and ddc2-S4 mutants were arrested with nocodazole at G2/ M and exposed to MMS. Cells were then analyzed by immunoblotting with anti-Rfa2 antibodies. We found that damage-induced Rfa2 phosphorylation was decreased in ddc2-S4 mutants.   The ddc2-S4 mutation does not affect Mec1 or RPA accumulation at sites of DNA damage [49] ( Fig 2C). Thus, the ddc2-S4 mutation impairs Rfa2 phosphorylation in vivo.
Mec1/ATR phosphorylates RPA efficiently in the presence of ssDNA in vitro [21,58,60]. However, whether RPA or ssDNA modulates ATR/Mec1 activity remains to be determined. To understand a role of Ddc2 in Mec1 activation, we reconstituted an in vitro system using purified Mec1-Ddc2 and RPA proteins. We have purified Mec1-Ddc2 and Mec1-Ddc2-S4 through a two-step affinity chromatography after overexpressing FLAG-tagged Mec1 and Histagged Ddc2 protein in yeast cells (S2 and S3 Figs). In agreement with the previous studies [21,58], Mec1-Ddc2 phosphorylated RPA efficiently in the presence of ssDNA (S4 Fig). The ddc2-S4 mutation contains two substation mutations (K263E, H382Y). K263 is implicated in Mec1-Ddc2 homodimerization [61] whereas H382 is in close proximity to the 177 KKRK 180 motif which is involved in DNA binding [61,62] (Fig 5B, S5 Fig). Ddc2/ATRIP itself interacts weakly with ssDNA but RPA stimulates ssDNA binding of Ddc2/ATRIP [17,62]. We determined the effect of ddc2-S4 mutation on the interaction of Mec1-Ddc2 with ssDNA or RPA-ssDNA by a pull-down assay. Mec1-Ddc2 and Mec1-Ddc2-S4 were found to bind similarly to ssDNA or RPA-ssDNA in vitro (Fig 5C), consistent with the observation that the ddc2-S4 mutation does not affect Mec1 accumulation at sites of DNA damage [49]. We further examined the effect of ddc2-S4 mutation on Ddc2-ssDNA binding. We prepared MBPfused Ddc2 or Ddc2-S4 from E. coli and further examined whether they interact with oligonucleotides in the presence or absence of RPA (Fig 5D). MBP-Ddc2 and MBP-Ddc2-S4 interacted similarly with ssDNA and RPA-ssDNA. MBP alone did not exhibit oligonucleotide binding (Fig 5D). Thus, the ddc2-S4 mutation does not affect its own ssDNA-or RPA-ssDNAbinding abilities.

ssDNA, but not RPA, stimulates Mec1 activity in a Ddc2-dependent manner
We addressed whether RPA or ssDNA regulates Mec1 activity using GST-Rad53 as a substrate (Fig 6). GST-Rad53 lacks the N-terminal kinase domain of Rad53; therefore, no phosphoincorporation into GST-Rad53 was observed without Mec1-Ddc2 [49] (Fig 6A and 6B). We first tested the effect of various concentrations of RPA on Mec1 kinase activity. RPA had no significant impact on Mec1 activity using GST-Rad53 as a substrate (Fig 6A).
We next investigated the effect of ssDNA on Mec1 activity using various lengths (20, 40, 60 or 80 mer) of oligo(dT) (Fig 6B and 6C). Although no apparent stimulation was observed with a 20 mer oligo(dT) (oligo(dT) 20 ), longer oligonucleotides, oligo(dT) 40 , oligo(dT) 60 and oligo (dT) 80 , were found to increase Mec1 activity more efficiently. Similar activation was observed with oligo(dT) 80 and an 80-mer biotinylated oligonucleotide containing all DNA bases (biooligo(dN) 80 ) (Fig 6C). We note that biotinylation of oligonucleotide has no impact on Mec1 activation (S6 Fig). High concentrations of 80-mer oligonucleotides (125 nM) were required to reach maximum activation compared with the concentration of Mec1 (5 nM). We further tested whether longer ssDNA stimulates Mec1 more strongly using 5 kb FX174 phage ssDNA (Fig 6D). A single-stranded form of FX174 phage stimulated Mec1 at much lower concentrations compared with 80-mer oligonucleotides. The maximum activation obtained with FX (3 fold) was slightly higher than that of 80-mer oligonucleotides (2.5 fold) (Fig 6C  and 6D). These results indicate that ssDNA stimulates Mec1 in a dosage-dependent and length-dependent manner.
