Activation of Rad53 kinase in response to DNA damage and its effect in modulating phosphorylation of the lagging strand DNA polymerase
1999; Springer Nature; Volume: 18; Issue: 22 Linguagem: Inglês
10.1093/emboj/18.22.6561
ISSN1460-2075
AutoresAchille Pellicioli, Chiara Lucca, Giordano Liberi, F. Marini, Massimo Lopes, Paolo Plevani, Alfredo Romano, Pier Paolo Di Fiore, Marco Foiani,
Tópico(s)Cancer-related Molecular Pathways
ResumoArticle15 November 1999free access Activation of Rad53 kinase in response to DNA damage and its effect in modulating phosphorylation of the lagging strand DNA polymerase Achille Pellicioli Achille Pellicioli Dipartimento di Genetica e di Biologia dei Microrganismi, Università degli Studi di Milano, via Celoria 26, 20133 Milan, Italy Search for more papers by this author Chiara Lucca Chiara Lucca Dipartimento di Genetica e di Biologia dei Microrganismi, Università degli Studi di Milano, via Celoria 26, 20133 Milan, Italy Search for more papers by this author Giordano Liberi Giordano Liberi Dipartimento di Genetica e di Biologia dei Microrganismi, Università degli Studi di Milano, via Celoria 26, 20133 Milan, Italy Search for more papers by this author Federica Marini Federica Marini Dipartimento di Genetica e di Biologia dei Microrganismi, Università degli Studi di Milano, via Celoria 26, 20133 Milan, Italy Present address: Imperial Cancer Research Fund, Clare Hall Laboratories, South Mimms, EN6 3LD UK Search for more papers by this author Massimo Lopes Massimo Lopes Dipartimento di Genetica e di Biologia dei Microrganismi, Università degli Studi di Milano, via Celoria 26, 20133 Milan, Italy Search for more papers by this author Paolo Plevani Paolo Plevani Dipartimento di Genetica e di Biologia dei Microrganismi, Università degli Studi di Milano, via Celoria 26, 20133 Milan, Italy Search for more papers by this author Alfredo Romano Alfredo Romano European Institute of Oncology, Milan, Italy Search for more papers by this author Pier Paolo Di Fiore Pier Paolo Di Fiore European Institute of Oncology, Milan, Italy Search for more papers by this author Marco Foiani Corresponding Author Marco Foiani Dipartimento di Genetica e di Biologia dei Microrganismi, Università degli Studi di Milano, via Celoria 26, 20133 Milan, Italy Search for more papers by this author Achille Pellicioli Achille Pellicioli Dipartimento di Genetica e di Biologia dei Microrganismi, Università degli Studi di Milano, via Celoria 26, 20133 Milan, Italy Search for more papers by this author Chiara Lucca Chiara Lucca Dipartimento di Genetica e di Biologia dei Microrganismi, Università degli Studi di Milano, via Celoria 26, 20133 Milan, Italy Search for more papers by this author Giordano Liberi Giordano Liberi Dipartimento di Genetica e di Biologia dei Microrganismi, Università degli Studi di Milano, via Celoria 26, 20133 Milan, Italy Search for more papers by this author Federica Marini Federica Marini Dipartimento di Genetica e di Biologia dei Microrganismi, Università degli Studi di Milano, via Celoria 26, 20133 Milan, Italy Present address: Imperial Cancer Research Fund, Clare Hall Laboratories, South Mimms, EN6 3LD UK Search for more papers by this author Massimo Lopes Massimo Lopes Dipartimento di Genetica e di Biologia dei Microrganismi, Università degli Studi di Milano, via Celoria 26, 20133 Milan, Italy Search for more papers by this author Paolo Plevani Paolo Plevani Dipartimento di Genetica e di Biologia dei Microrganismi, Università degli Studi di Milano, via Celoria 26, 20133 Milan, Italy Search for more papers by this author Alfredo Romano Alfredo Romano European Institute of Oncology, Milan, Italy Search for more papers by this author Pier Paolo Di Fiore Pier Paolo Di Fiore European Institute of Oncology, Milan, Italy Search for more papers by this author Marco Foiani Corresponding Author Marco Foiani Dipartimento di Genetica e di Biologia dei Microrganismi, Università degli Studi di Milano, via Celoria 26, 20133 Milan, Italy Search for more papers by this author Author Information Achille Pellicioli1, Chiara Lucca1, Giordano Liberi1, Federica Marini1,2, Massimo Lopes1, Paolo Plevani1, Alfredo Romano3, Pier Paolo Di Fiore3 and Marco Foiani 1 1Dipartimento di Genetica e di Biologia dei Microrganismi, Università degli Studi di