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Translational Science

A20/TNFAIP3 Regulates the DNA Damage Response and Mediates Tumor Cell Resistance to DNA-Damaging Therapy

Chuanzhen Yang, Weicheng Zang, Zefang Tang, Yapeng Ji, Ruidan Xu, Yongfeng Yang, Aiping Luo, Bin Hu, Zemin Zhang, Zhihua Liu and Xiaofeng Zheng
DOI: 10.1158/0008-5472.CAN-17-2143 Published February 2018

Abstract

A competent DNA damage response (DDR) helps prevent cancer, but once cancer has arisen, DDR can blunt the efficacy of chemotherapy and radiotherapy that cause lethal DNA breakage in cancer cells. Thus, blocking DDR may improve the efficacy of these modalities. Here, we report a new DDR mechanism that interfaces with inflammatory signaling and might be blocked to improve anticancer outcomes. Specifically, we report that the ubiquitin-editing enzyme A20/TNFAIP3 binds and inhibits the E3 ubiquitin ligase RNF168, which is responsible for regulating histone H2A turnover critical for proper DNA repair. A20 induced after DNA damage disrupted RNF168–H2A interaction in a manner independent of its enzymatic activity. Furthermore, it inhibited accumulation of RNF168 and downstream repair protein 53BP1 during DNA repair. A20 was also required for disassembly of RNF168 and 53BP1 from damage sites after repair. Conversely, A20 deletion increased the efficiency of error-prone nonhomologous DNA end-joining and decreased error-free DNA homologous recombination, destablizing the genome and increasing sensitivity to DNA damage. In clinical specimens of invasive breast carcinoma, A20 was widely overexpressed, consistent with its candidacy as a therapeutic target. Taken together, our findings suggest that A20 is critical for proper functioning of the DDR in cancer cells and it establishes a new link between this NFκB-regulated ubiquitin-editing enzyme and the DDR pathway.

Significance: This study identifies the ubiquitin-editing enzyme A20 as a key factor in mediating cancer cell resistance to DNA-damaging therapy, with implications for blocking its function to leverage the efficacy of chemotherapy and radiotherapy. Cancer Res; 78(4); 1069–82. ©2017 AACR.

Introduction

Mammalian cells are exposed to various physical and chemical agents that induce DNA damage. A single cell is likely to encounter tens of thousands of DNA lesions per day (1, 2). DNA double strand breaks (DSB) are among the most dangerous types of DNA damage, and unrepaired or incorrectly repaired DSBs lead to genome instability, cancer, and aging (3, 4). To maintain genomic integrity, cells have evolved a set of complex signaling cascades known as the DNA damage response (DDR; ref. 5). In response to DSBs, checkpoint kinase ATM (Ataxia Telangiectasia Mutated) phosphorylates H2AX (also designated as γH2AX) near the damage sites, leading to recruitment and phosphorylation of MDC1 (6, 7). In addition to kinase signaling, ubiquitination also plays an important role in the DDR. E3 ligase RNF8 is recruited by phosphorylated MDC1 and ubiquitinates histone H1 (8). Next, another E3 ligase, RNF168, is recruited to catalyze monoubiquitination of H2A and H2AX at Lys13 and Lys15, which initiates the subsequent formation of a lysine 63–linked polyubiquitin chain. The ubiquitin signaling catalyzed by the RNF8/RNF168 cascade promotes recruitment of downstream repair proteins such as 53BP1 (9, 10). Error-free homologous recombination (HR) and error-prone nonhomologous end-joining (NHEJ) are the major pathways to repair DSBs (1, 3). 53BP1 is a crucial effector that promotes DSB repair through NHEJ (3, 11). NHEJ is important for maintaining genome stability; however, overuse of NHEJ for repair leads to chromosomal translocation and genome instability (12, 13). Therefore, a proper cellular response to DNA damage is crucial for the maintenance of normal cell function. For instance, defective DNA repair results in a human immunodeficiency disorder called RIDDLE (radio sensitivity, immunodeficiency dysmorphic features, and learning difficulties) syndrome and enhanced DNA repair capacity renders cancer cells resistant to radiotherapy and chemotherapy (14, 15). This connection reveals the importance of delicate regulation of the DDR at DSBs.

In addition to E3 ligases, deubiquitinating enzymes (DUB) also participate in the DDR pathway through regulating H2A ubiquitination. The human genome encodes approximately 100 DUBs, which are divided into five families: UCH (ubiquitin C-terminal hydrolases), USP (ubiquitin specific proteases), OTU (ovarian tumor proteases), Josephin, and JAMM (JAB1/MPN/Mov34 metalloenzyme). So far, most of the identified DUBs that antagonize DSB-induced H2A ubiquitination belong to the USP family. USP3 and USP44 abolish accumulation of RNF168 and 53BP1 at DNA damage sites (16, 17). Recently, USP51 was demonstrated to specifically deubiquitinate H2A at Lys13 and Lys15 and fine-tune the DDR (18). USP16 deubiquitinates H2A at Lys 119 and represses gene transcription (19). OTUB1 is the only reported deubiquitinating enzyme in the OTU family that inhibits RNF168-mediated H2A ubiquitination. Independent of its DUB catalytic activity, OTUB1 antagonizes H2A ubiquitination via direct binding to and inhibition of E2 UBC13 (20). Although recent studies have revealed the importance of deubiquitinating enzymes in tightly controlling histone ubiquitination during the DDR, it remains unclear whether other DUBs from the OTU family can regulate DSB-induced H2A ubiquitination and the DDR.

TNFAIP3/A20, a member of the OTU deubiquitinase family, is a primary protein expressed in human venous endothelial cells in response to TNF, IL1, and LPS. Recent studies reveal that A20 is also expressed in other cell types in response to stimuli such as H2O2 and TPA (21). The most well-studied function of A20 is negative regulation of inflammation and immunity (22). In mice, knocking out A20 results in severe inflammation and cachexia, followed by death two weeks after birth (23). Moreover, several studies have identified somatic mutations, deletion, and aberrant expression of the TNFAIP3/A20 gene in various kinds of tumors (24–26). These studies reveal the importance of fully exploring the functions of A20 and its connection with tumors.

Here, we identify A20/TNFAIP3 as a negative regulator of RNF168-mediated ubiquitination of H2A Lys13 and Lys15 (H2AK13, 15ub). We find that NFκB is activated in response to DNA damage and binds to the A20 promoter, leading to upregulation of A20 expression. Subsequently, more A20 binds to chromatin and regulates the DDR. Deletion of A20 increases the persistence of RNF168 and 53BP1 foci at DNA damage sites and genome instability. Importantly, A20 is often upregulated in invasive breast carcinomas, and knockout of A20 increases the sensitivity of cancer cells to radiotherapy and chemotherapy, suggesting that A20 is a potential target for cancer therapy.

