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Mol Cell. Author manuscript; available in PMC Aug 14, 2010.
Published in final edited form as:
Mol Cell. Aug 14, 2009; 35(3): 384–393.
doi:  10.1016/j.molcel.2009.06.011
PMCID: PMC2756616

SOSS complexes participate in the maintenance of genomic stability


Proteins that bind to single-stranded DNA (ssDNA) are essential for DNA replication, recombinational repair and maintenance of genomic stability. Here we describe the characterization of a ssDNA-binding heterotrimeric complex, SOSS (Sensor Of Single-stranded DNA) in human, which consists of human SSB homologues hSSB1/2 (SOSS-B1/2), INTS3 (SOSS-A) and a previously uncharacterized protein C9orf80 (SOSS-C). We have shown that SOSS-A serves as a central adaptor required not only for SOSS complex assembly and stability, but also for facilitating the accumulation of SOSS complex to DNA ends. Moreover, SOSS-depleted cells display increased IR sensitivity, defective G2/M checkpoint and impaired HR repair. Thus, our study defines a pathway involving the sensing of ssDNA by SOSS complex and suggests that this SOSS complex is likely involved in the maintenance of genome stability.


DNA double-strand breaks (DSBs) are highly cytotoxic lesions that, if unrepaired or repaired incorrectly, can cause genome instability and promote tumorigenesis (Bartek and Lukas, 2007; Bartkova et al., 2005; Friedberg, 2003; Hoeijmakers, 2001). Cells possess two main DSB repair mechanisms: non-homologous end-joining (NHEJ) and homologous recombination (HR) (Kennedy and D’Andrea, 2006; Lukas and Bartek, 2004; Weinstock et al., 2006b). In vertebrates, NHEJ and HR differentially contribute to DSB repair, depending on the nature of the DSB and the phase of the cell cycle (Bartek et al., 2004; Sonoda et al., 2006). HR pathway is critical for the maintenance of genome stability through its involvement in the precise repair of DNA double-strand breaks and restarting stalled or collapsed DNA replication forks. It is believed that one of the initial steps during homologous recombination is the resection of DSBs to generate single-stranded DNA (ssDNA), which is bound by Single-stranded-DNA-binding proteins (SSBs) that play essential roles in DNA replication, recombination and repair in bacteria, archaea and eukarya (Borde, 2007; Buis et al., 2008; Clerici et al., 2005; Hopkins and Paull, 2008; Lavin, 2004; Lengsfeld et al., 2007; Petrini and Stracker, 2003; Sartori et al., 2007; Takeda et al., 2007; West, 2003; Williams et al., 2008; Wold, 1997).

The human SSB, known as human replication protein A (RPA), is a heterotrimer composed of subunits of 70, 32 and 14 kDa, each of which is conserved not only in mammals but also in all other eukaryotic species (Wold, 1997). RPA is generally believed to be the major SSB protein in eukaryotic cells given that it not only is critically important for DNA replication but also participates in various DNA repair or other cellular processes that involve DNA transaction.

This view was challenged by the recent identification of two additional human SSB homologues, hSSB1 and hSSB2 (Richard et al., 2008). Cells deficient in hSSB1 exhibit defective checkpoint activation, increased radiation sensitivity and defective homologous recombination repair, indicating that hSSB1 plays an important role in the cellular response to DNA damage (Richard et al., 2008). Unlike RPA, which exists as heterotrimeric complex, hSSB1 and hSSB2 were believed to be more similar to E. coli SSB, that exists as monomeric form or homo-oligomers (Richard et al., 2008). However, exactly how hSSB1 (or hSSB2) would specifically sense ssDNA regions during DNA damage repair but not be involved in normal DNA replication is still unknown. In this study, we used an affinity purification approach to isolate hSSB1/2-containing complex. Interestingly we identified a hetero-trimetric complex, which we refer to as SOSS (Sensor of Single-stranded DNA) complex that contains not only hSSB1/2, but also INTS3 and a previously uncharacterized protein C9orf80. We demonstrated that both SOSS complexes and CtIP/RPA act downstream of MRE11/RAD50/NBS1 complex and function in DNA damage repair.


