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1 Department of Molecular Pathology and 2 Center for the Development of Molecular Target Drugs, Cancer Research Institute, Kanazawa University; Ishikawa 920-0934, Japan
3 Department of Radiology, Graduate School of Medicine, Kanazawa University, Kanazawa, Ishikawa 920-0934, Japan
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Introduction |
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Genetic and biochemical studies in yeasts have established that Rad1 (Rad17), Rad9 (Ddc1) and Hus1 (Mec3) are all structurally related to proliferating cell nuclear antigen (PCNA) (Caspari et al. 2000; Venclovas & Thelen 2000) and are thought to function in the early phase of the cell-cycle checkpoint pathway as a hetero-trimeric (9-1-1) complex (Caspari et al. 2000; Kondo et al. 1999). Rad17 (Rad24) is related to replication factor C (RFC), and associates with four other RFC subunits (RFC2-5) (Naiki et al. 2000; Shimada et al. 1999). The RFC1-5 complex (called ‘clamp loader’) is known to recognize the primer-template junction during DNA replication and to load a homo-trimeric PCNA sliding clamp complex on to the DNA. Recent studies in yeasts have shown that the Rad17-containing complex recruits the 9-1-1 complex to DNA lesions at an early phase of the cell-cycle checkpoint pathway (Kondo et al. 2001; Melo et al. 2001), in a manner similar to PCNA complex loading by the RFC1-5 complex (Venclovas & Thelen 2000). Identification of mammalian homologs for Rad17, Rad1, Hus1 and Rad9 and subsequent biochemical studies showed that mammalian Rad9, Rad1 and Hus1 also form a hetero-trimer complex (Lindsey-Boltz et al. 2001; Onge et al. 1999; Volkmer & Karnitz 1999) and are localized to chromatin following DNA damage (Burtelow et al. 2000; Zou et al. 2002). Furthermore, it has been recently established that mammalian Rad17 is required for Rad9 recruitment to chromatin (Zou et al. 2002). Thus, the functions of the vertebrate Rad17-RFC and 9-1-1 complexes in cell-cycle checkpoint controls appear to be similar to those of their yeast homologs. However, the lethality of the gene disruption in mice precludes an extensive genetic evaluation of the roles of these vertebrate complexes (Weiss et al. 2000). Thus, the precise functions of the Rad17-RFC and 9-1-1 complexes in vertebrate cell-cycle checkpoint controls remain to be established.
To investigate the roles of vertebrate Rad17-RFC and 9-1-1 complexes in cell-cycle checkpoint controls, we disrupted the chicken Rad17 and Rad9 loci in the chicken B lymphocyte line DT40. Here we show that, in contrast to the Hus1 disruption in mice, which is lethal, Rad17- and Rad9-deficient DT40 cells are viable, and these mutant cells, unlike the ATM-deficient cells that we reported previously (Takao et al. 1999), are highly sensitive to UV irradiation, alkylating agents, and DNA replication inhibitors, such as hydroxyurea (HU). We further found that Rad17- and Rad9-deficient cells are defective in S-phase DNA damage checkpoint controls and in the cellular response to stalled DNA replication. These phenotypes are similar to those reported in mammalian cells expressing kinase-dead ATR or derived from ATR, Chk1 or Hus1 gene knockout mice (Brown & Baltimore 2000, 2003; Weiss et al. 2000, 2003; Heffernan et al. 2002; Takai et al. 2000; Cliby et al. 1998), suggesting a functional relationship between Rad17, Rad9, Hus1, ATR and Chk1 in vertebrate cell-cycle checkpoint controls.
