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Genetics. Feb 2007; 175(2): 553–566.
PMCID: PMC1800610

Sap1 Promotes the Association of the Replication Fork Protection Complex With Chromatin and Is Involved in the Replication Checkpoint in Schizosaccharomyces pombe

Abstract

Sap1 is involved in replication fork pausing at rDNA repeats and functions during mating-type switching in Schizosaccharomyces pombe. These two roles are dependent on the ability of Sap1 to bind specific DNA sequences at the rDNA and mating-type loci, respectively. In S. pombe, Swi1 and Swi3 form the replication fork protection complex (FPC) and play important roles in the activation of the replication checkpoint and the stabilization of stalled replication forks. Here we describe the roles of Sap1 in the replication checkpoint. We show that Sap1 is involved in the activation of the replication checkpoint kinase Cds1 and that sap1 mutant cells accumulate spontaneous DNA damage during the S- and G2-phases, which is indicative of fork damage. We also show that sap1 mutants have a defect in the resumption of DNA replication after fork arrest. Sap1 is localized at the replication origin ori2004 and this localization is required for the association of the FPC with chromatin. We propose that Sap1 is required to recruit the FPC to chromatin, thereby contributing to the activation of the replication checkpoint and the stabilization of replication forks.

ACCURATE DNA replication in eukaryotic genomes is a challenging task for the cellular machinery involved. Cells are constantly under stress from exposure to intrinsic and extrinsic agents that can interfere with the progression of the replication machinery (replisome). To manage and counter these events, cells are equipped with a quality control system, the DNA replication checkpoint, which is activated by stalled replication forks (Nyberg et al. 2002). Stalled forks threaten genomic integrity because they can potentially enable the rearrangement, breakage, or collapse through disassembly of the replisome complex (McGlynn and Lloyd 2002). In human cells, defects in the replication checkpoint can result in genomic instability, leading to both developmental and neurological defects and, significantly, a predisposition to cancer (Paulovich et al. 1997; Zhou and Elledge 2000; Abraham 2001; Nyberg et al. 2002; Kastan and Bartek 2004). In the fission yeast Schizosaccharomyces pombe, the Cds1 kinase enforces the replication checkpoint by coordinating both cell cycle arrest and DNA repair pathways (Boddy and Russell 2001). We have also shown in a previous study that Cds1 prevents fork collapse in response to deoxyribonucleotide triphosphate (dNTP) starvation (Noguchi et al. 2003), indicating that Cds1 is required to maintain replication forks in a replication-competent state. Likewise, in the budding yeast Saccharomyces cerevisiae, the inactivation of Rad53, a Cds1 homolog, is associated with replication fork collapse and loss of genomic integrity in response to dNTP starvation (Lopes et al. 2001; Tercero and Diffley 2001; Kolodner et al. 2002; Sogo et al. 2002).

Recently, we have shown that Swi1 and Swi3 form a “replication fork protection complex” (FPC) that is required for efficient Cds1 activation and the subsequent stabilization of replication forks in fission yeast (Noguchi et al. 2004). The Swi1–Swi3 complex is evolutionally conserved and is homologous to the Tof1–Csm3 complex in budding yeast and the Timeless–Tipin complex in humans (Gotter 2003; Lee et al. 2004; Mayer et al. 2004; Noguchi et al. 2004). In both budding and fission yeast, the FPC travels with the replication fork and probably acts during the very early stages of replication checkpoint signaling (Katou et al. 2003; Noguchi et al. 2004). The FPC has also been suggested to function in a novel S-phase checkpoint pathway that contributes to replication fork maintenance and survival following alkylation damage in fission yeast (Sommariva et al. 2005). In human cells, Timeless has been demonstrated to interact with Chk1 (a functional mammalian homolog of Cds1/Rad53) and ATR to control Chk1 activity (Unsal-Kacmaz et al. 2005). In addition, the downregulation of Timeless in human cells compromises replication and disrupts the intra-S-phase checkpoint (Unsal-Kacmaz et al. 2005), suggesting that there is functional conservation of the FPC across diverse species.

Swi1 and Swi3 are also required for programmed fork pausing near both the mating-type (mat1) locus and rDNA loci in fission yeast (Dalgaard and Klar 2000; Krings and Bastia 2004). This Swi1–Swi3-dependent fork pausing event at mat1 is required to initiate the gene conversion events that mediate mating-type switching (Dalgaard and Klar 2000). This mating-type switching process also requires switch-activating sites, SAS1 and SAS2, near the mat1 locus (Arcangioli and Klar 1991). SAS1 is known to interact with the Sap1 protein, another trans-acting factor that is also thought to be required for mating-type switching (Arcangioli et al. 1994a). Interestingly, recent studies have shown that Sap1 also binds the Ter1/RFB1 sequence, which has strong homology to SAS1 and is required for fork pausing at the rDNA loci (Krings and Bastia 2005; Mejia-Ramirez et al. 2005). The mechanisms by which the FPC and Sap1 contribute to programmed fork pausing are largely unknown, however. A previous structural study has shown that Sap1 acts as a dimer and bends DNA upon binding to its specific recognition site (Bada et al. 2000). Moreover, Sap1 appears to be essential for cell growth, and the overexpression of its C-terminal domain is associated with chromosomal fragmentation, the loss of minichromosomes, and the cut phenotype (Arcangioli et al. 1994a; de Lahondès et al. 2003), suggesting also that Sap1 plays important roles during DNA replication and in the maintenance of genomic integrity.

