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The protein products of several rad checkpoint genes of Schizosaccharomyces pombe (rad1+, rad3+, rad9+, rad17+, rad26+, and hus1+) play crucial roles in sensing changes in DNA structure, and several function in the maintenance of telomeres. When the mammalian homologue of S. pombe Rad9 was inactivated, increases in chromosome end-to-end associations and frequency of telomere loss were observed. This telomere instability correlated with enhanced S- and G2-phase-specific cell killing, delayed kinetics of γ-H2AX focus appearance and disappearance, and reduced chromosomal repair after ionizing radiation (IR) exposure, suggesting that Rad9 plays a role in cell cycle phase-specific DNA damage repair. Furthermore, mammalian Rad9 interacted with Rad51, and inactivation of mammalian Rad9 also resulted in decreased homologous recombinational (HR) repair, which occurs predominantly in the S and G2 phases of the cell cycle. Together, these findings provide evidence of roles for mammalian Rad9 in telomere stability and HR repair as a mechanism for promoting cell survival after IR exposure.
The DNA damage checkpoint pathways are highly conserved among a wide range of eukaryotes, from Saccharomyces cerevisiae to mammals. Human Rad9 (hRad9) is phosphorylated by ATM (ataxia-telangiectasia [A-T] mutated gene product) in response to DNA damage (7). Together with hRad1 and hHus1, hRad9 forms a nuclear complex that resembles PCNA and is believed to sense DNA damage (60, 65, 66). However, the cellular colocalization of hRad9 with the phosphorylated form of histone H2A variant X (γ-H2AX) after DNA damage is independent of ATM function (19), raising the possibility that hRad9 may also have a direct DNA damage-sensing function. In addition, Yin and coworkers (71) have reported that hRad9, much like p53, can transactivate p21. Loss of DNA damage response elements can result in cancer and other genetic diseases (22, 38). In fact, aberrant expression of Mrad9 or hRad9 has been linked to morphological transformation and cancer (8, 30, 59). Inactivation of Mrad9 in mouse embryonic stem (ES) cells results in an increased frequency of spontaneous chromosomal aberrations (23), and similar findings were observed in human cells with reduced levels of hRad9 (12). However, it is not known exactly how mammalian Rad9 influences genomic instability and whether the mechanism is related to defective DNA double-strand break (DSB) repair.
Interestingly, strains of Schizosaccharomyces pombe mutated in either the rad1+, the hus1+, the rad9+, or the rad17+ gene; S. cerevisiae with mutations in related orthologues; and nematodes altered in mrt-2 (encoding a Rad1 homolog) show telomere shortening; however, S. cerevisiae with mutant Mec3 shows telomere elongation, suggesting that these checkpoint proteins are also involved in telomere maintenance (1, 9, 11, 28, 36). Nabetani and coworkers (35) have found that hRad9, along with hHus1 and hRad1, is colocalized specifically at telomeric DNAs and promyelocytic leukemia bodies in ALT (alternative lengthening of telomeres) cells. Whether these observations are functionally significant in relation to telomere instability is unknown.
Genomic stability is maintained by telomeres since these chromosomal terminal structures protect chromosomes from fusion or degradation, as initially reported by Muller (34) and McClintock (32). Shortening or loss of telomeric repeats or altered telomere chromatin structure is correlated with chromosome end-to-end associations that could lead to genomic instability and gene amplification (10, 14, 39). Increased chromosome end-to-end associations, also known as telomeric associations typically seen at metaphase, have been reported in cells derived from tumor tissues, in senescent cells, in the Thiberge-Weissenbach syndrome, in A-T individuals, and following viral infections (45, 56). Chromosome end-to-end associations have been linked to genomic instability and carcinogenicity (10, 15, 38, 39). Both genetic and epigenetic factors can influence telomere stability (26, 55). For example, telomeric end-to-end fusions are enhanced in cells expressing dominant negative alleles of human telomeric protein TRF2 or overexpressing isoforms of human HP1 proteins (55, 64). Although recent data support a role for hRad9 in the cellular response to DNA damage, it is not clear whether this protein functions in telomere maintenance or DNA damage repair following ionizing radiation (IR) exposure. Here we report that inactivation or reduction in levels of mammalian Rad9 enhances telomere instability, increases cell killing by IR specifically in the S and G2 phases of the cell cycle, and reduces DNA DSB repair by homologous recombination (HR).
Human 293, GM5823+hTERT, MCF-7, and GM847 cells were maintained by previously published procedures (41, 55). Mouse ES cells were cultured by standard methods (23). Full-length hRad9 cDNA and DNA encoding the hRad9 C-terminal fragment (269 to 391 amino acids [aa]; ΔhRad9) were amplified by reverse transcription-PCR, cloned into pcDNA3.1, and transfected into cells (67). hRad9 small interfering RNA (siRNA) and control Luc siRNA were obtained from Dharmacon Research (Lafayette, CO). The siRNA sequence targeting hRad9 was 5′-AACCCUUGGAGGACGGGCUCU-3′.
Cells were plated onto 60-mm dishes in 5.0 ml of medium, incubated for 6 h, and subsequently exposed to IR. The number of cells per dish was chosen to ensure that about 50 colonies would survive a particular radiation dose treatment. Cells were exposed to IR in a dose range of 0 to 6 Gy at room temperature. Cells were incubated for 12 or more days and then fixed in methanol-acetic acid (3:1) prior to staining with crystal violet. Only colonies containing >50 cells were counted.