Materials and Methods

Antibodies, reagents, and plasmids

The antibodies and reagents used in this study were listed in the Supplementary Methods. OTUD3, OTUD5, and OTUD6B cDNAs were kindly provided by Dr. Lingqiang Zhang at the Beijing Institute of Radiation Medicine. Human wild-type TNFAIP3 (A20) and mutants, H2A-K118,119R mutant, RNF168 and deletion mutants, TAX1BP1, ITCH, and RNF11 were inserted into the 3Flag-pcDNA vector. A20, RNF8, and RNF168 were inserted into the 3Myc-pcDNA vector. A20-1-370 (A20-N) and A20-440-790 (A20-C) were amplified by PCR and cloned into the pET-28a vector. Human RNF168 and deletion mutants RNF168-1-249 and RNF168-249-571 were cloned into the pCMV-3HA vector. RNF168-1-249 was cloned into the pGEX-4T-1 vector. All expression plasmids were verified by DNA sequencing.

Cell culture and transfection

HeLa, MCF7, and U2OS cells were purchased from ATCC and HEK293T was acquired from the National Infrastructure of Cell Line Resource in 2014. The identities of all cell lines were authenticated by short tandem repeat analysis in 2016. Cell lines were tested for mycoplasma contamination by PCR. All cell lines were passaged for fewer than 2 months after resuscitation and were used at the fourth through twelfth passage in culture for this study. The cell lines were cultured in DMEM (Gibco) supplemented with 10% FBS (Gibco). HEK293T cells were transfected with PEI according to the manufacturer's instructions (Polyscience). HeLa cells were transfected with X-tremeGENE HP DNA Transfection Reagent according to the manufacturer's instructions (Sigma).

His-ubiquitin pull-down assay

HEK293T cells were transfected with his-ubiquitin and the indicated plasmids. Cells were harvested 48 hours after transfection. His-ubiquitin pull-down assays were performed following a method described in a previous study (27).

Mononucleosome purification

Mononucleosome purification was performed as described previously (28) with the following modifications: anti-Flag M2 beads binding with mononucleosomes were washed three times with washing buffer (10 mmol/L HEPES-KOH, pH 7.5; 1 mmol/L EDTA; 10 mmol/L KCl; 10% glycerol; protease inhibitors). Mononucleosomes were eluted using 400 μg/mL Flag peptide for 2 hours at 4°C.

Acid chromatin fractionation

Preparation of chromatin fractions was carried out following a procedure described in a previous study (29). Cell pellets were resuspended in NP-40 lysis buffer and incubated at 4°C for 30 minutes and nuclei were collected and resuspended in 0.2 mol/L HCl. The soluble fraction was neutralized with 1 mol/L Tris-HCl (pH 8.0).

IR treatment

IR treatment was performed following procedures described previously (30). After irradiation at 10 Gy, cells were incubated at 37°C for indicated time.

Immunofluorescence microscopy

HeLa cells were transfected with the indicated plasmids using PEI and treated with 10 Gy IR. At 24 hours after transfection, cells were collected and fixed in precooled methanol for 8 minutes at −20°C following a procedure described previously (30). Images were obtained using a confocal microscope (Zeiss LSM-710 NLO and DuoScan) using a 40 × or 63 × oil objective lens. Quantification analysis was performed using Imaris 7.6 software (Bitplane).

RT-PCR and quantitative real-time PCR

MCF7 cells were treated with 40 μmol/L etoposide (VP16) at different time points. Total RNA was extracted using TRIzol Reagent (Invitrogen) and subjected to reverse transcription to synthesize cDNA using the FastQuant RT Kit (TIANGEN). The primers used for the target genes are shown in Supplementary Table S1. Quantitative real-time PCR was performed using Fast Start Essential DNA Green Master (Roche).

Cell fractionation assay

The cell fractionation assay was performed as described previously (30) with modifications. Briefly, cells were lysed in buffer A on ice for 30 minutes. The supernatant was ultracentrifuged and collected as the cytosolic fraction. Cell pellets were washed in buffer A and resuspended in NP-40 lysis buffer as nuclei samples.

Chromatin immunoprecipitation

Chromatin immunoprecipitation (ChIP) was performed following procedures described previously (31) with modifications. MCF7 cells (1 × 107) were treated with DMSO or etoposide (VP16) for 1 hour before cross-linked. The input DNA and DNA from ChIP complexes were purified using the QIAquick PCR Purification Kit, and analyzed by quantitative real-time PCR. The primers used for the A20 promoter are shown in Supplementary Table S1. The detailed method was described in Supplementary information.

Chromatin extraction assay

Cells were lysed in chromatin extraction buffer A (10 mmol/L PIPES, pH 6.8; 100 mmol/L NaCl; 300 mmol/L sucrose; 3 mmol/L MgCl2; 1 mmol/L EGTA; 0.2% Triton X-100) on ice for 30 minutes and centrifuged at 3,000 × g for 5 minutes. The supernatant was removed, and the cell pellets were lysed in chromatin extraction buffer B (3 mmol/L EDTA, 0.2 mmol/L EGTA, 1 mmol/L DTT) and centrifuged at 3,000 × g for 5 minutes. The supernatant was completely removed, and the sediment was resuspended in buffer C (50 mmol/L Tris, pH 8.0; 150 mmol/L NaCl; 1 mmol/L EDTA; 0.1% SDS; 1% Triton X-100) and denatured with 2× SDS loading buffer.

Laser microirradiation

Laser microirradiation was carried out following procedures described previously (32). U2OS cells were grown on thin glass-bottom plates and irradiated with an ultraviolet laser (16 Hz pulse, 41% laser output). Images were taken using a Nikon A1 confocal imaging system every 30 seconds for 10 minutes.

Protein purification and in vitro assays

Recombinant proteins were purified and in vitro assays were performed following previously described procedures (20). For in vitro pull-down assays, GST-fusion proteins were incubated with His-tagged A20 in PBS buffer at 4°C for 1 hour. The beads were washed with PBS buffer and boiled with 2× SDS loading buffer, followed by immunoblotting. For in vitro ubiquitination assays, 0.0125 mmol/L UBE1 (E-305, Boston Biochem), 0.4 mmol/L UBC13/UEV1a (E2-664, Boston Biochem), 40 mmol/L ubiquitin (U-100H, Boston Biochem), 50 mmol/L Tris-HCl (pH 8.0), 5 mmol/L MgCl2, 2 mmol/L ATP, and 1 mmol/L DTT were incubated with recombinant OTUB1 or A20 for 16 hours at 37°C.

Coimmunoprecipitation

Cell lysate preparation, immunoprecipitation, and immunoblotting were performed as described previously (30).

Generation of RNF168−/− and A20−/− HEK293T cells by conventional CRISPR-Cas9 system

To generate a vector expressing pgRNA, two single-guide RNAs (sgRNAs 1–2) targeting different regions in the first exon of the human TNFAIP3/A20 and RNF168 genes were designed (Supplementary Table S2) and cloned into a lentiviral sgRNA vector containing the mCherry selection marker using the Golden Gate method (33). Cells cotransfected with the sgRNA vector and a Cas9 vector were selected by FACS (MOFLO, Cytomation). Single clones were obtained after 10 days of selection. Knockout efficiency was confirmed by immunoblotting. Mutations in the RNF168 and A20 genes were verified by PCR and sequencing.