INTS3, hSSB1/2 and C9orf80 form a heterotrimeric protein complex

In an attempt to understand what determines the specialized role of hSSBs in DNA repair, we performed tandem affinity purification using HEK293T cells stably expressing strepavidin-flag-S protein (SFB)-tagged wild-type hSSB1/2 for the identification of hSSB1/2-associated proteins. We repeatedly found INTS3 and a previously uncharacterized protein C9orf80 as major hSSB1/2-associated proteins (Fig. 1A). To further confirm that INTS3 and C9orf80 exist in the same complex with hSSB1 or hSSB2, we generated stable cells expressing triple-tagged INTS3 and C9orf80 respectively. Notably, mass spectrometry analyses of INTS3 or C9orf80-associated protein complexes revealed peptides that corresponded to hSSB1 and hSSB2 (data not shown), suggesting that these proteins likely form stable complex in vivo. INTS3 (integrator complex subunit 3) was originally identified as a subunit of Integrator (Baillat et al., 2005), however, there was no further characterization of this protein concerning its domain structure, activity, or biological function. C9orf80 is a hypothetical protein with no known function. Although INTS3 and C9orf80 were present in both hSSB1 and hSSB2 purification, we did not detect the presence of hSSB1 from several independent hSSB2 large-scale affinity purifications and vice versa (Fig. 1A and data not shown), indicating that hSSB1 and hSSB2 might exist in two complementary complexes which contain the common subunits INTS3 and C9orf80. Therefore in this study we named the complex containing INTS3/hSSB1/C9orf80 or INTS3/hSSB2/C9orf80 as SOSS1/2 (Sensor Of Single-stranded DNA complex 1/2), respectively. Accordingly, we designated INTS3, hSSB1/2 and C9orf80 as SOSS subunit A, B1/2 and C.

Fig. 1
Formation of a SOSS complex containing INTS3 (SOSS-A), hSSB1/2 (SOSS-B1/2), and C9orf80 (SOSS-C)

To verify the association among SOSS-A, SOSS-B1, SOSS-B2 and SOSS-C, we performed co-immunoprecipitation experiments. When immunoprecipitation experiments were conducted with anti-SOSS-A or anti-SOSS-C antibodies, all of the SOSS subunits could be detected (Figure 1B). However, while antibodies specifically recognizing SOSS-B1 or SOSS-B2 co-immunoprecipitated the common subunits SOSS-A and SOSS-C, no SOSS-B1 or 2 were present in each other immunocomplex (Figure 1B). These data agree with the data obtained from SOSS-B1/2 large-scale affinity purifications and support our hypothesis that SOSS-A, SOSS-B1/2 and SOSS-C might form two complementary hetrotrimetric complexes: SOSS1 (SOSS-A/B1/C) or SOSS2 (SOSS-A/B2/C) in vivo. Interestingly, neither RPA nor CTIP could interact with SOSS-A (Figure S1). Furthermore, the SOSS complex formation was DNA damage independent and these pre-existing complexes could be detected in HeLa cells as well as in other cell lines including HEK293T cells (Figure 1B and data not shown). Formation of the heterotrimeric complex was further ascertained by gel filtration analysis. Insect cells were coinfected with baculovirus expressing GST-SOSS-A, His-SOSS-B and SOSS-C and complex formation was studied by FPLC. As shown in Figure 1C, SOSS-A, SOSS-B and SOSS-C co-eluted as a heterotrimeric complex with a molecular mass of approximately 190 kDa. Previous study has already established that recombinant SOSS-B1/hSSB1 binds specifically to ssDNA substrates (Richard et al., 2008). As shown in Figure S2, recombinant SOSS heterotrimeric complex also specifically binds to ssDNA but not to dsDNA.