Telomeres in metaphase spreads were detected by fluorescent in situ hybridization (FISH) with a telomere-specific probe (55). To identify association between two human chromosomes, we used centromere and telomere probe combinations. Centromere-specific PNA probes were labeled with fluorescein isothiocyanate (green), and telomere-specific PNA probes were labeled with the fluorochrome Cy3 (red). For terminal restriction fragment (TRF) length analysis, DNA was isolated from exponentially growing cells, the DNA was digested with the restriction enzymes RsaI and HinfI (which do not cut the terminal TTAGGG repeat sequences), and the resulting DNA fragments were separated by agarose gel electrophoresis and then hybridized to a 32P-labeled (TTAGGG)5 probe (13). TRF detection and measurement of TRF lengths were performed as described previously, with ImageQuant version 1.2 (Molecular Dynamics) (13, 33, 54, 55). Nondenaturing in-gel hybridization to determine relative amounts of telomeric single-stranded DNA (G tails) was performed as described previously (33).
Telomerase activity was determined with the telomerase PCR enzyme-linked immunosorbent assay kit (Boehringer Mannheim) as previously described (52). Telomerase activity was determined in triplicate, and negative and positive controls were run with each experiment. As a negative control, an aliquot of each extract was heat inactivated for 10 min at 95°C.
To obtain cell samples enriched at different phases of the cell cycle, exponentially growing populations were fractionated by centrifugal elutriation (44). The cell cycle distribution of the fractionated samples was determined by flow cytometry measurements of DNA content (43, 44), as well as by using a premature chromosome condensation (PCC) protocol (43).
Metaphase chromosomes were prepared by standard procedures (45). Giemsa-stained chromosomes from metaphase spreads were analyzed for chromosome end-to-end associations and aberrations. PCC was used to visualize chromosome damage directly in G1- and G2-phase cell populations. However, after S-phase-specific PCC, chromosomes look pulverized and therefore could not be analyzed for chromosome breaks (43, 44).
G1-type chromosomal aberrations were assessed in cells exposed to 3 Gy of IR and incubated for 24 h. Cells were then subcultured, and metaphases were collected (24, 55). S-phase-specific chromosome aberrations were assessed after exponentially growing cells (pulse-labeled with bromodeoxyuridine [BrdU]) were irradiated with 2 Gy of IR. Metaphases were harvested following 4 h of irradiation, and S-phase types of chromosomal aberrations were scored. For G2-specific aberrations, cells were irradiated with 1 Gy and metaphases were collected 45 and 90 min posttreatment (13). Chromosome spreads were prepared after hypotonic treatment of cells, fixed in acetic acid-methanol, and stained with Giemsa (37). The categories of G1-type asymmetrical chromosome aberrations that were scored included dicentrics, centric rings, interstitial deletions/acentric rings, and terminal deletions. S-phase-type chromosome aberrations were assessed by counting both the chromosome and chromatid aberrations per metaphase as previously described (24, 55). G2-type chromosomal aberrations were assessed by counting chromatid breaks and gaps per metaphase as previously described (24, 55). The efficiency of the G1- or S-phase checkpoint was determined as previously described (24). The efficiency of G2 checkpoint control was evaluated by comparing different cell types for mitotic indices and chromatid-type aberrations at metaphase after irradiation (13). Fifty metaphases were scored for each postirradiation time point.
The technique of PCC was used to visualize chromosome damage directly in G1- and G2-phase cell populations (43, 44). The procedures employed for mitotic Chinese hamster ovary (CHO) cell accumulation, cell fusion, and chromosome preparations are the same as previously described (43). G1- and G2-phase cell populations were obtained by centrifugal elutriation, placed on ice, irradiated, and fused with mitotic CHO cells to induce PCC. Chromosome damage in G1-phase cells is seen in the G1-phase PCC as an increase in the number of chromosome fragments per cell as previously described (43). For treated cells in the G1 phase, the number of breaks per cell was determined by subtracting the number of chromosome fragments per cell in unirradiated cells from that found in irradiated cells. Chromosome damage in G2-phase cells is seen as an increase in breaks, gaps, and exchanges. For treated cells in the G2 phase, the numbers of chromatid gaps, breaks, and exchanges were determined by subtracting the number of such aberrations per cell in the unirradiated populations from that in irradiated cells.
Published protocols were used for immunoprecipitation and immunoblotting (21). The following antibodies were used: anti-hRad9 and anti-hRad51from Santa Cruz (Santa Cruz, CA) and ATM, ATM S-1981, Chk2, TRF2, and Chk2 P-T68 provided by K. D. Brown, T. deLange, S. Elledge, M. B. Kastan, and J. Chen.
Cells were grown in chamber slides, fixed in 2% paraformaldehyde for 15 min, washed in phosphate-buffered saline (PBS), permeabilized for 5 min on ice in 0.2% Triton X-100, and blocked in PBS with 1% bovine serum albumin. The procedure used for immunostaining is the same as that described previously (46, 53). Briefly, anti-γ-H2AX antibodies (Upstate Biotechnology) were applied to cells, which were then incubated for 1 h, washed with PBS containing 1% bovine serum albumin, and further incubated with Alexa Fluor 488-conjugated goat anti-rabbit secondary antibody (Molecular Probes) for 1 h at room temperature. Cells were rinsed with PBS and mounted in Vectashield mounting medium (Vector Laboratories). For detection of cells in S phase, cells were labeled with BrdU for 30 min. BrdU was detected by a monoclonal anti-BrdU antibody (Amersham Pharmacia Biotech) in concert with Alexa Fluor 594-conjugated goat anti-mouse secondary antibody (Molecular Probes). Foci were counted by microscopic and imaging processes. Fluorescence images were captured by using a Zeiss Axioskop 2 mot epifluorescence microscope equipped with a charge-coupled device camera and ISIS software (Metasystems, Altlussheim, Germany). Optical sections through nuclei were captured, and the images were obtained by projection of the individual sections. The results shown are from three independent experiments. Cells with a bubble-like appearance or micronuclei were not considered for γ-H2AX analysis.
hRad9 was immunodepleted from human 293 cell extracts by anti-hRad9 antibody (54), or cell extracts were made from 293 cells after hRad9 knockdown by siRNA. The end-joining assay was performed as previously described (5, 54).