In this study, we report the identification of the sap1+ gene as a dosage suppressor of the swi3-1 mutation. We further show that Sap1 plays an important role in the activation of the replication checkpoint and in the stabilization of replication forks. Sap1 appears to be associated with the replication origin ori2004 and to be required for the recruitment of the FPC to chromatin, which contributes to the preservation of genomic integrity.

MATERIALS AND METHODS

General techniques:

The methods used for genetic and biochemical analyses of fission yeast have been described previously (Moreno et al. 1991; Alfa et al. 1993). Immunoblotting, Cds1 kinase assay, and ultraviolet (UV) sensitivity assay were performed as described in our earlier studies (Noguchi et al. 2004). To visualize nuclear DNA, cells were fixed in 70% ethanol for 20 min, washed in PBS, and stained with 4′,6-diamidino-2-phenylindole (DAPI). Pulsed-field gel electrophoresis was performed as described (Nakamura et al. 2002; Noguchi et al. 2002).

Chromatin immunoprecipitation assay:

The cell extracts used for chromatin immunoprecipitation (ChIP) assays were prepared from 5 × 108 S. pombe cells as described previously (Ogawa et al. 1999; Noguchi et al. 2004). Briefly, S. pombe cells (5 × 108) were fixed in 1% formaldehyde for 20 min at room temperature and then quenched in 125 mm glycine for 5 min. Cells were washed in TBS and disrupted in lysis buffer (50 mm Hepes–KOH pH 7.5, 140 mm NaCl, 1 mm EDTA, and 1% Triton X-100) supplemented with protease inhibitors [0.2 mm p-(Amidinophenyl) methanesulfonyl fluoride (p-APMSF) and Roche protease inhibitor cocktail]. The broken cells were sonicated six times for 20 sec each with a Misonix Sonicator 3000 until chromatin DNA was sheared into 500- to 700-bp fragments. Cell lysate was clarified by two rounds of maximum-speed centrifugation in an Eppendorf 5415C microcentrifuge at 4°. Immunoprecipitations were performed in these cell extracts using either anti-FLAG M2 agarose (Sigma–Aldrich, St. Louis) or anti-GFP agarose (Santa Cruz Biotechnology, Santa Cruz, CA). PCR amplification conditions and the specific primers used in these studies have also been described previously (Ogawa et al. 1999).

Isolation of the sap1+ gene:

swi3-1 cells were transformed with an S. pombe genomic library cloned into a pUR18 vector (Barbet et al. 1992) with a ura4+ selection marker. Transformed cells were selected on medium lacking uracil and supplemented with 10 μm camptothecin (CPT). Library-derived genomic DNAs cloned into pUR18 were then isolated from CPT-resistant colonies. We isolated clone CR13, among others, that appeared to contain only one open reading frame encoding the Sap1 protein, which we designated as pSap1.

Isolation of sap1 temperature-sensitive mutants:

Genomic DNA was isolated from S. pombe cells containing the sap1-3FLAG-Kanr gene. The sap1-3FLAG fragment was then amplified from this genomic DNA preparation by EXtaq polymerase (TaKaRa, Ohtsu, Japan) using the primers P294 and P296 and subsequently cloned into the AhdI site of pBluescript II TKS (+), to generate the pTKS-Sap1-3FLAG-Kanr construct. The sap1-3FLAG-Kanr gene was mutagenized by error-prone PCR using pTKS-Sap1-3FLAG as the template and the primers P294 and P296. Error-prone PCR was performed using five- and threefold higher than recommended concentrations of EXtaq and dNTPs, respectively. The wild-type sap1+ gene was then replaced with the mutagenized sap1-3FLAG-Kanr gene at the sap1 locus by a standard transformation method. Kanamycin-resistant colonies were isolated and their growth was examined at both 25° and 35° to select for temperature-sensitive mutants.

Gene cloning, plasmids, primers, and S. pombe strain construction:

The S. pombe strains used in this study were constructed using standard techniques (Alfa et al. 1993), and their genotypes are listed in Table 1. sap1-3FLAG (sap1-3FLAG-Kanr) and sap1-GFP (sap1-GFP-Kanr) were generated by a two-step PCR method (Krawchuk and Wahls 1999) using primers P295, P296, P300, and P301 to construct a 3xFLAG and a GFP tag at the C terminus of sap1, respectively. pFA6a-3FLAG-KanMX6 (Noguchi et al. 2004) and pFA6a-GFP-KanMX6 (Bähler et al. 1998) were used as the templates for the PCR-based gene tagging. The primers used in these procedures are available on request. Mutations and epitope-tagged genes have previously been described for cdc25-22 (Fantes 1979), swi1Δ (swi1::Kanr) (Noguchi et al. 2003), swi3Δ (swi3::Kanr) (Noguchi et al. 2004), swi3-1 (Gutz and Schmidt 1985), cds1Δ (cds1::ura4+) (Boddy et al. 1998), chk1Δ (chk1::ura4+) (al-Khodairy et al. 1994), rad3Δ (rad3::ura4+), rad22-YFP (rad22-YFP-Kanr) (Noguchi et al. 2003), swi1-GFP (swi1-GFP-Kanr) (Noguchi et al. 2003), swi3-GFP (swi3-GFP-Kanr) (Noguchi et al. 2004), swi1-3FLAG (swi1-3FLAG-Kanr) (Noguchi et al. 2004), uve1Δ (uve1::LEU2) (Yonemasu et al. 1997), rad13Δ (rad13::ura4+) (Yonemasu et al. 1997), and orp1-5FLAG (orp1-5FLAG-Kanr) (Ogawa et al. 1999).