To find out exactly how the SOSS complex is assembled, we examined the association among SOSS-A, SOSS-B1 and SOSS-C in insect cells. As shown in Figure 1D, SOSS-A interacted strongly with SOSS-B or SOSS-C, whereas no direct binding was detected between SOSS-B and SOSS-C (data not shown), suggesting that SOSS-A likely serves as a central assembly factor which mediates the formation of this complex. Therefore we focused on this key subunit SOSS-A in this study.

We first sought to identify the regions on SOSS-A responsible for its interaction with SOSS-B or SOSS-C. Myc-tagged wild-type SOSS-A and a series of deletion mutants that span the entire SOSS-A open reading frame were subjected to co-immunoprecipitation with full-length SFB-tagged SOSS-B or SOSS-C. Results showed that while the SOSS-A N-terminus (residues 1-419) is responsible for SOSS-B binding, a larger N-terminal region (residues 1-628) is necessary for its binding to SOSS-C (Figure 1E and 1F). These data indicate that SOSS-B and SOSS-C share overlapping binding regions on SOSS-A.

SOSS-A is required for the proper localization and stability of SOSS-B1/2 in vivo

The absence of a critical subunit of a multicomponent protein complex often destabilizes the complex (Yin et al., 2005). Therefore, we depleted either SOSS-A or SOSS-B1/2 and examined the stability of the other subunits. As shown in Figure 2A, depletion of SOSS-B1 or SOSS-B2 did not result in any significant change in SOSS-A protein level, however depletion of SOSS-A by siRNA led to a dramatic decrease in SOSS-B1 and SOSS-B2 protein levels. This implies that SOSS-A may help to stabilize SOSS-B1 and SOSS-B2 in the cell. Interestingly, we noticed that the protein levels of SOSS-B1 and SOSS-B2 seem to have an inverse relationship, as depletion of one protein appears to increase the level of the other (Figure 2A), again indicating that SOSS-B1 and SOSS-B2 might play complementary role in DNA damage response pathway.

Fig. 2
SOSS-A regulates SOSS complex stability and focus formation

Like many DNA damage/repair proteins, SOSS-B1 was able to localize at sites of DNA breaks and form discrete foci that colocalize with the DNA double-strand break marker γ-H2AX (Richard et al., 2008). Given that SOSS-A and SOSS-C exist in a complex with SOSS-B1 or SOSS-B2, we would like to examine whether SOSS-A and SOSS-C could also form foci following DNA damage. Immunostaining experiments showed SOSS-A, SOSS-B2 and SOSS-C to be evenly distributed in the nucleoplasm in normal cells (data not shown). However, after exposure of cells to IR, SOSS-A, SOSS-B2 and SOSS-C relocalized to foci that costained with γ-H2AX (Figure 2B), indicating that the localization of SOSS-A, SOSS-B2 and SOSS-C, like that of SOSS-B1, is regulated in response to DNA damage. Since all the components of SOSS complexes form ionizing radiation-induced foci (IRIF), we next examined how they would influence each other’s foci formation ability. As shown in Figure 2C and 2D, IRIF of SOSS-B1 and SOSS-B2 were dramatically decreased in SOSS-A depleted cells. However, depletion of SOSS-B1 or SOSS-B2 did not lead to any significant change in SOSS-A IRIF formation (Figure 4B; Figure S6A), implying that SOSS-A may act upstream of SOSS-B1/2 and be required for SOSS-B1/2 focus formation. In agreement with our previous hypothesis that SOSS-B1 and SOSS-B2 might play complementary role in the DNA damage response, depletion of SOSS-B1 led to a modest increase of SOSS-B2 foci formation and vice verse (Figure 2D).

Fig. 4
Both SOSS and CTIP/RPA promote optimal DNA damage response

Generally, the DNA damage-induced focus formation reflects the assembly of proteins at the vicinity of DNA breaks. These proteins are recruited physically to the damaged DNA and become chromatin bound. Since SOSS-A is required for SOSS-B foci formation, we hypothesized that SOSS-A should also be required for the localization of SOSS-B to chromatin. Indeed, SOSS-A depletion results in abrogation of chromatin targeting of both SOSS-B1 and SOSS-B2 (Figure 2E). Together, these data suggest that SOSS-A not only is required for the assembly of this trimeric protein complex, but also plays an important role in stabilizing this protein complex at DNA damage sites.