To determine the chromosomal HR frequency, MCF-7 cells were transfected with 1 to 3 μg of linearized pDR-green fluorescent protein (pDR-GFP) plasmid using Lipofectamine 2000 (Invitrogen Corporation, Carlsbad, CA) (48, 72). Stably transfected cell lines were selected by growth in medium containing 1.0 μg/ml puromycin. Selected pDR-GFP-containing clones were shown to exhibit I-SceI-inducible HR, which produces GFP-positive cells detected by flow cytometry. Fifty-two hours posttransfection with either control siRNA or hRad9 siRNA, the cells were transfected with I-SceI expression vector pCMV3xn-1-SceI or a control cytomegalovirus vector. pGFP, containing the full-length GFP cDNA, was transfected into these cells as a calibration control for GFP-positive cells and to ensure that the transient transfection resulted in equal expression frequencies in the siRNA-treated cells. Seventy-two hours after I-SecI transfection, cells were trypsinized to form a single-cell suspension in Dulbecco modified Eagle medium and subjected to flow cytometric analysis. Two-color (green and propidium red) fluorescence analysis revealed the percentage of green fluorescent cells relative to the total cell number (48, 72). For each analysis, 100,000 cells were processed and each experiment was repeated three times.
hRad9 has been reported to localize on telomeres in cells that maintain telomeres by the telomerase-independent pathway known as ALT (35). To determine whether overexpression of hRad9 or its inactivation influences telomere stability, we overexpressed wild-type hRad9 and a mutant form of hRad9 (hRad9 C-terminal fragment [261 to 391 aa; ΔhRad9]) in cells and monitored telomere fusions at metaphase. We selected three different cell lines to study the effect of ΔhRad9 on telomeres, because in 293 and MCF-7 cells, telomeres are maintained by telomerase, whereas in GM847 cells, telomeres are maintained by the alternative mechanism (ALT). Cells ectopically expressing hRad9 had about threefold more of the encoded protein than did the parental cells (Fig. (Fig.1A1A and data not shown). Expression of ΔhRad9 did not influence levels of endogenous hRad9. Furthermore, the levels of endogenous hRad9 were similar in different phases of the cell cycle and were unaffected by expression of ΔhRad9 (Fig. (Fig.1B).1B). 293 cells overexpressing hRad9 exhibited telomere association (TA) frequencies similar to those of control cells, suggesting that an increase in the hRad9 level in these cells has no influence on telomere stability. However, as shown in Table Table1,1, in cells expressing ΔhRad9, there was a significant increase in TA per metaphase (range of 0.65 to 1.1) relative to the frequency observed for the parental cells (range of 0.09 to 0.13). These TAs were random. Mutant ΔhRad9 tagged with HA localized to the nucleus as detected by immunostaining (data not shown) had no influence on endogenous hRad9 levels (Fig. (Fig.1A),1A), suggesting that the truncated protein may have a dominant negative effect. Since chromosome end-to-end associations may lead to anaphase bridge formation, the same cells were analyzed for this aberration. For each case, 300 cells at anaphase were examined for bridges. Cells expressing ΔhRad9 displayed a two- to fourfold higher frequency of anaphase bridges in comparison to parental cells (Table (Table11).
The presence of ΔhRad9 may produce nonspecific effects that could complicate the interpretation of experimental results. Therefore, we used RNA interference-mediated hRad9 knockdown to confirm independently the dominant negative data. Cells transfected with hRad9-specific siRNA had an 80% reduction in hRad9 levels 48 h after transfection (Fig. (Fig.1C).1C). When examined for telomere end associations, these cells gave results similar to those obtained for cells with ΔhRad9 (Table (Table11).
The chromosome end-to-end association data described above suggest that, due to loss of telomere function, chromosome end associations are formed and that these associations are not resolved in cells devoid of normal levels of Rad9. To further examine how expression of ΔhRad9 or a reduction in hRad9 levels is linked to telomere dysfunction, we determined mean TRF sizes. Southern blotting indicated no significant difference in TRF or G-overhang sizes when comparing DNA derived from cells with or without expression of ΔhRad9 or reduced levels of hRad9 (Fig. (Fig.2A).2A). However, this analysis yields an appraisal only of the entire population of TRFs generated and does not monitor the ends of individual chromosomes. We therefore performed FISH for telomeric repeats on metaphase spreads by using a telomere-specific Cy3-labeled (CCCTAA)3 peptide nucleic acid probe. We applied centromere-specific Cy5-labeled FISH to detect human centromeres, which was used to monitor the hybridization for each chromosome and to visualize whether TA has occurred between two chromosomes. Fifty metaphase chromosome spreads (about 2,300 to 3,100 chromosomes) from each cell type were included and analyzed. A significantly higher proportion of chromatid ends in cells with aberrant functional hRad9 levels (about 10% of the telomeres per metaphase) lack telomere-specific fluorescent signals compared to the control cells, in which only <1% of the chromatids do not show telomere signals per metaphase (Fig. (Fig.2B).2B). The loss of telomere signals could be a result of breaks adjacent to telomeres or shortened telomere tracts. These observations indicate that some of the chromosome end-to-end fusions observed in Rad9-inactivated cells correlated with loss of telomeric repeats. However, telomere signals were seen in more than 52% of the fusion sites, indicating that a total loss of telomeres is not required for telomere fusions in such cell systems.
