Mrc1 phosphorylation in response to DNA replication stress is required for Mec1 accumulation at the stalled fork Maria L. Naylora, Ju-mei Lia,1, Alex J. Osbornb,1, and Stephen J. Elledgea,b,2 aDepartment
of Genetics, Harvard Medical School, Division of Genetics, Brigham and Women’s Hospital, Howard Hughes Medical Institute, Boston, MA 02115; and bDepartment of Biochemistry and Molecular Biology, Baylor College of Medicine, Houston, TX 77030
Contributed by Stephen J. Elledge, April 28, 2009 (sent for review April 13, 2009)
checkpoint 兩 DNA damage 兩 S phase 兩 replication stress response
T
he DNA damage and replication stress response consists of multiple interconnecting pathways involved in sensing damage and replication stress, transducing the signal, and effecting a cellular response. The response is particularly important in S phase, in which faithful and complete replication of the DNA is critical to a cell’s survival. Among the central signaling proteins involved in responding to DNA replication stress in Saccharomyces cerevisiae are a group of mediator proteins with roles in both DNA replication and signal transduction that include Mrc1 (ortholog of vertebrate Claspin), Tof1 (ortholog of vertebrate Timeless), and Csm3 (ortholog of vertebrate Tipin). Mrc1, a highly charged protein, was first identified in a screen for hydroxyurea (HU)-sensitive mutants that were also able to elongate their spindles in the presence of a replication block, which was indicative of a role in cell-cycle progression (1, 2), phenotypes similar to TOF1 mutants (3). Mrc1 and Tof1-Csm3 localize to replication forks (4–7), and during normal S phase Mrc1 promotes efficient replication (4, 8–10). Upon encountering a replication block, Mrc1 and Tof1 promote the formation of a stable pausing complex at the stalled fork. The deletion of either MRC1 or TOF1 is sufficient to uncouple the replisome from the replication fork in the presence of damage (5). Mrc1 and Tof1 also function independently of Rad53 to promote fork recovery in HU (9), but forks do not collapse in mrc1⌬ or mrc1AQ cells as they do in rad53 mutants (11–13). How Mrc1 carries out these different functions is not clear; however, it is known to associate with several replication proteins and protein complexes, including Cdc45 (5), the minichromosome maintenance protein helicase complex (6), and the GINS complex (14, 15). Mrc1 also interacts with DNA polymerase epsilon (Pol) (7), which itself has been implicated in DNA replication stress signaling (16). Additionally, Mrc1 and Tof1 are required to relay the DNA damage signal from Mec1 to activate Rad53 during DNA replication stress. In S. cerevisiae, Mrc1 is phosphorylated in a Mec1- and Rad53-dependent manner (1) and contains multiple SQ/TQ sites, the consensus sequence for Mec1 (4). Mutation of all 17 SQ/TQ sites in Mrc1 (mrc1AQ) successfully uncouples the replicative and signaling functions of Mrc1 (4). This Rad53 www.pnas.org兾cgi兾doi兾10.1073兾pnas.0904623106
activation is important because mrc1⌬ rad9⌬ double mutants are synthetically lethal, representing 2 separate branches that mediate Rad53 activation. Rad9 and Pol also function in parallel pathways to activate the DNA damage checkpoint (17). The inability to complete DNA replication leads to the loss of survival in S-phase checkpoint–deficient mutants. Little is known about the roles of the different domains of Mrc1 in chromatin binding and DNA replication stress signaling. In this study we performed a structural analysis of Mrc1 to identify regions required for its replication function, its recruitment to chromatin, and Rad53 activation. We find that the central core of Mrc1 is required for its critical functions and that in the absence of Mec1-dependent phosphorylation of Mrc1, Mec1 is no longer recruited to origins of replication. This helps explain Mrc1’s role in mediating Mec1 functions in replication fork restart and Rad53 activation. Results Identification of Key Functional Regions in Mrc1 Required for Viability in the Presence of Replication Stress. To further understand Mrc1’s