A20 linear donor construction and knockout HeLa cell selection

The linear donor was constructed following procedures described previously (34) using primers containing sgRNA-targeting regions (Supplementary Table S2) and protection sequences. HeLa cells were transfected with the purified A20 linear donor, A20 pgRNA, and Cas9. Two weeks after transfection, cells were treated with 1 μg/mL puromycin to obtain puromycin-resistant single clones. Knockout efficiency was confirmed by immunoblotting. A20 mutations were verified by PCR and sequencing.

Neutral comet assay

Neutral comet assays were performed using the Trevigen Comet Assay kit (Trevigen). Images were obtained using a fluorescence microscope (Olympus I ×73) with a 10× objective lens. Quantification was performed using Casp Lab software v1.2.2 (University of Wroclaw, Wroclaw, Poland). Approximately 100 cells were analyzed in each group.

NHEJ assay and HR assay

NHEJ assays were performed following a procedure described previously (35). For HR assays, A20WT and A20−/− cells were cotransfected with DR-GFP, an I-SceI expression vector, and a pCherry plasmid. At indicated time after transfection (36 hours for NHEJ assays and 48 hours for HR assays), cells were harvested and washed with 1× PBS. Green (EGFP) and red (Cherry) fluorescence was measured by FACS on an LSRFortessa instrument (BD Biosciences). The percentage of EGFP and pCherry double positive cells versus the percentage of pCherry-positive cells was taken as the repair efficiency. The results are normalized to those of the A20WT cells.

Ethics statement and tissue specimens

The study was approved by the Ethics Committee of the Chinese Academy of Medical Sciences and Peking University's Ethics Committee. Written informed consent was obtained from each individual based on the Declaration of Helsinki. Specimens from 60 breast invasive ductal carcinomas and 23 samples of adjacent normal tissue were analyzed. None of the patients had received radiotherapy or chemotherapy before surgery. Clinical specimens were obtained at the time of surgery. The specimens were immediately fixed in 4% polyformaldehyde and completely embedded in paraffin.

Tissue microarray and IHC

Tissue microarrays (10 mm tissue cores for each tissue) were constructed. IHC staining was carried out following the standard streptavidin–biotin–peroxidase complex method. Tissue microarrays were treated as described previously (36), followed by incubation with primary antibodies (anti-A20) overnight at 4°C in a humid chamber (1:50 dilution). For the negative controls, the primary antibody was replaced by nonimmune serum. After immunostaining, the sections were scanned by a single investigator who was not informed of their clinical characteristics. The value of the integral intensity was measured by Aperio Image Scope software (Aperio).

Statistical analysis

The experiments were repeated at least three times. Statistical analysis was performed using Student t test (two-tailed) or one-way ANOVA. All results are presented as mean ± SEM unless otherwise stated. (*, P < 0.05; **, P < 0.01; ***, P < 0.001).

Additional methods used in this study are described in the Supplementary Material.

Results

A20 inhibits DNA damage–induced H2AK13,15ub and 53BP1 focal accumulation

To identify new negative regulators of H2A ubiquitination upon DNA damage, we explored the effects of various DUBs from the OTU deubiquitinase family on H2A ubiquitination. HEK293T cells expressing his-ubiquitin and the indicated DUBs were treated with etoposide (VP16) to trigger DSBs, after which his-tagged ubiquitinated proteins were enriched through in vivo his-ubiquitin pull-down. Among the DUBs detected, a dramatic decrease in H2A ubiquitination was observed with overexpression of A20 or positive control OTUB1 (Fig. 1A). Histone H2A can be ubiquitinated at Lys118,119 or Lys13,15, and H2AK13,15ub induced by DNA damage triggers the recruitment of downstream repair proteins (10, 18). We therefore tested whether A20 inhibited DNA damage–induced H2AK13,15ub. We first purified H2A K118,119R-containing mononucleosomes using cells expressing a Flag-H2A (K118,119R) mutant, and examined the effect of A20 on H2AK13,15ub. Overexpression of A20 inhibited DNA damage–induced H2AK13,15ub (Fig. 1B). Furthermore, a recently generated H2AK15ub-specific antibody that recognizes monoubiquitinated H2A at Lys 15 was used to assess the inhibitory effect of A20 on H2AK15ub. HEK293T cells were treated with ionizing radiation (IR) and subjected to histone acid extraction. H2AK15ub abundance increased in response to IR, but this effect was significantly inhibited by A20 overexpression (Fig. 1C). Immunofluorescence assays were also performed using IR-treated HeLa cells to confirm this observation. Consistently, A20 suppressed IR-induced H2AK15ub (Fig. 1D). These observations suggest that A20 is a negative regulator of H2AK15ub.

Figure 1.
Figure 1.

A20 inhibits H2AK13,15ub and 53BP1 focal accumulation upon DNA damage. A, HEK293T cells transfected with his-ubiquitin and the indicated DUBs were treated with 40 μmol/L VP16 for 2 hours, after which his-tagged ubiquitinated proteins were enriched by his-ubiquitin pull-down assay and analyzed by immunoblotting using an anti-H2A antibody. WCL, whole-cell lysate. B, HEK293T cells expressing Flag-H2A K118,119R were transfected with Myc-empty vector or Myc-A20. After 48 hours, cells were treated with 40 μmol/L VP16 for 2 hours, after which mononucleosomes were extracted and analyzed by immunoblotting using an anti-Flag antibody. *, nonspecific bands. C, HEK293T cells transfected with Flag-empty vector or Flag-A20 were treated with or without IR (25 Gy). Histones were extracted by acid chromatin fractionation assay after 1 hour of incubation. The level of endogenous H2A ubiquitination at lysine 15 was detected using an anti-H2AK15ub antibody. D and E, HeLa cells transfected with Flag-empty vector or Flag-A20 were treated with or without IR (10 Gy). One hour later, immunofluorescence assays were performed using antibodies against Flag and H2AK15ub (D) or Flag and 53BP1 (E). The percentage of cells with ≥10 H2AK15ub foci or ≥15 53BP1 foci are shown. Data are shown as the mean ± SEM of three independent experiments. Statistical analysis was performed using Student t test (**, P < 0.01; ***, P < 0.001). Scale bar, 10 μm. Approximately 200 cells in each group were counted.

H2A/H2AX ubiquitination is a crucial step in the DDR, and DDR protein 53BP1 is recruited to DNA damage sites by recognizing H2AK15ub (9, 10). As A20 is a negative regulator of H2AK15ub, we detected the effect of A20 on IR-induced 53BP1 accumulation at DNA damage sites by performing immunofluorescence assays. A20 significantly abrogated IR-induced 53BP1 foci (Fig. 1E) to a degree similar to that of positive control OTUB1, whereas CYLD did not (Supplementary Fig. S1A). Moreover, it has been reported that A20 negatively regulates NFκB signaling in a complex with TAX1BP1, ITCH, and RNF11 (37). Thus, we assessed whether these three subunits are also involved in regulation of the DDR. In contrast with A20, TAX1BP1, ITCH, and RNF11 did not affect 53BP1 foci (Supplementary Fig. S1B); moreover, A20 inhibited 53BP1 foci independently of these three proteins (Supplementary Fig. S1C). These data demonstrate that A20 negatively regulates 53BP1 accumulation at DNA damage sites.

