|Home | About | Journals | Submit | Contact Us | Français|
The ataxia telangiectasia-mutated and Rad3-related (ATR) kinase functions as a central node in the DNA damage response signaling network. The mechanisms by which ATR activity is amplified and/or maintained are not understood. Here we demonstrate that BRIT1/microcephalin (MCPH1), a human disease-related protein, is dispensable for the initiation but essential for the amplification of ATR signaling. BRIT1 interacts with and recruits topoisomerase-binding protein 1 (TopBP1), a key activator of ATR signaling, to the sites of DNA damage. Notably, replication stress-induced ataxia telangiectasia-mutated or ATR-dependent BRIT1 phosphorylation at Ser-322 facilitates efficient TopBP1 recruitment. These results reveal a mechanism that ensures the continuation of ATR-initiated DNA damage signaling. Our study uncovers a previously unknown regulatory axis of ATR signaling in maintaining genomic integrity, which may provide mechanistic insights into the perturbation of ATR signaling in human diseases such as neurodevelopmental defects and cancer.
In the S phase of the cell cycle, cells need to duplicate the genome with both high efficiency and high fidelity (1). A variety of endogenous and environmental factors can cause DNA damage in S phase, including oxidative products, lack of sufficient deoxynucleotides, and genotoxins (2). When DNA replication forks encounter DNA lesions (e.g. nicked or broken DNA), the unexpected perturbation of the replication machinery can lead to replication stress, which poses risks of genome destabilization (1). In response to replication stress, cells have evolved an elaborate signaling pathway to maintain genomic integrity in S phase (3, 4). At the apex of the replication stress response is the protein kinase ataxia telangiectasia-mutated (ATM)3 and Rad3-related (ATR) (5, 6). When activated, ATR phosphorylates various protein substrates, including replication factor MCM2, replication protein A (RPA), checkpoint kinases Chk1 and Chk2, and apoptotic regulator p53. Through this cascade of phosphorylation events, ATR regulates DNA replication stability by activating the S phase checkpoint to arrest the cell cycle, promoting DNA repair, inhibiting late replication origin firing, preventing premature mitotic entry, and, when damage is severe, inducing programmed cell death. Therefore, elucidation of the mechanisms by which ATR signaling is regulated is central to our understanding of how genomic integrity is preserved during DNA replication.
Previous studies have revealed several steps in the process of ATR activation (1, 6, 7). In general, ATR, through ATR-interacting protein, is first recruited to DNA lesions by recognizing single-stranded DNA coated with RPA (8, 9). Second, the RAD9-RAD1-HUS1 (9-1-1) complex is loaded onto DNA damage sites by the clamp loader RAD17, which recruits the ATR activator TopBP1 to RPA-coated, single-stranded DNA (10, 11). Third, autophosphorylation of ATR on RPA-coated, single-stranded DNA enables interaction between ATR and TopBP1 to stimulate ATR kinase activity and facilitate recognition of the substrates of ATR (12, 13). However, it remains to be answered how ATR signaling is maintained/amplified when ATR is activated.
A parallel is often drawn between the molecular model of ATR signaling in response to replication stress and the molecular model of ATM signaling in response to DNA double strand breaks (DSBs) (5). Both ATR and ATM belong to the phosphoinositide 3-like protein kinase family. ATR and ATM share many common substrates in response to DNA damage (2, 14). When DSBs occur, ATM and/or ATR phosphorylate the histone H2A variant H2AX, which can spread thousands of base pairs around DSB sites (15, 16). The presence of phosphorylated H2AX (γ-H2AX) provides docking sites to recruit DNA damage-responsive sensors, such as NBS1 and MDC1, through their phosphoprotein-interacting BRCA1 C terminus (BRCT) domains, and recruitment of these sensor proteins further activates or maintains ATM kinase activity and amplifies ATM signaling (17,–19). Therefore, a deficiency of H2AX does not affect the initiation of DSB-induced ATM signaling but does impair maintenance of the DNA damage response (20,–22). Whether a similar positive phosphorylation feedback loop exists for ATR signaling and what molecules might function as ATR signaling amplifiers remains elusive.