SOSS complex participates in cellular response to DNA double-strand breaks

Cells deficient in hSSB1 display enhanced genomic instability including defective G2/M checkpoint activation, increased IR sensitivity and deficient homologous recombination repair (Richard et al., 2008). We examined whether the loss of the SOSS-A would result in similar defects in the DNA damage response. Using a previously established G2/M checkpoint assay (Xu et al., 2001), we showed defective G2/M checkpoint control in SOSS-A-depleted cells (Figure 3A). SOSS-A-depleted cells were also more sensitive to radiation than control cells (Figure 3B). Moreover, we performed a gene conversion assay to examine HR efficiency using the DR-GFP reporter system (Weinstock et al., 2006a). Indeed, HR repair efficiency was reduced by ~2-to-2.5-fold in SOSS-A-depleted cells (Figure 3C and 3D). The recombination protein RAD51 is the key component of the homologous recombination repair machinery and the formation of Rad51 foci can be used as another indicator of HR repair. In agreement with the results from gene conversion assay, DNA damage-induced Rad51 foci formation was also impaired in SOSS-A-depleted cells (Figure 3E and 3F; Figure S3). Together these data indicated that SOSS complexes play an important role in DNA damage response.

Fig. 3
SOSS-A is required for IR-induced G2/M checkpoint, cell survival and efficient DNA repair

MRN is required for efficient SOSS foci formation in S/G2 cells

It has been shown that the MRE11/RAD50/NBS1 (MRN) complex promote DNA end resection and the generation of single-stranded DNA (ssDNA), which is critically important for recruitment of replication protein A (RPA) and HR repair (Borde, 2007; Buis et al., 2008; Hopkins and Paull, 2008; Lavin, 2004; Petrini and Stracker, 2003; Williams et al., 2008). Given that SOSS complex bound to ssDNA, we expected that MRN might be required for SOSS complex foci formation. Strikingly, like RPA2, SOSS complex foci formation was significantly reduced upon MRN depletion, indicating that MRN complex are involved in the generation of not only RPA but also SOSS-coated-ssDNAs (Figure 4A; Figure S4A-4C). This requirement of MRN complex for the formation of SOSS foci appears to be restricted to S or G2 cells, since the damage-induced SOSS focus formation in G1 arrested cells could still occur independent of MRN complex (Figure S4D and 4E).

Because SOSS-A was absolutely required for SOSS complex chromatin targeting and focus formation, we tested the possibility that MRN complex might bring SOSS complex to ssDNAs via a direct interaction with SOSS-A. Indeed, we found that SOSS-A specifically interacted with NBS1, but not with Mre11 or RAD50 (Figure S5A). Moreover, SOSS-A interacted with NBS1 in insect cells (Figure S5B). This interaction between SOSS-A and NBS1 suggests that SOSS and MRN complexes may at least in part act together and participate in DNA damage response.

Both SOSS complex and CTIP function in DNA damage repair

More recently, the proposed mammalian Sae2 homolog, CTIP, was also shown to interact with MRN complex and be required for the generation of RPA-coated ssDNAs (Clerici et al., 2005; Lengsfeld et al., 2007; Mimitou and Symington, 2008; Sartori et al., 2007; Takeda et al., 2007). We thus examined whether CTIP, like MRN complex, would also be involved in SOSS complex focus formation. When cells were treated with CTIP siRNA, only RPA focus formation, but not the focus formation of SOSS complex, was disrupted (Figure 4B; Figure S6A-6D). Conversely, in SOSS-depleted-cells, CTIP or RPA2 foci formation was not obviously altered (Figure 4B; Figure S6A-6D and data not shown), indicating that the foci formation of RPA and SOSS can arise independently of each other.