Milano, via Celoria 26, 20133 Milan, Italy 2Present address: Imperial Cancer Research Fund, Clare Hall Laboratories, South Mimms, EN6 3LD UK 3European Institute of Oncology, Milan, Italy *Corresponding author. E-mail: [email protected] The EMBO Journal (1999)18:6561-6572https://doi.org/10.1093/emboj/18.22.6561 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info The Saccharomyces cerevisiae Rad53 protein kinase is required for the execution of checkpoint arrest at multiple stages of the cell cycle. We found that Rad53 autophosphorylation activity depends on in trans phosphorylation mediated by Mec1 and does not require physical association with other proteins. Uncoupling in trans phosphorylation from autophosphorylation using a rad53 kinase-defective mutant results in a dominant-negative checkpoint defect. Activation of Rad53 in response to DNA damage in G1 requires the Rad9, Mec3, Ddc1, Rad17 and Rad24 checkpoint factors, while this dependence is greatly reduced in S phase cells. Furthermore, during recovery from checkpoint activation, Rad53 activity decreases through a process that does not require protein synthesis. We also found that Rad53 modulates the lagging strand replication apparatus by controlling phosphorylation of the DNA polymerase α-primase complex in response to intra-S DNA damage. Introduction In response to genotoxic agents, cell cycle blocks or alterations of particular cellular structures, all eukaryotic cells activate a set of surveillance mechanisms, called checkpoints, (for reviews, see Carr and Hoekstra, 1995; Elledge, 1996; Paulovich et al., 1997; Weinert, 1998). A subset of these mechanisms is represented by the DNA damage checkpoint, which is triggered by DNA lesions. The activation of this signal transduction pathway leads to a delay of cell cycle progression to prevent replication and segregation of damaged DNA molecules, and to induce transcription of several DNA repair genes (Paulovich et al., 1997; Weinert, 1998). Increasing evidence indicates that defects in the cellular response to DNA damage are relevant for the early stages of carcinogenesis as they cause increased mutagenesis and genomic instability (reviewed in Hartwell and Kastan, 1994; Weinert, 1997). The mechanisms that control these responses have been studied in different organisms, including the yeast Saccharomyces cerevisiae and Schizosaccharomyces pombe, which have been invaluable in providing the genetic tools to dissect the checkpoint response pathways. In S.cerevisiae, the DNA damage checkpoint acts at three stages of the cell cycle: at the G1/S transition, during S phase and at the G2/M boundary, and the genetic requirements for proper DNA damage response in G1, S and G2 are at least partially distinct from each other (Paulovich et al., 1997; Weinert, 1998). These differences may be linked to the recognition of the DNA lesions. In fact, it is possible that DNA damage is more accessible or can only be processed in certain periods of the cell cycle, and the options for the different types of repair processes might be different in G1, S and G2 cells. Two essential genes, MEC1 and RAD53, play a central role in the DNA damage checkpoint at all the cell cycle stages. Mec1 is a member of the evolutionarily conserved subfamily of phosphatidylinositol 3-kinase (PI3-kinase), which includes budding yeast Tel1, fission yeast Rad3, mammalian ATM and ATR and DNA-dependent protein kinase (DNA-PK) (Elledge, 1996). The similarity to DNA-PK, a protein kinase activated by binding to DNA and stimulated by the association of the Ku complex (Smith and Jackson, 1999), has suggested that Mec1 can also act as a protein kinase whose activity might be influenced by the association with other checkpoint proteins (Longhese et al., 1998). However, the biochemical properties and protein–protein interactions of Mec1 remain speculative in the absence of any biochemical data on the MEC1 gene product. Rad53 is an essential protein kinase required for cell cycle arrest in response to replication blocks and DNA damage (Allen et al., 1994; Weinert et al., 1994). Rad53 is phosphorylated in response to genotoxic treatments and this phosphorylation depends on the function of several DNA damage checkpoint genes (Sanchez et al., 1996; Sun et al., 1996). Rad53 has two FHA (forkhead-associated) domains, and Rad9, another checkpoint protein, interacts with the C-terminal FHA domain of Rad53 (Sun et al., 1998), raising the possibility that Rad53 kinase activity might be influenced by association with other proteins. Besides Rad9, other factors involved in the DNA damage response pathway include Mec3, Ddc1, Rad17 and Rad24. Rad17 exhibits a structural similarity to the proliferating cell nuclear antigen (PCNA) (Thelen et al., 1999), while Rad24 is related to replication factor C (RF-C) (Lydall and Weinert, 1997), a protein complex that during DNA replication binds to template–primer junctions and loads PCNA onto the DNA, thereby recruiting replicative DNA polymerases (Waga and Stillman, 1998). Even though the function of these checkpoint proteins is still unknown, it has been suggested that they might participate in DNA damage recognition and/or processing (Weinert, 1998). Significant progress has been made in identifying many of the factors involved in the DNA damage checkpoint pathway, but very little is known about the physiological downstream targets. It has been suggested that the DNA replication machinery might be the final target of the DNA damage checkpoint response, which expands the length of S phase in the presence of genotoxic agents (Paulovich and Hartwell, 1995). Indeed, some replication factors have been implicated in the checkpoints, including DNA polymerase ϵ (Pol2), replication protein A (RP-A) and the DNA polymerase α–primase complex (pol–prim) (Navas et al., 1995; Longhese et al., 1996; Marini et al., 1997). Pol2, whose role in DNA replication is still uncertain (Kesti et al., 1999), is thought to be required for an early step in checkpoint activation, possibly as a sensor of DNA damage (Navas et al., 1995). RP-A is involved in replication, recombination and repair (Wold, 1997) and, in response to DNA damage, is phosphorylated through a mechanism dependent upon Mec1, but not on Rad53 (Brush et al., 1996). Moreover, the pol–prim complex, which is absolutely required for initiation of DNA synthesis at origins of replication and for lagging strand DNA synthesis (Foiani et al., 1997), acts downstream of Rad53 and may represent one of the targets of the checkpoint pathway (Marini et al., 1997). It has been shown that checkpoint activation prevents the firing of late replication origins in the presence of genotoxic agents (Santocanale and Diffley, 1998; Shiraige et al., 1998), suggesting that either initiation at late origins or some other step of DNA synthesis is controlled by Mec1 and Rad53. Although the checkpoint targets that prevent late origin firing are still unknown, it has been suggested that Mec1 and Rad53 might modulate the activity of the Cdc28 and Cdc7 kinases that regulate initiation of DNA synthesis at the origins (Santocanale and Diffley, 1998). However, it is not clear yet whether the checkpoint negatively regulates S phase or rather promotes a specialized intrinsically slow replication process, or both. In fact, there are some analogies between the slow replication process occurring in the presence of genotoxic agents and the process that couples replication to recombination, which has been described in prokaryotes and in yeast (Malkova et al., 1996; Kogoma, 1997; Foiani et al., 1998). Intriguingly, all the replication factors implicated so far in the checkpoints are also involved in a repair pathway activated by a double strand break (DSB) and which couples replication to recombination (Holmes and Haber, 1999). Here we have analysed the activity of the Rad53 protein kinase in response to genotoxic agents using an in situ autophosphorylation assay. Our data demonstrate that Rad53 activation does not require physical association with other proteins or nucleic acids and depends upon a functional Mec1. We show that activation of Rad53 requires the Rad9, Mec3, Ddc1, Rad17 and Rad24 checkpoint factors in G1, while this dependency is strongly reduced when cells progress through S phase. Moreover, cells recovering from checkpoint activation inactivate Rad53 through a process that does not require protein synthesis. Finally, we show that the lagging strand replication apparatus is controlled by Rad53 that prevents phosphorylation of the pol–prim complex in response to intra-S DNA