In line with the essential role of ATR in maintaining DNA replication stability, loss-of-function mutations in ATR are not compatible with cell viability (23). Reduced ATR function caused by hypomorphic mutations in patients leads to Seckel syndrome, in which microcephaly is a characteristic clinical feature (1). This clinical feature of impaired ATR function is also a clinical feature of deficiency of another DNA damage-responsive protein, BRIT1/microcephalin (MCPH1), the first gene identified as causative of primary recessive autosomal microcephaly (24, 25). This fact led us to investigate whether BRIT1 plays a role in regulating ATR signaling. In the study reported here, we found that BRIT1 functionally interacts with the ATR activator TopBP1 and is required for the continuation of ATR signaling.
U2OS osteosarcoma cells and MCF10A normal breast epithelial cells were purchased from the ATCC. U2OS cells were maintained in McCoy's 5A medium (Cellgro) supplemented with 10% fetal bovine serum. MCF10A cells were maintained in mammary epithelial cell growth medium (Clonetics), a proprietary serum-free medium containing insulin, hydrocortisone, epidermal growth factor, and bovine pituitary extract with 5% horse serum. A lymphoblastoid control cell line and two microcephaly cell lines (MCPH#1 (C74G) and MCPH#2 (G321C)) (27) were grown as a suspension culture in RPMI 1640 medium supplemented with 20% fetal bovine serum. Cells were incubated at 37 °C in a humidified incubator with 5% CO2.
The p3×FLAG-CMV vector encoding full-length BRIT1 has been generated previously in our laboratory. The deletions of BRIT1 were generated from FLAG-BRIT1 plasmids via polymerase chain reaction using primers with restriction sites and subcloned into the N-terminal p3×FLAG-CMV plasmids in-frame. The TopBP1 wild-type and deletion plasmids Δ1-Δ8 and ΔAD were provided by Dr. Junjie Chen (The University of Texas MD Anderson Cancer Center). FLAG-tagged ATM, ATR, ATM-KD (catalytically dead) and ATR-KD plasmids were provided by Dr. M. Kastan (St. Jude Children's Research Hospital), Dr. K. Cimprich (Stanford University), and Dr. L. Zou (Harvard University). Cell culture transfection was performed using Lipofectamine 2000 (Invitrogen), FuGENE6 (Roche), and Oligofectamine (Invitrogen) following the protocols of the manufacturers.
BRIT1 siRNAs, TopBP1 siRNA#1 and #2, and control siRNA have been described previously (26). BRIT1 and TopBP1 transient knockdown was performed with Lipofectamine 2000 or Oligofectamine (Invitrogen) following the protocols of the manufacturers.
Rabbit anti-BRIT1 antibody was generated as described previously. Anti-FLAG M2-agarose affinity gel, anti-FLAG M2, and anti-β-actin were purchased from Sigma-Aldrich. Anti-p53 (DO-1) and anti-p53-HRP were purchased from Santa Cruz Biotechnology. Anti-TopBP1 and anti-phospho-RPA32 were purchased from Bethyl Laboratories. Anti-H2AX was purchased from Calbiochem. Anti-phospho-H2AX was purchased from Millipore. Anti-RPA (Ab-1) was purchased from NeoMarkers. Anti-phospho-p53 (Ser-15), anti-phospho-CHK1 (Ser-345), anti-Ser(P)/Thr(P)) ATM/ATR substrate, anti-ATM, anti-ATR, anti-phospho-histone H3 (Ser-10), and anti-GST were purchased from Cell Signaling Technology. Anti-phospho-MCM (BM28) was purchased from BD Transduction Laboratories. Hydroxyurea (HU) was obtained from Sigma-Aldrich and used at a concentration of 1–2 mm. Nocodazole was obtained from Sigma-Aldrich and used at a concentration of 1 μg/ml.