It is now apparent that there are at least two sets of heterotrimeric ssDNA-binding complexes, RPA and SOSS, involved in DNA damage response. The observation that the association of SOSS or CtIP/RPA with ssDNA occurs independently of each other raises the possibility that these two different complexes may each be responsible for a part of this DNA damage repair process. To test this possibility, we compared their relative contributions to HR repair through siRNA-mediated deletion of these proteins either individually or in combination. As shown in Figure 4C and Figure S6E, although HR repair efficiency is impaired in the absence of SOSS-A or CTIP, we noticed that co-depletion SOSS-A and CTIP decreased homologous recombination efficiency further than that achieved by SOSS-A or CTIP depletion alone. Consistently, simultaneous ablation of SOSS-A and CTIP by siRNA resulted in a further increase of cellular sensitivity to ionizing radiation (Figure 4D). Together, these results indicate that SOSS and CtIP/RPA likely represent two independent sub-pathways, which act at least in part downstream of MRN complex and function in DNA damage repair (Figure 4E).

In summary, we identified a trimeric complex, which we refer to as SOSS complex in this study. The existence of two independent ssDNA-binding complexes, RPA and SOSS, in mammalian cells underscores the importance of this process in DNA repair. We believe that further study of this second human ssDNA-binding heterotrimeric complex will provide insight in the repair of DNA double-strand breaks, especially the poorly understood homologous recombination repair process.



Rabbit polyclonal anti-SOSS-B1, SOSS-B2 and SOSS-C antibodies were generated by immunizing rabbits with MBP-SOSS-B1, MBP-SOSS-B2 and MBP-SOSS-C recombinant protein expressed and purified from E. Coli respectively. These antibodies were further affinity purified using columns containing corresponding GST-fusion proteins. Antibodies against the myc epitope, γ-H2AX, and RAD51 were previously described (Chen et al., 1998; Huen et al., 2007). The anti-SOSS-A and anti-H3 antibodies were purchased from Bethyl and Millipore, respectively. Anti-CHK1 and GST antibodies were obtained from Santa Cruz. Anti-Flag (M2) and anti-β-actin antibodies were purchased from Sigma. Anti-MRE11, RAD50 and NBS1 antibodies were purchased from Novus Biologicals, Abcam and Calbiochem, respectively.

Cell cultures and Transfection

293T and HeLa Cells were maintained in RPMI supplemented with 10% fetal bovine serum and 1% penicillin and streptomycin. SF9 insect cells were maintained in Grace’s medium supplemented with 10% fetal bovine serum. Cell lines of human origin were maintained in 37°C incubator with 5% CO2, whereas insect cells were maintained at 27°C. U2OS cells with DR-GFP integration were kindly provided by Maria Jasin at Memorial Sloan-Kettering Cancer Center (New York). Cell transfection was performed using Lipofectamine 2000 (Invitrogen) following manufacture’s protocol.


The full-length and deletion/point mutants of human SOSS-A, SOSS-B, SOSS-C and NBS1 were generated by PCR and subcloned into the pDONR201 vector using Gateway Technology (Invitrogen). The corresponding fragments in entry vectors were transferred into a Gateway compatible destination vector, which harbors an N terminal triple-epitope tag (S protein tag, Flag epitope tag and Streptavidin binding peptide tag) or Myc epitope tag for expression in mammalian cells.

Baculoviruses and protein purification from insect cells

DNA fragment containing full-length SOSS-A, SOSS-B1, SOSS-C or NBS1 in pDONR201 vector were transferred to pDEST20, pDEST10, pDEST8 and SFB-tagged vectors for the expression of GST-SOSS-A, His-SOSS-B1, SOSS-C and SFB-fusion proteins in insect cells, respectively. Transposition occurred in DH10Bac competent cells and correct bacmids confirmed by PCR were transfected into SF9 cells for baculovirus production. After viral amplification, SF9 cells were infected with desired baculovirus and cell lysates were collected 48 hours later.

