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Mrc1 Is Required for Normal Progression of Replication Forks throughout Chromatin in S. cerevisiae
Molecular Cell, Vol. 19, 691–697, September 2, 2005, Copyright ©2005 by Elsevier Inc. DOI 10.1016/j.molcel.2005.06.037
Mrc1 Is Required for Normal Progression of Replication Forks throughout Chromatin in S. cerevisiae Shawn J. Szyjka, Christopher J. Viggiani, and Oscar M. Aparicio* Molecular and Computational Biology Program Department of Biological Sciences University of Southern California Los Angeles, California 90089
Summary Mrc1 associates with replication forks, where it transmits replication stress signals and is required for normal replisome pausing in response to nucleotide depletion. Mrc1 also plays a poorly understood role in DNA replication, which appears distinct from its role in checkpoint signaling. Here, we demonstrate that Mrc1 functions constitutively to promote normal replication fork progression. In mrc1D cells, replication forks proceed slowly throughout chromatin, rather than being specifically defective in pausing and progression through loci that impede fork progression. Analysis of genetic interactions with Rrm3, a DNA helicase required to resolve paused forks, indicates that Mrc1 checkpoint signaling is dispensable for the resolution of stalled replication forks and suggests that replication forks lacking Mrc1 create DNA damage that must be repaired by Rrm3. These findings elucidate a central role for Mrc1 in normal replisome function, which is distinct from its role as a checkpoint mediator, but nevertheless critical to genome stability. Introduction The accurate duplication of chromosomal DNA presents a significant challenge to the cell. Replication forks must contend with the presence and activities of various chromatin structures, transcription complexes, and recombination machinery, which often cause replication forks to pause (reviewed in Rothstein et al., 2000). For example, replication forks pause at tRNA genes when the direction of transcription is opposite that of the replication fork (Deshpande and Newlon, 1996). Although the exact function of fork pausing at tRNA genes is not known, it may help prevent fork dysfunction that might result from collision with the transcription machinery, and thus, facilitate resumption of normal fork progression. Multiple mechanisms contribute to replication fork stability and faithful genome duplication. For example, the Rrm3 DNA helicase has been implicated as playing an important role in the progression of replication forks through natural impediments such as tRNA genes and chromatin complexes (Ivessa et al., 2003, 2002, 2000). In addition, surveillance mechanisms called checkpoints detect and respond to problems that arise during DNA replication. A critically important pathway that *Correspondence: [email protected]
Short Article
operates during S phase is the replication stress checkpoint (reviewed in Osborn et al., 2002). This pathway acts by maintaining adequate deoxyribonucleotide (dNTP) levels, increasing transcription of DNA repair genes, slowing cell cycle progression, and maintaining stable replication forks that are poised to resume replication following perturbations. Mediator of replication checkpoint protein 1 (Mrc1) localizes with replication forks and acts in the replication stress response by transducing signals of stress from the “sensor” kinase Mec1 to the “effector” kinase Rad53 (Alcasabas et al., 2001; Katou et al., 2003; Osborn and Elledge, 2003; Tanaka and Russell, 2001). Thus, in cells challenged with an inhibitor of DNA synthesis such as hydroxyurea (HU), which depletes deoxyribonucleotide levels, Mrc1 is required for effective activation of Rad53 by Mec1. Remarkably, in mrc1D cells undergoing DNA replication in the presence of HU, the progression of DNA synthesis halts, while replication fork proteins, including Mcm proteins, Cdc45 and DNA polymerases, appear to progress farther along the chromatin, suggesting that uncoupling of the replication apparatus from the site of DNA synthesis has occurred (Katou et al., 2003). This has led to the conclusion that Mrc1 functions in a pausing complex that maintains replisome integrity when exogenous stresses are encountered. Mrc1 also functions during normal DNA synthesis in cells that have not received exogenous perturbation. This is evidenced by the somewhat-slower rate of bulk chromosomal DNA synthesis in mrc1D cells, which is accompanied by a Rad9-dependent DNA damage response (Alcasabas et al., 2001). The reason for the slower replication of mrc1D cells remains unclear but does not depend on checkpoint signaling by either Rad9 or Mrc1 (Alcasabas et al., 2001; Osborn and Elledge, 2003). It is also unknown whether the defective replisome pausing in HU reflects loss of the replication or checkpoint-signaling function of Mrc1. In this study, we have investigated the role of Mrc1 in DNA replication of unperturbed cells, by examining origin initiation, fork elongation, and fork pausing in cells lacking Mrc1 function. We also have explored the possible role of Mrc1 checkpoint signaling in response to fork pausing by analyzing genetic interactions with Rrm3. Together, our findings demonstrate that Mrc1 plays a central and constitutive role in the efficient function of the replisome. Our results also suggest that Rrm3 is vital to repair DNA damage created by defective replication forks in mrc1D cells. Results and Discussion Mrc1 Is Required for a Normal Rate of Replication Fork Progression throughout Chromatin The replication defect of cells lacking Mrc1 could be due to a defect in the initiation of replication, in elongation, or in replisome pausing. To distinguish between these possibilities, we monitored origin initiation and
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Figure 1. Mrc1 Is Required for Normal Replication Fork Progression along Chromosome VI (A) Position of probes (thin bars beneath the chromosome depiction) and relevant ClaI (C) and EcoRI (E) restriction sites used in the 2D gel analysis of modified chromosome VI are shown approximately to scale. Thick bars indicate the positions of sequences analyzed for BrdU incorporation and Cdc45 association shown in Figures 2 and 4. The bent arrows indicate the positions and transcriptional orientations of the tRNA genes tA(AGC)F in region A and SUP6 in region B. (B) Wild-type (SSy48) and mrc1D (SSy47) cells were blocked in G1 with α factor at 23°C and released synchronously into S phase at 20°C. Cells were collected at the indicated times for 2D gel analysis by digestion with EcoRI and ClaI. Blots were probed for ARS607 and stripped and reprobed for regions A, B, and C. Filled and unfilled arrowheads indicate small and large bubble structures, respectively; open and double arrowheads indicate large and small fork structures, respectively; the complete arrows indicate pause sites. The greater signal intensity of replication structures in mrc1D cells is reproducible and appears to reflect the longer period of time that forks are present within each analyzed fragment.