Cells were washed in PBS, and whole cellular extracts were prepared with urea buffer (8 m urea, 50 mm Tris-HCl (pH 7.4), and 150 mm 2-mercaptoethanol) or modified radioimmune precipitation assay (RIPA) buffer (50 mm Tris-HCl (pH 7.4), 1% Nonidet P-40, 150 mm NaCl, 1 mm EDTA, and 0.25% sodium deoxycholate freshly added to 1 mm PMSF, 1 mm Na3VO4, 1 mm NaF, 1 μg/ml aprotinin, 1 μg/ml leupeptin, and 1 μg/ml pepstatin) for 30 min on ice. Lysates were cleared by centrifugation, and proteins were separated by gel electrophoresis. Membranes were blocked in Tris-buffered saline, 0.1% Tween 20 (TBST) with 5% (w/v) nonfat dry milk for 1 h at room temperature. Membranes were then incubated with primary antibodies diluted in PBS, 5% bovine serum albumin for 2 h at room temperature. Subsequently, membranes were washed with TBST and incubated with horseradish peroxidase secondary antibody (1:5000) (Sigma-Aldrich) diluted in TBST with 5% nonfat dry milk. Membranes were washed in TBST, and bound antibody was detected by enhanced chemiluminescence (GE Healthcare). For coimmunoprecipitation, U2OS cells were transiently cotransfected with HA-BRIT1 wild-type plasmids and FLAG-TopBP1 wild-type/deletion plasmids or with TopBP1 wild-type plasmids and FLAG-BRIT1 wild-type/deletion plasmids. Cellular proteins were extracted in RIPA buffer and immunoprecipitated with anti-FLAG M2 affinity gel (Sigma-Aldrich) overnight. Bead-bound immunocomplexes were eluted with 3× FLAG peptide (Sigma-Aldrich) and subjected to SDS-PAGE. For reciprocal immunoprecipitation, whole cellular extracts were prepared in RIPA buffer and subjected to incubation with antibody overnight, followed by incubation with Protein A/G Plus-agarose beads (Santa Cruz Biotechnology) for 4 h at 4 °C. The precipitates were washed three times with RIPA buffer, eluted in 3× loading buffer by boiling at 95 °C for 5 min, and resolved by SDS-PAGE, followed by immunoblotting. For DNA digestion prior to immunoprecipitation, samples were treated with 10 units/ml DNase I in 1× reaction buffer (Promega) overnight at 4 °C, followed by clarification using centrifugation to remove debris.
Cells were lysed in 10 mm HEPES (pH 7.9), 1.5 mm MgCl2, 10 mm KCl, 0.34 m sucrose, 10% glycerol, 1 mm dithiothreitol, and 0.1% Triton X-100, and nuclear extracts were lysed in 3 mm EDTA, 0.2 mm EGTA, and 1 mm dithiothreitol. Samples were clarified, and pellets were resuspended in SDS sample buffer.
250–500 cells of different treatment groups were seeded in 6-well plates, and 0.001% crystal violet was used to stain the colonies 10–14 days later. The formed colonies were counted and analyzed.
BL21 bacteria containing the indicated plasmids were allowed to grow for 6 h after addition of isopropyl 1-thio-β-d-galactopyranoside. Cell pellets were resuspended in lysis buffer and sonicated. The supernatant was incubated with glutathione-Sepharose 4B beads (GE Healthcare) at 4 °C overnight. After washing, GST fusion proteins were eluted with glutathione. For pulldown, GST and GST-BRCT5 expressed in BL21 were purified using glutathione-Sepharose 4B beads and incubated with 293T lysate containing FLAG-BRIT1. The mixture was incubated at 4 °C with gentle rotation for 1 h, and the beads were washed three times with 1× PBS before addition of SDS sample buffer.