Thus, ssDNA appears to stimulate Mec1 activity through a Ddc2-dependent mechanism.

RPA modulates ssDNA-dependent Mec1 activation in vitro
We next determined the combination effect of RPA and ssDNA on Mec1 kinase activity using GST-Rad53 as a substrate. RPA prompts ssDNA binding of Mec1-Ddc2 or Ddc2 (Fig 5C and  5D) whereas ssDNA stimulates Mec1 kinase activity (Fig 6). We thus expected that RPA promotes ssDNA-dependent Mec1 activation. However, Mec1-Ddc2 was found to interact with RPA independently of ssDNA in vitro, in agreement with the current view that the N-terminus of Rfa1 interacts directly with the N-terminus of Ddc2 [63,64] (Fig 7A). Therefore, RPA by itself could compete with ssDNA-bound RPA for Mec1-Ddc2 binding (S7 Fig). Moreover, RPA is a good substrate of Mec1 (S4 Fig); that is, RPA could compete with GST-Rad53 as a Mec1 substrate [65]. Hence, high RPA concentrations could have negative impacts on ssDNAdependent Mec1 activation in vitro.
We first examined the effect of RPA on Mec1 activation with a low concentration of 80-mer oligonucleotides (12.5 nM) (Fig 7B). We note that only weak Mec1 activation was observed at this concentration (Fig 6B and 6C). We incubated oligonucleotides with various concentrations of RPA and subsequently with Mec1-Ddc2 to initiate the kinase reaction. Lower concentrations of RPA enhanced Mec1 kinase activity whereas higher concentrations of RPA attenuated (Fig 7B and 7D). We further tested the effect of RPA on Mec1 activation with a higher concentration of oligonucleotide (125 nM) ( Fig 7C). Again, lower concentrations of RPA stimulated Mec1 activity whereas higher concentrations of RPA attenuated. These results are consistent with the hypothesis that RPA promotes ssDNA-dependent Mec1 activation although high RPA concentrations have negative impacts on ssDNA-dependent Mec1 activation in vitro. The stimulatory effect of RPA was less pronounced when Mec1-Ddc2 was incubated with a higher concentration of oligonucleotides (Fig 7C and 7D), consistent with the observation that ssDNA, but not RPA, stimulates Mec1-Ddc2 activity (Fig 6).

Discussion
Previous studies have established the model in which ATRIP/Ddc2 interacts with RPA-coated ssDNA and recruits ATR/Mec1 to sites of DNA damage [4,5]. However, Ddc2 appears to by ssDNA. Kinase reactions were carried out with Mec1-Ddc2 or Mec1-Ddc2-S4 (5 nM) using various concentrations of bio-oligo(dN) 80 . Incorporation of 32 P into GST-Rad53 was analyzed as in A. The error bars indicate standard deviation from three independent experiments. https://doi.org/10.1371/journal.pgen.1008294.g006 ssDNA-dependent activation of Mec1  [49]. In this study we have further characterized the ddc2-S4 mutation by carrying out genetic and biochemical experiments. The ddc2-S4 mutation causes defects in phosphorylation and accumulation of the Rad9 checkpoint mediator at sites of DNA damage. The ddc2-S4 mutation also confers a defect in phosphorylation of the S-phase specific Mrc1 checkpoint mediator. The Ddc1-Dpb11 axis and Dna2 contribute to Mec1 activation in S phase [26]. Notably, the ddc2-S4 mutation causes a more significant defect in S phase checkpoint signaling than the ddc1Δ dna2-AA mutation. Thus, Ddc2 controls Mec1 activation through a Ddc1/Dpb11/Dna2-independent mechanism. We further examined the effect of ddc2-S4 mutation on Mec1 kinase activity using an in vitro reconstitution system. ssDNA, but not RPA, stimulates Mec1-Ddc2 kinase activity. However, RPA can promote ssDNA-dependent Mec1 activation. Neither ssDNA nor RPA-ssDNA stimulates Mec1-Ddc2-S4 effectively. Our data support a model in which Ddc2 increases Mec1 kinase activity upon ssDNA recognition.