role in maintaining fork stability and signaling during replication stress, we created a series of mutants using a loxP-based recombination system (18). A cassette, encoding 13 aa constituting the loxP sequence and 12 additional flanking amino acids, was used to replace blocks of 25 aa at the endogenous Mrc1 locus, either maintaining its full 1096 residue length (internal replacements, I) or creating truncations in the C terminus (C) or N terminus (N) of the protein (Fig. 1). Replaced or deleted regions were targeted along regularly spaced intervals of the protein, as well as at regions of high sequence conservation [see supporting information (SI) Fig. S1]. Because Mrc1 has a documented role in the S-phase stress response and mrc1⌬ mutants are sensitive to HU, all mutants were first assayed for growth on plates containing 150 mM HU (Fig. S2 A–C). Mutants were spotted on rich media (YPD) to control for any growth defects, of which there were none (data not shown). Selected mutants were also assayed for survival in response to transient, acute HU exposure (Fig. S2 D and E), results of which mirrored the data for chronic exposure. Of all internal replacements, only mrc1-I6 (433–457) displays sensitivity to HU, indicating a high level of redundancy within Mrc1 for functions related to its response to replication stress. We also identified the smallest N-terminal fragment, mrc1-C14 (1–843), and C-terminal fragment, mrc1-N4 (266–1096), to still confer Author contributions: M.L.N., J.L., A.J.O., and S.J.E. designed research; M.L.N., J.L., and A.J.O. performed research; M.L.N., J.L., A.J.O., and S.J.E. analyzed data; and M.L.N. and S.J.E. wrote the paper. The authors declare no conflict of interest. 1J.L.
and A.J.O. contributed equally to this article.
2To
whom correspondence should be addressed. E-mail:
[email protected]. harvard.edu.
This article contains supporting information online at www.pnas.org/cgi/content/full/ 0904623106/DCSupplemental.
PNAS 兩 August 4, 2009 兩 vol. 106 兩 no. 31 兩 12765–12770
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DNA replication stress activates a response pathway that stabilizes stalled forks and promotes the completion of replication. The budding yeast Mec1 sensor kinase, Mrc1 mediator, and Rad53 effector kinase are central to this signal transduction cascade in S phase. We report that Mec1-dependent, Rad53-independent phosphorylation of Mrc1 is required to establish a positive feedback loop that stabilizes Mec1 and the replisome at stalled forks. A structure–function analysis of Mrc1 also uncovered a central region required for proper mediator function and association with replisome components. Together these results reveal new insight into how Mrc1 facilitates checkpoint signal amplification at stalled replication forks.
A
B
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Fig. 1. Identification of key internal, C-terminal, and N-terminal regions for normal Mrc1 S-phase checkpoint function. Schematic depiction of the (A) internal replacements (Y2500 –Y2523), (B) C-terminal truncations (Y2524 –Y2548), and (C) N-terminal truncations (Y2549 –Y2557) used in this study. Blue bars indicate mutants with normal growth on HU plates compared with MRC1 (Y1134), green bars indicate compromised growth, and yellow bars indicate compromised growth with synthetic lethality with rad9⌬. C8.5 (Y2536) is green/yellow to reflect low viability with rad9⌬. Black with triangles symbolize a 25-aa cassette containing loxP. The numbers in parentheses indicate either the regions targeted for replacement (A) or the intact length of Mrc1 (B and C).