To provide more evidence for the influence of mammalian Rad9 on telomere stability, we evaluated mouse ES cells in which both alleles of Mrad9 were inactivated (23). About 4% of Mrad9 knockout cells showed chromosomal aneuploidy, as well as polyploidy (Fig. (Fig.2C).2C). Consistent with the influence of hRad9 on telomere stability in human cells, we also found that Mrad9 knockout mouse ES cells had a significantly higher frequency of telomere end associations, as well as loss of telomere signals (Fig. (Fig.2C2C and Table Table1).1). These cells frequently showed Robertsonian fusions, supporting the argument that loss of telomeres results in fusions that lead to metacentric chromosomes (Fig. (Fig.2C).2C). Telomere signals were seen in more than 20% of the fusion sites, indicating that total loss of telomeres is not required for telomere fusions (Fig. (Fig.2C2C).
Telomere end association has been correlated with the inactivation of TRF2 (64). Therefore, we tested whether hRad9 interacts with TRF2. Analysis of anti-hRad9 immunoprecipitates with anti-TRF2 antibody demonstrated interaction between the two (Fig. (Fig.2D).2D). When cells were exposed to IR, the interaction of hRad9 with TRF2 was unaffected (data not shown). These observations suggest that interactions of hRad9 with TRF2 occur prior to induction of DNA damage by the application of exogenous agents. Since expression of mutant TRF2ΔBΔM is known to induce telomere fusions (60), we further tested whether knockdown of hRad9 in cells expressing mutant TRF2ΔBΔM will affect telomere function. Cells expressing mutant TRF2ΔBΔM had more than 60% of their telomeres dysfunctional. However, cells expressing mutant TRF2ΔBΔM and reduced levels of hRad9 had levels of dysfunctional telomeres similar to those seen in cells expressing only mutant TRF2ΔBΔM (Fig. (Fig.2B).2B). We failed to observe any additional telomere dysfunction measured by chromosome end fusions in cells expressing the mutant TRF2ΔBΔM and knocked-down hRad9 levels. These results suggest that hRad9 cannot rescue TRF2-mediated telomere stability.
We next determined whether telomere dysfunction in Rad9-deficient cells was due to abnormal telomerase activity. This activity was compared in vitro in extracts of cells with or without reduced levels of hRad9, and no significant difference was observed (data not shown). Mrad9+/+ and Mrad9−/− cell extracts also had similar levels of telomerase activity (data not shown), suggesting that mammalian Rad9 does not influence telomerase activity, as determined by an in vitro assay.
Recent studies have revealed that Mrad9 knockout ES cells, relative to Mrad9+/+ cells, have a higher frequency of spontaneous chromosomal aberrations (23), which correlates with their telomere instability phenotype. Thus, we determined whether the frequency of spontaneous chromosome and chromatid aberrations was increased in human cells with inactivated hRad9. Again, cells expressing ΔhRad9, as well as those with reduced levels of hRad9 due to the presence of siRNA, displayed a higher frequency of chromatid, as well as chromosome-type, aberrations compared to parental cells (Table (Table1).1). No significant increase in these frequencies was observed in cells overexpressing hRad9 (Table (Table1).1). Thus, mammalian cells with reduced levels of Rad9 or expression of ΔhRad9 or inactivated Rad9 expressed had frequent loss of telomere signals, enhanced telomere fusions, and higher frequencies of chromatid breaks and gaps, but not of chromosomal breaks and gaps, compared to control cells. Based on these observations of telomere dysfunction and chromosome aberrations in cells with inactivated mammalian Rad9, such genomic instability might be due to defective DNA repair or damage response checkpoints.
Rad9 forms a complex with Hus1 and Rad1 (hRad9-hHus1-hRad1) called the 9-1-1 complex (73). This complex is loaded onto DNA in response to many different types of genotoxic stress, including IR, UV light, replication inhibitors, and alkylation, suggesting that Rad9 has a role in cellular responses activated by multiple types of DNA damage (73). Although it is known that Mrad9 inactivation results in enhanced cell killing after IR exposure (23), it was not known whether Rad9 inactivation affects IR sensitivity in a specific cell cycle phase.
Telomere instability in mammals can correlate with organismal hypersensitivity to IR (18). Since inactivation of mammalian Rad9 influences telomere stability, we determined whether overexpression or inactivation of wild-type hRad9 influences cell survival after exposure to IR at a specific cell cycle phase. Cell survival after IR exposure was determined by using a colony formation assay described previously (13). Cells overexpressing hRad9 did not exhibit any altered sensitivity to IR exposure in comparison to the parental cells; however, cells with hRad9 levels reduced by siRNA, as well as cells expressing ΔhRad9, demonstrated enhanced cell killing after IR exposure (Fig. (Fig.3A).3A). To define the precise phase of enhanced cell death after IR exposure, we enriched cells in different phases of the cell cycle by centrifugal elutriation. The G1-phase-enriched cell population had about 91% such cells, the S-phase-enriched population contained greater than 81% cells in S phase, and the G2/M-phase-enriched population contained 72 to 76% cells in that phase (data not shown). The survival characteristics of the cells varied through the cell cycle. Expression of ΔhRad9 did not alter the IR sensitivity of cells in G1 (Fig. (Fig.3B).3B). However, S- and G2-phase cells with inactivated hRad9 showed enhanced IR-induced cell killing relative to control cells (Fig. 3C and D). These studies indicate that hRad9 has a specific role in S- and G2-phase cell survival after IR exposure. We further established that enhanced cell killing in S- and G2-phase cells was not due to the change in hRad9 levels since the abundance of this protein was similar in control radioresistant hRad9+ cells in different phases of the cell cycle (Fig. (Fig.1B1B).