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Results |
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We isolated the chicken rad17 cDNA (submitted to the DDBJ database under accession number ‘AB105453’), which encodes a protein of 694 amino acids with a calculated molecular weight of 78 124 kD (Fig. 1A). The chicken Rad17 protein shows 
73% identity to the human Rad17 protein and 40% to the S. pombe Rad17 protein. Sequence comparison showed that functional domains identified in NTPases are all well conserved in S. pombe, chicken and human Rad17 proteins (Fig. 1A): these include a potential nucleotide-binding segment called ‘P loop’ (a sequence motif known as ‘Walker A’), an acidic domain that bears similarity to a metal binding catalytic site (so-called ‘DExx’ or ‘Walker B’ motif), and possible sensor domains for ATP binding or hydrolysis (called ‘Sensor-1’ and ‘Sensor-2’ motifs) (Guenther et al. 1997). In addition, ATM/ATR phosphorylation sites (Bao et al. 2001) are conserved in chicken and human Rad17 proteins (Fig. 1A). The chicken Rad9 protein sequence deduced from the chicken rad9 cDNA sequence (submitted to the DDBJ database under accession number ‘AB105452’) is also highly homologous to its human (63% identity) and S. pombe (50% identity) homologs (Fig. 1B). In particular, the domains (Komatsu et al. 2000), c-Abl phosphorylation sites (Yoshida et al. 2002), ATM/ATR phosphorylation sites (Chen et al. 2001), and nuclear localization sequences (Hirai & Wang 2002) are all well conserved in the chicken and human Rad9 proteins (Fig. 1B).
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We first analysed the ability of Rad17–/– and Rad9–/– DT40 cells to arrest their cell cycle in response to X-ray. We previously reported that, following X-ray irradiation, wild-type DT40 cells accumulate in the G2/M phase, while ATM–/– DT40 cells do not (Takao et al. 1999). In a similar fluorescence-activated cell sorting (FACS) cell cycle analysis with propidium iodide (PI) staining, we could not detect a significant G2/M checkpoint abnormality in Rad17–/– and Rad9–/– cells: Rad17–/– and Rad9–/– cells accumulate in the G2/M phase following X-ray irradiation (data not shown). We therefore further analysed G2/M checkpoint responses to X-ray in Rad17–/– and Rad9–/– cells by two different assays. In the first, mitotic entry was monitored on FACS after staining cells for phospho-histone H3, a mitotic entry marker. While radiation-induced transient mitotic delay was not clearly detectable in ATM–/– cells, consistent with the results of the previous report (Takao et al. 1999), radiation-induced mitotic delay appeared to be normally functional in Rad17–/– and Rad9–/– cells (data not shown). However, when mitotic entry was assessed by measurement of the accumulating number of cells in mitosis and interphase in mitotic spreads, radiation-induced transient mitotic delay as observed in wild-type cells was not clearly recognized in Rad17–/– and Rad9–/– cells (data not shown), indicating that Rad17–/– and Rad9–/– cells have a minor defect in radiation-induced mitotic delay which can not be clearly detected by phospho-histone H3 staining. These results are in agreement with those reported very recently for conditional Rad17 gene-knockout human colon epithelial cells (Wang et al. 2003), but not with those for Hus1–/–/p21–/– MEFs (Weiss et al. 2003). However, it is possible that a minor radiation-induced G2/M checkpoint defect in Hus1–/–/p21–/– MEFs may not be clearly detected in the analysis with phospho-histone H3 staining, as shown in the present study. These results are also consistent with those derived from radiation sensitivity analysis: while ATM–/– DT40 cells were highly X-ray sensitive, as we reported previously (Morrison et al. 2000; Takao et al. 1999, 2000), Rad17–/– and Rad9–/– cells were only moderately sensitive to X-rays (Fig. 2).
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We then analysed the sensitivity of Rad17–/– and Rad9–/– DT40 cells to other types of DNA damage, such as that caused by UV irradiation (254 nm UV light) or an alkylating agent, methyl methanesulphonate (MMS). As shown in Fig. 3, the Rad9–/– and Rad17–/– cells were highly sensitive to low doses of UV and MMS, although ATM–/– DT40 cells did not show hypersensitivity to UV or MMS. Since the DNA damage induced by UV irradiation or MMS are known to stall DNA replication, we examined whether chicken Rad17 and Rad9 might be involved in the regulation of S-phase progression in response to DNA damage. Asynchronously growing wild-type and mutant DT40 cells were treated with UV irradiation or MMS, and the S-phase progression was monitored by FACS after PI staining. The untreated asynchronous cell cycle distribution of wild-type and the various mutant cells examined were similar (Fig. 4), and these FACS patterns remained similar in further incubation when untreated (data not shown). However, as shown in Fig. 4, the S-phase progression became slower in wild-type cells when treated with UV irradiation or MMS; while most of the Rad17–/– and Rad9–/– cells had completed the S phase 4 h after treatment with UV irradiation or MMS, the wild-type cells did not complete the S phase even 6 h after UV or MMS treatment. ATM–/– DT40 cells behaved similarly, indicating that Rad17 and Rad9 but not ATM are required for the UV/MMS-induced slowing of the S-phase progression in DT40 cells.