The ddc2-S4 mutation causes a more significant defect in S phase checkpoint activation than the ddc1Δ dna2-AA mutation. Notably, the residual checkpoint activation in ddc1Δ dna2-AA cells largely depends on Mec1 function. These results suggest that Ddc2 promotes Mec1 activation through a Ddc1/Dbp11/Dna2-independent mechanism. Mec1 phosphorylates the Mrc1 checkpoint mediator and activates Rad53 in S phase [32,35]. Notably, Mrc1-dependent Rad53 activation does not require Rad9 or Ddc1/Dpb11 function [55]. Recent evidence suggests that the dna2-AA mutation affects DNA replication or repair function rather than checkpoint activation [27]. Thus, Ddc2-dependent Mec1 activation appears to play a key role in the stimulation of the Mrc1 checkpoint pathway during S phase.
The ddc2-S4 mutation causes a defect in ssDNA-dependent Mec1 activation. Then how does ssDNA stimulate Mec1 kinase activity? Mec1-Ddc2 forms a dimer of heterodimers through multiple interfaces including the PIKK regulatory domain (PRD) [67]. Interestingly, the PRD is closely positioned near the catalytic and activation loop at the kinase RPA or ssDNA. Experiments were carried out three times and the representative result is shown. (D) Effects of RPA concentrations on ssDNA-dependent Mec1 activation. Kinase activities of Mec1-Ddc2 using various concentrations of RPA in the presence of bio-oligo (dN) 80 (12.5 nM or 125 nM) were normalized to that observed with Mec1-Ddc2 alone (See Fig 7B or 7C, respectively). Relative kinase activities with various concentrations of RPA in the absence of ssDNA are also included (see Fig 6A). The error bars indicate standard deviation from three independent experiments. (E) Effect of ddc2-S4 mutation on Mec1 activation by RPA and ssDNA. Kinase reactions were carried out with Mec1-Ddc2 or Mec1-Ddc2-S4 (5 nM) using various concentrations of RPA in the absence or the presence of biooligo(dN) 80 (12.5 nM). Incorporation of 32 P into GST-Rad53 was analyzed as in Fig 6A. The error bars indicate standard deviation from three independent experiments. https://doi.org/10.1371/journal.pgen.1008294.g007 ssDNA-dependent activation of Mec1 domain, thereby blocking kinase activity and substrate entry [61,67]. We propose that ssDNA binding of Ddc2 triggers conformation changes of the Mec1-Ddc2 homodimer, which could open up the catalytic active site (Fig 8A). The ddc2-S4 mutation carries two substation mutations (K263E, H382Y). K263 contributes to Mec1-Ddc2 homodimerization [61] (S5 Fig) whereas H382 is positioned near the putative DNA binding ( 177 KKRK 180 ) region [61] (Fig 5B). Thus, the ddc2-S4 mutation may affect conformation changes of Mec1-Ddc2 homodimer upon ssDNA binding. Mec1 phosphorylates Mec1 activators (Ddc1, Dpb11 or Dna2) after DNA damage [66,68,69]. Conformation changes of the kinase domain could ssDNA-dependent activation of Mec1 therefore enhance binding of Mec1 activators to its own kinase domain. Consistent with the view, Dpb11 has been shown to activate Mec1 more strongly in the presence of RPA and ssDNA [22].