H2AX is associated with nuclear foci containing factors that are essential for DNA repair, replication, and cell cycle regulation (17, 49, 50). H2AX phosphorylation is critical for protecting the genome from spontaneous DSBs, as well as those induced by IR or V(D)J recombination (3, 4). H2AX phosphorylation coincides with the sites of DSBs, irrespective of their origin (6, 29, 47, 51, 69). Since inactivation of Rad9 influences cell survival after IR exposure and hRad9 colocalizes with γ-H2AX at DNA DSBs (19), we examined whether a reduction in hRad9 abundance or activity can influence the appearance of IR-induced γ-H2AX foci. Human 293 cells with reduced hRad9 levels had about three γ-H2AX foci per cell 2 min after irradiation with 1 Gy. Control cells with normal levels of hRad9 had about seven foci per cell under the same conditions (Fig. (Fig.4A).4A). Similar results were observed in Mrad9−/− ES cells, where the appearance of γ-H2AX foci was delayed compared to that in control Mrad9+/+ ES cells (data not shown).
To assess the disappearance of γ-H2AX foci, we analyzed human cells 120 and 360 min postirradiation. Furthermore, we determined whether differences in the appearance of γ-H2AX foci after IR exposure are due to cell cycle distribution. To address this question, we labeled the cells for 30 min with BrdU, followed by irradiation, and then examined them for γ-H2AX foci at different times after treatment. Specifically, S-phase Mrad9−/− cells, as well as human cells with inactivated hRad9, have a lower number of γ-H2AX foci compared to parental control cells (Fig. (Fig.4B4B and data not shown). In general, the number of γ-H2AX foci in cells with inactivated mammalian Rad9 was significantly lower until 5 min postirradiation. However, the γ-H2AX focus number was higher at 120 min postirradiation, suggesting that this protein plays a role in DNA damage repair (Fig. (Fig.4A4A).
Telomere instability and IR sensitivity are linked to defective DNA repair (27, 38, 54, 55, 70). To determine whether the S- and G2-phase-specific increased cell killing by IR in cells with reduced levels of hRad9 correlated with DNA repair, we compared cell cycle stage-specific IR-induced chromosomal aberrations in cells with or without reduced levels of this protein. We first set out to determine frequencies of chromosomal aberrations induced by IR in G1-, S-, and G2-phase cells. Cell cycle phase-specific chromosome aberrations were ascertained based on the frequency of chromosomal and chromatid-type aberrations observed at metaphase. G1-specific aberrations detected at metaphase are mostly of the chromosomal type and include a high frequency of dicentrics (55). S-phase-type aberrations detected at metaphase are chromosomal, as well as those involving chromatids. G2-type aberrations detected at metaphase are predominantly the chromatid type with the least number of dicentrics. To determine G1-type chromosome damage, cells were treated with 3 Gy and replated 24 h after irradiation, and aberrations were scored at metaphase as previously described (24, 55). No difference in residual IR-induced G1 chromosomal aberrations was seen in metaphase cells with or without reduced levels of hRad9 (Fig. (Fig.5A).5A). Furthermore, cells overexpressing hRad9 did not show any difference in G1-phase-type chromosome aberrations relative to parental cells after IR exposure (data not shown). To determine whether defective repair can be documented in cells overexpressing hRad9 or with ΔhRad9 in phases of the cell cycle other than G1, we evaluated S-phase-specific chromosome aberrations. We first determined the time needed for S-phase cells to reach metaphase after IR treatment. Exponentially growing 293 cells were labeled with BrdU for 30 min as previously described (55) and then irradiated with 2 Gy. Anti-BrdU immunostaining was performed to determine when metaphase chromosomes contain BrdU. In these experiments, BrdU-labeled metaphases appeared approximately 3 h postirradiation (data not shown). Thus, cells producing hRad9 protein were treated with 2 Gy of IR and metaphases were collected 3 to 5 h postirradiation. Cells overexpressing hRad9 and collected 3 h postirradiation displayed frequencies of metaphases with chromatid and chromosomal aberrations similar to those of parental cells (data not shown). In contrast, cells with reduced levels of hRad9 had significantly higher frequencies of chromosome and chromatid aberrations after IR exposure compared to parental cells (Fig. (Fig.5B).5B). Furthermore, when cells were treated with 1 Gy of gamma rays, hRad9 overexpression had no effect on G2-phase-specific chromosome repair (data not shown), while hRad9 inactivation increased G2-phase-specific chromosome aberrations (Fig. (Fig.5C).5C). These observations suggest a role for Rad9 in repairing chromosome damage in the S, as well as in the G2, phase of the cell cycle.