Gel filtration chromatography

Sf9 cells were coinfected with baculovirus stocks expressing GST-SOSS-A, His-SOSS-B1 and untagged SOSS-C. 48 hours later, cells were harvested, washed with 1X PBS and resuspended in 10 ml Lysis Buffer (10% (v/v) glycerol, 20 mM Hepes, pH7.6, 0.3M KCl, 0.01% NP-40, 1mM DTT, 0.2 mM PMSF, 1 μg/mL each of leupeptin, aprotinin and pepstatin). Cells were homogenized with 10 strokes with Dounce homogenizer on ice. The lysate was centrifuged for 15 min at 10,000 rpm. The supernatant was incubated at 4°C with 300 μl of Glutathione Sepharose 4B for 4 hours. The resin was washed with wash buffer (lysis buffer containing 0.5M KCl). Protein was eluted with Elution buffer (lysis buffer containing 20 mM Glutathione). Eluted protein was dialyzed in buffer B (10% (v/v) glycerol, 20 mM Sodium Phosphate, pH7.6, 0.3M KCl, 0.01% NP-40, 1mM DTT) and incubated with 200 μl Ni-NTA beads at 4°C for 4 hours. Ni-NTA beads were washed with wash buffer (buffer B containing 20 mM imidazole) and eluted with elution buffer (buffer B containing 300 mM imidazole). The eluted protein was resolved on Superdex 200 gel filtration column against buffer C (10% (v/v) glycerol, 20 mM Hepes, pH7.6, 0.3M KCl, 0.01% NP-40, 1mM DTT). Indicated fractions were analyzed on 12.5% SDS-PAGE.

Electrophoretic mobility shift assay

Reaction mixtures (20 μl) contained 20 mM Tris-HCl (pH 7.5), 50 mM NaCl, 1 mM MgCl2, 0.5 mM EDTA, 0.2 mM DTT, 100 pmol 32P-labeled ssDNA (d60T) or dsDNA (d60A-T; d60T annealed to its complimentary d60A) were incubated with increasing concentrations of SOSS-B1 (0, 0.5, 1, 2, 4, 6 μM) or SOSS complex (0, 0.5, 1, 2, 4, 6, 8, 10 μM) for 60 min at 37°C. Reaction was terminated by the addition of 2 μl of gel loading dye [0.1 % (w/v) bromophenol blue and 0.1 % (w/w) xylene cyanol in 20 % glycerol] and transferred on ice. Samples were separated by PAGE in an 8 % gel at 10V/cm for 6 hours at 4°C using 45 mM Tris-borate (pH 8.3) and 1 mM EDTA as the running buffer. The gels were visualized by phosphor imaging.

RNA interference

All siRNA duplexes were purchased from Dharmacon Research (Lafayette, CO). The following sequences were used in HeLa Cells: SOSS-A#1: GAUGAGAGUUGCUAUGACAdTdT; SOSS-A#2: CCAAGCGAGCUGUGACGAAdTdT; SOSS-B1: CGACGGAGACCUUUGUGAAdTdT; SOSS-B2: CGUGCAAAGUAGCAGAUAAdTdT; MRE11: GGAGGUACGUCGUUUCAGAdTdT; RAD50: ACAAGGAUCUGGAUAUUUAUU; and NBS1: CCAACUAAAUUGCCAAGUAUU. The following sequences were used in U2OS cells: SOSS-A#1: CGUGAUGGCAUGAAUAUUGdTdT; SOSS-A#2: GUAGUCCACCCUUCUAAUGdTdT; and RAD51: CUAAUCAGGUGGUAGCUCAUU. The siRNA for CtIP was previously described (Yu and Chen, 2004). The siRNAs transfection was performed using Oligofectamine (Invitrogen) following manufacturer’s instruction.

The establishment of stable cell lines and Affinity Purification of S-Flag-SBP(SFB)-tagged protein complexes

293T cells were transfected with plasmids encoding SFB-tagged proteins. Cell lines stably expressing tagged proteins were selected by culturing in the medium containing puromycin (2 μg/ml) and confirmed by immunoblotting and immunostaining. For affinity purification, 293T cells stably expressing tagged proteins were lysed with NETN buffer for twenty minutes. Crude lysates were removed by centrifugation at 14,000 rpm at 4 °C for ten minutes, and pellet was sonicated for 40 seconds in high-salt solution (20 mM HEPES at pH 7.8, 0.4 M NaCl, 1 mM EDTA, 1 mM EGTA, protease inhibitor) to extract chromatin-bound proteins fractions. The supernatants were cleared at 14,000 rpm to remove debris and then incubated with streptavidin-conjugated beads (Amersham) for 2 hours at 4°C. The immunocomplexes were washed three times with NETN buffer and then bead-bound proteins were eluted with NETN buffer containing 1 mg/mL biotin (Sigma). The elutes were incubated with S protein beads (Novagen). The immunocomplexes were again washed three times with NETN buffer and subjected to SDS-PAGE. Protein bands were excised, digested and the peptides were analyzed by mass spectrometry.

Immuofluoresence staining procedure

To visualize IRIF, cells cultured on coverslips were treated with 10 Gy of gamma irradiation (1 Gy = 100 Rads) followed by recovery for six hours. Cells were then fixed using 3% paraformaldehyde solution for ten minutes at room temperature and then extracted with buffer containing 0.5% triton X-100 for five minutes. Samples were blocked with 5% goat serum and incubated with primary antibody for thirty minutes. Samples were washed and incubated with secondary antibody for thirty minutes. Cells were then counter-stained with DAPI to visualize nuclear DNA.

Cell synchronization

HeLa cells were treated with 2 mM thymidine for 19 hours and then released in fresh medium for 9 hours. 2 mM thymidine was added again and cells were incubated for another 16 hours to arrest cells in G1 phase. Cell cycle distributions were confirmed by FACS analysis.

Chromatin fractionation

Preparation of chromatin fractions were described previously with some modifications (Yu et al., 2006). Briefly, cells were collected 2 hours after treatment with 10 Gy of ionizing radiation, and washed once with PBS. Cell pellets were subsequently resuspended in NETN buffer (10 mM HEPES pH7.4, 10 mM KCl, 0.05% NP-40 and protease inhibitors) and incubated on ice for twenty minutes. Crude lysates were removed by centrifugation at 14,000 rpm at 4 °C for ten minutes, and pellet was recovered and resuspended in 0.2 M HCl for twenty minutes. The soluble fraction was then neutralized with 1 M Tris-HCl pH 8.0 for further analysis.

G2/M checkpoint assay

G2/M checkpoint assay was performed as described previously (Lou et al., 2003). Briefly, cells were treated with 2 Gy IR. One hour later, cells were fixed with 70% (v/v) ethanol overnight, and then stained with anti-phospho-histone H3 (Ser10) antibody and propidium iodide. Samples were analyzed by flow cytometry to determine the percentages of cells in mitosis.

Cell survival assays

1×103 cells were seeded onto 60 mm dish in triplicates. Twenty-four hours after seeding, cells were irradiated with IR and then incubated for fourteen days. Resulting colonies were fixed and stained with Coomassie blue. Numbers of colonies were counted using a GelDoc with Quantity One software (BIORAD). Results were the averages of data obtained from three independent experiments.

Gene conversion assay

A U2OS cell clone stably expressing HR reporter DR-GFP was described previously (Weinstock et al., 2006a). 1×106 U2OS-DR-GFP cells were electroporated with 12 μg of pCBASce plasmid at 270V, 975uF using a BioRad genepulsar II. Cells were plated onto 10cm dishes and incubated for 48 hours prior to FACS analyses using a Becton-Dickinson FACScan on a green (FL1) versus orange (FL2) fluorescence plot. Results were the averages of data obtained from three independent experiments.

Supplementary Material



We thank all colleagues in Chen’s laboratory for insightful discussion and technical assistance, especially Dr. Jingsong Yuan and Michael S.Y. Huen. This work was supported in part by grants from the National Institutes of Health to J.C. (CA089239, CA092312 and CA100109). J.C is a recipient of an Era of Hope Scholar award from the Department of Defense and a member of the Mayo Clinic Breast SPORE program.


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