NFκB binds to the A20 promoter and upregulates A20 upon induction of the DDR

Previous studies have demonstrated that A20 is transcriptionally upregulated by NFκB (p50/p65 dimer) after treatment with TNF, IL1, LPS, or other stimuli (21). However, little attention has been paid to whether A20 senses DNA damage, especially DNA DSBs. To investigate the capacity of A20 to sense DNA damage, we measured the abundance of A20 mRNA and protein in MCF7 cells treated with VP16 by quantitative real-time PCR and Western blot analyses, respectively. A20 transcription was upregulated after VP16 treatment (Fig. 2A). In contrast, the level of OTUB1 showed no obvious change (Supplementary Fig. S2). In addition, increased abundance of A20 protein and decreased protein abundance of IκBα, an NFκB inhibitor, were observed (Fig. 2B, lane 1–4). To clarify whether DNA damage–induced upregulation of A20 transcription is dependent on NFκB, cells were treated with NFκB inhibitor PDTC before VP16 treatment. In the presence of PDTC, A20 expression was no longer increased, and IκBα was not degraded after VP16 treatment (Fig. 2B). Moreover, both fractionation and immunofluorescence assays showed that VP16 treatment induced translocation of NFκB subunit p65 from the cytosol to the nucleus (Fig. 2C and D), and PDTC inhibited translocation of p65 and upregulation of A20 (Supplementary Fig. S3A). We also found that ATM kinase inhibitor Ku55933 blocked translocation of p65 (Supplementary Fig. S3B), suggesting that NFκB activation upon DNA damage is ATM-dependent. Next, we performed ChIP-qPCR to determine whether NFκB binds to the A20 promoter and induces augmented expression of A20 in response to DSBs. Indeed, VP16 induced binding of p65 to the A20 promoter (Fig. 2E), indicating that p65 upregulates A20 expression at the transcriptional level in response to DSBs. Furthermore, to examine whether A20 associates with chromatin in response to DNA damage, cells were treated with VP16 for the indicated time periods, after which the chromatin fraction was extracted. Consistent with the results shown in Fig. 2B, A20 protein abundance increased in the whole-cell lysates. More importantly, a more obvious time-dependent increase in chromatin-bound A20, but not TAX1BP1, ITCH, or RNF11, was observed after VP16 treatment (Fig. 2F; Supplementary Fig. S1D). Furthermore, we examined the subcellular location of endogenous A20, which revealed that DNA damage promoted expression and nuclear localization of A20 (Fig. 2G). These observations drove us to investigate whether A20 is recruited to DSB sites. We observed moderate accumulation of A20 at DSBs by performing laser microirradiation assays (Fig. 2H). This phenomenon could be a result of the transient presence of A20 at damage sites, which is in accordance with observations regarding USP16 and OTUB2 (38, 39). Taken together, these results suggest that NFκB-induced expression of A20 is involved in the DDR.

Figure 2.
Figure 2.

p65 upregulates A20 expression in response to DNA damage. A, MCF7 cells were treated with 40 μmol/L VP16 for the indicated time periods. The mRNA level of A20 was analyzed by qRT-PCR. Data are shown as the mean ± SEM of three independent experiments. Statistical analysis was performed using Student t test (***, P < 0.001). B, MCF7 cells were treated with DMSO or 300 nmol/L of NFκB inhibitor PDTC before VP16 treatment. Whole-cell lysates were analyzed by immunoblotting using the indicated antibodies. C, MCF7 cells were treated with 40 μmol/L VP16 for the indicated time periods, after which cytoplasmic and nuclear fractions were extracted and analyzed by immunoblotting using the indicated antibodies. D, MCF7 cells were treated with DMSO or 20 μmol/L VP16 for 1 hour, after which immunofluorescence assays were performed using an anti-p65 antibody. Scale bar, 10 μm. E, MCF7 cells were treated with DMSO or 40 μmol/L VP16 for 1 hour, after which ChIP assays were performed to detect the recruitment of p65 to the A20 promoter using an anti-p65 antibody. IgG served as a negative control. The A20 promoter sequences in the input DNA and DNA from ChIP complexes were detected by quantitative PCR. The results were normalized using the IgG abundance of cells treated with DMSO. The results are shown as the mean ± SEM of three experimental replicates. Statistical analysis was performed using Student t test (***, P < 0.001). F, MCF7 cells were treated with 40 μmol/L VP16 for the indicated time periods, after which chromatin was isolated and analyzed using the indicated antibodies. The intensity of A20 was normalized against that of H3. G, MCF7 cells were treated with or without IR (10 Gy) and after 24 hours of incubation, immunofluorescence assays were performed using antibodies against A20. Quantification analysis was performed using Volocity software. Statistical analysis was performed using Student t test (***, P < 0.001). About 100 cells were counted in each group. H, U2OS cells transfected with GFP-A20 were subjected to a laser microirradiation assay. Accumulation of GFP-A20 was detected by fluorescent microscopy at different time points. The red line indicates the positions for laser microirradiation.

A20 affects the DDR independently of its DUB catalytic activity

A20 contains an N-terminal OTU domain and seven zinc finger (ZnF) domains in the C-terminus mediating the interaction between A20 and its substrates/partners (40). A catalytic triad (Asp70, Cys103, and His256) within the OTU domain of A20 is responsible for its deubiquitinating activity, among which Cys103 is a critical residue and forms a region responsible for interacting with other proteins (21). In addition, zinc finger 4 of A20 binds ubiquitin and possesses E3 ligase activity (41). To detect whether the inhibitory effect of A20 on H2A ubiquitination is dependent on its DUB catalytic activity, we generated a series of A20 point mutation constructs, including D70A, C103A, H256A, and the catalytic triad mutant D70A/C103A/H256A (designated as 3A; Fig. 3A). HEK293T cells expressing the Flag-H2A (K118,119R) mutant and A20 point mutants were treated with VP16 and subjected to chromatin extraction. VP16 treatment induced an increase in H2AK13, 15ub, and, interestingly, all of the tested mutants showed a decrease in H2AK13,15ub (Fig. 3B). Moreover, the results of the immunofluorescence assays showed that A20-C103A and A20-3A mutants inhibited 53BP1 foci accumulation as efficiently as did wild-type A20 (Fig. 3C and D).

Figure 3.
Figure 3.