The underlying reasons for the increased aberrations at mitosis observed in cells with inactivated Rad9 could be the inability to modulate cell cycle progression following IR exposure, increased initial levels of genetic damage, and/or decreased efficiency of DNA damage repair. To address whether IR-induced increased initial levels of genetic damage and/or decreased efficiency of repair is responsible, we used PCC to visualize directly chromosome events in interphase cells following irradiation (40). G1 chromosomes are rodlike and unreplicated, whereas G2 chromosomes are replicated and have two chromatids (43). Prematurely condensed chromosomes from S-phase cells appear fragmented (pulverized) even without irradiation; therefore, S-phase cells cannot be compared. G1- and G2-phase cells, with or without hRad9, were enriched by centrifugal elutriation, gamma irradiated, and then allowed to repair the resulting DNA damage for increasing intervals of time prior to the induction of PCC. Subsequently, residual levels of chromosome damage were assessed directly. Cells with or without reduced levels of hRad9 were irradiated with 3 Gy of gamma rays on ice and then incubated at 37°C for different periods of time prior to fusion with mitotic cells to visualize chromosomes. Chromosome fragments were counted at different time points, and the mean additional number of fragments seen immediately after irradiation was about 12 higher than the control. No difference in initial chromosome damage (12 breaks per cell) or the kinetics of chromosome repair was observed in G1-phase cells with or without normal Rad9 function (Fig. (Fig.5D),5D), consistent with results observed at metaphase (Fig. (Fig.5A).5A). Interestingly, G2-phase cells with Rad9 inactivated had initial chromosome damage (10 breaks) similar to that of control cells; however, hRad9-deficient cells had a higher frequency of unrepaired chromosome breaks and gaps compared to parental cells (Fig. (Fig.5E).5E). Such results were consistent with the observations made for cells at metaphase (Fig. (Fig.5C)5C) and thus support the argument that hRad9 influences the repair of G2-phase-specific chromosome damage.
To determine whether the increase in S-phase-specific IR-induced chromosomal damage is due to defective cell cycle checkpoints, we first determined whether mammalian Rad9 inactivation results in loss of the G1 checkpoint. We found that 293 cells with reduced levels of hRad9 caused by siRNA or ΔhRad9 expression showed a significant increase in G1-phase cells with a decrease in S-phase entry after IR exposure, similar to control cells that showed a significant decrease in S-phase entry, indicating G1 arrest (Fig. (Fig.6A6A and data not shown). Similar results were found for Mrad9−/− and Mard9+/+ cells. These results suggest that Rad9 has no (detectable) influence on the G1 checkpoint in these cell systems.
Exposure to IR causes a transient inhibition of DNA replication. Cells deficient in ATM function exhibit radioresistant DNA synthesis, an indicator of loss of the S-phase checkpoint (38). Cells expressing ΔhRad9 or cells with siRNA-reduced Rad9 levels, as well as Mrad9−/− cells, did show inhibition of DNA synthesis after IR exposure which was similar to that of the control cells (Fig. (Fig.6B6B and data not shown). To determine whether mammalian Rad9 influences the G2 checkpoint, the efficiency of G2 delay was evaluated by measuring the proportion of cells in mitosis (mitotic index) after irradiation. Cells expressing ΔRad9 or having reduced levels of hRad9 showed no significant difference in the mitotic index after irradiation with 2 Gy of IR compared to the control, suggesting that inactivation of mammalian Rad9 does not lead to a defective G2 checkpoint (Fig. (Fig.6C).6C). These results suggest that Rad9 has a critical role in DNA damage repair rather than in IR-induced cell cycle checkpoint control. Such results are consistent with previous studies of Mrad9−/− mouse ES cells, which demonstrate that there is no major problem with IR-induced G2 checkpoint control initiation, only a defect in the prolonged maintenance of the checkpoint, consistent with a defect in DNA repair (23). Thus, the enhanced chromosomal damage seen in mammalian cells with inactivated Rad9 cannot be attributed to defective cell cycle checkpoints, again suggesting that altered DNA repair is the more likely explanation.
In mammalian cells, in response to DNA damage, ATM gets autophosphorylated and phosphorylates various proteins that control G1-, S-, and G2-phase cell cycle arrest (16, 38, 57). We examined whether inactivation of hRad9 influences IR-induced ATM phosphorylation at Ser-1981 in 293 cells. ATM phosphorylation by IR was found in cells with or without knockdown of hRad9 (Fig. (Fig.7).7). We also examined whether inactivation of hRad9 influences ATM-mediated, IR-induced phosphorylation of Chk2 in 293 cells. ATM phosphorylates Chk2 on Thr-68 in response to DNA damage (31). Cells with or without reduced levels of hRad9 showed no difference in the IR-induced phosphorylation of Chk2 at Thr-68 (Fig. (Fig.7).7). Since hRad9 function does not influence IR-induced ATM or ATM-mediated Chk2 phosphorylation, such results argue further against a role for hRad9 in cell cycle checkpoint control but thus support a role in DNA damage repair.
There are two major ways to maintain the ends of chromosomes in yeast and mammalian nuclei: mechanisms related to telomerase and recombination. Without telomerase or recombination, cells enter a state of growth arrest. Since mammalian Rad9 inactivation did not influence telomerase activity, we tested whether hRad9 was involved in NHEJ or HR.