damage. Results An in situ renaturation assay (ISA) to measure Rad53 autophosphorylation activity To monitor Rad53 kinase activation directly and to begin to understand how Rad53 is able to transduce incoming signals, we have developed a renaturation assay to measure Rad53 autophosphorylation in situ (see Materials and methods for details). As shown in Figure 1, when extracts prepared from wild-type exponentially growing cells were tested with the ISA, four major bands ranging in size from 45 to 60 kDa were detected and the intensity of these active kinase bands did not change when extracts were prepared from cells previously treated with the genotoxic agents hydroxyurea (HU) and methymethanesulfonate (MMS). Figure 1.In situ autophosphorylation assay for Rad53 protein kinase. Log-phase (L) cultures of isogenic strains K699 (RAD53), CY2817 (rad53 [RAD53-MYC]) or CY2034 (rad53) were treated for 3 h with 0.2 M HU (H) or 0.02% MMS (M). A 25 μg aliquot of total protein was prepared from the indicated strains and analysed by ISA or Western blotting using anti-Rad53 antibodies as described in Materials and methods. Download figure Download PowerPoint It has been shown that yeast cells respond to HU and MMS treatment by activating a signal transduction pathway that leads to Rad53 phosphorylation (Sanchez et al., 1996; Sun et al., 1996; and Figure 1). We found that an autophosphorylation activity with the same electrophoretic mobility as Rad53 was barely detectable in untreated wild-type extracts, but the intensity of this active band dramatically increased after HU and MMS treatment (Figure 1). Two control experiments indicate that the active kinase band indeed corresponds to Rad53: first, HU and MMS treatment of the kinase-defective rad53-K227A mutant strain caused phosphorylation of the Rad53 mutant protein without a concomitant increase in the autophosphorylation activity (Figure 1); and second the active Rad53 band was shifted to the expected higher molecular weight form when extracts were prepared from rad53 mutant cells carrying a 9Myc-RAD53 gene on a plasmid (Figure 1). Both the Myc-tagged and the mutant Rad53 proteins were phosphorylated, but only the Myc-Rad53 protein was active in the ISA. We failed to detect any fluctuation in the basal level of Rad53 autophosphorylation activity in untreated wild-type cycling cells synchronized by α-factor treatment (data not shown). We have also followed the kinetics of Rad53 activation in response to DNA damage. In wild-type cells released from a G1 block in the presence of MMS, Rad53 underwent a significant mobility shift during the approximate time of S phase, which also correlated with the increase in Rad53 autophosphorylation activity (Figure 2A). When the same experiment was performed with the kinase-defective rad53-K227A mutant allele, Rad53 modification started 10 min earlier than in wild-type cells, but the autophosphorylation activity was barely detectable (Figure 2B). The anticipated modification timing can be related to the faster cell cycle progression observed by fluorescence activated cell sorting (FACS) of rad53-K227A cells under damaging conditions and is probably the result of in trans phosphorylation events. Wild-type cells released from a G1 block in the presence of HU started to accumulate more slowly migrating Rad53 isoforms 30 min after the release, which correlates with the increase in autophosphorylation activity (Figure 2C). Again, Rad53 phosphorylated isoforms were detectable in rad53-K227A mutant cells, but no autophosphorylation activity was measured by the ISA (Figure 2D). Figure 2.Rad53 activity in response to DNA damage and during recovery. (A–D) Log-phase (Log) cultures of isogenic strains K699 (RAD53) (A and C) or CY2034 (rad53) (B and D) were pre-synchronized by α-factor treatment (αF) and released from the G1 block in YPD with 0.02% MMS (A and B) or 0.2 M HU (C and D). Samples were taken at the time points indicated and analysed by FACS. A 25 μg aliquot of total protein was prepared and analysed by ISA (Kin) or Western blotting. (E and F). Log-phase (Log) cultures of the RAD53 strain were treated with 0.2 M HU for 3 h (HU), washed with the same volume of YPD to remove HU, and released from the HU block in YPD with (F) or without (E) 10 μg/ml cycloheximide. Samples