A20 affects the DNA damage response independently of its DUB catalytic activity. A, Schematic of A20 point mutants. B, HEK293T cells expressing the Flag-H2A (K118,119R) mutant were transfected with the indicated A20 mutants. At 48 hours after transfection, cells were treated with 40 μmol/L VP16 for 2 hours and subjected to chromatin extraction. *, nonspecific bands. C, Schematic for the A20 point and deletion mutants. D, HeLa cells were transfected with different A20 mutants. At 24 hours after transfection, cells were irradiated at 10 Gy and incubated for 1 hour. Immunofluorescence assays were performed using the indicated antibodies. Immunofluorescence images and the percentage of cells with ≥15 53BP1 foci are shown. The results are shown as the mean ± SEM of three independent experiments. Statistical analysis was performed using Student t test (***, P < 0.001). Scale bar, 10 μm. Approximately 200 cells in each group were counted. E, The partial coding sequences of human A20 exon 1 and the sequencing results for the mutated alleles of A20−/− clone 1 are shown. Knockout efficiency was determined using an anti-A20 antibody. F and G, A20WT, A20−/− HeLa cells (clone 1), and A20−/− cells, to which the A20 or A20 mutant were reintroduced, were treated with IR (10 Gy). Immunofluorescence assays were performed at the indicated time points after DNA damage (12 hours for H2AK15ub and 24 hours for 53BP1). The percentages of cells with ≥10 H2AK15ub foci or ≥15 53BP1 foci are shown. Data are shown as the mean ± SEM of three independent experiments. Statistical analysis was performed using Student t test (***, P < 0.001). Scale bar, 10 μm. Approximately 200 cells in each group were counted. n.s., nonsignificant.

To explore the defective functionality of mutant A20, we made N-terminal or C-terminal truncation mutants of A20 and tested their effects on H2A ubiquitination. Surprisingly, both the N-terminal and C-terminal were critical for inhibition of H2A ubiquitination by A20 (Supplementary Fig. S4A). Next, we constructed more mutants with altered N-terminal and C-terminal regions (Supplementary Fig. S4B and S4C). The catalytic triad mutant in the N-terminal, A20 N-3A, lost its inhibitory effect on H2A ubiquitination. However, the mutant lacking ZnF4 in the C-terminus did not completely lose its function (Supplementary Fig. S4B and S4C). Assessments using additional constructed mutants showed that, in addition to ZnF4, ZnF5 and ZnF7 were also responsible for H2A deubiquitination (Supplementary Fig. S4D). Moreover, the results in Supplementary Fig. S4E and Fig. 3D show that only A20 mutant 3A-ΔZnF4–7, lacking ZnFs 4–7 and the catalytic triad, revealed complete abolition of the normal inhibitory effect of A20 on H2A ubiquitination and accumulation of 53BP1 foci.

Next, we constructed A20−/− HeLa cells using a recently reported linear donor insertion system (Fig. 3E; ref. 34) and performed immunofluorescence assays to examine the effect of A20 deletion on the DDR. The results showed that A20−/− cells contained more H2AK15ub and 53BP1 foci than did wild-type cells, and reintroducing wild-type A20 inhibited foci accumulation significantly, while A20-deficient mutant 3A-ΔZnF4–7 showed no effect (Fig. 3F and G). Together, these observations indicate that zinc fingers 4–7 and the integrity of the OTU domain, rather than deubiquitinating activity, are crucial for the negative regulatory effect of A20 on the DDR.

A20 does not affect UBC13 stability in response to DNA damage

It has been reported that A20 interacts with UBC13 (E2) and mediates the degradation of UBC13 upon stimulation by TNF, IL1, and LPS (42). As RNF8–UBC13 complex-catalyzed H1 ubiquitination is important for recruitment of RNF168 and activation of downstream repair signaling (8), we assessed whether A20-triggered UBC13 degradation followed VP16 treatment for the indicated time periods. We did not observe degradation of UBC13 associated with the DNA damage–induced increase in A20 protein abundance (Fig. 4A). Next, we treated cells with DMSO or proteasome inhibitor MG132 before VP16 treatment and prolonged the VP16 exposure time to 240 minutes. However, no degradation of UBC13 was observed in cells exposed to VP16 (Fig. 4B). Moreover, to explore the direct effect of A20 on UBC13 stability, A20WT and A20−/− HeLa cells were treated with or without IR (10 Gy), harvested at the indicated times, and subjected to measurement of UBC13 abundance. A20 deletion did not affect the stability of UBC13 following IR treatment (Fig. 4C). Furthermore, to investigate whether A20 inhibits the E2 activity of UBC13 as OTUB1 does, we performed in vitro ubiquitination assays and found that A20 did not affect UBC13-dependent ubiquitination (Fig. 4D; Supplementary Fig. S5A). These results suggest that UBC13 stability is not affected by A20 in response to DNA damage.

Figure 4.
Figure 4.

A20 does not affect UBC13 stability in response to DNA damage. A, MCF7 cells were treated with 40 μmol/L VP16 for the indicated time periods. Whole-cell lysates were analyzed by immunoblotting using the indicated antibodies. B, MCF7 cells were treated with DMSO or 20 μmol/L MG132 before VP16 treatment, after which whole-cell lysates were analyzed by immunoblotting. C,A20WT and A20−/− HeLa cells (clone 1) were treated with or without IR (10 Gy) and harvested at the indicated time. Whole-cell lysates were analyzed by immunoblotting using the indicated antibodies. D,In vitro ubiquitination assays were performed using combinations of UBE1, UBC13/UEV1a, and ubiquitin with recombinant OTUB1 or A20. The reaction mixtures were analyzed by immunoblotting using an anti-ubiquitin antibody.

A20 directly interacts with RNF168 and abrogates RNF168 accumulation at DNA damage sites

As A20 does not affect the stability of UBC13, we next determined whether A20 inhibits H2A ubiquitination by interacting with the E3 ligases responsible for H2A ubiquitination upon DNA damage, with the goal of elucidating the mechanism underlying the effect of A20 on H2A ubiquitination. HEK293T cells were transfected with Flag-A20 and E3 ligases RNF8 or RNF168, and coimmunoprecipitation (co-IP) analysis was performed. A20 interacted only with RNF168, which catalyzes H2A and H2AX monoubiquitination at Lys13 and Lys15 (Fig. 5A). Moreover, we confirmed the endogenous interaction between A20 and RNF168, and found that IR- or VP16-induced DNA damage enhanced this interaction (Fig. 5B), which is dependent on ATM (Supplementary Fig. S3C). We also found that wild-type A20, but not A20-deficient mutant 3A-ΔZnF4–7, interacted with RNF168 (Fig. 5C).

Figure 5.
Figure 5.