The aforementioned results established a role for hRad9 in S- and G2-phase-specific DNA damage repair. It has been suggested that the cell cycle could influence DSB repair since NHEJ acts predominantly in G1 and HR acts in S and G2 (61). To determine whether the hRad9 protein is directly involved in DNA end joining, we used a well-characterized in vitro assay that involves the rejoining of DNA ends of a plasmid in 293 cell extracts as previously described (5, 54). Plasmid pUC19 was digested with SalI, and DNA end joining was performed as previously described (5, 54). hRad9 in 293 cell extracts was depleted by immunoprecipitation with hRad9 antibody. Complete immunodepletion of hRad9 was confirmed by Western analysis (Fig. (Fig.8A).8A). hRad9-immunodepleted extracts or extracts from Rad9 siRNA-treated cells were used to assay DNA end-joining activity. No difference in this activity was found in extracts with or without hRad9 (Fig. (Fig.8B).8B). These results suggest that hRad9 does not have a role in the DNA end-joining process. However, these observations did not rule out a role for hRad9 in chromosomal recombination.
The defect in chromosome repair found in hRad9 knockdown cells in S and G2 suggested a role for hRad9 in HR since this process predominates in cells in these phases of the cell cycle, when sister chromatids are available as a template for repair. It is known that the hRad9-hHus1-hRad1 complex is loaded onto damaged DNA in an hRad17-dependent manner (73) and hRad9 colocalizes with γ-H2AX at those sites (19). hRad9 also colocalizes with telomeric DNAs and promyelocytic leukemia bodies in ALT cells (35). ALT cells have abundant telomeric circles, suggesting that frequent t-loop HR events could promote rolling-circle replication of telomeres in the absence of telomerase. Wang and coworkers (68) demonstrated that t-loop deletion by HR influences mammalian telomeres. A key feature of HR is DNA strand invasion, which is catalyzed by the Rad51 protein coating single-stranded DNA; however, the exact roles of the many accessory proteins that facilitate this process are not known. Rad51D is involved in the maintenance of telomeres. Rad51 can form distinct nuclear foci in response to IR exposure (63). These foci can be visualized by microscopy and are thought to be sites where repair reactions take place (62). Since hRad9 influences telomere stability, as well as DNA damage repair, we determined whether hRad9 interacts with Rad51.
Analysis of anti-hRad9 immunoprecipitates with anti-hRad51 antibody demonstrated interaction between the two (Fig. (Fig.2D).2D). When cells were exposed to IR, the association of hRad9 with hRad51 was unaffected (data not shown). These observations suggest that interaction of hRad9 with hRad51, as well as with TRF2, occurs prior to induction of DNA damage by the application of exogenous agents.
The interaction of hRad9 with hRad51 suggests that hRad9 may have a role in HR. Therefore, we measured HR by examining the frequency of reconstitution of a GFP reporter gene within a chromosomally integrated plasmid substrate in cells with or without reduced levels of hRad9. Following I-SceI transfection, cells containing the pDR-GFP substrate demonstrated a >80-fold increase in the number of GFP-positive members compared with cells containing the control vector, indicating that almost all of the GFP-positive cells resulted from DSB-induced recombination (Fig. (Fig.9).9). pDR-GFP cells treated with hRad9 siRNA had a three- to fourfold reduction in the number of GFP positives in comparison to control transfected cells (t test, P < 0.001), providing strong evidence that hRad9 affects the HR pathway for DNA DSB repair.
The function of mammalian Rad9, a gene involved in sensing DNA damage, was examined in the context of chromosome stability and DNA repair in mammalian cells. Human cells in which hRad9 was inactivated by expression of ΔhRad9, or levels of the protein were reduced by siRNA, or mouse cells with Mrad9 deleted (23) were used. A cDNA fragment encoding aa 269 to 391 of hRad9 was expressed in different human cells, which were subsequently examined for various end points. We found that the C-terminal fragment of hRad9 (aa 269 to 391) demonstrated dominant negative activity for almost all of the assays employed. Thus, expression of this fragment might block the interaction of wild-type hRad9 with other proteins and thus has a dominant negative function. This function of the hRad9 fragment is consistent with the previously described observation of Wang and coworkers (67), who reported that the C-terminal region of hRad9 interacts with other proteins. We found that, indeed, the frequencies of chromosomal abnormalities, such as chromosome end-to-end associations and chromosomal breaks, were higher in cells with inactivated mammalian Rad9. These chromosomal aberrations appear to involve dysfunctional telomeres, as a significant number of cells showed TAs. Thus, the results suggest that abrogating the function of mammalian Rad9, which is implicated in mediating DNA damage-sensing checkpoints, increases the frequency of unrepaired chromosomal aberrations that can lead to an increase in the number of telomere fusions. This mechanism might contribute to the inviability of mouse embryonic fibroblasts when Mrad9 is inactivated (23). The frequencies of spontaneous, as well as IR-induced, chromosomal aberrations were dramatically higher in cells with inactivated Rad9 relative to those in controls, suggesting that loss of telomere function occurred by breakage near telomeres, rather than by a telomerase-based mechanism. This is supported by our finding that knockdown of hRad9 does not influence telomerase activity (data not shown). hRad9 forms a heterotrimer with Hus1 and Rad1, which is loaded onto DNA by Rad17-mediated mechanisms, and these proteins are evolutionarily conserved. Interestingly, studies with S. pombe, S. cerevisiae, and Caenorhabditis elegans orthologues also indicate roles for these proteins in telomere metabolism (1, 11, 25, 28, 36).