293T cells were transfected with 8 μg of FLAG-tagged ATM, ATR, ATM-KD, and ATR-KD plasmids. Cell extracts were prepared in lysis buffer (50 mm Tris (pH 7.5), 150 mm NaCl, 1% Tween 20, 0.3% Nonidet P-40, 1 mm sodium fluoride, 1 mm Na3VO4, 1 mm phenylmethylsulfonyl fluoride, 50 mm glycerophosphate, 1 mm DTT, 1 mm EGTA, 10% glycerol, and protease inhibitor mixture (Roche Molecular Biochemicals). Cleared supernatants were immunoprecipitated with anti-FLAG M2 antibody (Sigma). After washing with lysis buffer and kinase buffer (20 mm HEPES (pH 7.5), 50 mm NaCl, 10 mm MgCl2, 1 mm dithiothreitol, and 10 mm MnCl2) five times, the immunoprecipitates were resuspended in 50 μl of kinase buffer containing 10 μCi of [-32P]ATP, 10 μm ATP, 1 mm sodium fluoride, 1 mm Na3VO4, 20 mm glycerophosphate, and 1 μg of GST fusion substrate. The kinase reaction was performed at 30 °C for 20 min and stopped by the addition of SDS sample buffer. Proteins were separated on SDS-PAGE gel and transferred to a PVDF membrane. Radiolabeled proteins were visualized by autoradiography.
For HU treatment followed by nocodazole, 2 days after siRNA transfection and rescue transfection, cells were incubated in medium with or without HU for 24 h. Nocodazole was added to the medium when the HU was removed, and cells were harvested 0, 8, and 16 h after release. Harvested cells were fixed in ethanol, stained with propidium iodide, and analyzed by FACS. For HU treatment followed by Taxol, cells were left untreated or treated with HU (2 mm) for 24 h. Cells were then fixed and stained using phospho-histone H3 (Ser-10)-specific antibody (p-H3) and propidium iodide.
Cells cultured on coverslips were washed twice in PBS, incubated in cytoskeleton buffer (PIPES (pH 6.8), 100 mm NaCl, 300 mm sucrose, 3 mm MgCl2, 1 mm EGTA, and 0.5% Triton X-100) for 3 min on ice. The cells were then washed with ice-cold PBS three times and incubated in stripping buffer (10 mm Tris-HCl (pH 7.4), 10 mm NaCl, 3 mm MgCl2, 1% Tween 20, and 0.25% sodium deoxycholate) for 3 min on ice. After another three washes with ice-cold PBS, cells were fixed with 4% paraformaldehyde at 4 °C for 30 min, permeabilized in 1% Triton X-100 and 0.5% Nonidet P-40 for another 30 min, blocked with 1% bovine serum albumin, and incubated with primary antibody for 2 h and secondary antibody (fluorescein isothiocyanate or rhodamine) for 1 h. Cells were then stained with DAPI to visualize nuclear DNA. The coverslips were mounted on glass slides with antifade solution and visualized using a fluorescence microscope.
Immunofluorescent images were captured using a microscope (Nikon, Eclipse E800) equipped with a ×60 oil objective lens (Plan Fluor, numerical aperture 1.3), and a cooled charge-coupled device camera (QImaging, Qiclick F-M-12). Images were acquired using NIS-Elements (Nikon). The fluorochromes used include Alexa Fluor 488 and 594 antibody conjugates (Invitrogen) and DAPI.
Wild-type BRIT1 cDNA cloned in the p3×FLAG-CMV vector was used to generate BRIT1 mutants by using the QuikChange II site-directed mutagenesis kit (Stratagene). Mutant clones were sequenced to confirm the changed nucleotides in BRIT1 cDNA.
A BrdU incorporation assay was performed using a kit from Calbiochem following the protocol of the manufacturer. Briefly, cells were knocked down with BRIT1 siRNA and reconstituted with the indicated BRIT1 constructs resistant to BRIT1 siRNA in U2OS cells. The next day, cells were split and seeded at 104-106 cells/ml into a 96-well plate in a 100-μl volume and exposed to ionizing radiation treatment at the indicated dosages. Then cells were incubated with BrdU (5 mm) for 24 h, fixed, and denatured as described in the manual for the kit. Cells were washed and incubated with anti-BrdU antibody for 1 h and peroxidase goat anti-mouse IgG horseradish peroxidase conjugate for 30 min. Then cells were incubated with the fluorogenic substrate working solution. The horseradish peroxidase catalyzes the conversion of the fluorogenic substrate to a blue fluorescent product. The blue fluorescent product was quantified using a fluorometer.