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Because Rad17–/– and Rad9–/– DT40 cells were also highly sensitive to HU (Fig. 3), we examined whether chicken Rad17 and Rad9 are involved in DNA replication checkpoint control, which coordinates the completion of DNA replication and the onset of mitosis. As shown in Fig. 5A, in the presence of 10 mM HU, DNA replication was effectively inhibited in wild-type as well as in ATM–/– cells, and most of the cells were accumulated in the early S phase as early as 1 h after HU treatment. The results shown in Fig. 5A indicate that these HU-arrested cells were stable and viable for as long as 4 h (see also Figs 7 and 8). In addition, this HU-induced arrest in wild-type and ATM–/– cells was a reversible process, given that these cells resumed cell-cycle progression upon the removal of HU (Fig. 5B). In contrast to these results in wild-type and ATM–/– cells, however, the HU treatment of Rad17–/– and Rad9–/– DT40 cells resulted in the accumulation of these cells in sub-G1 fractions, presumably representing apoptotic cell populations (see also Fig. 7). Furthermore, these cells could not resume cell-cycle progression and instead accumulated in sub-G1 fractions after HU removal (Fig. 5B). Similar results were obtained when cells were treated with 1 mM HU (data not shown).
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The results shown in Fig. 5C demonstrate that Rad17 and Rad9 are required for the chicken DNA replication checkpoint control. Since Chk1 plays an essential role in the vertebrate replication checkpoint (Takai et al. 2000; Zachos et al. 2003), and since several recent studies showed that Rad17 and Rad9 are required for Chk1 activation induced by genotoxic agents such as IR or UV (Roos-Mattjus et al. 2003; Wang et al. 2003; Zou et al. 2002), we studied whether chicken Rad17 and Rad9 are involved in Chk1 activation during replication arrest. Wild-type, various mutant cells were treated with HU for various times, and cells were harvested for Western blot analysis with antibodies specific to phospho-serine 345 (Ser-345) (Liu et al. 2000). As shown in Fig. 5D, treatment of wild-type DT40 cells with 10 mM HU led to activation of Chk1 as evidenced by increased phosphorylation of Ser-345 as well as altered electrophoretic mobility of total Chk1. However, these HU-induced Chk1 phospholylation was almost completely eliminated in both Rad17–/– and Rad9–/– cells, though HU-induced Chk1 phospholylation was not affected by deletion of ATM. These results indicate that the Rad17-RFC and 9-1-1 complexes are required for Chk1 activation in the chicken DNA replication checkpoint control.
Although we found that Rad17–/– and Rad9–/– cells are defective in the response to DNA replication stall induced by HU (Fig. 5C), a very recent study using conditional ATR gene-knockout MEFs indicate that an initial mitotic delay is functional in absence of ATR and ATM when replication stall is induced by aphidicolin, though aphidicolin-induced Chk1 phosphorylation is defective in the absence of ATR (Brown & Baltimore 2003). We therefore analysed mitotic entry in Rad17–/– and Rad9–/– cells by phosph-histone H3 staining following aphidicolin treatment. Surprisingly, mitotic entry was effectively prevented in both Rad17–/– and Rad9–/– cells as in wild-type cells when replication arrest was induced by aphidicolin (Fig. 6A), though Rad17–/– and Rad9–/– cells resumed cell-cycle progression and entered mitosis upon the removal of aphidicolin, as effectively as wild-type cells did (data not shown). We also examined whether Rad17 and Rad9 are involved in Chk1 activation during replication arrest induced by aphidicolin. As shown in Fig. 6B, aphidicolin treatment led to Chk1 phosphorylation on Ser-345 and decreased electrophoretic mobility of Chk1 in wild-type cells. However, aphidicolin-induced Chk1 phosphorylation was almost completely eliminated in both Rad17–/– and Rad9–/– cells (Fig. 6B), indicating that Rad17 and Rad9 are also required for aphidicolin-induced Chk1 activation.