It is important to note that in cells with both alleles of Rad9 inactivated, frequent chromosome breakage with complete loss of telomeric repeats in some chromosomes, as well as chromosome end fusions, occurred under normal growth conditions in the absence of exogenous genotoxic challenges (Fig. (Fig.22 and Table Table1).1). The fact that the G1 checkpoint was operative in cells with reduced levels of Rad9 is supported by the observations that irradiation induced comparable frequencies of cells in G1 and S phase compared to those cells with normal levels of hRad9 (Fig. (Fig.6A).6A). Furthermore, cells with reduced levels of Rad9 did not show any defect in the S-phase-specific cell cycle checkpoint, as such cells did not differ from control cells for radioresistant DNA synthesis (Fig. (Fig.6B).6B). The mitotic index after irradiation was identical in cells with or without functional Rad9, indicating that mammalian Rad9 inactivation does not influence the G2 checkpoint in the cell lines studied (Fig. (Fig.6C).6C). hRad9 knockdown had no influence on IR-induced ATM Ser-1981 phosphorylation or Chk2 Thre-68 phosphorylation, supporting the argument that mammalian Rad9 has a minimum role in cell cycle checkpoint control postirradiation. In contrast, there was a clear and significant difference in the chromosome aberration frequency if log-phase cells were irradiated, and mitoses shortly thereafter were monitored; at 4 h postirradiation, there was a 1.8-fold difference in the frequencies of S-phase aberrations/metaphase between cells with and without Rad9 fully functional (Fig. (Fig.5B).5B). Furthermore, when G2-induced aberrations were monitored after irradiation, cells with reduced levels of Rad9 showed higher frequencies of aberrations in the same time span compared to controls, indicating very inefficient repair (Fig. (Fig.5C).5C). Utilizing the PCC technique, we found that there was no difference in the kinetics of repair or the residual number of chromosome breaks in G1-phase cells with or without reduced levels of hRad9 (Fig. (Fig.5D);5D); however, cells deficient in Rad9 had a higher frequency of chromosome gaps and breaks in the G2 phase (Fig. (Fig.5E).5E). Consistent with these data, survival studies demonstrated that cells in the S and G2 phases were sensitive to irradiation if Rad9 was inactivated (Fig. 3C and D). These data, taken together, reinforce the idea that in cells with hRad9 knockdown, chromosomal damage that might occur during the S or G2 phase could be due to defective chromosome repair (Fig. 5B, C, and E). In cells lacking Rad9 protein, cell cycle arrest after DNA damage is operative and the cells do not enter mitosis exceptionally early. Our study reveals that as a consequence of this inability to appropriately repair, such cells enter mitosis with a higher frequency of chromosomal aberrations.
hRad9 is rapidly retained at sites of damaged DNA and colocalizes with the phosphorylated form of H2AX (γ-H2AX) (19). In addition, we found quantitative differences in the kinetics of appearance and disappearance of γ-H2AX foci and kinetics of repair of G2 chromosomes (Fig. (Fig.44 and and5E),5E), dependent upon whether cells do or do not have functional mammalian Rad9. These results provide strong evidence that mammalian Rad9 plays a critical role in repair of DNA damage.
The association of hRad9 with TRF2 provides further evidence that hRad9 may have a role in maintaining the stability of telomeres. Although knockdown of hRad9 has a minimal role in TRF2 association with telomeres (data not shown) and inactivation of hRad9 and TRF2 did not rescue a telomere instability phenotype (Fig. (Fig.2),2), it is possible that TRF2 might be mediating DNA damage signaling through Rad9. This notion is further supported by the data indicating that inactivation of hRad9 influences HR (Fig. (Fig.9)9) but not NHEJ (Fig. (Fig.88).
Our results demonstrate that hRad9 can influence telomere stability and HR. However, it is not clear exactly how the protein functions in these processes. It is possible that hRad9 modulates chromatin structure, an early essential step for HR. This is supported by our demonstration that hRad9 inactivation delays the appearance of IR-induced γ-H2AX foci. Although the data support a role for Rad9 in chromatin remodeling, especially at telomeres, it is not clear whether the protein can influence chromatin structure beyond this limited region. Such a general chromatin alteration should influence repair in all phases of the cell cycle, as has been observed in ATM null cells (42). However, our results do not support this activity for hRad9 since inactivation of the protein resulted in an enhanced defect in chromosome repair confined only to the S and G2 phases of the cell cycle. Alternatively, Rad9 may interact with and influence the activity of proteins that have a role in HR repair, which occurs predominantly in S- and G2-phase cells. This is supported by our demonstration of a physical interaction between hRad9 and hRad51. In fact, Shinohara and coworkers (58) demonstrated that S. cerevisiae orthologues of some of the hRad9 checkpoint pathway-related proteins interact with Rad51 for repair of DSBs in meiosis via recombination. Furthermore, Grushcow and coworkers (20), as well as Aylon and Kupiec (2), showed that the S. cerevisiae equivalents of hRad9, hRad1, hHus1, and hRad17 pathway members are involved in recombination partner choice. In addition, we observed the delayed appearance of γ-H2AX foci in S-phase cells after IR exposure of Mrad9−/− relative to Mrad9+/+ cells. Moreover, the high frequency of chromatid gaps and breaks in hRad9-inactivated cells supports a role for the protein in HR repair. Inactivation of mammalian Rad9 could thus contribute, at least in part, to the radiosensitivity, genomic instability, and loss of telomeres observed by mediating the abrogation of HR repair.
This investigation was supported by NIH grants (NS34746, CA10445), the Department of Defense, the A-T Children's Society, and funds from Radiation Oncology, Washington University School of Medicine, to T.K.P.; NIH grants CA89816 and GM52493 to H.B.L.; and a Canadian Cancer Society grant (013235) to R.J.W.
We thank W. Wright for critical comments on and suggestions for the manuscript. We thank K. D. Brown, M. B. Kastan, S. J. Elledge, T. deLange, and J. Chen for providing several reagents that were used in this study.