BRIT1 contains BRCT domains, which are often involved in DNA damage response, with tandem BRCT repeats promoting phosphopeptide binding and direct protein-protein interactions (28). To understand how BRIT1 might regulate protein-protein interactions in response to DNA damage, we conducted a proteomic analysis to systematically identify its binding partners (27). Interestingly, we found that BRIT1 interacted with TopBP1, a major ATR signaling activator. To confirm this observation, we showed that FLAG-BRIT1 could pull down endogenous TopBP1 (Fig. 1A) and that there is a physical interaction between endogenous TopBP1 and BRIT1 (Fig. 1B). Interestingly, the endogenous interaction was elevated after HU treatment, suggesting that replication stress promotes the BRIT1-TopBP1 interaction. We also showed that DNA did not mediate this interaction because BRIT1 was coimmunoprecipitated with TopBP1 even after DNase I treatment (Fig. 1C). To further confirm that the BRIT1-TopBP1 interaction is replication stress-dependent, we used FLAG-BRIT1 to conduct reciprocal immunoprecipitation and found that FLAG-BRIT1 could be coimmunoprecipitated with endogenous TopBP1 and that their interaction was enhanced after exposure to low-dose UV radiation (Fig. 1D). Furthermore, we found that BRIT1 accumulated at DNA lesions and formed nuclear foci in response to UV treatment and that these foci colocalized with TopBP1 foci (Fig. 1E). These results reveal that BRIT1 is a TopBP1-associated protein. Furthermore, these results suggest that BRIT1-TopBP1 interaction might play a functional role in regulating cellular responses to replication stress.
Next we sought to determine whether BRIT1 regulates ATR signaling. In response to replication stress, ATR phosphorylates the RPA32 subunit of RPA, which is recognized as an important upstream signal induced by ATR activation (10, 29). As shown in Fig. 2A, TopBP1 knockdown abolished Ser-33-RPA32 phosphorylation under UV treatment conditions. In contrast, BRIT1 knockdown only modestly reduced Ser-33-RPA32 phosphorylation 1 h after UV or HU treatment (Fig. 2B). Interestingly, when we assessed the kinetics of Ser-33-RPA32 phosphorylation, we observed that BRIT1 depletion with siRNA significantly impaired amplification of RPA phosphorylation after its initial activation (Fig. 2C). Consistent with these observations, we found a significantly decreased number of Ser(P)-33-RPA32-positive cells at later time points after HU treatment in BRIT1-depleted cells (Fig. 2D). Because previous work shows that BRIT1 depletion does not impact the percentage of cells in S phase (27, 30), we infer that the reduction in Ser(P)-33-RPA32 signal at later time points is not due to cell cycle changes resulting from BRIT1 depletion. Together, these data indicate that BRIT1 is not required for initial ATR activation but may be required for establishing a positive feedback loop to amplify ATR signaling.
Because we observed colocalization of TopBP1 and BRIT1 at DNA lesions (Fig. 1E), we asked whether BRIT1 regulates TopBP1 recruitment to sites of replication stress lesions. When TopBP1 was depleted, the formation of BRIT1 foci was not affected (Fig. 3A). In contrast, TopBP1 focus formation induced by HU or UV was reduced significantly in BRIT1-mutant MCPH1#1 cells (Fig. 3, B and C). In line with reduced TopBP1 foci, BRIT1-specific siRNA in U2OS cells led to decreased phospho-MCM2 (Fig. 3D). These results indicate that BRIT1 facilitates the efficient recruitment of TopBP1 to replication stress lesions and promotes the activation of proteins in the ATR pathway.