the progression of replication forks using two-dimensional gel electrophoresis (2D gel) across regions of chromosomes III and VI. In each of these regions, one or more active replication origins has been deleted, so that in wild-type cells, the replication of these regions is unidirectional, facilitating analysis of fork progression (Figures 1A and 3A) (Tercero and Diffley, 2001). In wild-type cells released synchronously into S phase, initiation of ARS607 occurred at about 30 min. By 36 min, primarily larger replication structures were present due to fork elongation (Figure 1B). In mrc1D cells, initiation of ARS607 occurred with timing like that of wild-type cells, as bubbles were first observed at 30 min (Figure 1B). The shapes of the bubble arcs and their greater intensities suggested a delay in the elongation of initiation bubbles in mrc1D cells. At 30 min, small bubbles predominated (Figure 1B), and at 36 min, small and large bubbles were present, as well as large forks (Figure 1B). These larger structures persisted near ARS607 at 48–60 min. The presence of small bubbles at 30–36 min together with their absence at later times, as in wild-type cells, indicates that initiation timing of these origins is not altered by the absence of Mrc1 (also, see below). Furthermore, the efficiency of ARS607 is not altered by loss of Mrc1 based on the virtual absence of smaller fork structures in these gels and on the analysis of unsynchronized cultures (data not shown). These results suggest that replication forks
emanating from ARS607 elongated more slowly in mrc1D cells than in wild-type cells. Next, we analyzed the progression of replication forks from ARS607 into adjacent chromosome VI sequences, restriction fragments “A,” “B,” and “C,” whose midpoints lie at 4, 12, and 20 kb from ARS607 (Figure 1A). In addition, regions A and B each contain a tRNA gene that permitted analysis of replication fork pausing (discussed below). In wild-type cells, the peak signals for forks at A, B, and C occurred at 36, 48, and 60 min, respectively (Figure 1B). Based on the initial appearance of replication structures at ARS607 at 30 min and at region C at 48 min, the data indicate an average fork rate of w1 kb per minute in wild-type cells. In the absence of Mrc1, the progression of replication forks into and throughout the adjacent chromosomal regions was significantly delayed. The peak signals for forks at regions A, B, and C occurred at about 48, 72, and 84 min, and forks were present within each region for a longer period, consistent with their slower movement through each region (Figure 1B). Furthermore, the preponderance of smaller forks earlier and larger forks later in the interval during which forks move through each region is consistent with slow movement of replication forks throughout the region (Figure 1B). Based on the timing of initial appearance of replication structures at ARS607 and at region C, we estimate that replication forks progress at about half the wild-type rate in
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Figure 2. Mrc1 Is Required for Normal Progression of DNA Synthesis Wild-type (CVy39), mrc1-AQ (CVy40), and mrc1D (CVy29) cells, which all express GPD-TK, were blocked in G1 with α factor at 23°C and released synchronously into S phase at 20°C. (A) DNA content analysis. (B) Cells were collected at the indicated times for analysis of BrdU incorporation. Immunoprecipitated DNA sequences at ARS607 and at distances of 5, 15, and 20 kb (see Figure 1A) were detected by PCR amplification. (C) PCR products in (B) and of input DNA (data not shown) were quantified and plotted as the ratio of immunoprecipitated DNA/ input DNA (% BrdU Incorporated).
mrc1D cells. Together with the slow progression of total DNA synthesis in mrc1D cells (Figure 2A) (Alcasabas et al., 2001), these findings suggest that Mrc1 is required for normal replication fork progression throughout much or all of the genome. Whether replication forks in mrc1D cells progress at a uniformly slow rate or whether the slow progression is due to occasional stalling at random sites (requiring restart) is not clear. However, the perdurance of some replication forks at later
times (e.g., Figure 1B, ARS607 at 72–96 min) suggests that some forks have stalled but remain stable. We also analyzed initiation and fork progression in a strain harboring the mrc1-AQ allele, in which all hypothetical Mec1 phosphorylation sites have been mutated (Osborn and Elledge, 2003). This allele is defective in checkpoint signaling to activate Rad53, but appears to retain its function in DNA replication, based on its approximately normal rate of replication as measured by DNA content analysis (Figure 2A) (Osborn and Elledge, 2003). 2D gel analysis indicated that initiation and fork movement in the mrc1-AQ strain was similar to wildtype cells (Figure S1 in the Supplemental Data available with this article online). Thus, replication forks in mrc1AQ cells progress with similar kinetics as in wild-type cells, and the slower replication fork movement of mrc1D cells is not due to loss of Mrc1’s checkpoint function. To confirm our conclusion that normal fork progression depends on Mrc1, but not its checkpoint function, we analyzed nascent DNA synthesis by measuring incorporation of the thymidine analog bromodeoxyuridine (BrdU) along chromosome VI (Figures 2B and 2C). In wild-type mrc1-AQ and mrc1D cells, BrdU incorporation occurred with similar timing and efficiency at the origin ARS607, primarily between 24 and 36 min, reinforcing our conclusion that initiation is not affected by loss of Mrc1 function. The kinetics of BrdU incorporation into DNA sequences at distances of 5, 15, and 20 kb from ARS607 was indistinguishable between wildtype and mrc1-AQ cells. However, BrdU incorporation into DNA sequences at distances of 5, 15, and 20 kb from ARS607 was delayed in mrc1D cells. The degree of delay is consistent with the results of the 2D gel analysis, with a delay of about 24 min at a distance of 20kb. Thus, in the absence of Mrc1, the replication timings of DNA sequences proximal to ARS607 were delayed in relation to their distance from the origin, with more distal sequences experiencing a greater delay. These data are fully consistent with the conclusion that replication forks progress more slowly in cells lacking Mrc1 function, whereas the checkpoint signaling function of Mrc1 appears to be dispensable for normal fork progression in unperturbed cells. To extend our analysis to other chromosomal regions and further test the idea that Mrc1 plays a general role in replication fork progression, we analyzed replication of a region of chromosome III adjacent to the efficient origin ARS306 (Figure 3A). In wild-type cells, initiation of ARS306 occurred at 30 min and replication forks were mainly detected at a distance of 23 kb from this origin at 48–60 min (Figure 3B, “Region D”). In mrc1D cells, ARS306 initiated at 30 min as in wild-type cells, and as observed with ARS607, replication bubbles from ARS306 appeared to elongate slowly in mrc1D cells (Figure 3B). Most replication forks in mrc1D cells did not reach region D until at least 72 min, thus having required at least 42 min to travel 23 kb. Together with the results of the chromosome VI analysis and total genomic DNA replication, these findings support the conclusion that normal progression of replication forks through much if not all of the genome requires Mrc1. We also analyzed replication of ARS304, an inefficient origin at a distance of 45 kb from ARS306. Unex-
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Figure 3. Mrc1 Is Required for Normal Replication Fork Progression along Chromosome III (A) Position of probes (thin bars beneath the chromosome depiction) and relevant ClaI (C) and EcoRI (E) restriction sites used in the 2D gel analysis of modified chromosome III are shown approximately to scale. (B) Blots from the experiments in Figure 1 were stripped and reprobed for ARS306, Region D, and ARS304. See Figure 2A for DNA content analysis.
pectedly, we observed increased initiation of ARS304 in mrc1D cells (Figure 3B). Because the level of ARS304 initiation in mrc1-AQ cells was similar to wild-type (Figure S1), we conclude that the increased initiation frequency of ARS304 in mrc1D cells is due primarily to the slower progression of replication forks from ARS306, which permits more time for ARS304 initiation before it is passively replicated by a fork from ARS306. We also have observed evidence for increased initiation of the inefficient origin near the VI-R telomere in mrc1D cells (data not shown). The increased initiation of some origins due to their delayed passive replication helps explain why genome duplication in mrc1D cells does not appear to require twice as long as wild-type cells (Figure 2A; and Tourrière et al., 2005 [this issue of Molecular Cell]), although our data indicate that forks progress at about half the wild-type rate. Progression of Cdc45 Is Slower in the Absence of Mrc1 The possibility that Mrc1 plays a role in coupling replication fork components to the site of DNA synthesis prompted us to examine whether the progression of Cdc45 along chromatin also was slowed in the absence of Mrc1 or whether it might move rapidly ahead of the slow-moving replication forks. To examine the movement of Cdc45, we analyzed its association with chromosome VI by chromatin immunoprecipitation (ChIP) (Figure 4). This analysis was performed at 16°C to enhance the resolution of the assay. In both wild-type and mrc1D cells, Cdc45 association with ARS607 peaked at 60 min and diminished soon after. The similar timings
in wild-type and mrc1D cells reinforce our conclusion that initiation timing of ARS607 is not altered by loss of Mrc1 function. However, the progression of Cdc45 along chromatin was altered in the absence of Mrc1. In wild-type cells, Cdc45 moved rapidly, associating maximally with sequences 5, 15, and 20 kb distant from ARS607 at 72, 72, and 84 min, respectively. In mrc1D cells, Cdc45 progressed more slowly, associating maximally with the sequences 5, 15, and 20 kb from ARS607 at 72, 108, and 120 min, respectively. Hence, the progression of Cdc45 along chromatin during S phase, like that of replication forks, is slowed in the absence of Mrc1. Although the slowed progression of Cdc45 may be consistent with Cdc45 remaining coupled with the site of DNA synthesis in unperturbed mrc1D cells, it is possible that dissociation of Cdc45 from the site of DNA synthesis does occur, but can be resolved by our analysis only when DNA synthesis is inhibited strongly with HU. Whether or not a defect in coupling exists, the results make clear that Mrc1 is required for the normal progression of Cdc45 along chromatin during DNA replication. Resolution of Naturally Paused Replication Forks Does Not Require Mrc1 Next, we examined whether replication forks lacking Mrc1 were defective in progression through endogenous pause sites, which might contribute to the slower progression of replication forks that we have observed in mrc1D cells. We analyzed pausing at two tRNA genes within regions A and B adjacent to ARS607 (Figure 1A). Fork pausing at each tRNA gene was apparent
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fork pausing (Figure 3). We also note that mrc1D cells are not highly sensitive to HU, suggesting these cells are able to restart replication effectively (Alcasabas et al., 2001). We conclude that Mrc1 is not required for the stability and restart of naturally stalled forks.