RPA at lower concentrations promotes ssDNA-dependent Mec1 activation in vitro. Ddc2 and RPA recognize ssDNA through different mechanisms; the KKRK motif of Ddc2 is implicated in DNA binding [62] whereas RPA utilizes its own DNA binding domain (DBD) [8] (Fig  8B). However, the N-terminus of Ddc2 interacts with RPA independently of ssDNA although other domains of Ddc2 may be involved in RPA interaction [49,63,64] (Fig 8B). Thus, RPA-Ddc2 interaction could stimulate Ddc2-ssDNA binding by providing an additional ssDNA-binding interface, thereby boosting ssDNA-dependent Mec1 activation. However, we cannot fully exclude the possibility that RPA, once coated on ssDNA, acquires the ability to directly stimulate Mec1 activity. RPA at higher concentrations attenuates ssDNA-dependent Mec1 activation in vitro. One explanation is that RPA by itself competes with RPA-ssDNA for Mec1-Ddc2 binding. Alternatively, there would be substrate competition between RPA and in vitro substrates. At the moment, it remains to be determined which property of RPA downregulates ssDNA-dependent Mec1 activation in vitro. Previous in vitro studies have shown that RPA-ssDNA has no apparent impact on ATR-ATRIP/Mec1-Ddc2 kinase activity; however, the effect of different RPA concentrations has not been extensively investigated [21,22,63,70]. RPA not only stabilizes ssDNA but also stimulates various repair processes [8,[71][72][73][74][75]. Moreover, RPA binding to ssDNA is highly dynamic using different binding modes [76,77]. Interestingly, RPA depletion modulates ssDNA generation and Mec1 activation differently [74]. Thus, dynamic interactions between ssDNA, RPA and Mec1-Ddc2 might be important for efficient Mec1 activation. A high-resolution structure of Mec1-Ddc2 has been recently reported [61]. However, the structure and dynamics of how Mec1-Ddc2 and RPA assemble on ssDNA remain to be elucidated.
In summary, we have shown that Ddc2 promotes Mec1 activation independently of Ddc1, Dpb11 and Dna2. We have also provided evidence supporting that Ddc2 promotes Mec1 activation through ssDNA recognition. ATR/Mec1 recognizes RPA-ssDNA and controls many cellular activities during DNA replication and repair [4,5]. Our studies thus provide insight into how RPA-containing DNA structures modulate ATR/Mec1 activation, and suggest that ATR/Mec1, like DNA-PK and ATM/Tel1, is activated upon the recruitment to sites of DNA damage.

Strains and plasmids
pRS424-GAL-FLAG-MEC1 is a high-copy plasmid version of YCp/pRS316-GAL-FLAG-MEC1 [78]. The GAL1-GAL10 promoter region was amplified by PCR with the primer pair 3016 and 3017, fusing a sequence encoding MEHHHHHH to the GAL1 promoter. The PCR product was cleaved with EcoRI and MluI. The DDC2 or ddc2-S4 coding sequence [49] was amplified by PCR with the primer pair KS460 and KSX001, fusing a HA epitope to the N-terminus of Ddc2 or Ddc2-S4, respectively. The PCR product was cleaved with MluI and SalI. The EcoRI-MluI and the MluI-SalI fragments were cloned into YEplac195, generating YEp195-GAL-His-HA-Ddc2 or YEp195-GAL-His-HA-Ddc2-S4, respectively. The YCp-T-Rad53-HA plasmid has been described [49]. The dna2-W128A, Y130A (dna2-AA) mutation [26] was integrated into the own locus after PCR fusion [79] using primers KS2943, KS2944, KS2955 and KS2946. The MRC1-HA::TRP1 construct was generated by a PCR-based method [80] using the primer pair KS3649 and KS3650. The strains used in this study are listed in S1 Table. DNA end resection assay Quantitative PCR analysis of DNA end