To mechanistically understand how BRIT1 modulates TopBP1 accumulation at DNA damage sites, we characterized the BRIT1-TopBP1 interaction. We first analyzed the critical regions in BRIT1 that might mediate this interaction. We made a series of deletion mutants of BRIT1 (Fig. 4A). We found that the BRIT1 region from bp 250–500 was required for TopBP1 binding (Fig. 4B). We then generated a deletion mutant specifically lacking this TopBP1-interacting region (BΔ7) (Fig. 4C). As expected, this mutant could not pull down TopBP1 (Fig. 4D), suggesting that the BRIT1-TopBP1 interaction is indeed dependent on this BRIT1 bp 250–500.
In response to DNA damage, ATM and ATR are central kinases regulating the phosphorylation-dependent signaling cascade (5). Therefore, we asked whether BRIT1 is a substrate of ATM/ATR whose phosphorylation might mediate its interaction with the fifth BRCT domain of TopBP1. ATM/ATR substrates share a common S/TQ motif. Therefore, we first tested whether BRIT1 could be pulled down using the Ser(P)/Thr(P) antibody. As shown in Fig. 5A, BRIT1 could be recognized by the Ser(P)/Thr(P) antibody, which was induced markedly by HU treatment. Phosphatase treatment abolished BRIT1 pulldown by the Ser(P)/Thr(P) antibody (Fig. 5A), suggesting phosphorylation-specific recognition of BRIT1 by the Ser(P)/Thr(P) antibody. This result indicates that BRIT is a potential ATM/ATR substrate.
To identify which serine or threonine site on BRIT1 is potentially phosphorylated by ATM/ATR, we analyzed the protein sequence of BRIT1. Interestingly, we found that sequences around BRIT1 Ser-322 are very similar to the sequences around p53 (Ser-15), a known ATM/ATR target site. On the basis of this observation, we used the antibody against p-p53 (Ser-15) to detect BRIT1 phosphorylation. Consistent with Ser(P)/Thr(P) antibody data, BRIT1 could be pulled down by p-p53 (Ser-15) antibody even without DNA damage stimuli, suggesting a basal level of BRIT1 phosphorylation, possibly induced by endogenous DNA damage lesions (Fig. 5B). Nevertheless, BRIT1 phosphorylation was strongly induced by HU treatment and abolished by phosphatase treatment or by the serine-to-alanine mutation on BRIT1 Ser-322 (Fig. 5B). To further confirm that HU induced BRIT1 phosphorylation at Ser-322, we used the anti-Ser(P)/Thr(P) antibody to pull down wild-type and mutant BRIT1. As expected, the S322A mutation abolished replication stress-induced phosphorylation of BRIT1, although a basal level of BRIT1 phosphorylation might be maintained (Fig. 5C). Together, these data show that BRIT1 is potentially phosphorylated by ATM/ATR at Ser-322 in response to replication stress. To further demonstrate that BRIT1 is a substrate of ATM/ATR, we conducted in vitro kinase assays. We cloned 100 amino acids around Ser-322 of BRIT1 into a GST vector. As shown in Fig. 5, D and E, the wild-type BRIT1 fragment could be phosphorylated by ATM or ATR but not by ATM kinase-dead or ATR kinase-dead mutant fragments. The BRIT1 S322A mutation abolished phosphorylation of BRIT1 by ATM (Fig. 5D), whereas the same mutation significantly reduced phosphorylation of BRIT1 by ATR but did not completely block phosphorylation of BRIT1 by ATR (Fig. 5E). This result suggests that multiple sites of BRIT1 might be targeted by ATR, which is consistent with the fact that S322A did not abolish the basal-level phosphorylation of BRIT1 in the absence of DNA damage stimuli. Taken together, our findings indicate that BRIT1 is phosphorylated by ATM/ATR at Ser-322, which suggests that the damage-enhanced interaction between BRIT1 and TopBP1 (Fig. 1, B and C) may be through BRCT-mediated phosphopeptide recognition.