Figure 4. Slower Progression of Cdc45 in mrc1D Cells Wild-type (CVy31) and mrc1D (CVy30) cells, which both express Cdc45-HA, were blocked in G1 with α factor at 23°C and released synchronously into S phase at 16°C. Cells were collected at the indicated times for ChIP analysis with anti-HA antibody. DNA sequences at ARS607 and at distances of 5, 15, and 20 kb (see Figure 1A) were quantified by PCR amplification. The ratio of immunoprecipitated to input DNA is plotted as “% Cdc45 Bound.”
as a dark spot within each fork arc. Compared to wildtype cells, the pause signals appeared reduced in mrc1D cells (Figure 1B, arrows). However, we suspected that the decreased pause signal resulted at least in part from the slower fork movement of mrc1D cells, which increased the length of time spent by replication forks throughout regions A and B, and thus decrease the relative intensity of the pause signal. Consistent with this idea, careful quantification of the pause at the tRNA gene in region A relative to total replication structures indicated that pausing was reduced by about half (Figure S2). Because replication forks progress at about half the wild-type rate, these data suggest that the duration of pausing at this tRNA gene is not significantly altered in the absence of Mrc1. In addition, these results strongly suggest that Mrc1 is not required for resolution of paused forks, which would appear as an increased pause signal. It was possible that paused forks in mrc1D cells were unstable and hence interfered with our ability to detect the paused structure itself and/or a defect in its resolution. To address directly the possibility that defective pausing somehow contributes to fork dysfunction in mrc1D cells, we analyzed fork progression along the 20 kb chromosome VI region in mrc1D cells in which we deleted both tRNA genes. Deletion of the tRNA genes did not significantly affect the rate of fork progression across this region (data not shown), indicating that the slow fork movement of mrc1D cells is not due to defective pausing or instability of forks encountering these sites. In support of this conclusion, forks appeared to progress slowly within all the regions that we analyzed, including the chromosome III region, which does not contain elements known to be associated with
The Replication Defect of mrc1D Cells Creates a Dependence on Rrm3 for Viability The Rrm3 DNA helicase enables the resolution of stalled replication forks. In the absence of Rrm3, fork pausing at natural sites is prolonged, and cells become dependent on MRC1 for viability (Ivessa et al., 2003, 2002; Ooi et al., 2003; Tong et al., 2004; Torres et al., 2004). These findings have suggested that the repair or restart of stalled forks in rrm3D cells is dependent on the replication stress response, which is mediated by Mrc1 (Torres et al., 2004). To determine whether the requirement of rrm3D cells for MRC1 reflects the role of Mrc1 in checkpoint signaling or in fork progression, we assessed the viability of rrm3D mrc1-AQ cells by dissection of a diploid strain heterozygous for both alleles. Haploid rrm3D mrc1-AQ cells emerged at a frequency equivalent to either single mutant strain and grew at a similar rate (Figure 5A). Thus, Mrc1’s checkpoint function is dispensable in cells lacking Rrm3. As expected, rrm3D mrc1D cells were inviable (Figure 5B). These results suggest that the lethality of rrm3D mrc1D cells reflects a dependence on Rrm3 to resolve replication fork defects incurred by loss of Mrc1 function, rather than a vital replication checkpoint response to prolonged fork stalling in rrm3D cells. To examine further the function of the replication checkpoint in response to fork stalling, we analyzed fork pausing at tRNA genes in wild-type and rrm3D cells with either an intact or defective replication checkpoint. Pausing was not altered in mrc1-AQ cells, indicating that the replication checkpoint is not required for the resolution of paused forks (Figure 5C, data not shown for SUP6). As expected, rrm3D cells exhibited greatly increased stalling at the tRNA gene (Figures 5C and 5D). Furthermore, we observed a similar level of fork pausing in rrm3D mrc1-AQ cells as in rrm3D cells (Figures 5C and 5D). Thus, checkpoint signaling by Mrc1 does not appear to participate in replication fork pausing or resolution, either in the presence or absence of Rrm3. Because mrc1D cells are defective in fork progression throughout chromatin and require Rrm3 for viability, we propose that Rrm3 acts throughout chromatin in response to replication fork defects and impediments. A Central Role for Mrc1 at Replication Forks? Previous analysis of mrc1D cells treated with HU has led to the conclusion that Mrc1 is required for normal fork pausing in response to replication stress by maintaining coupling of replication proteins to the site where DNA synthesis is inhibited (Katou et al., 2003). Our demonstration that lack of Mrc1’s replication function, but not its checkpoint function, causes defective fork progression throughout chromatin in untreated cells is consistent with a coupling role for Mrc1 in the replisome and further suggests this role is constitutive rather than checkpoint induced. Mrc1 coimmunopreci-
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The slower rate of fork progression in cells lacking Mrc1 may reflect uncoupling of replication factors. Studies of bacterial and viral eukaryotic DNA replication have demonstrated synergistic stimulation of helicase and polymerase activities that depend upon their physical association (Dong et al., 1996; Kim et al., 1996; Yuzhakov et al., 1996). If loss of Mrc1 function disrupts similar interactions in the eukaryotic replisome, slower fork progression might result. In addition to compromising the efficacy of DNA synthesis at the replication fork, loss of Mrc1 from the site of DNA synthesis may expose normal replication structures to DNA damage sensors, DNA repair proteins, and recombination factors. This could explain some of the defects associated with loss of Mrc1, such as activation of the DNA damage response and increased levels of homologous recombination (Alcasabas et al., 2001; Xu et al., 2004). These phenotypes may reflect the presence of excess unwound DNA, or DNA structures that may serve as effective recombination substrates such as doublestrand breaks, or 3# ends. The presence of such structures in mrc1D cells is suggested by their vital dependence on the Srs2 and Rrm3 DNA helicases (Ooi et al., 2003; Tong et al., 2004; Torres et al., 2004; Xu et al., 2004), which appear to function in the repair of stalled or damaged replication intermediates, perhaps by preventing their subversion into recombination pathways. The elucidation of a general role of Mrc1 in the function of replication forks provides insight into the mechanisms that promote genome stability during DNA replication. Experimental Procedures
Figure 5. Rrm3 Is Required for Viability of mrc1D Cells but Not mrc1-AQ Cells Diploid strains SSy108 and SSy109, in (A) and (B), respectively, were induced to sporulate at 23°C; germination was at 30°C. The relevant genotypes are indicated in the panels below as follows: WT = wild-type, AQ = mrc1-AQ, m1 = mrc1D, and r3 = rrm3D. The genotypes of inviable spores were inferred assuming 2:2 segregation of genetic loci within each tetrad. (C) Haploid strains W3031a (WT), SSy72 (mrc1-AQ), SSy69 (rrm3D), and SSy108 (mrc1-AQ rrm3D) were subjected to 2D gel analysis of tA(AGC)F by digestion with SpeI (see Figure S2A for position of probe and restriction sites). (D) Three sets of blots like those in (C) were quantified by measuring the pause signal and dividing by the signal for all replication intermediates along the fork arc (excluding the 1N spot). The data was plotted relative to wild-type, which was arbitrarily assigned the value 1. Error bars denote standard deviation.