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The above results suggest that stalled replication eventually induce apoptotic cell death in Rad17–/– and Rad9–/– cells. These severe cell-cycle checkpoint phenotypes observed in HU-treated Rad17–/– and Rad9–/– cells are similar to those reported for mammalian cells expressing kinase-dead ATR or derived from ATR and Hus1 gene knockout mice (Brown & Baltimore 2000; Cliby et al. 1998; Nghiem et al. 2001; Weiss et al. 2000). To analyse further these abnormal phenotypes in Rad17–/– and Rad9–/– cells, we first examined whether the sub-G1 cell populations observed in Rad17–/– and Rad9–/– cells after HU treatment (Fig. 5A) represent apoptotic cell populations. We quantified the initial phase of apoptosis following treatment with X-rays, UV irradiation, or HU, using annexin V FACS analysis. The results shown in Fig. 7 clearly indicate that HU is highly effective in inducing apoptosis in Rad17–/– and Rad9–/– cells, but not in ATM–/– cells. By contrast, UV irradiation was far less effective in inducing apoptosis, consistent with the finding that significant sub-G1 cell populations were not detectable in UV-treated Rad17–/– and Rad9–/– cells (Fig. 4A). The results with annexin V FACS analysis for apoptosis were further confirmed with the results of the assay detecting the later phase of apoptosis, i.e. DNA degradation: HU treatment resulted in DNA degradation in Rad17–/– cells (data not shown).
We next examined whether HU induces chromatin fragmentation in Rad17–/– and Rad9–/– cells. After HU treatment, cells were fixed and stained with Hoechst33258. Representative chromatin fragmentation patterns observed in Rad17–/– cells after HU treatment are shown in Fig. 8A, and were similar to patterns previously reported by Schlegel & Pardee (1986). The quantitative measurement of chromatin fragmentation in wild-type and various mutant cells after HU treatment indicated that, while significant chromatin fragmentation was not detectable in HU-treated wild-type and ATM–/– cells, as much as 30–40% of Rad17–/– and Rad9–/– cells showed chromatin fragmentation after HU treatment for 4 h (Fig. 8B). To analyse further chromosome integrity, mitotic spreads were prepared from wild-type, Rad17–/–, and Rad9–/– cells after a 2-h HU treatment. While more than 95% of the HU-treated wild-type cells displayed intact chromosomes, a significant fraction (30–50%) of the metaphase spreads from HU-treated Rad17–/– and Rad9–/– cells showed chromosome fragmentation patterns (Fig. 8C) similar to those reported in mammalian cells expressing kinase-dead ATR or derived from ATR or Hus1 gene knockout mice (Brown & Baltimore 2000; Nghiem et al. 2001; Weiss et al. 2000).
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Discussion |
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In the present study we presented evidence that the chicken Rad17-RFC and 9-1-1 complexes play an essential role in DNA replication checkpoint control; Rad17–/– and Rad9–/– DT40 cells are highly sensitive to HU (Fig. 3) and are defective in HU-induced mitotic delay (Fig. 5C); HU causes irreversible damage to the DNA replication checkpoint mechanisms in these cells, resulting in chromatin and chromosome fragmentation (Fig. 8), and finally apoptotic cell death (Figs 5A and 7). These severe cell-cycle checkpoint phenotypes observed in HU-treated Rad17–/– and Rad9–/– cells are similar to those reported for mammalian cells expressing kinase-dead ATR or derived from ATR gene knockout mice (Brown & Baltimore 2000; Cliby et al. 1998; Nghiem et al. 2001), and are likely due to mitotic catastrophe/premature chromatin condensation. In contrast, as shown in the present study, ATM–/– cells are not sensitive to HU (Fig. 3), show normal DNA replication checkpoint control (Fig. 5A,C), and do not undergo irreversible damage when treated with HU (Figs 5B, 7 and 8). We further found that HU-induced Chk1 phospholylation on Ser-345 is defective in Rad17–/– and Rad9–/– cells, but not in ATM–/– cells (Fig. 5D), consistent with the facts that mouse Chk1 plays an essential role in the replication checkpoint (Takai et al. 2000). These results therefore indicate that the Rad17-RFC and 9-1-1 complexes are required for ATR-mediated Chk1 activation in the chicken DNA replication checkpoint control.
A very recent study with conditional ATR gene-knockout MEFs has shown that, when replication arrest is induced by aphidicolin, a delayed mitotic entry occurs even in the absence of ATM and ATR, although aphidicolin-induced Chk1 phosphorylation and CDC2 tyrosine phosphorylation are impaired in the absence of ATR. This study also has shown that ATR is essential for preventing the generation of DSBs upon stalled DNA replication (Brown & Baltimore 2003). Another study using Chk1–/– DT40 cells has shown that Chk1 is dispensable for an initial mitotic delay induced by aphidicolin, but is required for recovery from replication stall (Zachos et al. 2003). We also found that an aphidicolin-induced mitotic delay is functional in Rad17–/– and Rad9–/– DT40 cells (Fig. 6A), though Chk1 is not phosphorylated by aphidicolin in the absence of Rad17 and Rad9 (Fig. 6B). In contrast, it has been shown that an aphidicolin-induced mitotic delay is impaired in Chk1–/– early embryonic mouse cells (Takai et al. 2000). These findings therefore suggest a possibility that, while the vertebrate Rad17-RFC/9-1-1/ATR/Chk1 checkpoint machinery plays essential roles in the DNA replication checkpoint, ill-defined somatic vertebrate systems may play a role in the absence of these checkpoint proteins when DNA replication is inhibited by aphidicolin. Another interesting possibility is that aphidicolin induces an initial mitotic arrest independently of DNA replication inhibition. Further work will be required to resolve these important questions.
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Experimental procedures |
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Chicken EST clones homologous to human Rad17 and Rad9 proteins were searched for using the BLAST program in the chicken bursal EST database (Buerstedde et al. 2002). Oligonucleotide PCR primers were designed based on the identified chicken Rad17 (riken1 16b18r1.blsp) and Rad9 (dkfz426 5C5r1.blsp) EST clones, and partial chicken rad17 and rad9 cDNAs were amplified by RT-PCR using mRNA extracted from DT40 cells. Using these cDNA fragments as probes, chicken rad17 and rad9 cDNA clones were isolated from a chicken DT40 cDNA library (kindly provided by S. Takeda), and chicken rad17 and rad9 genomic clones from a spleen lambda genomic library (Stratagene), respectively. The identity of these clones was confirmed by DNA sequencing. Rad17 targeting vectors were constructed by inserting neo- or his-selection marker gene cassettes under the control of the ß-actin promoter into exon 3 of the rad17 gene. Rad9 disruption constructs were made by replacing exon 1, exon 2, and parts of exon 3 of the rad9 gene with selection-drug-resistance gene cassettes. Cells were cultured in RPMI-1640 supplemented with 10–5 M ß-mercaptoethanol, 10% foetal calf serum, 1% chicken serum, 2 mM L-glutamine, and penicillin/streptomycin at 39.5 °C. For gene targeting of each chicken rad17 and rad9 locus, wild-type DT40 cells were sequentially transfected with pRAD17-neo and pRAD17-his, or with pRAD9-neo and pRAD9-his, respectively, as previously described (Takao et al. 1999). RT-PCR analysis of chicken rad17 and rad9 mRNA expression was performed using the following primers: Rad17, 5'-gAgCAggTAACggATTggTTCgCCCAT-3' and 5'-gCgCTTCTTATATCACCTgAACAACC-3'; Rad9, 5'-gTgAAAgCCCTCggCCACgCCgT-3' and 5'-CAACCACAgCTCTgTCACCA-3'.
Colony survival assay
X-ray irradiation was performed using an MBR-1520R radiator (Hitachi) set at 150-kVp, 20 mA, 0.5 mm aluminium, and 0.9 mm copper filtration with a dose rate of 1 Gy/min. UV irradiation at 254 nm was performed with a germicidal lamp at a fluorescence rate of 0.5 J/m2/s. Serially diluted cells were plated in 60-mm dishes with 5 ml of 1.5% methylcellulose plates containing D-MEM/F-12, 15% FCS, 1.5% chicken serum, penicillin/streptomycin, 2 mM L-glutamine and 10–5 M ß-mercaptoethanol after irradiation. To determine sensitivities to MMS (Aldrich Chemical Company Inc.) and HU (Sigma), serially diluted cells were treated with MMS or HU as indicated, washed with PBS to remove the MMS or HU, and plated into methylcellulose plates. Colonies were counted 7–10 days after irradiation or treatment. Percentage survival was determined relative to the numbers of colonies from untreated cells.