resection was performed as described previously [51]. HO cleaves the HO cut site and generate a DSB. The DNA was digested with the EcoRI restriction enzyme that cleaves the amplicons at 0.8 kb and 5.8 kb from the DSB, but not in the SMC2 control region. The ssDNA percentage over total DNA was calculated using the following formula: % ssDNA = [100/[(1+2 ΔCt )/2]]/f, in which ΔCt values are the difference in average cycles between digested template and undigested template of a given time point and f is the HO cut efficiency [51]. HO cutting efficiency was determined as described [81]. The oligonucleotides used are listed in S2 Table. Protein purification of Mec1-Ddc2 and Mec1-Ddc2-S4 The yeast strain (mec1Δ ddc2Δ sml1Δ; KSC3218) was transformed with pRS424-GAL-FLAG-MEC1 and YEp195-GAL-His-HA-DDC2 or YEp195-GAL-His-HA-DDC2-S4. Transformed cells were grown in sucrose media (2% sucrose 0.05% glucose) to a log-phase and incubated with 2% galactose for 5 hr to induce expression from the GAL promoter. Crude extracts were prepared from 10 gram of cells in 50 ml of buffer A (20 mM Tris-HCl [pH 8], 10% glycerol, 3 mM DTT, 0.1% Triton X-100) containing 1 mM EDTA, 100 mM NaCl and inhibitors (1 mM phenylmethylsulfonyl fluoride (PMSF), 1 μg/ml leupeptin, 1 mM benzamidine, 1 mM Na 3 VO 4 ) by vortexing with 600 μl of glass beads. After clearing by centrifugation, supernatant was incubated with 2 ml of ANTI-FLAG-M2 affinity agarose (Sigma) for 2 hr. Resin was washed with 20 ml of buffer A containing 400 mM NaCl, 10 ml of buffer A containing 100 mM NaCl, 20 ml of buffer A containing 100 mM NaCl, 5 mM MgCl 2 and 1mM ATP, and 20 ml of buffer A containing 100 mM NaCl. FLAG-tagged protein was eluted with 4 ml of buffer A containing 100 mM NaCl, 300 μg /ml of FLAG-peptide (Sigma), 2.5 mM MgCl 2 , 5U of Benzonase (Millipore). The FLAG-eluate was incubated with 1 ml of Ni-NTA-agarose (Clontech) for one hour, washed with 5 ml of buffer A containing 100 mM NaCl. Bound protein was eluted with 1.5 ml of buffer A containing 150 mM NaCl and 300 mM imidazole and then concentrated using a Vivaspin 500 column (GE Healthcare) with buffer A containing 150 mM NaCl. All the protein purification procedures were performed at 4˚C.

Purification of MBP-Ddc2 and MBP-Ddc2-S4 protein
The coding sequences for DDC2 and ddc2-S4 were amplified by PCR using YCpT-myc-DDC2 [82] and YCp-myc-DDC2-S4 [49] with the primer pair KS3620 and KSX001, and cloned into the BamHI and SalI sites of pMAL-c2X (New England Biolabs) to generate the plasmid pMAL-Ddc2 and pMAL-Ddc2-S4, respectively. Proteins were expressed in E. coli Rosetta (Novagen) after the incubation with 1 mM IPTG at 30˚C for 4 hr. The cell pellet from one liter of culture was suspended in 50 ml of buffer M (25 mM Tris-HCl pH 7.5], 10% glycerol, 0.5 mM EDTA, 1 mM DTT) containing 300 mM NaCl and protease inhibitors (leupeptin and pepstatin A at 5 μg/ml each, 1 mM PMSF). After sonication, crude cell lysates were clarified by centrifugation and then incubated with pre-equilibrated 1 ml of amylose resin (New England Biolabs) for 2 hr. After washing with buffer M containing 1 M NaCl, bound proteins were eluted with 2 ml of buffer M containing 300 mM NaCl and 10 mM maltose. Eluates were pooled and concentrated using Vivaspin 500 columns.

Interaction of Mec1-Ddc2 with RPA
Mec1-Ddc2 and RPA were incubated with or without oligonucleotides in the binding buffer B containing ANTI-FLAG-M2 affinity agarose for 30 min at 30˚C. Beads were subjected to immunoblotting analysis.