pitates with Cdc45, Mcm2, and Mcm3 (Katou et al., 2003; Nedelcheva et al., 2005). Furthermore, recent biochemical analyses of S. pombe Mrc1 and its apparent human homolog, Claspin, indicate that Mrc1/Claspin can bind branched DNA structures in vitro (Sar et al., 2004; Zhao and Russell, 2004). Together, these data suggest that Mrc1 links Cdc45 and/or Mcm proteins to replication forks in vivo. Depletion of Claspin from Xenopus replication extracts also modestly reduced the rate of DNA synthesis suggesting that Claspin may play a similar role at replication forks in vertebrates (Lee et al., 2003).
Plasmid and Strain Constructions Strains are described in Table S1. Methods for strain construction have been described previously (Gibson et al., 2004). pRS306Mrc1-AQ contains the NotI-KpnI mrc1-AQ fragment from pRS405Mrc1-AQ (Gibson et al., 2004) inserted into pRS306. mrc1-AQ was introduced at its native locus by a pop-in-pop-out strategy, or by a simple pop-in. Yeast Methods YEPD medium was used for all experiments. Cell culturing, synchronization, DNA content analysis, spore analysis, and ChIP have been described (Aparicio et al., 2004; Gibson et al., 2004). Twentytwo cycles of PCR were performed for analysis of the ChIP experiment. Analysis of Replication Structures 2D gel analysis was performed as described previously, except that 30 g of DNA was used for each sample (Gibson et al., 2004). Quantitative comparisons were performed on samples run in the same gels and blotted and hybridized together to minimize any errors that might result from differential transfer or hybridization efficiencies. Analysis of BrdU Incorporation Strains expressing seven copies of HSV-TK were grown in the presence of 400 g/ml BrdU as described (Lengronne et al., 2001). Immunoprecipitation of BrdU-substituted DNA was carried out as described in (Katou et al., 2003) and references therein, with the following modifications: DNA was extracted by glass bead beating, sheared by sonication to an average size of w500 bp, and isolated by phenol/chloroform extraction and ethanol precipitation. DNA was treated with RNaseA and Proteinase K and further purified on Qiaquick PCR purification spin-columns (Qiagen). 500 ng of DNA was combined with 20 g of sheared salmon sperm DNA, dena-
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tured, and incubated with anti-BrdU antibody (Oxford Biotechnology) followed by G-Sepharose beads (Amersham). 22 cycles of PCR were performed on the immunoprecipitated material and on a 1:10 dilution of the input material. Sequences of primers used for PCR analyses are available upon request.
incorporation in TK(+) yeast strains. Nucleic Acids Res. 29, 1433– 1442.
Supplemental Data Supplemental Data include two additional figures and a table and can be found with this article online at http://www.molecule.org/ cgi/content/full/19/5/691/DC1/.
Ooi, S.L., Shoemaker, D.D., and Boeke, J.D. (2003). DNA helicase gene interaction network defined using synthetic lethality analyzed by microarray. Nat. Genet. 35, 277–286.
Acknowledgments We thank: N. Arnheim and M. Goodman for sharing equipment; J. Diffley, S. Elledge, and E. Schwob for strains and plasmids; P. Pasero for communicating unpublished data; and J. Aparicio, J. Campbell, J. Li, and R. Lipford for critical reading of the manuscript. This work was supported by NIH grant 1RO1GM65494 to O.M.A.
Received: February 23, 2005 Revised: May 26, 2005 Accepted: June 28, 2005 Published: September 1, 2005 References Alcasabas, A.A., Osborn, A.J., Bachant, J., Hu, F., Werler, P.J., Bousset, K., Furuya, K., Diffley, J.F., Carr, A.M., and Elledge, S.J. (2001). Mrc1 transduces signals of DNA replication stress to activate Rad53. Nat. Cell Biol. 3, 958–965. Aparicio, J.G., Viggiani, C.J., Gibson, D.G., and Aparicio, O.M. (2004). The Rpd3-Sin3 histone deacetylase regulates replication timing and enables intra-S origin control in Saccharomyces cerevisiae. Mol. Cell. Biol. 24, 4769–4780. Deshpande, A.M., and Newlon, C.S. (1996). DNA replication fork pause sites dependent on transcription. Science 272, 1030–1033. Dong, F., Weitzel, S.E., and von Hippel, P.H. (1996). A coupled complex of T4 DNA replication helicase (gp41) and polymerase (gp43) can perform rapid and processive DNA strand-displacement synthesis. Proc. Natl. Acad. Sci. USA 93, 14456–14461. Gibson, D.G., Aparicio, J.G., Hu, F., and Aparicio, O.M. (2004). Diminished S-Phase Cyclin-Dependent Kinase Function Elicits Vital Rad53-Dependent Checkpoint Responses in Saccharomyces cerevisiae. Mol. Cell. Biol. 24, 10208–10222. Ivessa, A.S., Lenzmeier, B.A., Bessler, J.B., Goudsouzian, L.K., Schnakenberg, S.L., and Zakian, V.A. (2003). The Saccharomyces cerevisiae helicase Rrm3p facilitates replication past nonhistone protein-DNA complexes. Mol. Cell 12, 1525–1536. Ivessa, A.S., Zhou, J.Q., Schulz, V.P., Monson, E.K., and Zakian, V.A. (2002). Saccharomyces Rrm3p, a 5# to 3# DNA helicase that promotes replication fork progression through telomeric and subtelomeric DNA. Genes Dev. 16, 1383–1396. Ivessa, A.S., Zhou, J.Q., and Zakian, V.A. (2000). The Saccharomyces Pif1p DNA helicase and the highly related Rrm3p have opposite effects on replication fork progression in ribosomal DNA. Cell 100, 479–489. Katou, Y., Kanoh, Y., Bando, M., Noguchi, H., Tanaka, H., Ashikari, T., Sugimoto, K., and Shirahige, K. (2003). S-phase checkpoint proteins Tof1 and Mrc1 form a stable replication-pausing complex. Nature 424, 1078–1083. Kim, S., Dallmann, H.G., McHenry, C.S., and Marians, K.J. (1996). Coupling of a replicative polymerase and helicase: A tau-DnaB interaction mediates rapid replication fork movement. Cell 84, 643– 650. Lee, J., Kumagai, A., and Dunphy, W.G. (2003). Claspin, a Chk1regulatory protein, monitors DNA replication on chromatin independently of RPA, ATR, and Rad17. Mol. Cell 11, 329–340. Lengronne, A., Pasero, P., Bensimon, A., and Schwob, E. (2001). Monitoring S phase progression globally and locally using BrdU
Nedelcheva, M.N., Roguev, A., Dolapchiev, L.B., Shevchenko, A., Taskov, H.B., Stewart, A.F., and Stoynov, S.S. (2005). Uncoupling of unwinding from DNA synthesis implies regulation of MCM helicase by Tof1/Mrc1/Csm3 checkpoint complex. J. Mol. Biol. 347, 509–521.
Osborn, A.J., and Elledge, S.J. (2003). Mrc1 is a replication fork component whose phosphorylation in response to DNA replication stress activates Rad53. Genes Dev. 17, 1755–1767. Osborn, A.J., Elledge, S.J., and Zou, L. (2002). Checking on the fork: the DNA-replication stress-response pathway. Trends Cell Biol. 12, 509–516. Rothstein, R., Michel, B., and Gangloff, S. (2000). Replication fork pausing and recombination or “gimme a break”. Genes Dev. 14, 1–10. Sar, F., Lindsey-Boltz, L.A., Subramanian, D., Croteau, D.L., Hutsell, S.Q., Griffith, J.D., and Sancar, A. (2004). Human claspin is a ringshaped DNA-binding protein with high affinity to branched DNA structures. J. Biol. Chem. 279, 39289–39295. Tanaka, K., and Russell, P. (2001). Mrc1 channels the DNA replication arrest signal to checkpoint kinase Cds1. Nat. Cell Biol. 3, 966–972. Tercero, J.A., and Diffley, J.F. (2001). Regulation of DNA replication fork progression through damaged DNA by the Mec1/Rad53 checkpoint. Nature 412, 553–557. Tong, A.H., Lesage, G., Bader, G.D., Ding, H., Xu, H., Xin, X., Young, J., Berriz, G.F., Brost, R.L., Chang, M., et al. (2004). Global mapping of the yeast genetic interaction network. Science 303, 808–813. Torres, J.Z., Schnakenberg, S.L., and Zakian, V.A. (2004). Saccharomyces cerevisiae Rrm3p DNA helicase promotes genome integrity by preventing replication fork stalling: viability of rrm3 cells requires the intra-S-phase checkpoint and fork restart activities. Mol. Cell. Biol. 24, 3198–3212. Tourrière, H., Versini, G., Cordón-Preciado, V., Alabert, C., and Pasero, P. (2005). Mrc1 and Tof1 Promote Replication Fork Progression and Recovery Independently of Rad53. Mol. Cell 19, 699–706. Xu, H., Boone, C., and Klein, H.L. (2004). Mrc1 is required for sister chromatid cohesion to aid in recombination repair of spontaneous damage. Mol. Cell. Biol. 24, 7082–7090. Yuzhakov, A., Turner, J., and O'Donnell, M. (1996). Replisome assembly reveals the basis for asymmetric function in leading and lagging strand replication. Cell 86, 877–886. Zhao, H., and Russell, P. (2004). DNA binding domain in the replication checkpoint protein Mrc1 of Schizosaccharomyces pombe. J. Biol. Chem. 279, 53023–53027.
Mrc1 Is Required for Normal Progression of Replication Forks throughout Chromatin in S. cerevisiae Shawn J. Szyjka, Christopher J. Viggiani, and Oscar M. Aparicio* Molecular and Computational Biology Program Department of Biological Sciences University of Southern California Los Angeles, California 90089
Summary Mrc1 associates with replication forks, where it transmits replication stress signals and is required for normal replisome pausing in response to nucleotide depletion. Mrc1 also plays a poorly understood role in DNA replication, which appears distinct from its role in checkpoint signaling. Here, we demonstrate that Mrc1 functions constitutively to promote normal replication fork progression. In mrc1D cells, replication forks proceed slowly throughout chromatin, rather than being specifically defective in pausing and progression through loci that impede fork progression. Analysis of genetic interactions with Rrm3, a DNA helicase required to resolve paused forks, indicates that Mrc1 checkpoint signaling is dispensable for the resolution of stalled replication forks and suggests that replication forks lacking Mrc1 create DNA damage that must be repaired by Rrm3. These findings elucidate a central role for Mrc1 in normal replisome function, which is distinct from its role as a checkpoint mediator, but nevertheless critical to genome stability. Introduction The accurate duplication of chromosomal DNA presents a significant challenge to the cell. Replication forks must contend with the presence and activities of various chromatin structures, transcription complexes, and recombination machinery, which often cause replication forks to pause (reviewed in Rothstein et al., 2000). For example, replication forks pause at tRNA genes when the direction of transcription is opposite that of the replication fork (Deshpande and Newlon, 1996). Although the exact function of fork pausing at tRNA genes is not known, it may help prevent fork dysfunction that might result from collision with the transcription machinery, and thus, facilitate resumption of normal fork progression. Multiple mechanisms contribute to replication fork stability and faithful genome duplication. For example, the Rrm3 DNA helicase has been implicated as playing an important role in the progression of replication forks through natural impediments such as tRNA genes and chromatin complexes (Ivessa et al., 2003, 2002, 2000). In addition, surveillance mechanisms called checkpoints detect and respond to problems that arise during DNA replication. A critically important pathway that *Correspondence: [email protected]
Short Article
operates during S phase is the replication stress checkpoint (reviewed in Osborn et al., 2002). This pathway acts by maintaining adequate deoxyribonucleotide (dNTP) levels, increasing transcription of DNA repair genes, slowing cell cycle progression, and maintaining stable replication forks that are poised to resume replication following perturbations. Mediator of replication checkpoint protein 1 (Mrc1) localizes with replication forks and acts in the replication stress response by transducing signals of stress from the “sensor” kinase Mec1 to the “effector” kinase Rad53 (Alcasabas et al., 2001; Katou et al., 2003; Osborn and Elledge, 2003; Tanaka and Russell, 2001). Thus, in cells challenged with an inhibitor of DNA synthesis such as hydroxyurea (HU), which depletes deoxyribonucleotide levels, Mrc1 is required for effective activation of Rad53 by Mec1. Remarkably, in mrc1D cells undergoing DNA replication in the presence of HU, the progression of DNA synthesis halts, while replication fork proteins, including Mcm proteins, Cdc45 and DNA polymerases, appear to progress farther along the chromatin, suggesting that uncoupling of the replication apparatus from the site of DNA synthesis has occurred (Katou et al., 2003). This has led to the conclusion that Mrc1 functions in a pausing complex that maintains replisome integrity when exogenous stresses are encountered. Mrc1 also functions during normal DNA synthesis in cells that have not received exogenous perturbation. This is evidenced by the somewhat-slower rate of bulk chromosomal DNA synthesis in mrc1D cells, which is accompanied by a Rad9-dependent DNA damage response (Alcasabas et al., 2001). The reason for the slower replication of mrc1D cells remains unclear but does not depend on checkpoint signaling by either Rad9 or Mrc1 (Alcasabas et al., 2001; Osborn and Elledge, 2003). It is also unknown whether the defective replisome pausing in HU reflects loss of the replication or checkpoint-signaling function of Mrc1. In this study, we have investigated the role of Mrc1 in DNA replication of unperturbed cells, by examining origin initiation, fork elongation, and fork pausing in cells lacking Mrc1 function. We also have explored the possible role of Mrc1 checkpoint signaling in response to fork pausing by analyzing genetic interactions with Rrm3. Together, our findings demonstrate that Mrc1 plays a central and constitutive role in the efficient function of the replisome. Our results also suggest that Rrm3 is vital to repair DNA damage created by defective replication forks in mrc1D cells. Results and Discussion Mrc1 Is Required for a Normal Rate of Replication Fork Progression throughout Chromatin The replication defect of cells lacking Mrc1 could be due to a defect in the initiation of replication, in elongation, or in replisome pausing. To distinguish between these possibilities, we monitored origin initiation and
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Figure 1. Mrc1 Is Required for Normal Replication Fork Progression along Chromosome VI (A) Position of probes (thin bars beneath the chromosome depiction) and relevant ClaI (C) and EcoRI (E) restriction sites used in the 2D gel analysis of modified chromosome VI are shown approximately to scale. Thick bars indicate the positions of sequences analyzed for BrdU incorporation and Cdc45 association shown in Figures 2 and 4. The bent arrows indicate the positions and transcriptional orientations of the tRNA genes tA(AGC)F in region A and SUP6 in region B. (B) Wild-type (SSy48) and mrc1D (SSy47) cells were blocked in G1 with α factor at 23°C and released synchronously into S phase at 20°C. Cells were collected at the indicated times for 2D gel analysis by digestion with EcoRI and ClaI. Blots were probed for ARS607 and stripped and reprobed for regions A, B, and C. Filled and unfilled arrowheads indicate small and large bubble structures, respectively; open and double arrowheads indicate large and small fork structures, respectively; the complete arrows indicate pause sites. The greater signal intensity of replication structures in mrc1D cells is reproducible and appears to reflect the longer period of time that forks are present within each analyzed fragment.
the progression of replication forks using two-dimensional gel electrophoresis (2D gel) across regions of chromosomes III and VI. In each of these regions, one or more active replication origins has been deleted, so that in wild-type cells, the replication of these regions is unidirectional, facilitating analysis of fork progression (Figures 1A and 3A) (Tercero and Diffley, 2001). In wild-type cells released synchronously into S phase, initiation of ARS607 occurred at about 30 min. By 36 min, primarily larger replication structures were present due to fork elongation (Figure 1B). In mrc1D cells, initiation of ARS607 occurred with timing like that of wild-type cells, as bubbles were first observed at 30 min (Figure 1B). The shapes of the bubble arcs and their greater intensities suggested a delay in the elongation of initiation bubbles in mrc1D cells. At 30 min, small bubbles predominated (Figure 1B), and at 36 min, small and large bubbles were present, as well as large forks (Figure 1B). These larger structures persisted near ARS607 at 48–60 min. The presence of small bubbles at 30–36 min together with their absence at later times, as in wild-type cells, indicates that initiation timing of these origins is not altered by the absence of Mrc1 (also, see below). Furthermore, the efficiency of ARS607 is not altered by loss of Mrc1 based on the virtual absence of smaller fork structures in these gels and on the analysis of unsynchronized cultures (data not shown). These results suggest that replication forks
emanating from ARS607 elongated more slowly in mrc1D cells than in wild-type cells. Next, we analyzed the progression of replication forks from ARS607 into adjacent chromosome VI sequences, restriction fragments “A,” “B,” and “C,” whose midpoints lie at 4, 12, and 20 kb from ARS607 (Figure 1A). In addition, regions A and B each contain a tRNA gene that permitted analysis of replication fork pausing (discussed below). In wild-type cells, the peak signals for forks at A, B, and C occurred at 36, 48, and 60 min, respectively (Figure 1B). Based on the initial appearance of replication structures at ARS607 at 30 min and at region C at 48 min, the data indicate an average fork rate of w1 kb per minute in wild-type cells. In the absence of Mrc1, the progression of replication forks into and throughout the adjacent chromosomal regions was significantly delayed. The peak signals for forks at regions A, B, and C occurred at about 48, 72, and 84 min, and forks were present within each region for a longer period, consistent with their slower movement through each region (Figure 1B). Furthermore, the preponderance of smaller forks earlier and larger forks later in the interval during which forks move through each region is consistent with slow movement of replication forks throughout the region (Figure 1B). Based on the timing of initial appearance of replication structures at ARS607 and at region C, we estimate that replication forks progress at about half the wild-type rate in
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Figure 2. Mrc1 Is Required for Normal Progression of DNA Synthesis Wild-type (CVy39), mrc1-AQ (CVy40), and mrc1D (CVy29) cells, which all express GPD-TK, were blocked in G1 with α factor at 23°C and released synchronously into S phase at 20°C. (A) DNA content analysis. (B) Cells were collected at the indicated times for analysis of BrdU incorporation. Immunoprecipitated DNA sequences at ARS607 and at distances of 5, 15, and 20 kb (see Figure 1A) were detected by PCR amplification. (C) PCR products in (B) and of input DNA (data not shown) were quantified and plotted as the ratio of immunoprecipitated DNA/ input DNA (% BrdU Incorporated).
mrc1D cells. Together with the slow progression of total DNA synthesis in mrc1D cells (Figure 2A) (Alcasabas et al., 2001), these findings suggest that Mrc1 is required for normal replication fork progression throughout much or all of the genome. Whether replication forks in mrc1D cells progress at a uniformly slow rate or whether the slow progression is due to occasional stalling at random sites (requiring restart) is not clear. However, the perdurance of some replication forks at later
times (e.g., Figure 1B, ARS607 at 72–96 min) suggests that some forks have stalled but remain stable. We also analyzed initiation and fork progression in a strain harboring the mrc1-AQ allele, in which all hypothetical Mec1 phosphorylation sites have been mutated (Osborn and Elledge, 2003). This allele is defective in checkpoint signaling to activate Rad53, but appears to retain its function in DNA replication, based on its approximately normal rate of replication as measured by DNA content analysis (Figure 2A) (Osborn and Elledge, 2003). 2D gel analysis indicated that initiation and fork movement in the mrc1-AQ strain was similar to wildtype cells (Figure S1 in the Supplemental Data available with this article online). Thus, replication forks in mrc1AQ cells progress with similar kinetics as in wild-type cells, and the slower replication fork movement of mrc1D cells is not due to loss of Mrc1’s checkpoint function. To confirm our conclusion that normal fork progression depends on Mrc1, but not its checkpoint function, we analyzed nascent DNA synthesis by measuring incorporation of the thymidine analog bromodeoxyuridine (BrdU) along chromosome VI (Figures 2B and 2C). In wild-type mrc1-AQ and mrc1D cells, BrdU incorporation occurred with similar timing and efficiency at the origin ARS607, primarily between 24 and 36 min, reinforcing our conclusion that initiation is not affected by loss of Mrc1 function. The kinetics of BrdU incorporation into DNA sequences at distances of 5, 15, and 20 kb from ARS607 was indistinguishable between wildtype and mrc1-AQ cells. However, BrdU incorporation into DNA sequences at distances of 5, 15, and 20 kb from ARS607 was delayed in mrc1D cells. The degree of delay is consistent with the results of the 2D gel analysis, with a delay of about 24 min at a distance of 20kb. Thus, in the absence of Mrc1, the replication timings of DNA sequences proximal to ARS607 were delayed in relation to their distance from the origin, with more distal sequences experiencing a greater delay. These data are fully consistent with the conclusion that replication forks progress more slowly in cells lacking Mrc1 function, whereas the checkpoint signaling function of Mrc1 appears to be dispensable for normal fork progression in unperturbed cells. To extend our analysis to other chromosomal regions and further test the idea that Mrc1 plays a general role in replication fork progression, we analyzed replication of a region of chromosome III adjacent to the efficient origin ARS306 (Figure 3A). In wild-type cells, initiation of ARS306 occurred at 30 min and replication forks were mainly detected at a distance of 23 kb from this origin at 48–60 min (Figure 3B, “Region D”). In mrc1D cells, ARS306 initiated at 30 min as in wild-type cells, and as observed with ARS607, replication bubbles from ARS306 appeared to elongate slowly in mrc1D cells (Figure 3B). Most replication forks in mrc1D cells did not reach region D until at least 72 min, thus having required at least 42 min to travel 23 kb. Together with the results of the chromosome VI analysis and total genomic DNA replication, these findings support the conclusion that normal progression of replication forks through much if not all of the genome requires Mrc1. We also analyzed replication of ARS304, an inefficient origin at a distance of 45 kb from ARS306. Unex-
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Figure 3. Mrc1 Is Required for Normal Replication Fork Progression along Chromosome III (A) Position of probes (thin bars beneath the chromosome depiction) and relevant ClaI (C) and EcoRI (E) restriction sites used in the 2D gel analysis of modified chromosome III are shown approximately to scale. (B) Blots from the experiments in Figure 1 were stripped and reprobed for ARS306, Region D, and ARS304. See Figure 2A for DNA content analysis.
pectedly, we observed increased initiation of ARS304 in mrc1D cells (Figure 3B). Because the level of ARS304 initiation in mrc1-AQ cells was similar to wild-type (Figure S1), we conclude that the increased initiation frequency of ARS304 in mrc1D cells is due primarily to the slower progression of replication forks from ARS306, which permits more time for ARS304 initiation before it is passively replicated by a fork from ARS306. We also have observed evidence for increased initiation of the inefficient origin near the VI-R telomere in mrc1D cells (data not shown). The increased initiation of some origins due to their delayed passive replication helps explain why genome duplication in mrc1D cells does not appear to require twice as long as wild-type cells (Figure 2A; and Tourrière et al., 2005 [this issue of Molecular Cell]), although our data indicate that forks progress at about half the wild-type rate. Progression of Cdc45 Is Slower in the Absence of Mrc1 The possibility that Mrc1 plays a role in coupling replication fork components to the site of DNA synthesis prompted us to examine whether the progression of Cdc45 along chromatin also was slowed in the absence of Mrc1 or whether it might move rapidly ahead of the slow-moving replication forks. To examine the movement of Cdc45, we analyzed its association with chromosome VI by chromatin immunoprecipitation (ChIP) (Figure 4). This analysis was performed at 16°C to enhance the resolution of the assay. In both wild-type and mrc1D cells, Cdc45 association with ARS607 peaked at 60 min and diminished soon after. The similar timings
in wild-type and mrc1D cells reinforce our conclusion that initiation timing of ARS607 is not altered by loss of Mrc1 function. However, the progression of Cdc45 along chromatin was altered in the absence of Mrc1. In wild-type cells, Cdc45 moved rapidly, associating maximally with sequences 5, 15, and 20 kb distant from ARS607 at 72, 72, and 84 min, respectively. In mrc1D cells, Cdc45 progressed more slowly, associating maximally with the sequences 5, 15, and 20 kb from ARS607 at 72, 108, and 120 min, respectively. Hence, the progression of Cdc45 along chromatin during S phase, like that of replication forks, is slowed in the absence of Mrc1. Although the slowed progression of Cdc45 may be consistent with Cdc45 remaining coupled with the site of DNA synthesis in unperturbed mrc1D cells, it is possible that dissociation of Cdc45 from the site of DNA synthesis does occur, but can be resolved by our analysis only when DNA synthesis is inhibited strongly with HU. Whether or not a defect in coupling exists, the results make clear that Mrc1 is required for the normal progression of Cdc45 along chromatin during DNA replication. Resolution of Naturally Paused Replication Forks Does Not Require Mrc1 Next, we examined whether replication forks lacking Mrc1 were defective in progression through endogenous pause sites, which might contribute to the slower progression of replication forks that we have observed in mrc1D cells. We analyzed pausing at two tRNA genes within regions A and B adjacent to ARS607 (Figure 1A). Fork pausing at each tRNA gene was apparent
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fork pausing (Figure 3). We also note that mrc1D cells are not highly sensitive to HU, suggesting these cells are able to restart replication effectively (Alcasabas et al., 2001). We conclude that Mrc1 is not required for the stability and restart of naturally stalled forks.
Figure 4. Slower Progression of Cdc45 in mrc1D Cells Wild-type (CVy31) and mrc1D (CVy30) cells, which both express Cdc45-HA, were blocked in G1 with α factor at 23°C and released synchronously into S phase at 16°C. Cells were collected at the indicated times for ChIP analysis with anti-HA antibody. DNA sequences at ARS607 and at distances of 5, 15, and 20 kb (see Figure 1A) were quantified by PCR amplification. The ratio of immunoprecipitated to input DNA is plotted as “% Cdc45 Bound.”
as a dark spot within each fork arc. Compared to wildtype cells, the pause signals appeared reduced in mrc1D cells (Figure 1B, arrows). However, we suspected that the decreased pause signal resulted at least in part from the slower fork movement of mrc1D cells, which increased the length of time spent by replication forks throughout regions A and B, and thus decrease the relative intensity of the pause signal. Consistent with this idea, careful quantification of the pause at the tRNA gene in region A relative to total replication structures indicated that pausing was reduced by about half (Figure S2). Because replication forks progress at about half the wild-type rate, these data suggest that the duration of pausing at this tRNA gene is not significantly altered in the absence of Mrc1. In addition, these results strongly suggest that Mrc1 is not required for resolution of paused forks, which would appear as an increased pause signal. It was possible that paused forks in mrc1D cells were unstable and hence interfered with our ability to detect the paused structure itself and/or a defect in its resolution. To address directly the possibility that defective pausing somehow contributes to fork dysfunction in mrc1D cells, we analyzed fork progression along the 20 kb chromosome VI region in mrc1D cells in which we deleted both tRNA genes. Deletion of the tRNA genes did not significantly affect the rate of fork progression across this region (data not shown), indicating that the slow fork movement of mrc1D cells is not due to defective pausing or instability of forks encountering these sites. In support of this conclusion, forks appeared to progress slowly within all the regions that we analyzed, including the chromosome III region, which does not contain elements known to be associated with
The Replication Defect of mrc1D Cells Creates a Dependence on Rrm3 for Viability The Rrm3 DNA helicase enables the resolution of stalled replication forks. In the absence of Rrm3, fork pausing at natural sites is prolonged, and cells become dependent on MRC1 for viability (Ivessa et al., 2003, 2002; Ooi et al., 2003; Tong et al., 2004; Torres et al., 2004). These findings have suggested that the repair or restart of stalled forks in rrm3D cells is dependent on the replication stress response, which is mediated by Mrc1 (Torres et al., 2004). To determine whether the requirement of rrm3D cells for MRC1 reflects the role of Mrc1 in checkpoint signaling or in fork progression, we assessed the viability of rrm3D mrc1-AQ cells by dissection of a diploid strain heterozygous for both alleles. Haploid rrm3D mrc1-AQ cells emerged at a frequency equivalent to either single mutant strain and grew at a similar rate (Figure 5A). Thus, Mrc1’s checkpoint function is dispensable in cells lacking Rrm3. As expected, rrm3D mrc1D cells were inviable (Figure 5B). These results suggest that the lethality of rrm3D mrc1D cells reflects a dependence on Rrm3 to resolve replication fork defects incurred by loss of Mrc1 function, rather than a vital replication checkpoint response to prolonged fork stalling in rrm3D cells. To examine further the function of the replication checkpoint in response to fork stalling, we analyzed fork pausing at tRNA genes in wild-type and rrm3D cells with either an intact or defective replication checkpoint. Pausing was not altered in mrc1-AQ cells, indicating that the replication checkpoint is not required for the resolution of paused forks (Figure 5C, data not shown for SUP6). As expected, rrm3D cells exhibited greatly increased stalling at the tRNA gene (Figures 5C and 5D). Furthermore, we observed a similar level of fork pausing in rrm3D mrc1-AQ cells as in rrm3D cells (Figures 5C and 5D). Thus, checkpoint signaling by Mrc1 does not appear to participate in replication fork pausing or resolution, either in the presence or absence of Rrm3. Because mrc1D cells are defective in fork progression throughout chromatin and require Rrm3 for viability, we propose that Rrm3 acts throughout chromatin in response to replication fork defects and impediments. A Central Role for Mrc1 at Replication Forks? Previous analysis of mrc1D cells treated with HU has led to the conclusion that Mrc1 is required for normal fork pausing in response to replication stress by maintaining coupling of replication proteins to the site where DNA synthesis is inhibited (Katou et al., 2003). Our demonstration that lack of Mrc1’s replication function, but not its checkpoint function, causes defective fork progression throughout chromatin in untreated cells is consistent with a coupling role for Mrc1 in the replisome and further suggests this role is constitutive rather than checkpoint induced. Mrc1 coimmunopreci-
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The slower rate of fork progression in cells lacking Mrc1 may reflect uncoupling of replication factors. Studies of bacterial and viral eukaryotic DNA replication have demonstrated synergistic stimulation of helicase and polymerase activities that depend upon their physical association (Dong et al., 1996; Kim et al., 1996; Yuzhakov et al., 1996). If loss of Mrc1 function disrupts similar interactions in the eukaryotic replisome, slower fork progression might result. In addition to compromising the efficacy of DNA synthesis at the replication fork, loss of Mrc1 from the site of DNA synthesis may expose normal replication structures to DNA damage sensors, DNA repair proteins, and recombination factors. This could explain some of the defects associated with loss of Mrc1, such as activation of the DNA damage response and increased levels of homologous recombination (Alcasabas et al., 2001; Xu et al., 2004). These phenotypes may reflect the presence of excess unwound DNA, or DNA structures that may serve as effective recombination substrates such as doublestrand breaks, or 3# ends. The presence of such structures in mrc1D cells is suggested by their vital dependence on the Srs2 and Rrm3 DNA helicases (Ooi et al., 2003; Tong et al., 2004; Torres et al., 2004; Xu et al., 2004), which appear to function in the repair of stalled or damaged replication intermediates, perhaps by preventing their subversion into recombination pathways. The elucidation of a general role of Mrc1 in the function of replication forks provides insight into the mechanisms that promote genome stability during DNA replication. Experimental Procedures
Figure 5. Rrm3 Is Required for Viability of mrc1D Cells but Not mrc1-AQ Cells Diploid strains SSy108 and SSy109, in (A) and (B), respectively, were induced to sporulate at 23°C; germination was at 30°C. The relevant genotypes are indicated in the panels below as follows: WT = wild-type, AQ = mrc1-AQ, m1 = mrc1D, and r3 = rrm3D. The genotypes of inviable spores were inferred assuming 2:2 segregation of genetic loci within each tetrad. (C) Haploid strains W3031a (WT), SSy72 (mrc1-AQ), SSy69 (rrm3D), and SSy108 (mrc1-AQ rrm3D) were subjected to 2D gel analysis of tA(AGC)F by digestion with SpeI (see Figure S2A for position of probe and restriction sites). (D) Three sets of blots like those in (C) were quantified by measuring the pause signal and dividing by the signal for all replication intermediates along the fork arc (excluding the 1N spot). The data was plotted relative to wild-type, which was arbitrarily assigned the value 1. Error bars denote standard deviation.
pitates with Cdc45, Mcm2, and Mcm3 (Katou et al., 2003; Nedelcheva et al., 2005). Furthermore, recent biochemical analyses of S. pombe Mrc1 and its apparent human homolog, Claspin, indicate that Mrc1/Claspin can bind branched DNA structures in vitro (Sar et al., 2004; Zhao and Russell, 2004). Together, these data suggest that Mrc1 links Cdc45 and/or Mcm proteins to replication forks in vivo. Depletion of Claspin from Xenopus replication extracts also modestly reduced the rate of DNA synthesis suggesting that Claspin may play a similar role at replication forks in vertebrates (Lee et al., 2003).
Plasmid and Strain Constructions Strains are described in Table S1. Methods for strain construction have been described previously (Gibson et al., 2004). pRS306Mrc1-AQ contains the NotI-KpnI mrc1-AQ fragment from pRS405Mrc1-AQ (Gibson et al., 2004) inserted into pRS306. mrc1-AQ was introduced at its native locus by a pop-in-pop-out strategy, or by a simple pop-in. Yeast Methods YEPD medium was used for all experiments. Cell culturing, synchronization, DNA content analysis, spore analysis, and ChIP have been described (Aparicio et al., 2004; Gibson et al., 2004). Twentytwo cycles of PCR were performed for analysis of the ChIP experiment. Analysis of Replication Structures 2D gel analysis was performed as described previously, except that 30 g of DNA was used for each sample (Gibson et al., 2004). Quantitative comparisons were performed on samples run in the same gels and blotted and hybridized together to minimize any errors that might result from differential transfer or hybridization efficiencies. Analysis of BrdU Incorporation Strains expressing seven copies of HSV-TK were grown in the presence of 400 g/ml BrdU as described (Lengronne et al., 2001). Immunoprecipitation of BrdU-substituted DNA was carried out as described in (Katou et al., 2003) and references therein, with the following modifications: DNA was extracted by glass bead beating, sheared by sonication to an average size of w500 bp, and isolated by phenol/chloroform extraction and ethanol precipitation. DNA was treated with RNaseA and Proteinase K and further purified on Qiaquick PCR purification spin-columns (Qiagen). 500 ng of DNA was combined with 20 g of sheared salmon sperm DNA, dena-
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tured, and incubated with anti-BrdU antibody (Oxford Biotechnology) followed by G-Sepharose beads (Amersham). 22 cycles of PCR were performed on the immunoprecipitated material and on a 1:10 dilution of the input material. Sequences of primers used for PCR analyses are available upon request.
incorporation in TK(+) yeast strains. Nucleic Acids Res. 29, 1433– 1442.
Supplemental Data Supplemental Data include two additional figures and a table and can be found with this article online at http://www.molecule.org/ cgi/content/full/19/5/691/DC1/.
Ooi, S.L., Shoemaker, D.D., and Boeke, J.D. (2003). DNA helicase gene interaction network defined using synthetic lethality analyzed by microarray. Nat. Genet. 35, 277–286.
Acknowledgments We thank: N. Arnheim and M. Goodman for sharing equipment; J. Diffley, S. Elledge, and E. Schwob for strains and plasmids; P. Pasero for communicating unpublished data; and J. Aparicio, J. Campbell, J. Li, and R. Lipford for critical reading of the manuscript. This work was supported by NIH grant 1RO1GM65494 to O.M.A.
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