Assessment of cell-cycle checkpoint functions and apoptosis
For cell-cycle distribution analysis, cells were fixed in 70% ethanol and stored at –20 °C for more than 8 h. Before sorting, cells were washed twice in PBS, resuspended in PBS supplemented with 0.5 µg/µl RNaseA, and incubated for 30 min at room temperature. Cells were then stained in the same buffer supplemented with 50 µg/µl PI, incubated for 1 h at room temperature, and immediately analysed using a FACS Calibur (Becton Dickinson). Fluorescence data were displayed as peaks using the Cell Quest software (Becton Dickinson). For quantification of phospho-histone H3, fixed cells were incubated with anti-phospho-histone H3 antibody (Upstate Biotechnology) in 1%BSA/PBS at room temperature. The cells were then incubated with FITC-conjugated goat anti-rabbit antibody (Santa Cruz Biotechnology) for 1 h at room temperature, and were stained with PI for 1 h at 37 °C for FACS analysis. The extent of radiation-induced mitotic delay was determined by measuring mitotic indices following incubation of cells with colcemid for indicated time periods after X-ray irradiation. In brief, cells were cultured in medium containing 0.1 µg/µl colcemid (Gibco-BRL) after irradiation, and collected at 1 h intervals. The collected cells were treated in 0.9% sodium citrate, fixed in methanol/acetic acid (3 : 1), and stained with Giemsa. The percentage of mitotic cells (mitotic index) was determined from counts of at least 200 cells. Apoptosis was quantified by FACS using an annexin V apoptosis Kit (Clontech). To assay DNA degradation, cells were treated with 10 mM HU for the indicated times, and DNA was prepared for separation on a 2% agarose gel.
Western blot analysis
Cell extracts were prepared by lysing in 20 mM Tris-HCl (pH 8.0), 150 mM NaCl, 1 mM EDTA, 0.5% NP-40, containing 10 mMß-glycerophosphate, 1 mM NaF, 0.1 mM Na3VO4, 1 mM PMSF. Proteins were separated by SDS-PAGE, followed by blotting to nitrocellulose membrane (Pall Corporation). For primary antibodies, monoclonal anti-Chk1(G-4) (Santa Cruz Biotechnology) or polyclonal anti-phospho-Ser-345 Chk1 (Cell Signalling Technology) were used.
Analysis of chromatin and chromosome fragmentation
Cells were harvested by centrifugation at 1000 r.p.m. (200 x g) for 2 min, and were fixed twice in 5 ml of ice-cold Carnoy's fixative (1 : 3 v/v acetic acid:methanol). After fixation, the cell pellets were resuspended and applied dropwise on to the surface of glass slides. The slides were stained with Hoechst33258 (1 µg/µl) (Sigma) for 10 min and mounted in 90% glycerol using glass coverslips. At least 200 cells were counted using an IX70 fluorescence microscope (OLYMPUS) to determine the percentage of cells containing chromatin fragmentation. For chromosome fragmentation analysis, after a 4-h treatment with 10 mM HU, cells were treated with 0.1 µg/µl colcemid for 2 h and subsequently processed for the preparation of mitotic spreads as follows. Cells were harvested and fixed twice in 5 ml of ice-cold Carnoy's fixative (1 : 3 v/v acetic acid:methanol). The cells were then spun, and the supernatant was aspirated except for 100 µL above the pellet. The cells were resuspended and applied dropwise on to the surface of glass slides. The slides were dried and stained with DAPI (0.1 µg/µl) (Sigma) for 10 min, and mounted in 90% glycerol using glass coverslips. Fluorescence-labelled DNA was analysed using an IX70 fluorescence microscope (OLYMPUS).
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Acknowledgements |
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Footnotes |
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*Correspondence: E-mail: kyamamot{at}kenroku.kanazawa-u.ac.jp
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References |
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Received: 24 October 2003
Accepted: 15 December 2003
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