A20 directly interacts with RNF168 and abrogates the accumulation of RNF168 at DNA damage sites. A, HEK293T cells transfected with Flag-A20 and the indicated Myc-tagged E3 ligases were subjected to co-IP using an anti-Myc antibody. B, HeLa cells were treated with VP16 (40 μmol/L) for 2 hours or IR (10 Gy), followed by 1 hour of incubation. Co-IP assays were then performed to examine the endogenous interaction between A20 and RNF168. C and D, HEK293T cells transfected with different plasmids were subjected to co-IP assays with the indicated antibodies. E, HEK293T cells transfected with Flag-RNF168 wild-type or mutants were treated with VP16 (40 μmol/L) for 2 hours before they were subjected to a co-IP assay using an anti-Flag antibody. F, The N-terminus (1–370 aa) and C-terminus (440–790 aa) of His-A20 and GST-RNF168 (1–249 aa) were purified from E. coli. In vitro pull-down analysis was performed with GST protein as a negative control. G,A20WT and A20−/− HEK293T cells were transfected with the indicated plasmids. At 48 hours after transfection, cells were treated with VP16 for 2 hours, followed by co-IP assays with the indicated antibodies. H, HEK293T cells transfected with HA-RNF168 (1–249 aa) with or without Myc-A20 were treated with VP16 for 2 hours and then subjected to a co-IP assay. I, HeLa cells transfected with the indicated plasmids were treated with or without IR (10 Gy) and subjected to immunofluorescence assays after 1 hour of incubation. Immunofluorescence images and the percentage of cells with ≥10 RNF168 foci are shown. The results are shown as the mean ± SEM of three experimental replicates. Statistical analysis was performed using Student t test (***, P < 0.001). Scale bar, 10 μm. Approximately 200 cells in each group were counted. J, HeLa cells transfected with the indicated plasmids were irradiated at 10 Gy. Immunofluorescence assays were performed using antibodies against Flag/MDC1 or Flag/RNF8 after 1-hour incubation. The percentage of cells with ≥10 MDC1 and RNF8 foci are shown. Data are shown as the mean ± SEM of three independent experiments. Statistical analysis was performed using Student t test. Scale bar, 10 μm. Approximately 200 cells in each group were counted. n.s., nonsignificant.

To map the critical domain of RNF168 responsible for its interaction with A20, we constructed truncations, including RNF168 1–249 and RNF168 249–571, and assessed their interactions with Flag-A20. The N-terminus, rather than the C-terminus, of RNF168 was essential for its binding to A20 (Fig. 5D). Furthermore, the co-IP results showed that both the RING and the UDM1 domain at the N-terminus of RNF168 are important for its interaction with A20 (Fig. 5E; Supplementary Fig. S6A). Moreover, the results of in vitro pull-down assays using purified proteins (Supplementary Fig. S5B) demonstrated that RNF168 is a direct target of A20, while both the N-terminus and C-terminus of A20 bind to RNF168 (Fig. 5F).

The RING domain of RNF168 is important for recognizing its target H2A (Supplementary Fig. S6B and S6C; ref. 43), while the UDM1 domain is responsible for identification of ubiquitinated H1 (8). As both the RING and UDM1 domains of RNF168 are essential for its binding to A20 (Fig. 5E), we speculated that A20 might disrupt the binding of RNF168 to H2A and ubiquitinated H1. Indeed, co-IP assays showed that A20 deletion promoted the RNF168–H2A interaction, which was inhibited by reexpression of wild-type A20, but not by reexpression of the A20-deficient mutant (Fig. 5G; Supplementary Fig. S7A). In addition, A20 disrupted the interaction between RNF168 and ubiquitinated H1 (Fig. 5H), but it did not affect H1 ubiquitination (Supplementary Fig. S8). Moreover, we determined whether A20 abrogated the accumulation of RNF168 at DNA damage sites. Immunofluorescence assays showed that A20 significantly reduced the number of IR-induced RNF168 foci (Fig. 5I). To exclude the possibility that A20 might affect upstream regulators of RNF168, we examined the effect of A20 on MDC1 and RNF8 foci by immunofluorescence assays. The results showed that A20 did not affect MDC1 and RNF8 foci (Fig. 5J; Supplementary Fig. S9). Overall, these results suggest that A20 regulates the DDR by directly binding to RNF168 and impeding accumulation of RNF168 at damage sites.

Deletion of A20 results in persistent accumulation of DNA damage foci

To further elucidate the effect of A20 on the dynamic regulation of DNA damage foci, the abundance of RNF168 and 53BP1 foci in A20WT and A20−/− HeLa cells was assessed at different time points following IR treatment. Knockout of A20 increased the abundance of chromatin-bound RNF168 and 53BP1 under normal conditions (Fig. 6A and B). The numbers of RNF168 and 53BP1 foci were increased significantly in wild-type and A20−/− cells 1 hour after IR treatment, and A20−/− cells contained slightly more RNF168 and 53BP1 foci than did wild-type cells (Fig. 6A and B). This phenomenon is in accordance with previous reports that DDR proteins are recruited dramatically during the early phase of DNA damage to repair DNA lesions (18, 38). Consistently, the abundance of A20 was slightly increased at 1 hour after IR treatment (Fig. 4C), suggesting that A20 moderately regulates IR-induced DNA damage foci during the early period of DNA repair (1 hour after IR). Strikingly, at 24 hours after IR treatment, A20−/− cells contained more RNF168 and 53BP1 foci than did wild-type cells (Fig. 6A and B). In accordance with this observation, after DNA lesions were repaired (24 hours after IR treatment), the abundance of A20 protein increased significantly (Fig. 4C), and knockout of A20 resulted in RNF168 and 53BP1 retention at damage sites (Fig. 6A and B). Taken together, these results indicate that A20 fine-tunes DNA damage–induced foci during the early phase of the DDR and is essential for the disassembly of 53BP1 at DNA damage sites after repair.

Figure 6.
Figure 6.

Deletion of A20 results in persistent accumulation of DNA damage foci. A and B,A20WT and A20−/− HeLa cells were treated with or without IR (10 Gy). Immunofluorescence assays were performed at the indicated time points after DNA damage. Immunofluorescence images and the percentages of cells with ≥10 RNF168 foci or ≥15 53BP1 foci from three independent experiments are shown. Data are shown as mean ± SEM. Statistical analysis was performed using Student t test (*, P < 0.05; **, P < 0.01; ***, P < 0.001). Approximately 400 cells in each group were counted. C and D, The partial sequences of human RNF168 exon 1 and the sequencing results for the mutated alleles are shown. Knockout efficiency was assessed using an anti-RNF168 antibody. E,RNF168WT and RNF168−/− HEK293T cells transfected with or without Flag-A20 were treated with VP16 for 2 hours and then subjected to an acid chromatin fractionation assay. The level of endogenous H2A ubiquitination was detected using an anti-H2A antibody. F,RNF168WT and RNF168−/− HEK293T cells transfected with or without Flag-A20 were irradiated at 10 Gy and subjected to a chromatin extraction assay.

Moreover, to investigate whether the inhibitory effect of A20 on DDR is dependent on RNF168, we generated RNF168−/− 293T cells using a CRISPR-Cas9 system (Fig. 6C and D; ref. 33) and compared the level of H2A ubiquitination upon DNA damage in RNF168WT and RNF168−/− cells with or without A20. The results in RNF168−/− cells showed that H2A ubiquitination decreased obviously, while A20 was unable to inhibit H2A ubiquitination (Fig. 6E). In addition, RNF168WT and RNF168−/− 293T cells transfected with or without Flag-A20 were irradiated at 10 Gy and subjected to chromatin separation. Deletion of RNF168 impeded recruitment of 53BP1 to chromatin, which was consistent with a published report (10); moreover, A20 no longer functioned in RNF168−/− cells (Fig. 6F). These results suggest that A20 relies upon RNF168 to finely regulate the DDR.

A20 knockout cells exhibit increased sensitivity to ionizing radiation and DNA-damaging agents

Next, we determined whether A20 affects HR and NHEJ efficiency by performing reporter assays. We found that A20 deletion resulted in decreased HR efficiency and increased NHEJ efficiency (Fig. 7A). NHEJ is an error-prone repair mechanism with a tendency to produce chromosome translocation, leading to genome instability (15). Thus, we performed neutral comet assays to investigate the effect of A20 on genome stability, which showed that A20 deletion slightly increased comet tail length under normal conditions. Moreover, A20−/− cells possessed more obvious comet tails than did wild-type cells at 24 hours after IR treatment (Fig. 7B), suggesting that the loss of A20 resulted in impaired DNA repair kinetics and increased genome instability. These results suggest that A20 plays an important role in guaranteeing proper DNA repair.

Figure 7.
Figure 7.

Loss of A20 expression sensitizes cancer cells to ionizing radiation. A,A20WT and A20−/− cells were subjected to HR and NHEJ assays. The experiments were performed three times. The results were normalized to those of the A20 wild-type cells. Data are shown as mean ± SEM. Statistical analysis was performed using Student t test (*, P < 0.05). B,A20WT and A20−/− HeLa cells were subjected to the neutral comet assay. About 100 cells were counted in each group. Images and quantified data are shown. Statistical analysis was performed using Student t test (*, P < 0.05; ***, P < 0.001). n.s., nonsignificant. C–E, Cells were treated with the indicated IR doses and subjected to clonogenic survival assays. Endogenously and exogenously expressed A20 were confirmed by immunoblotting using an anti-A20 antibody. Statistical analysis was performed using Student t test (**, P < 0.01; ***, P < 0.001). F, A model for the role of A20 in the DDR. In response to DNA damage, A20 is upregulated by NFκB and binds to chromatin, where it terminates H2A ubiquitination by disrupting the binding of RNF168 to H2A and ubiquitinated H1, thereby reducing the accumulation of RNF168 and facilitating disassembly of 53BP1 at DNA damage sites.

In addition, to explore the effect of A20 on cell survival after DNA damage, a clonogenic survival analysis was performed. A20WT and A20−/− cells (Fig. 3E; Supplementary Fig. S7B) were treated with different doses of IR and allowed to grow for 12 days, after which the number of colonies was counted. A20−/− cells were more sensitive to IR treatment than were wild-type cells (Fig. 7C). In addition, expression of wild-type A20 but not A20-deficent mutant in A20−/− cells rescued cell viability (Fig. 7D). Similarly, in response to VP16 treatment, A20−/− cells also showed reduced survival in comparison with wild-type cells (Supplementary Fig. S10A and S10B). Moreover, overexpression of A20 rendered cancer cells resistant to IR (Fig. 7E) and VP16 treatment (Supplementary Fig. S10C), suggesting that increased A20 conferred resistance to DNA damage therapy in cancer cells. These results suggest that A20 is required for cell survival following DNA damage.

A20 is highly expressed in breast carcinoma

Radiotherapy and chemotherapy resistance are obstacles of cancer treatments. As increased A20 could confer resistance to IR- and DNA-damaging agents, we wanted to understand the biological relevance of A20 in cancer and to explore whether it can serve as a therapeutic target. First, we analyzed A20 expression in specimens from 60 breast invasive ductal carcinomas and 23 samples of adjacent normal tissue by performing tissue microarrays and IHC. We found that A20 expression was significantly higher in tumor tissue in comparison with its expression level in adjacent normal breast tissue (Supplementary Fig. S11A and S11B). Moreover, we used a gene expression dataset (GSE70905) from the NCBI-GEO for differential expression analysis between the tumor group and adjacent normal group, which showed that A20 was highly expressed in tumor tissue in comparison with its expression level in adjacent normal tissue (Supplementary Fig. S11C). Analysis of dataset GSE65194 from the NCBI-GEO confirmed this conclusion (Supplementary Fig. S11D). A20 expression in breast cancer patients with different histologic grades was also analyzed using the breast cancer dataset from the work by Calda and colleagues (44, 45). The results of this analysis showed that A20 expression was increased significantly in high-grade breast cancers relative to that of the low-grade group (Supplementary Fig. S11E). These results demonstrate that A20 is highly expressed in breast carcinomas, particularly in patients afflicted with high-grade tumors.

Discussion

This study reveals that chromatin-bound A20 plays an important role in connecting NFκB signaling and the DDR. Specifically, we show that, in response to DSBs, NFκB translocates from the cytosol to the nucleus and binds to the A20 promoter. Once A20 is transcriptionally upregulated, more A20 binds to chromatin, where it directly binds to RNF168 and disrupts the binding of RNF168 to H2A and ubiquitinated H1, thereby inhibiting accumulation of RNF168 at DNA damage sites. Thus, A20 inhibits RNF168-mediated H2AK13,15ub and impairs accumulation of downstream repair proteins at DNA damage sites (Fig. 7F).

Previous studies have shown that A20 exerts an anti-inflammatory effect by downregulating NFκB signaling (22). In addition, A20 is an NFκB-responsive gene upon various types of stimulation (21). The role of A20 in the cytoplasm is well characterized, but little is known about its function in the nucleus. Although DNA damage can trigger NFκB activation, the consequences of NFκB activation for the DDR have not been elucidated. Here, we demonstrated that A20 expression is induced by NFκB when DSBs occur. Moreover, we showed that, in the nucleus, A20 inhibits RNF168-mediated H2AK13,15ub and accumulation of repair protein 53BP1 at DNA damage sites. This study reveals a connection between NFκB signaling and the DDR.

It has been shown that OTUB1 inhibits RNF168-dependent H2A ubiquitination and suppresses the DDR. OTUB1 binds to and inhibits E2 UBC13 independently of its DUB catalytic activity (20). Interestingly, here we found that A20 negatively regulated H2A ubiquitination in a DUB activity–independent manner (Fig. 3B). The noncatalytic role of DUBs such as Ubp6 has also been observed (46). In addition, it has been shown that, together with TAX1BP1, A20 interacts with UBC13 and triggers ubiquitin–proteasome degradation in response to TNF, IL1, and LPS stimulation (42). Distinct from this mechanism, here we found that A20 does not affect UBC13 stability in response to DNA DSBs (Fig. 4). Instead, we showed that A20 affects the DDR through direct interaction with RNF168 and attenuates the accumulation of RNF168 at DNA damage sites, which is independent of TAX1BP1, ITCH, or RNF11. These studies suggest that A20 utilizes different mechanisms to function in different pathways.

Moreover, we demonstrated that the integrity of the A20 OTU domain and ZnFs 4–7, rather than its deubiquitinating activity, are important for its effect on the DDR. This observation is similar to previous studies, in which A20 has been shown to bind target proteins either through the conserved surface patch formed by its catalytic triad (21) or via its seven zinc finger domains (40). In addition to directly removing the ubiquitin chain from its target, A20 also affects ubiquitination in an indirect manner. For example, the OTU and ZnF4 domains of A20 are important for its inhibitory effect on E3 ligase activity, which is accomplished by blocking the interaction between E2 and E3 enzymes (42). Other studies also demonstrate that the OTU domain collaborates with ZnF4 and ZnF7 at the C-terminus of A20 to negatively regulate NFκB activation (47, 48). Therefore, further research should focus on studying the structure of full length A20 to explain the coordination between its OTU domain and zinc finger domains.

Recently, several studies have shown that a number of DUBs in the USP family may target H2AK15ub and thus affect the DDR, but these DUBs may function in different stages of the DDR. For instance, USP51 directly targets ubiquitinated H2A on K13 and K15 and modulates the DDR (18). USP3 and USP44 affect recruitment of RNF168 at DNA damage sites and inhibit DNA damage–induced H2A ubiquitination (16, 17). In this study, we demonstrated that A20 inhibited H2AK13,15ub and regulated the DDR. Independently of its deubiquitinase activity, A20 directly interacts with RNF168 and disrupts the binding of RNF168 to H2A and ubiquitinated H1. Unlike other deubiquitinating enzymes, the abundance of A20 is regulated by NFκB in response to DNA damage. A20 is significantly upregulated at 12 and 24 hours after IR treatment (Fig. 4C), suggesting that it mainly functions at the late stage of DNA repair. Therefore, we conclude that A20 is essential for the disassembly of RNF168 foci after DNA repair to avoid hyperaccumulation of RNF168 at DNA damage sites and prevent excessive ubiquitination. Loss of A20 leads to increased NHEJ activity and decreased HR, which may be a result of persistent 53BP1 accumulation at DSB sites and disruption of the balance among DNA repair pathways, thereby contributing to impaired DNA repair kinetics and increased sensitivity of cancer cells to DNA damage. These observations reveal the importance of an appropriate DNA damage response and repair process.

Interestingly, we also found that A20 deletion resulted in persistent BRCA1 accumulation at DNA damage sites (Supplementary Fig. S12). However, A20 deletion decreased HR efficiency. Although these observations may seem contradictory, they can be explained by results from other studies. RAP80 has been shown to recognize RNF168-generated K63–ubiquitin chains and recruits the BRCA1-A complex (2, 49). A model has been proposed in which BRCA1 functions together with RAP80 in the BRCA1-A complex to reduce HR by restricting DSB end processing, while it promotes resection when interacting with other complexes (50, 51). Moreover, other groups have reported that accumulation of 53BP1 at DSBs promotes NHEJ while suppressing HR (39), and RNF168 inhibits HR similar to 53BP1 (52). Accordingly, the results in this study suggest that A20 may affect the DNA repair pathway choice by modulating DNA end resection.

A20 has been reported to be a crucial regulator in many types of cancer. It plays an oncogenic role in some solid tumors including gliomas (26), hepatocellular carcinoma (53), poorly differentiated head and neck cancers, and undifferentiated nasopharyngeal carcinoma (54). Meanwhile, other studies have revealed that A20 is also a tumor suppressor that is frequently deleted and inactivated in B-cell lymphoma, Hodgkin lymphomas, and non-Hodgkin lymphomas (24). These findings suggest that the function of A20 in tumors is cell-type–dependent. In our study, we found that the abundance of A20 protein is upregulated in invasive breast carcinomas (Supplementary Fig. S11A and S11B), which is in accordance with a previous study showing that the mRNA level of A20 is higher in more aggressive breast cancers (25), as well as another recently published article (55). Moreover, A20−/− cells are more sensitive to IR and VP16 treatment than are wild-type cells, whereas wild-type cells with A20 overexpression show resistance to IR and VP16 (Fig. 7C–E; Supplementary Fig. S10). These findings indicate that A20 influences chemotherapy and radiation resistance, suggesting the potential of A20 as a target in breast cancer treatment.

In summary, our finding that A20 functions in the nucleus as an inhibitor of DNA damage-induced H2A ubiquitination provides new insights into the connection between NFκB signaling and the DDR. Our results indicate that A20 regulates the DDR by inhibiting the binding of RNF168 to H2A and ubiquitinated H1, thereby playing an important role in guaranteeing proper DNA repair and maintaining genome stability. A20 might be a promising clinical target for new strategies to prevent resistance to conventional radiotherapy and chemotherapy in breast cancer.

Disclosure of Potential Conflicts of Interest

No potential conflicts of interest were disclosed.

Authors' Contributions

Conception and design: C. Yang, X. Zheng

Development of methodology: C. Yang, W. Zang, Y. Ji, Z. Liu

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): C. Yang, W. Zang, Z. Tang, Y. Ji, R. Xu, Y. Yang, A. Luo, B. Hu, Z. Liu

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): C. Yang, W. Zang, Z. Tang, Z. Zhang, Z. Liu, X. Zheng

Writing, review, and/or revision of the manuscript: C. Yang, Z. Zhang, X. Zheng

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): C. Yang, X. Zheng

Study supervision: X. Zheng

Acknowledgments

This work was supported by the National Science Foundation of China (81730080, 31470754, 31670786) and the National Key Research and Development Program of China (2016YFC1302401). We sincerely thank Prof. Lingqiang Zhang for providing DUB plasmids, Prof. Wensheng Wei for providing CRISPR/Cas9-related plasmids, Dr. Qinzhi Xu for assistance with neutral comet assays, Prof. Huadong Pei for providing the HR and NHEJ systems, and Prof. Xingzhi Xu for helping with the laser microirradiation assays. We also appreciate the assistance of Xiaochen Li, Guopeng Wang, Liying Du, and Hongxia Lv from the Core Facilities of Life Sciences at Peking University for their assistance with microscopic imaging and cell flow cytometry.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Footnotes

  • Note: Supplementary data for this article are available at Cancer Research Online (http://cancerres.aacrjournals.org/).

  • Received July 18, 2017.
  • Revision received November 2, 2017.
  • Accepted December 1, 2017.

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A20/TNFAIP3 Regulates the DNA Damage Response and Mediates Tumor Cell Resistance to DNA-Damaging Therapy
Chuanzhen Yang, Weicheng Zang, Zefang Tang, Yapeng Ji, Ruidan Xu, Yongfeng Yang, Aiping Luo, Bin Hu, Zemin Zhang, Zhihua Liu and Xiaofeng Zheng
Cancer Res February 15 2018 (78) (4) 1069-1082; DOI: 10.1158/0008-5472.CAN-17-2143
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