were taken at the indicated time points and analysed by FACS. A 25 μg aliquot of total protein was prepared and analysed by ISA (Kin) or Western blotting. Download figure Download PowerPoint When HU was removed from the medium, cells recovered from the cell cycle block and restored a normal cell cycle progression. To test whether during recovery the cells adapt to the high level of Rad53 activity or, instead, turn Rad53 off, we treated wild-type cells with HU to induce checkpoint activation and then we removed it to allow recovery. As shown in Figure 2E, Rad53 activity began to decrease rapidly after HU removal and, concomitantly, cells started to proceed through S phase. However, it should be pointed out that even 3 h after the release from the HU block, when the cells were already in the next cell cycle, Rad53 activity was well above the basal level. Western blot analysis showed that the Rad53 phosphorylated isoforms progressively disappeared during recovery with the same kinetics observed for Rad53 activity with the ISA. These data indicate that recovery from HU correlates with a decrease in Rad53 activity. Moreover, residual Rad53 activity observed at late time points after the HU release is not sufficient to cause a cell cycle block, suggesting either that checkpoint activation requires a threshold of Rad53 activity or that the cell is able to adapt to a low level of Rad53 activity during recovery. We then addressed whether the decrease of Rad53 activity observed during recovery was dependent upon protein synthesis. As shown in Figure 2F, wild-type cells released from the HU block in the presence of cycloheximide were able to proceed through S phase even though with slightly slower kinetics compared with the control cells, but failed to enter mitosis. Therefore, all the factors required to proceed through S phase during recovery have already been synthesized at the HU block, while the failure to execute mitosis in the presence of cycloheximide has already been described (Burke and Church, 1991). Since the decrease of Rad53 activity during recovery occurs with the same kinetics with or without cycloheximide treatment, the factor(s) required to turn Rad53 off seems to be already present at the HU block. Altogether, the data above suggest that Rad53 is phosphorylated in trans in response to DNA damage and that this phosphorylation event activates an autophosphorylation reaction that can be measured by the ISA. The peculiarity of such an in situ renaturation assay also indicates that Rad53 autophosphorylation activity does not require structural association with other proteins, and that the very low Rad53 autophosphorylation activity observed under unperturbed conditions is not due to the presence of putative inhibitors. Moreover, our finding that Rad53 activity decreases during recovery suggests the existence of a cellular pathway required to turn Rad53 off, probably to allow cell cycle recovery after checkpoint activation. Rad53 activity in different checkpoint- and cell cycle-defective mutants The finding that phosphorylation of Rad53 caused by treatment with genotoxic agents is the cumulative result of in trans and autophosphorylation events prompted us to re-examine the genetic requirements of Rad53 activation by using the ISA. We first tested the capacity of cdc mutants defective in the execution of critical cell cycle transition steps to activate Rad53 after shift to the restrictive temperature in the presence of HU or MMS (Figure 3A). The cdc28-13 allele causes arrest at START at the restrictive temperature (Hereford and Hartwell, 1974) and, as shown in Figure 3A, in cdc28-13-arrested cells HU treatment failed to cause Rad53 activation, while fully active Rad53 was detectable in MMS-treated cells. The same result was obtained with cdc4 mutant cells which arrest post-START in late G1 (Schwob et al., 1994). cdc7-1 mutant cells fail to initiate DNA synthesis at the restrictive temperature (Hereford and Hartwell, 1974). MMS treatment in cdc7-1 cells led to Rad53 activation, while, in the presence of HU, Rad53 activity was significantly lower. POL1/CDC17, CDC2, CDC9 and CDC8 respectively code for the catalytic subunits of DNA polymerase α, DNA polymerase δ, DNA ligase and thymidylate kinase (Murray and Hunt, 1993), and mutants in the corresponding genes arrest in S phase at the restrictive temperature. As shown in Figure 3A, in response to either HU or MMS treatment, Rad53 was fully active in pol1, cdc2, cdc9 and cdc8 mutant cells at the restrictive temperature. Moreover, in these mutant backgrounds, Rad53 was partially active in the absence of any treatment probably due to defective DNA synthesis resulting in checkpoint activation in the absence of genotoxic agents. Finally, in cdc5 mutant cells that arrest in M phase at the non-permissive temperature (Murray and Hunt, 1993), MMS but not HU treatment led to Rad53 activation. Figure 3.Rad53 activity in yeast strains defective in cell cycle progression and checkpoint response. (A) Log-phase cultures of strains K699 (wt), CY1884 (cdc28), K4083 (cdc4), DMP2537/5C (cdc7), YLL9 (pol1), H2C2A1 (cdc2), H9CIB1 (cdc9), H8CIA1 (cdc8) and H5C1B1 (cdc5) grown at 23°C were shifted to the restrictive temperature (37°C) for 4 h (0). The terminal phenotype of the cdc mutants was monitored microscopically. Then 0.2 M HU (H) or 0.02% MMS (M) was added for an additional 4 h at 37°C. A 25 μg aliquot of total protein was prepared and analysed by ISA (Kin) or Western blotting. (B) Log-phase cultures (L) of strains K699 (wt), CY2034 (rad53), DMP2541/8A (mec1), CY427 (rad9), DMP1913/11C (rad17), DMP1913/20B (rad24), YLL244 (ddc1), YLL134 (mec3), CY387 (pri1), K699-M2 (rfa) and SS111-2-11 (pol2) were treated with 0.2 M HU (H) or 0.02% MMS (M) for 3 h at 25°C. Cell cycle progression was analysed by FACS (data not shown). In the case of pol2 mutants, the experiment was repeated at 30 and 37°C with analogous results also in strains TC102-2-11 and TC102-2-12 (data not shown). A 25 μg aliquot of total protein was prepared and analysed by ISA (Kin) or Western blotting. (C) Log-phase cultures of the same strains indicated in (B) were pre-synchronized in G1 by adding 20 μg/ml of α-factor. Each culture was kept in G1 for an additional hour with (+) or without (−) 2 μg/ml 4NQO. The incubations were carried out at 25°C. A 25 μg aliquot of total protein was prepared and analysed by ISA (Kin) or Western blotting. Download figure Download PowerPoint In conclusion, this analysis indicates that MMS treatment leads to Rad53 activation at any stage of the cell cycle, while Rad53 activation in response to HU treatment appears to be S phase specific. However, it should be pointed out that Rad53 activation was very low when the MMS treatment was carried out at 25°C in wild-type G1-arrested cells compared with cells experiencing S phase (data not shown), suggesting that the capacity of MMS to activate Rad53 is enhanced at 37°C and is maximal during S phase (Marini et al., 1997; Vialard et al., 1998). Several checkpoint factors have been placed upstream of Rad53 in the pathway leading to Rad53 phosphorylation (Weinert, 1998), and mutations in the corresponding genes result in the inability to slow down cell cycle progression in response to HU or MMS treatment. As shown in Figure 3B, Rad53 activity in response to HU or MMS treatment was greatly reduced in a mec1-1 mutant background. Conversely, the Rad53 autophosphorylation reaction in response to HU treatment was unaffected in rad9Δ, rad17Δ, rad24Δ, ddc1Δ or mec3Δ cells, although a partial reduction in the activity was detectable in the same mutant cells after MMS treatment. Other DNA replication proteins have also been implicated in the DNA damage checkpoint pathway, including DNA polymerase ϵ, pol–prim and RP-A (Navas et al., 1995; Longhese et al., 1996; Marini et al., 1997). Figure 3B shows that in pri1, rfa1 and pol2 checkpoint-defective mutants, Rad53 could carry out the autophosphorylation reaction in response to HU and MMS treatment, although in MMS-treated pol2 cells the Rad53 activity was lower than in wild-type cells. In pri1, rfa1 and pol2 mutant cells, Rad53 activity was slightly higher in untreated cells compared with wild-type cells, probably as a consequence of defective DNA replication. These results indicate that, in response to HU treatment, Mec1 is absolutely required for Rad53 activation, while Rad9, Rad17, Rad24, Mec3 and Ddc1 appear to be dispensable. Conversely, Rad9, Rad17, Rad24, Mec3 and Ddc1 seem to be partially required for proper Rad53 activation in response to MMS treatment, which is again fully dependent on Mec1. Moreover, our finding that Rad53 is fully active in response to HU and MMS treatment in pri1 and rfa1 cells suggests that the checkpoint defect exhibited by these replication mutants is unlikely to be related to their inability to activate Rad53. Pol2 is thought to be required for checkpoint activation and Rad53 phosphorylation in response to HU treatment, and pol2-12 mutants fail to delay cell cycle progression in the presence of HU (Navas et al., 1995). It was somewhat surprising to find that in HU-treated pol2-12 cells Rad53 was fully active and did not accumulate elongated mitotic spindles to a significant extent (data not shown). Since it has been suggested that pol2 checkpoint defects are enhanced at higher temperatures (Navas et al., 1996), we also tested the effect of the pol2-11 and pol2-12 mutations at 30 and 37°C. We found that Rad53 was fully active in HU-treated pol2 mutant cells even at 37°C (data not shown) and, therefore, we conclude that, in our experimental conditions, the pol2 checkpoint defect in the presence of HU is not due to a failure in activating Rad53. However, Rad53 activation was partially reduced in MMS-treated pol2 cells. G1-arrested cells treated with a variety of genotoxic agents are able to delay entry into S phase by activating the Mec1- and Rad53-dependent checkpoint pathway. We therefore tested whether Rad53 activation in response to DNA damage in G1 was affected in the mutant backgrounds analysed in Figure 3B. As shown in Figure 3C, we found that Rad53 was fully active in wild-type cells arrested in G1 and treated with 4-nitro-quinoline-N-oxide (4NQO) throughout the G1 block, while Rad53 activity was not detectable in treated mec1, rad9, rad17, rad24, mec3 and ddc1 cells. Conversely pri1, rfa1 and pol2 mutants were still able to activate Rad53 under the same experimental conditions. Analogous results were obtained using MMS instead of 4NQO (data not shown). These data indicate that Rad53 activation in G1-arrested cells strongly depends upon functional Mec1, Rad9, Rad17, Rad24, Mec3 and Ddc1. Relationship between Rad53 activation and S phase progression We have found recently that the pol–prim complex may be one of the final targets of the intra-S DNA damage checkpoint pathway mediated by Rad53 (Marini et al., 1997). Accordingly, the data presented in Figure 3 seem to exclude a role for pol–prim and possibly other replication complexes upstream of Rad53. However, so far, there is no evidence to exclude that these replication proteins may represent a direct substrate of Rad53. Pol–prim is a highly regulated enzyme that undergoes cell cycle-dependent phosphorylation and dephosphorylation (Foiani et al., 1997). The B subunit of the complex is phosphorylated early in S phase and dephosphorylated while cells are exiting from mitosis (Foiani et al., 1995), and recent evidence indicates that its phosphorylation is dependent upon a functional Clb–Cdc28 complex (Desdouets et al., 1998; G.Liberi and M.Foiani, unpublished data). In order to understand how pol–prim (and possibly the whole replication machinery) is regulated by the Rad53-dependent pathway in response to DNA damage, we have analysed the phosphorylation state of the pol–prim B subunit in wild-type cells released from a G1 block in the presence of HU or MMS. As shown in Figure 4A and B, B subunit phosphorylation, which usually occurs 20–30 min after release from α-factor (data not shown; Foiani et al., 1995), was greatly delayed in the presence of MMS or HU, and this delay was reduced in a rad53 mutant background. Figure 4.Rad53 delays pol–prim B subunit phosphorylation in response to DNA damage. (A and B) Log-phase cultures of isogenic strains K699 (RAD53) or CY2034 (rad53) were pre-synchronized by α-factor treatment (αF) and released from the G1 block in YPD containing 0.02% MMS or 0.2 M HU. Samples were taken at the times points indicated and 25 μg of total protein were prepared and analysed by Western blotting using antibodies against the B subunit. The slower migrating form represents phosphorylated B subunit. Cell cycle progression was analysed by FACS (data not shown). (C) A log-phase culture (Log) of strain CY947, carrying GAL1-rad53-D339A on a plas
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