WT resistance to HU. As expected, we observed that increasingly shorter C-terminal and N-terminal mutants display a gradation of increasing sensitivities. Cells deleted for MRC1 require RAD9 for survival, presumably to mediate Rad53 activation in response to the DNA damage that accumulates in the absence of Mrc1 during normal S phase (1). To further assess Mrc1 functionality, each mutant was also crossed into a rad9⌬ background (Fig. 1 and Table S1). None of the internal replacement mutants are synthetically lethal with rad9⌬. mrc1-C8.5 (1–584) is the smallest N-terminal fragment to remain viable (20%) in a rad9⌬ background, mrc1-C9 (1–635) is fully viable with rad9⌬, and mrc1-N4 (266–1096) is the smallest C-terminal fragment to maintain viability with rad9⌬. The synthetic lethality with rad9⌬ thus allowed us to identify the minimal regions required for a functional Mrc1. The above analysis shows that the amino acids between 266 and 635, which contain the region replaced in the mrc1-I6 mutant 12766 兩 www.pnas.org兾cgi兾doi兾10.1073兾pnas.0904623106
Fig. 2. mrc1-I6, mrc1-K438E, and mrc1-C13 exhibit severe defects in activating Rad53 in response to replication stress. (A) Amino acid alignment of S. cerevisiae Mrc1 region 6 (aa 433– 457) with Candida glabrata, Kluyveromyces lactis, Debaryomyces hansenii, and S. pombe homologous sequences. Arrow indicates a conserved lysine residue. Mutants (Y2507, Y2566 –Y2569, and Y2633) and controls (Y1134 and Y2558) were spotted onto YPD or 150-mM HU plates and grown at 30 °C for 3 days. (B) Mrc1 mutants in a rad9⌬ background (Y2570 –Y2576) were synchronized in G1, released into 200 mM HU at 30 °C, and samples collected at indicated times. Rad53 phosphorylation was assessed by Western blot. (C) Cells (Y1134, Y2507, Y2558, Y2566, and Y2620) were synchronized in G1 and released into YPD at 30 °C. Samples were collected at indicated time points and analyzed with FACS. The first and second vertical lines indicate the 1n and 2n DNA content peaks, respectively.
that confers HU sensitivity upon the cell, are critical for Mrc1 function. Focusing specifically on region 6, we identified a highly conserved lysine (Fig. 2A). Positively charged lysines are important for a number of lysine-specific posttranslational modifications, such as acetylation and ubiquitylation. We mutated the endogenous K438 to either a negatively charged glutamic acid (E) or a positively charged arginine (R) and assayed the point mutants for growth on 150 mM HU (Fig. 2 A). Only the mrc1-K438E mutant displays an HUS phenotype, intermediate to the mrc1-I6 mutant and WT. The normal growth of mrc1-K438R on HU rules out K438 as a site for acetylation or ubiquitylation that may play a role in HU resistance. The mrc1-K438E mutant Naylor et al.
also displays HU sensitivity when assayed for survival in response to acute HU exposure (Fig. S2D). Although mrc1-K438E does not display the same level of HU sensitivity as mrc1-I6, K438 is clearly a residue crucial for maintaining full Mrc1 activity.
A
Loss of Viability in mrc1 Mutants Results from Impaired Rad53 Activation. Mrc1 was identified as a part of the replication stress
C-Terminus of Mrc1 Is Necessary for Normal S-Phase Progression.
Mrc1 plays an important role in normal S-phase progression, loading onto the replisome after origin firing and traveling with the fork during elongation (1, 4, 8, 9). mrc1⌬ cells display a prolonged S phase, with elongation estimated at approximately half the WT rate, but S phase does not take twice as long, owing to the eventual firing of late and inefficient origins (8, 9). Therefore, we examined our mutants for DNA replication defects using FACS. The mrc1-I6, -K438E, -N4, -N5, and -C16 mutants show a slight 5-min delay through S phase compared with WT, whereas mrc1-C17 and -C18 display WT progression (Fig. 2C and Fig. S4 A and B). Importantly, the mrc1-C13, -C13.5, -C14, and -C15 mutants all exhibit a delayed S-phase progression similar to that of mrc1⌬ mutants. Thus, mrc1-C14 represents a mutant in which Mrc1’s replicative and signaling functions have been largely uncoupled, but opposite to that of the mrc1AQ mutant—a mostly intact checkpoint but defective S phase. Although Rad53 activation in the mrc1-C14 mutant is mildly defective, this uncoupling suggests that neither Mrc1’s replication nor its Rad53 activation functions are completely dependent on the full capabilities of the other. mrc1⌬ mutants incur DNA damage during normal S phase, presumably a result of impaired replication that requires Rad9mediated Rad53 phosphorylation for survival (1). We noted a correlation in the C-terminal truncation mutants between a slow S phase and Rad9-dependent Rad53 phosphorylation (Fig. S4C). Rad53 activation during normal S phase, in the absence of external damaging or blocking agents, was observed in mrc1C13, -C13.5, -C14, and -C15, which all exhibit impaired S-phase progression. mrc1-C16, -C17, and -C18 exhibit neither slow replication nor phosphorylated Rad53, thus strongly correlating the slow S phase in mrc1 mutants with the presence of DNA damage. Moreover, synthetic lethality of mrc1-C13, -C13.5, -C14, and -C15 mutants with rrm3⌬ (Table S1), a helicase that is vital for repairing the damage generated by replication forks in mrc1⌬ cells (8), further suggests that the slow S phase in these mutants Naylor et al.
B
Fig. 3. Mrc1-I6 and Mrc1AQ are depleted at stalled forks. (A and B) Cells with MYC-tagged Mrc1 (Y1134, Y2298, and Y2507) and control (Y300) were synchronized in G1 and released into 200 mM HU at 18 °C. Samples were collected at the indicated time points, and immunoprecipitated DNA was analyzed for the presence of ARS305 and late-replicating sequences on ChrVII by qPCR. Relative enrichment reflects detected levels of ARS305 to ChrVII.
stems from damage generated in the absence of fully functional Mrc1. The C-terminal region we identified as important for DNA replication overlaps with the region found to interact with the C terminus of Pol2 (7), providing a possible biochemical explanation for the defect. Mrc1-Mediated Stress Response Is Necessary for Maintaining the Replisome at Stalled Forks. Mrc1 travels with the replisome during
normal S phase (4) and remains localized to the site of stalled forks, in which it forms a stable pausing complex with the replisome and prevents uncoupling from the site of DNA synthesis (5). To establish whether our mutants can stably associate with the paused fork in HU, we performed ChIP with quantitative PCR (qPCR) as a read-out (19), which determines the relative enrichment of these proteins at an early firing origin, ARS305, relative to unreplicated sequences located on ChrVII. Although MRC1 and the mrc1-I6 and mrc1-K438E mutants all have comparable levels of protein localized at the early origin during a normal S phase (Fig. S5A), localization of Mrc1-I6 is significantly lower in the presence of HU compared with WT (Fig. 3A). Levels of Mrc1-K438E are only slightly affected (data not shown), mirroring the relative HU sensitivities observed in these mutants. The correlation between HU sensitivity and levels of Mrc1 associated with the stalled fork is also observed in the mrc1-N4, -N5, -C13, -C13.5, and -C14 mutants (Fig. S5 B and C), in which the HUS mrc1-N5 and -C13 mutants display significantly less accumulation at the stalled early origin. This is PNAS 兩 August 4, 2009 兩 vol. 106 兩 no. 31 兩 12767
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signaling pathway for its role in mediating the activation of Rad53 and was found to be phosphorylated in a Mec1- and Rad53-dependent manner (1). To determine the extent to which the HUS phenotype of mrc1 mutants might be a result of impaired Rad53 regulation, we examined Rad53 phosphorylation in response to HU (Fig. 2B). Because the RAD9 pathway acts as a back-up to activate Rad53 in late S phase in mrc1⌬ mutants, we studied the mrc1 mutants in a rad9⌬ background. We found that Rad53 activation in the mrc1-N4, -C13.5, and -C14 mutants is slightly delayed and less intense compared with WT and that phosphorylated Rad53 is not maintained over longer periods of HU incubation (100 and 120 min), all indicating subtle defects in the replication stress response. Mrc1 phosphorylation is also surprisingly normal in mrc1-N4 (Fig. S3), with only 6 of 17 SQ/TQ sites retained in this mutant, which suggests either redundancy in Mec1-targeted phosphorylation sites or efficient phosphorylation of other sites by activated Rad53. In contrast, the mrc1-I6, -K438E, and -C13 mutants display profound defects in the timing and intensity of Rad53 activation. Interestingly, Mrc1 phosphorylation is not significantly affected in the mrc1-I6 and mrc1-K438E mutants, which suggests that the HUS phenotype of these mutants is not due to a defect in Mec1-mediated phosphorylation of Mrc1.
A
during normal S phase and discovered a decreased interaction between Mrc1-I6 and Tof1 (Fig. 4B). mrc1-K438E again displays a moderately decreased level of interaction with Tof1 (data not shown). This association does not seem to be important for efficient replication, given that mrc1-I6 exhibits mostly normal S-phase progression. Instead, the decreased interaction between Mrc1-I6 and Cdc45 and Tof1 could play a crucial role during replication stress when the replisome is stalled. Mrc1 Phosphorylation, but Not Rad53 Activation, Is Required for Mec1 Recruitment at Stalled Forks. Mrc1’s association with the replisome
B
Fig. 4. Mrc1-I6 has decreased interaction with Cdc45 and Tof1 during normal S phase. (A) Mrc1–5MYC Cdc45–3HA (Y2577–Y2578) and (B) Mrc1–5MYC Tof1–3HA (Y2617–Y2618) double-tagged cells and controls (YBO469 and Y2616) were synchronized in G1 and released into YPD at 30 °C. Samples were collected after 45 min and immunoprecipitated for Mrc1–5MYC. Levels of Mrc1, Cdc45, and Tof1 were analyzed by Western blot.
not due to differences in mutant protein expression (data not shown). Because Mrc1 has been shown to be necessary for replisome stability at stalled forks, we sought to determine whether association of the Mrc1-I6 mutant protein with the fork in HU is correlated with Cdc45 and Pol2 maintenance. As expected, levels of Cdc45 and Pol2 associated with the stalled fork in mrc1-I6 mutants are significantly lower relative to MRC1 suggesting that the Mrc1-I6 protein is unable to maintain replisome coupling in the presence of replication stress (Fig. S5 D and E). Levels of Cdc45 and Pol2 in mrc1-K438E were more comparable to WT (data not shown). The depletion of Mrc1, Cdc45, and Pol2 in the mrc1-I6 mutant might be explained by replisome uncoupling from the site of DNA replication rather than a physical dissociation from the fork, a possibility that cannot be ruled out. Mrc1 Interaction with Replisome Components Is Required for an Efficient Stress Response. As demonstrated previously, Mrc1 in-
teracts with Cdc45 during S phase and is required to keep it coupled to the site of DNA synthesis in the presence of HU (5). To determine whether Mrc1-I6 and Cdc45 fully associate during normal S phase, we performed a coimmunoprecipitation (coIP) of MYC-tagged Mrc1 with HA-tagged Cdc45 (Fig. 4A). Compared with MRC1, mrc1-I6 shows a significantly decreased level of interaction between Mrc1 and Cdc45, even in the absence of exogenous damage. mrc1-K438E displays a moderately decreased level (data not shown). Tof1 also specifically interacts with Cdc45 during S phase and is required for the formation of a stable pausing complex in the presence of HU (5). Moreover, tandem affinity purification of Tof1 and Mrc1 has revealed an association between these 2 replisome components (6). We carried out a coIP of MYC-tagged Mrc1 with HA-tagged Tof1 12768 兩 www.pnas.org兾cgi兾doi兾10.1073兾pnas.0904623106
uniquely positions it at the site of the stalled fork and in close proximity to localized Mec1. Data in both S. cerevisiae and Schizosaccharomyces pombe suggest that the Rad9 mediator and Mec1 independently associate to sites of damage (20). Therefore, we sought to determine whether Mrc1 has a role in Mec1 recruitment. We used ChIP to determine the ability of mrc1-I6 and mrc1⌬ mutants to stabilize Mec1 at the stalled fork and found a significant depletion of Mec1 levels in mrc1⌬ (Fig. 5A). In contrast, only a modest decrease in the mrc1-I6 mutant was identified. Near-WT levels of Mec1 at the stalled fork were detected in mrc1-K438E (data not shown). To analyze in more depth the effect of mrc1⌬ on Mec1 recruitment to the chromosome, a ChIP-on-chip assay was performed in Mec1–18MYC– tagged cells. We observed reduced levels of Mec1 at stalled early origins on ChrIII (ARS305, -306, and -307) in mrc1⌬ mutants compared with WT cells (Fig. 5C), suggesting that Mrc1 is important for Mec1 recruitment to and/or maintenance at the stalled fork. In response to DNA replication stress, Mrc1 is phosphorylated by Mec1, which leads to activation of Rad53 and further phosphorylation of Mrc1 by Rad53. To determine whether accumulation of Mec1 at the stalled fork requires phosphorylated Mrc1, the mrc1AQ mutant was examined. A ChIP analysis was carried out in Mec1–18MYC–tagged mrc1AQ cells (Fig. 5B). Surprisingly, the accumulation of Mec1 is substantially reduced at stalled forks in mrc1AQ mutants, similar to that in mrc1⌬ cells, indicating that Mec1-dependent Mrc1 phosphorylation is crucial for recruitment and/or maintenance of Mec1 at these sites. Phosphorylated Mrc1 is also important for maintaining Mrc1 and other replisome components at the stalled fork. ChIP performed in mrc1AQ mutants demonstrates that whereas mrc1AQ is present at the fork during normal S phase at WT levels (ref. 4 and Fig. S5F), it is depleted at stalled forks in HU (Fig. 3B). Moreover, Cdc45 and Pol2 also show reduced levels at the fork in the presence of HU in mrc1AQ mutants (Fig. S5 G and H). Mrc1 phosphorylation is required for Rad53 activation. To determine whether Rad53 activation has a role in Mec1 retention at sites of DNA replication stress, we carried out a ChIPon-chip assay in MEC1–18MYC–tagged sml1⌬ and sml1⌬rad53⌬ mutants (Fig. 5D). Mec1 is recruited to early origins on ChrIII at comparable levels in both mutants, strongly suggesting that Rad53 activity is not required for early Mec1 recruitment and maintenance. These data are further supported by ChIP analysis of Mec1 recruitment in sml1⌬ and sml1⌬rad53⌬ cells, which reveals no significant difference in the levels of Mec1 enrichment in HU (Fig. S6A). Therefore, levels of Mec1 stabilized at stalled forks seem to be specifically controlled by Rad53-independent, Mec1-dependent phosphorylation of Mrc1. ChIP performed in tof1⌬ mutants, which exhibit timely Mrc1 phosphorylation but delayed and attenuated Rad53 activation in HU, also show Mec1 enrichment at the early ARS305 origin, comparable to WT cells (Fig. S6 B and C), further supporting the dispensability of Rad53 for Mec1 stability. This gives rise to the hypothesis of a positive feedback loop, in which Mec1 is recruited to the stalled fork and phosphorylates Mrc1, which in turn allows Mrc1 to retain Mec1. How Mrc1 phosphorylation precisely stabilizes Mec1 is unclear. Naylor et al.
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D
Discussion It has been well established that Mrc1 has key functions both as a mediator of S-phase checkpoint and as a facilitator of normal replication elongation. Our data suggest that Mrc1 has dual functions as a mediator of the DNA replication stress response pathway, both to establish initial Mec1 accumulation at the stalled fork and to provide a platform for eventual Rad53 recruitment and phosphorylation. A diagram of Mrc1 functional domains (Fig. S7) summarizes established and newly defined regions identified in this study. During a normal replication stress response (Fig. S8A), Mec1 is recruited to the site of the stalled fork and phosphorylates Mrc1. We propose that this phosphorylation event maintains Mrc1 localization, which in turn helps maintain Mec1 at the stalled fork. Mrc1 establishes a stable stalled replisome that remains coupled to the replication fork and provides the platform for later checkpoint events, including the recruitment and phosphorylation of Rad53. The activation of Rad53 and its additional downstream phosphorylation of Mrc1 allow for the maintenance of the replisome and fork until replication can be resumed. mrc1AQ mutants are not phosphorylated by Mec1, therefore the positive feedback loop required for Mec1 stability at the fork is not established (Fig. S8B). Rad53 fails to be activated in a timely fashion, and the replisome becomes uncoupled from the fork. mrc1AQ is less HU sensitive than mrc1⌬, Naylor et al.
highlighting an additional role in maintaining cell viability, potentially through supporting Tof1 mediator function or another mechanism for recovering stalled forks. mrc1-I6 is phosphorylated by Mec1 and is therefore largely competent for establishing the positive feedback loop for initial Mec1 accumulation (Fig. S8C). However, Mrc1-I6’s compromised structure and/or interactions with other replisome components, such as Cdc45 and Tof1, independent of its phosphorylation state, prevent it from establishing a stable stalled replisome and creating a competent platform for appropriate Rad53 phosphorylation. Delayed Rad53 activity likely compromises Mec1 maintenance and replisome stability, affecting their localization to the fork. On the basis of the ChIP data, as well as Mrc1-I6 and Mrc1-K438E phosphorylation kinetics in HU, it is clear that Mec1 phosphorylation of Mrc1 alone is insufficient to maintain fork stability in HU and facilitate downstream checkpoint events. The model proposed by Lou et al. (7), which predicts a conformational change during the replication stress response that alters Mrc1 interactions with replisome components, could account for the phosphorylation-dependent and -independent checkpoint functions that are both required for efficient Mrc1 mediator activity. It is also supported by data demonstrating that Mrc1 and Mec1 are required to stabilize Pol␣ and Pol before Rad53 activation (5, 21, 22). However, rad53K227A cells show PNAS 兩 August 4, 2009 兩 vol. 106 兩 no. 31 兩 12769
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Fig. 5. Mrc1 is required for Mec1 accumulation at stalled replication forks. (A) Mec1–18MYC–tagged cells (Y2612–Y2614) and (B) Mec1–18MYC mrc1⌬ cells with either pMRC1 (Y2609), pmrc1AQ (Y2610), or an empty vector (Y2611) were processed and analyzed as in Fig. 3. (C) Mec1–18MYC mrc1⌬ cells with either pMRC1 (Y2609) or an empty vector (Y2611) and (D) Mec1–18MYC–tagged sml1⌬ (Y2624) and sml1⌬rad53⌬ (Y2625) mutants were synchronized in G1 and released into 200 mM HU at 18 °C. Samples were collected at 144 min, and immunoprecipitated DNA was hybridized to an Agilent Yeast Whole Genome ChIP-on-chip Microarray Kit. Data were plotted for relative Mec1 enrichment on ChrIII vs. non-immunoprecipitated input.
depleted levels of Pol␣, Pol␦, and Pol on chromatin (23). The investigators also detected a significant increase in Ddc2, as well as Ddc1 and RFA stability, in response to HU in rad53K227A mutants, whereas we found relatively equal levels of Mec1 at stalled forks in both sml1⌬ and sml1⌬rad53⌬. The differences can perhaps be explained by a difference in kinetics (time courses in all 3 studies were conducted at different temperatures) and/or differences in the nature of the different Rad53 mutations used in each. Recent work has also demonstrated that Mec1 and Rad53 operate in separate pathways to stabilize stalled forks, in which Rad53 has Exo1-dependent and -independent functions required for maintaining functional forks (24, 25). Future experiments addressing how phosphorylated Mrc1 mediates Mec1 stability during replication stress will be key in furthering our understanding of how the S-phase stress response signal is amplified at stalled forks. Materials and Methods Media and Growth Conditions. Yeast cells were grown at 30 °C unless otherwise noted. In ChIP experiments, Mec1–18MYC–tagged strains and controls were grown at 25 °C before being shifted to 18 °C. YPD and SC media were glucose based and made as per Kaiser et al. (26). Log-phase cells were synchronized in G1 by ␣-factor arrest, as previously described (11). Strains and Plasmids. See Table S2 for a complete list of the strains and plasmids used in this study. See SI Methods for details on plasmid and strain construction. HU Viability Assays. For chronic HU exposure, cells were grown to log phase and normalized to a concentration of 107 cells/mL. Cultures were serially diluted by factors of 10, spotted onto YPD and 150-mM HU plates, and grown at 30 °C for 3 days. For acute HU exposure, cells were grown to log phase, arrested in G1, and released into 200 mM HU at 30 °C. Samples were removed at indicated time points, plated onto YPD plates, and grown at 30 °C for 3 days.
(Santa Cruz Biotechnology). Mrc1-Cdc45 and Mrc1-Tof1 coIP were prepared as described by Osborn and Elledge (4) and fractionated by 3– 8% Tris-acetatePAGE. Samples were transferred to PVDF membranes and detected by monoclonal 9E10 ␣-MYC (Covance) or ␣-HA.11 (Covance). FACS Analysis. Cells were grown to log phase, arrested in G1, and released into YPD at 30 °C. Samples were collected at indicated time points and processed as described previously (1). ChIP Assays. Chromatin immunoprecipitation was performed essentially as described previously (4), with the following exceptions. Cells were fixed with 1% formaldehyde at 30 °C for 30 min. Whole-cell lysates were sonicated for 4 ⫻ 12 sec each and clarified by centrifugation for 15 min at 22,000 ⫻ g. Immunoprecipitations were carried out using 9E10 ␣-MYC or ␣-HA.11. PCR Analysis. For quantitative PCR analysis, 10% of immunoprecipitated DNA was used in reactions prepared with 0.2 M primers and Platinum SYBR Green qPCR SuperMix-UDG with ROX (Invitrogen). Reactions were carried out in triplicate or quadruplicate in optical 96-well reaction plates (Applied Biosystems) using the 7500 Fast Real-Time PCR System (Applied Biosystems). See SI Methods for primer sequences. Immunoprecipitated DNA in Fig. S6 B and C was processed as previously described (4). Immunoprecipitation Assays. Cells were grown to log phase, arrested in G1, and released into YPD. Samples were collected at 45 min, when the majority of cells were in S phase, and processed essentially as previously described (7), with the following exceptions. Whole-cell extracts were incubated with 9E10 ␣-MYC and precipitated with prewashed agarose protein A/G beads (Santa Cruz Biotechnology). DNase I, type II (Sigma) was used to degrade chromosomal DNA in extracts. ChIP-on-chip. Experiments were carried out as for ChIP. Immunoprecipitated DNA was hybridized to a Yeast Whole Genome ChIP-on-chip Microarray Kit, 4 ⫻ 44K (Agilent). Data were analyzed using Agilent ChIP Analytics 1.3 and plotted for relative enrichment on ChrIII vs. non-immunoprecipitated input.
Western Blotting. Mrc1 and Rad53 time-course samples were prepared by trichloroacetic acid precipitation (27) for Western blotting. Precipitated protein was prepared as described previously (4), transferred to nitrocellulose membranes, and detected by polyclonal ␣-Mrc1 (1) or ␣-Rad53 antibodies
ACKNOWLEDGMENTS. We thank C. Cotta-Ramusino, K. Hurov, D. Chou, and D. Lee for helpful discussions. This work is supported by a grant from the National Institutes of Health to S.J.E. S.J.E. is a Howard Hughes Medical Institute Investigator. M.L.N. was supported by National Research Service Award Training Grants 2 T32 GM07196 –29 and 5 T32 GM07196 –30.
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