To determine whether BRIT1 phosphorylation and/or BRIT1-TopBP1 interaction is functionally required for accumulation of TopBP1 at replication stress-induced DNA lesions, we depleted BRIT1 and reconstituted in these cells either wild-type BRIT1, the BRIT1 deletion mutant BΔ7 (which lacks the TopBP1-interacting domain), or the BRIT1 phospho-mutant S322A (which blocks BRIT1 phosphorylation induced by replication stress). Our results show that the recruitment of TopBP1 to DNA damage sites induced by HU could be restored by reconstitution of wild-type BRIT1 but not by the BRIT1 mutants (BΔ7 or S322A) (Fig. 6, A and B). As expected, reconstitution of BRIT1-depleted cells with BΔ7 or S322A was unable to restore RPA phosphorylation induced by HU (Fig. 6C). These data further support the idea that BRIT1 regulates ATR signaling and that this regulation is dependent on ATM/ATR-mediated BRIT1 phosphorylation and the interaction of BRIT1 with TopBP1.
Having determined a role for BRIT1 in coordinating TopBP1 recruitment in the ATR pathway, we next tested whether BRIT1 is required for checkpoint control in response to replication stress. As shown in Fig. 7, A and B, depletion of BRIT1 led to increased cellular sensitivity to HU and UV treatment compared with the sensitivity of control cells. Moreover, BRIT1-deficient cells showed radioresistant DNA synthesis when cells were exposed to ionizing radiation (Fig. 7C), indicating that BRIT1 depletion led to a defective intra-S phase checkpoint as a result of impaired ATR signaling. Reconstitution of BRIT1-depleted cells with wild-type BRIT1 could rescue the intra-S phase defect. However, reconstitution of BRIT1-depleted cells with the TopBP1 binding-defective mutant of BRIT1 (BΔ7) was unable to rescue the defect (Fig. 7C). These results support the conclusion that the BRIT1-TopBP1 interaction promotes the ATR signaling-mediated intra-S phase checkpoint. In addition to being essential for intra-S phase checkpoint control, ATR signaling is essential for cells to recover and complete DNA replication after replication stress (7, 31). Therefore, we tested whether BRIT1-dependent regulation of TopBP1 is required for replication stress recovery. We first showed that BRIT1 deficiency did not affect normal S phase progression (Fig. 7D). Next, we compared replication stress recovery between cells expressing WT or mutant BRIT1 where the endogenous BRIT1 was depleted by siRNA transfection. As shown in the propidium iodide staining profile, BRIT1 knockdown significantly delays cell cycle progression after replication stress (Fig. 8, A and B). Furthermore, BrdU labeling shows that BRIT1 knockdown impairs progression through S phase and reduces the number of cells entering G2-M phase (Fig. 8, A and B). The defective restart of replication after replication stress can be rescued by WT BRIT1 but not mutant BRIT1 (Fig. 8, A and B). These data show that BRIT1 is required for proficient recovery from replication stress and that the BRIT1-TopBP1 interaction is required for this function. Collectively, these data demonstrate that the BRIT1-TopBP1 regulatory axis is involved in executing proper ATR signaling.
ATR signaling is essential for the replication stress response. Previous studies have identified detailed mechanisms by which ATR signaling is activated. Here we have shown evidence that BRIT1 interacts with TopBP1 and is phosphorylated by ATM/ATR in response to replication stress and that BRIT1 phosphorylation is required for recruiting TopBP1 to DNA lesions, thereby further enhancing phosphorylation of RPA for proper ATR signaling maintenance and amplification.