TABLE 1
The S. pombe strains used in this study

RESULTS

Identification of sap1+ as a dosage suppressor of the swi3-1 mutation:

To understand how the FPC is regulated during its role in the stabilization of replication forks, we performed a genetic screen to isolate multicopy suppressors of the swi3-1 mutation. The swi3-1 mutation introduces an extra adenine at nucleotide position 126 in the open reading frame of the gene, causing a frameshift and a truncation of the Swi3 protein (Noguchi et al. 2004). However, overproduction of the swi3-1 allele can still partially suppress the mating-type switching defect of swi3Δ cells (data not shown), suggesting that this mutant gene might express reduced levels of functional Swi3 protein by translational readthrough or that the truncated protein product has weak biological activity. Nevertheless, the swi3-1 mutation renders fission yeast cells sensitive to a variety of genotoxic agents including CPT and hydroxyurea (HU) (Figure 1, A and B). CPT traps topoisomerase I by forming a single-strand nick in DNA. This causes breaks in the replication forks when they collide with the CPT-stabilized topoisomerase I–DNA complex (Tsao et al. 1993; Pommier 1996, 2004). HU depletes the dNTP pool by inhibiting ribonucleotide reductase and thereby leads to a replisome progression arrest.

Figure 1.
The isolation of sap1+ as a dosage suppressor of swi3-1. (A) Fivefold serial dilutions of swi3-1 cells harboring the indicated plasmids were incubated on yeast extract with supplements (YES) agar medium supplemented with 0 or 5 μm CPT ...

We transformed swi3-1 mutant cells with an S. pombe genomic library to screen for clones that could grow in the presence of CPT and identified the sap1+ gene (Figure 1A). Similar levels of suppression were also observed in the presence of HU (Figure 1B). To determine the mechanism underlying the suppression of swi3-1 mutation by sap1+, we investigated whether a dosage increase in the sap1+ gene could also suppress the CPT sensitivities of the swi1Δ or swi3Δ mutants. As shown in Figure 1C, sap1+ partially rescued the CPT sensitivity of swi1Δ and suppressed swi3Δ only very weakly, indicating that the presence of Swi3 is important for this partial suppression. These results together suggest that Sap1 may physically interact with Swi3 to facilitate a damage response to the effects of CPT. However, we performed a number of immunoprecipitation studies but did not detect any physical interaction between Swi3 and Sap1 (data not shown). Moreover, we did not identify any Sap1 peptides in our previous mass spectrometric analysis of Swi1-associated proteins (Noguchi et al. 2004). These results suggest that the Swi3–Sap1 interaction may be unstable and/or transient in the cell or that the association between these proteins is indirect.

Isolation of temperature-sensitive mutants of sap1:

To facilitate genetic analysis of sap1+, we isolated a number of temperature-sensitive (ts) mutants of the sap1 gene using the error-prone PCR method. Among these mutants, sap1-1 and sap1-48 show severe growth defects at 32°, whereas sap1-27 fails to grow at 35° (Figure 2A). We determined mutation sites of the sap1 ts alleles and found that sap1-1 contained a substitution at the 181st codon (TTA to TCA), leading to the amino acid change from leucine to serine (L181S). sap1-27 contained a substitution at the 41st codon (ATG to ACG), resulting in the amino acid change from methionine to threonine (M41T). sap1-48 contained a substitution at the 188th codon (ATG to AAG), leading to the amino acid change from methionine to lysine (M188K). An earlier study has shown that Sap1 has functional domains (domains I, II, and III) essential for SAS1-specific DNA interaction (supplemental Figure S1B at http://www.genetics.org/supplemental/). This study has also identified two additional domains (IV and V) required for efficient Sap1–DNA interaction and has found that domain IV is involved in Sap1 oligomerization (Arcangioli et al. 1994b) (supplemental Figure S1B). Interestingly, our mutational analysis revealed that the sap1-27 (M41T) mutation resides in domain I. This is identical to the mutation shown to have a defect in DNA binding reported in an earlier study (Arcangioli et al. 1994b). sap1-1 (L181S) and sap1-48 (M188K) mutations are novel and are both in domain IV required for oligomerization and efficient DNA binding (supplemental Figure S1B).

Figure 2.
Sap1 is required for cell survival after replication fork arrest. (A) Temperature-sensitive phenotypes of sap1-1, sap1-27, and sap1-48 mutants. Fivefold serial dilutions of cells were plated on YES agar medium and incubated at the indicated temperatures ...

Sap1 plays an important role in survival during S-phase following either replication fork arrest or DNA damage:

We first investigated whether Sap1 is involved in promoting cell survival following the generation of DNA lesions that block replication forks. UV irradiation causes the formation of cyclobutane dimers and other lesions that ultimately lead to an arrest in replisome progression. Serial dilutions of cells plated on agar growth medium were exposed to increasing doses of UV irradiation. These experiments were done at 25°, a permissive temperature for the sap1 ts mutations, and we observed that sap1-1 and sap1-27 cells were not sensitive to UV irradiation (data not shown), whereas sap1-48 cells display only weak sensitivity to UV irradiation (Figure 2C, top). This suggests that Sap1 does not have a direct role in the repair of UV lesions. To confirm this prediction, we next examined whether Sap1 contributes to cell survival in a mutant strain defective for both nucleotide excision repair (rad13Δ) and UV damage excision repair (uve1Δ). These two pathways are responsible for all detectable UV damage repair in fission yeast (Yonemasu et al. 1997). Moreover, strains that are defective for both pathways are acutely sensitive to UV irradiation, and their survival in these conditions depends upon either homologous recombination or translesion synthesis pathways that can bypass the UV damage during DNA replication (Friedberg et al. 1995; Woodgate 1999; Boddy et al. 2000). sap1-48 mutants were not sensitive to UV irradiation at low doses but this mutation reproducibly enhanced the UV sensitivity of a rad13Δ uve1Δ double mutant (Figure 2B). These results suggest that Sap1 is critical when UV-damaged DNA is not repaired and is therefore important for UV damage tolerance in S. pombe.

We also performed additional UV survival assays to detect genetic interactions between sap1-48 and mutations in checkpoint kinases. The checkpoint kinases Cds1 and Chk1 have overlapping roles during the S-phase stress responses (Rhind and Russell 2000; Boddy and Russell 2001). We found from our current experiments that sap1-48 chk1Δ cells were much more sensitive to UV irradiation than either of the single mutant cells. In contrast, there were no evident genetic interactions between sap1-48 and cds1Δ. The sap1-48 cds1Δ cells were equally sensitive to UV irradiation as the sap1-48 single mutant (Figure 2C, top). Chk1 is an effector kinase that functions during the G2-M DNA damage checkpoint and is activated by DNA damage during G2. Hence, our current findings suggest that the sap1-48 mutation leads to an increased level of DNA damage following UV exposure, creating an increased requirement for Chk1 activity during the G2-M checkpoint. These data also lend support to the idea that Sap1 is important for cellular tolerance to replication fork arrest caused by UV lesions. We obtained similar results using another ts mutant, sap1-1 (Figure 2C, top).

We next assessed whether Sap1 contributes to the response to stalled replication forks caused by HU or methylmethanesulfonate (MMS). HU is a ribonucleotide reductase inhibitor leading to an arrest in replisome progression. MMS causes the alkylation of template DNA bases and thereby disrupts the replication process. sap1-48 shows significant sensitivity to HU and weak sensitivity to MMS (Figure 2C, middle and bottom). These sensitivities were further strengthened by introduction of the chk1Δ background into the sap1-48 strain. In contrast, the sap1-48 mutation did not show synergistic interaction with cds1Δ in HU and MMS sensitivity assays (Figure 2C, middle and bottom). Taken together, our present data suggest that Sap1 is important for survival during a replication fork arrest that is provoked by exposure to HU and MMS. Interestingly, we also found synergistic interactions involving rad3Δ and either sap1-1 or sap1-48 in HU and MMS sensitivity assays. Rad3 is responsible for cell cycle arrest that is controlled by checkpoint kinase Chk1 and Cds1 (Boddy and Russell 2001; Nyberg et al. 2002). Therefore, the synergistic interaction between rad3Δ and sap1 suggests that Sap1 has an important function that is independent of cell cycle arrest, and this function may contribute to cell survival following a replication fork arrest induced by HU or MMS.

We also examined whether Sap1 is involved in the S-phase DNA damage response. For this purpose, we induced fork breakage during S-phase by exposure to CPT. sap1-48 displays significant sensitivity to CPT, suggesting that Sap1 does play an important role in the tolerance to DNA damage during S-phase. In addition, sap1-48 again showed a genetic interaction with chk1Δ but not with cds1Δ (Figure 2C, bottom).

In all of the above sensitivity assays for sap1 mutants, we performed similar experiments in h and h+ backgrounds and observed no significant difference between the two mating types (data not shown), indicating that the mating type has little or no effect on Sap1's function in tolerance to DNA damage and fork block.

Sap1 is involved in the replication checkpoint:

In all of our current sensitivity assays that we have already described, we observed synergistic genetic interactions between the sap1 and chk1Δ mutations. In contrast, sap1 cds1Δ mutants did not display a stronger sensitivity to S-phase stress agents, compared with either single mutant. Therefore, it is possible that Sap1 is involved in a Cds1-dependent checkpoint pathway. To address this possibility, we assessed the role of Sap1 in HU-induced division arrest. Because Cds1 and Chk1 have redundant roles in this arrest pathway (Boddy and Russell 2001; Rhind and Russell 2000), the role of Sap1 was evaluated in both a cds1Δ and a chk1Δ background. Interestingly, in the absence of HU, sap1-48 chk1Δ cells show a slight increase in the frequency of aberrant mitoses (6.3%) when compared with sap1-48 (2.1%) or sap1-48 cds1Δ (2.8%) (Figure 3, A and C). Moreover, when cells are treated with HU, the wild-type and sap1-48, cds1Δ and chk1Δ mutant strains undergo cell division arrest, whereas rad3Δ or cds1Δ chk1Δ mutant cells, which are unable to activate either Chk1 or Cds1, display a mitotic catastrophe or “cut phenotype” that is indicative of checkpoint failure (Figure 3, B and C). In addition, a significantly increased population of double-mutant sap1-48 chk1Δ cells fails to undergo cell division arrest following exposure to HU (19.2% aberrant mitoses), whereas this arrest pathway was functional in sap1-48 cds1Δ mutant cells (1.6% aberrant mitoses) (Figure 3, B and C). The data from these experiments thus suggest that Sap1 is involved in the HU-induced cell cycle arrest pathway that is mediated by Cds1, thus contributing to activation of the replication checkpoint. To further address the role of Sap1 in the activation of the replication checkpoint, Cds1 was immunoprecipitated and assayed using myelin basic protein as a substrate. HU was found to cause potent activation of Cds1 in wild-type cells, whereas Cds1 activation was strongly diminished in swi1Δ cells. A significant reduction in Cds1 activation was also observed in sap1-48 cells (Figure 3D), supporting our conclusion that Sap1 is involved in activation of the replication checkpoint in fission yeast.

Figure 3.
Sap1 is required for HU-induced cell cycle arrest enforced by Cds1. The indicated strains were incubated in YES liquid medium supplemented with 0 (A) or 12 mm HU (B) for 6 hr at 25° and then stained with DAPI to visualize nuclear DNA. A differential ...

To provide a molecular explanation of how the plasmid-borne Sap1 suppresses the drug sensitivities of the swi3-1 mutation, we examined whether the Sap1 plasmid is able to reactivate Cds1 kinase activity in swi3-1 and swi1Δ strains. However, we did not observe significant reactivation of Cds1 (data not shown). This result suggests that the suppression of swi3-1 by the Sap1 plasmid might not be due to reactivation of the Cds1 kinase. It is also possible that Cds1 kinase reactivation is not readily detectable in our kinase assay system. Nevertheless, Sap1 overexpression may be sufficient to protect replication forks when they are arrested in the presence of HU.

Sap1-48 mutant cells accumulate spontaneous Rad22 DNA repair foci:

Our data thus far show that Sap1 plays an important role in the replication checkpoint pathway. Interestingly, mutations in Sap1 were also found to significantly elevate the sensitivity levels to HU and MMS in a rad3Δ mutant that is defective in the activation of both the Cds1 and the Chk1 pathways (Figure 2C), suggesting that Sap1 has a checkpoint-independent role in the cellular tolerance to replication fork arrest following HU or MMS treatment. Moreover, we observed a slight increase in the frequency of aberrant mitoses in the unperturbed sap1-48 chk1Δ cells (Figure 3, A and C), suggesting that sap1-48 mutant cells may undergo spontaneous DNA damage, which should be repaired by the Chk1-dependent G2-M DNA damage checkpoint pathway. To address this possibility, we monitored the formation of Rad22 DNA repair foci in sap1-48 cells. The design of this experiment was based on previous studies using FPC-deficient cells, which display a large increase in the numbers of spontaneous Rad22 foci (Noguchi et al. 2003, 2004). The sap1-48 strain was engineered to express Rad22-YFP under its endogenous promoter. Fission yeast Rad22 is homologous to Rad52 in budding yeast, which is a protein that binds to single-stranded DNA (ssDNA) during homologous recombination (Paques and Haber 1999). Moreover, Rad22 is known to form nuclear foci at double-strand breaks and other sites that have exposed ssDNA segments (Kim et al. 2000; Du et al. 2003). We predicted therefore that if the sap1-48 cells were to incur spontaneous DNA damage as the result of malfunctioning replication forks, Rad22 would be recruited to the abnormal DNA structures containing ssDNA regions. As expected, a large increase in spontaneous Rad22-YFP foci was detectable in sap1-48 cells at 25°, a permissive temperature for sap1-48 (Figure 4A). In wild-type cells, 10.7% of the nuclei contained a single Rad22-YFP focus and only 0.4% contained two or more foci (Figure 4, A and B). In contrast, we observed a dramatic increase in the number of Rad22-YFP foci in sap1-48 cells whereby 37.7% of the sap1-48 mutant nuclei displayed at least one focus, and 6.3% harbored two or more foci (Figure 4, A and B). To explore whether Rad22 foci arose from replication abnormalities, the cell cycle position of cells containing Rad22-YFP foci was estimated by analyzing cell length, nuclei number and position, and the presence of a division plate (Figure 4B). This cell cycle analysis demonstrated that the Rad22-YFP foci formed during S-phase and were maintained during G2 phase in the sap1-48 cells (Figure 4B). These results suggest that sap1-48 cells may experience replication fork abnormalities during S-phase. Our data may also indicate that the repair of fork abnormalities is delayed in the sap1-48 cells, although further investigations will be needed to confirm this.

Figure 4.
Sap1 is involved in the stabilization of replication forks. (A) Cells expressing Rad22-YFP were grown in YES medium at 25° until midlog phase. A large increase in Rad22-YFP foci accumulation was observed in the sap1-48 cells. All images shown ...

Sap1 is known to be involved in mating-type switching. To address whether this function at the mating-type locus affects the formation of Rad22 foci, we examined Rad22-YFP localization in the smt0 background. In this background, S. pombe cells do not experience a double-strand break (DSB) at the mating-type locus. We compared smt0 sap1+, h sap1+, and h+ sap1+ cells and found no significant difference in Rad22 foci formation among these strains (supplemental Figure S2A at http://www.genetics.org/supplemental/). As expected, smt0 sap1-48 cells displayed a significant increase in Rad22-YFP formation compared to smt0 cells. However, this increase was further enhanced in h sap1-48 and h+ sap1-48 cells. Considering the fact that cells experience a DSB at the mating-type locus in h and h+ cells, our data suggest that Sap1 may be required for repair of DSB at the mating-type locus.

We have previously shown that inactivation of Swi1 or Swi3 causes a large increase in Rad22-YFP foci formation (Noguchi et al. 2003, 2004). Therefore, to rationalize the suppression of swi3-1 by the plasmid-borne Sap1 (Figure 1), we examined whether the Sap1 plasmid was able to suppress the accumulation of Rad22-YFP DNA repair foci formed in swi3-1 and swi1Δ cells. Although the effect is not dramatic, we observed a significant reduction of Rad22-YFP foci formation in swi3-1 and swi1Δ cells by the plasmid-borne Sap1 (supplemental Figure S2B at http://www.genetics.org/supplemental/). This result suggests that Sap1 contributes to fork protection by facilitating the function of Swi3.

Sap1 is required for the resumption of the replication fork after arrest:

To further explore the role of Sap1 in the maintenance of replication forks, we examined the recovery of DNA replication after fork arrest induced by HU exposure. Wild-type and sap1-48 cells were cultured at a permissive temperature (25°) and chromosome samples were prepared prior to (log) and at 5 hr post-HU treatment and also at different times during recovery after the removal of HU. These chromosomes were then resolved by pulsed-field gel electrophoresis (PFGE). Chromosomes from logarithmically growing cells in both wild type and sap1-48 were observed to have migrated into the gel, indicating that the sap1-48 cells have no significant defect in replicating DNA at 25° (Figure 4C, log). HU treatment caused an arrest of DNA replication, which in turn reduced the quantity of the chromosomes that could enter in the gel for both strains (Figure 4C, HU, 5 hr). Wild-type chromosomes migrated into the gel at 2 hr after the HU removal due to the completion of DNA synthesis. However, chromosomes from the sap1-48 cells did not migrate at this time point and displayed a reduced capacity to enter the gel at either 4 or 6 hr during recovery (Figure 4C). This indicates that Sap1 functions during the recovery of DNA replication after fork arrest. Hence, given the fact that sap1-48 cells also accumulate spontaneous DNA damage during DNA replication, we concluded that Sap1 is involved in the stabilization of replication forks.

It is noteworthy that we repeatedly observed that the sap1-48 strains harbor a shorter chromosome III (Figure 4C, log), which in S. pombe contains rDNA repeats. This result suggests that there is an increased mitotic recombination rate at the rDNA repeats in the sap1-48 cells. Similar findings have been reported in a number of replication mutants including swi1Δ and swi3Δ (Sommariva et al. 2005), further suggesting that Sap1 plays an important role in maintaining proper DNA replication and genomic integrity.

Sap1 is associated with the replication origin ori2004:

FPC has been shown previously to be a component of replication forks (Katou et al. 2003; Noguchi et al. 2004; Calzada et al. 2005). To determine whether Sap1 also functions at the replication forks, we examined its localization by ChIP analysis. The cdc25-22 strain was engineered to express a FLAG-tagged Sap1 protein (Sap1-3FLAG) via its endogenous gene promoter. All sap1-3FLAG strains showed normal growth rates and no detectable sensitivity to HU, MMS, CPT, or UV irradiation, indicating that Sap1-3FLAG is functional. The cdc25-22 allele was utilized to synchronize the cells at the G2-M boundary. We subsequently examined the localization of Sap1-3FLAG at the well-characterized replication origin ori2004 on S. pombe chromosome II (Ogawa et al. 1999). The movement of Sap1 along the chromosome was monitored at ori2004 and at two positions 14 and 30 kb away (Figure 5, A and H). Sap1 showed only a weak association with ori2004 in G2-arrested cells. Upon release from the cdc25-22 arrest at the G2-M boundary, Sap1-3FLAG was observed to strongly associate with this replication origin at 60–80 min, which subsequently declined by 180 min. There was no significant Sap1 association with the chromatin at the distal 14- and 30-kb positions throughout the experiment (Figure 5A).

Figure 5.
Sap1 associates with the replication origins ori2004 and is required for the recruitment of the FPC to chromatin. (A) ChIP assays of Sap1-3FLAG were performed at ori2004 and at sites located 14 or 30 kb away from this origin. cdc25-22 cells were synchronized ...

We also examined septation to monitor cell cycle progression, which in fission yeast occurs in S-phase. The level of Sap1 association with ori2004 was found to correlate with an increase in the septation index, which also coincided with the onset of S-phase, and this interaction was found to decline as the septation index decreased (Figure 5A), indicating that Sap1 tightly associates with ori2004 during S-phase. As controls, we performed ChIP analyses of Orp1-FLAG and Rad11-3FLAG. Orp1 is a component of the origin recognition complex and associates with replication origins throughout the cell cycle (Ogawa et al. 1999), whereas Rad11 is the large subunit of the replication protein A (RPA) complex that migrates with the replication forks (Noguchi et al. 2004). Consistent with previous reports (Ogawa et al. 1999), Orp1-5FLAG was also found to localize at ori2004, and this association persisted throughout the cell cycle (Figure 5C). In contrast, Rad11 associated with ori2004 at the onset of S-phase and then relocated from ori2004 to the 14- and 30-kb positions during the course of the experiment (Figure 5D). The association of Rad11 with ori2004 peaks between 60 and 80 min. Rad11 most strongly associates with the 14-kb position at 80 min and with the 30-kb position between 80 and 120 min, confirming that Rad11 travels with replication forks. This is in striking contrast with Orp1 and Sap1, both of which localize only at ori2004. These results thus strengthen our conclusion that Sap1 tightly associates with the replication origin ori2004 during S-phase.

To confirm Sap1 association with ori2004, we reduced the rate of fork movement by releasing cdc25-22 sap1-3FLAG cells from G2-M block in the presence of hydroxyurea. As shown in Figure 5B, Sap1 association with ori2004 again becomes prominent as the septation index increased. This association persisted throughout the extended S-phase due to the hydroxyurea treatment. Again, we found no significant increase in the level of Sap1 association with the 14- and 30-kb positions. Thus, we concluded that Sap1 tightly associates with ori2004 throughout S-phase.

Since Sap1 has only a weak association with ori2004 outside of S-phase, we further examined localization of Sap1-GFP within the cells. Strains carrying the sap1-GFP gene from its native promoter showed normal growth rate and no significant sensitivity to HU and CPT, suggesting that Sap1-GFP is functional. Sap1-GFP colocalized with DAPI-stained DNA in cells at all stages of the cell cycle and in HU-treated cells (data not shown). In situ chromatin-binding assays (Noguchi et al. 2003) were used to determine whether Sap1-GFP interacts with chromatin. After Triton X-100 extraction of soluble nuclear proteins, Sap1-GFP was still colocalized with DNA in all cells (data not shown). This might represent the localization of Sap1 to nonorigin DNA sequences such as the rDNA and mating-type loci as previously reported (Arcangioli et al. 1994a; Krings and Bastia 2005; Mejia-Ramirez et al. 2005). However, we speculate that Sap1 has a weak affinity to origins outside of S-phase, but that this association becomes very tight during S-phase as ChIP analysis clearly showed that Sap1's association with ori2004 increased during S-phase.

Sap1 is required for the association between FPC and chromatin:

To further understand how the Sap1 protein contributes to the stabilization of replication forks, we examined whether Sap1 mutants can also associate with ori2004. FLAG-tagged Sap1 temperature-sensitive mutant proteins were expressed from the endogenous sap1 promoter and ChIP analyses were performed with cells expressing these mutants at a permissive temperature. As shown in Figure 5E, wild-type Sap1 displayed a tight association with ori2004 in an asynchronous culture. However, all three Sap1 mutant proteins displayed a strongly reduced level of association with ori2004, suggesting that the chromatin association of Sap1 is important for the stabilization of the replication fork. This result is consistent with our mutational analysis showing that sap1 mutants contain mutations in domains required for Sap1–DNA interaction (supplemental Figure S1B at http://www.genetics.org/supplemental/).

We then determined whether the association of Sap1 with chromatin is required for FPC localization to chromatin. A sap1-48 strain was engineered to express either Swi1-GFP or Swi3-GFP from the endogenous gene promoter. Moreover, the cells expressing these fusion products were found to be resistant to S-phase stressing agents, indicating that Swi1-GFP and Swi3-GFP are fully functional. When ChIP was performed with Swi1-GFP or Swi3-GFP in a sap1+ wild-type background, both the GFP-tagged proteins associated strongly with the 14- and 30-kb positions and only weakly with ori2004 in asynchronous cultures (Figure 5F). These results indicate that, under these conditions, Swi1 and Swi3 have already relocated away from ori2004 as the replication fork progresses. In contrast, when ChIP was performed with Swi1-GFP or Swi3-GFP in a sap1-48 background, we observed a dramatic decrease in the levels of Swi1–Swi3 associated with the ori2004 flanking region (Figure 5F). We also obtained similar results in the presence of HU (Figure 5G). In this condition most of the replication origins have already fired, explaining why Swi1 and Swi3 were not observed to be concentrated at ori2004. Similarly, Swi1 and Swi3 failed to associate with chromatin in the sap1-48 background (Figure 5G). In contrast, the association of Sap1 with ori2004 was unaffected by the absence of Swi1 or Swi3 (data not shown). Taken together with the evidence that the Sap1-48 protein fails to associate with ori2004 (Figure 5E), our results suggest that the association between Sap1 and chromatin is required for FPC recruitment to the replication forks.

DISCUSSION

In our study, we have described the results of a series of experiments that establish the involvement of Sap1 in replication (S-phase) stress response pathways in fission yeast. These analyses include the assessment of the role of Sap1 in the stabilization of replication forks and in the Cds1-dependent replication checkpoint pathway. We demonstrate from these experiments that Sap1 associates with ori2004 and is required for FPC recruitment to the ori2004-flanking region. On the basis of our present data, we thus propose that Sap1 assists in the maintenance of genomic integrity probably by recruiting the replication fork protection complex to the origins of replication.

The role of Sap1 in the activation of the replication checkpoint and in fork stabilization:

Recent studies have identified a group of proteins that are required for activation of the replication checkpoint and the stabilization of replication forks in fission yeast. Mrc1, a mediator of the replication checkpoint, is known to be essential for Cds1 activation in a Rad3-dependent manner (Tanaka and Russell 2001; Zhao et al. 2003). Swi1, a Tof1/Timeless-related protein, which is required for replication fork pausing at both mating-type and rDNA loci, is also important for proper activation of Cds1 (Noguchi et al. 2003). In addition, Swi1 forms a FPC with Swi3 and is involved in replication fork stabilization (Noguchi et al. 2004). Hsk1-Dfp1, the Cdc7-Dbf4-related kinase that interacts with FPC, is also important for the activation of Cds1 (Takeda et al. 2001; Matsumoto et al. 2005). In our present study, we describe an additional factor, Sap1, which is required for the activation of the replication checkpoint and the stabilization of replication forks. Sap1-48 mutant cells show decreased levels of Cds1 activation in response to HU and further show increased sensitivities to a variety of agents that lead to a replication fork arrest (Figures 2 and and3).3). Furthermore, in a similar manner to the FPC (Noguchi et al. 2004), Sap1 is required for the resumption of DNA replication after HU-induced fork arrest (Figure 4C). Interestingly, our data also suggest that Sap1 may associate with the replication origins (Figure 5) and that this is a prerequisite for the localization of FPC to the replication forks (Figure 5, F and G), suggesting that Sap1 recruits FPC to these forks. This function requires DNA binding and oligomerizing activities of Sap1 since mutations in DNA-binding and oligomerization domains greatly reduced the level of the Sap1-ori2004 association (supplemental Figure S1B at http://www.genetics.org/supplemental/ and Figure 5E). In our current model, upon the initiation of replication, Sap1 remains localized at the replication origins while the FPC migrates with the replication forks. Hence, Sap1 may not have any direct activity in fork stabilization. Although we have not successfully shown that there is a physical interaction between Sap1 and FPC, our present genetic data suggest that there may well be a weak or transient association between these factors. Taken together, the results of our current study suggest that Sap1 recruits the FPC to the replication forks and plays an important role in mediating the timely activation of the replication checkpoint. Our findings also indicate that Sap1 functions in replication fork stabilization.

The role of Sap1 in replication fork pausing:

To date, Sap1 has been reported to interact with at least two cis-elements, SAS1 of the mating-type locus and Ter1/RFB1 of the rDNA array (Arcangioli et al. 1994a; Krings and Bastia 2005; Mejia-Ramirez et al. 2005). Sap1 has also been shown to be involved in the induction of a stalled replication fork at rDNA repeats (Krings and Bastia 2005; Mejia-Ramirez et al. 2005). In addition, the stalled fork at the Ter1/RFB1 site also requires Swi1 and Swi3 (Krings and Bastia 2004), both of which are components of the replication forks (Noguchi et al. 2004). Hence, we speculate that the FPC associates with Sap1 at the Ter1/RFB1 site to halt fork progression. However, it has been shown that Sap1 is not required for the stalled replication fork at SAS1 of the mating-type locus (Dalgaard and Klar 2000), suggesting that more complicated mechanisms control the halting of replication forks, involving Swi1–Swi3 and Sap1.

Fork pausing may inevitably involve the stabilization of replication forks, which would be required to avoid collapse or rearrangements at programmed pausing sites, which are spread throughout the genome. In our present study, we provide evidence that Sap1 may also associate with replication origins. In addition, since Sap1 recruits the FPC to chromatin, we propose that the Sap1–FPC interaction is critical for the recruitment of the FPC to facilitate fork stabilization and fork pausing.

Sap1's roles in cell viability:

Earlier studies have suggested that Sap1 acts downstream of Swi1 and Swi3 during the process of mating-type switching in fission yeast (Dalgaard and Klar 2000). However, Sap1 is essential for cell viability, indicating its important roles in other general cellular functions (Arcangioli et al. 1994a; de Lahondès et al. 2003). Moreover, we have consistently found putative Sap1 homologs in a variety of other organisms that do not employ mating-type switching. For example, Sap1 shows significant similarity to Girdin (also known as APE, GIV, or Hook-related protein) in humans and mice (Anai et al. 2005; Enomoto et al. 2005; Le-Niculescu et al. 2005; Simpson et al. 2005) and also to the Xenopus LOC431973 protein (supplemental Figure S1A at http://www.genetics.org/supplemental/), indicating its conservation throughout evolution. However, we could not identify a Sap1-like protein in budding yeast. Sap1 comprises a DNA-binding domain, a dimerization domain, and a C-terminal domain (Arcangioli et al. 1994b) (supplemental Figure S1A) and it is significant that the regions of greatest homology reside in the DNA-binding and oligomerization domains, indicating that the functions of Sap1 in the replication checkpoint and during fork stabilization may be conserved. Interestingly, in human, Girdin has been implicated in the regulation of DNA synthesis (Anai et al. 2005), suggesting its possible role in the replication checkpoint. In S. pombe, de Lahondès et. al. have reported that the overexpression of the Sap1 C-terminal domain during DNA replication strongly induces minichromosome loss, which is associated with the cut phenotype (de Lahondès et al. 2003). This suggests that Sap1 may play an important conserved role in DNA replication per se. In this study, we have provided evidence that Sap1 tightly associates with the replication origin ori2004 during S-phase. Since Sap1 appears to be located on chromatin outside the S-phase, it is possible that Sap1 is an ancillary component of the prereplicative complex (pre-RC). Taken together, our data suggest that Sap1 might be essential for the initiation of DNA replication. It would therefore be interesting to examine in a future study whether Sap1 is a component of pre-RC that is required for the initiation of DNA replication.

Acknowledgments

We thank T. Wang for generously providing Cds1 polyclonal antisera and H. Masukata and P. Russell for donating the S. pombe strains. We also thank B. Arcangioli, T. Nakamura, J. Nickels, and P. Russell for their helpful discussions. This work was supported by a Leukemia Research Foundation grant (to E.N.) and a Drexel University College of Medicine start-up fund (to E.N.).

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