Our study reveals that a complex protein-protein interaction network is required for a multistep regulation of ATR signaling in response to replication stress. Our study shows that, following initial phosphorylation of RPA at DNA replication lesions, BRIT1 recruits TopBP1, which is required for amplifying RPA phosphorylation. It is worth noting that BRIT1 knockdown has been reported to reduce TopBP1 expression in HEK293 cells, which was caused by loss of BRIT1-mediated regulation of E2F (32). However, in the cell lines used in our study, we did not observe that BRIT1 knockdown decreased TopBP1 protein expression, whereas Fig. 1, A and D, shows that BRIT1 overexpression did not increase TopBP1 protein expression. Therefore, our data identify BRIT1 as a new regulatory step of this protein-protein interaction network that provides cells with a signal circuit linking two important ATR signaling activation events, namely TopBP1 recruitment and RPA phosphorylation, to maintain ATR signaling.
Posttranslational modifications play a vital role in regulating DNA damage signaling. BRIT1 has been reported to regulate cellular responses to DSBs, including ATM signaling, chromatin remodeling, DNA repair, and G2/M checkpoint activation (26, 27, 33). In this study, we identify a previously unknown ATM/ATR phosphorylation site on BRIT1. This modification may provide a recognition site for the BRCT domain and facilitate additional TopBP1 recruitment when ATR signaling is initiated. It is worth noting that BRIT1 contains three BRCT domains, which can also recognize phosphorylated proteins in the replication stress response. Therefore, it is possible that BRIT1 is recruited to replication stress-associated DNA lesions via ATR-mediated phosphorylation of proteins involved in DNA replication and/or the replication stress response. Indeed, our proteomic analysis of BRIT1-interacting proteins identified multiple DNA replication factors as its binding partners, including MCM2. It is very likely that BRIT1 is recruited to DNA lesions via different mechanisms depending on the form of DNA damage: DSBs or replication lesions. Overall, BRIT1 might provide a positive feedback loop via the interaction between BRCT domains and phosphorylation modifications, which is important for maintaining/amplifying ATR signaling.
In response to replication stress stimuli, it is essential that cells not only activate ATR signaling but also maintain the signaling in a temporal and spatial manner. Inefficient maintenance of ATR signaling could lead to premature deactivation of checkpoint activation and premature entry into mitosis, which is, in fact, the scenario observed in BRIT1-deficient cells of patients with microcephaly and BRIT1 knockout mouse cells (34, 35). The genetic disease microcephaly has been proposed to link closely with ATR signaling defects, and BRIT1 has been found to regulate Chk1 protein expression and activation (26, 34,–36). Consistent with these findings, we also observed reduced expression of Chk1 phosphorylation (Ser-317), suggesting that the role of TopBP1 in the regulation of Rad17/9-1-1 complex-dependent Chk1 phosphorylation is also compromised by BRIT1 knockdown.
Our findings indicate that BRIT1 recruits TopBP1 and participates in ATR signaling maintenance and amplification, which constitutes a major undiscovered mechanism potentially underlying the pathological changes of microcephaly. In addition, aberrations of BRIT1 have been found in several major human cancers, including breast cancer, prostate cancer, and ovarian cancer (33, 37, 38). Recent studies propose that the replication stress response provides an anticancer barrier (39, 40). In this model, the replication stress response induced by excessive growth signals because of oncogene activation or loss of tumor suppressors leads to cellular senescence or apoptosis, which prevents precancerous lesions from progressing to overtly malignant lesions (41, 42). On the basis of our results, we speculate that loss of BRIT1 can lead to defective ATR signaling, which likely results in a breach of this anticancer barrier. In summary, our findings may provide mechanistic insights into the pathological relevance of BRIT1 in human diseases.
We thank S. Deming for proofreading the manuscript; Dr. Gregory Ira and Dr. Shiaw-Yih Lin for scientific discussions and comments on the manuscript. Dr. Junjie Chen for TopBP1 plasmids; and the Flow Cytometry and Cellular Imaging Facility at The University of Texas MD Anderson Cancer Center for FACS analysis.
*This work was supported, in whole or in part, by National Institutes of Health Grant R00CA149186 (to G. P.) and National Institutes of Health Cancer Center Support Grant CA016672 (to The University of Texas MD Anderson Cancer Center).
3The abbreviations used are: