Nucleotide Excision Repair Factor XPC Enhances DNA Damage–Induced Apoptosis by Downregulating the Antiapoptotic Short Isoform of Caspase-2

DNA lesions induced by UV light and the chemotherapeutic agent cisplatin are mainly removed by the nucleotide excision repair (NER) consisting of 2 subpathways termed transcription-coupled repair (TCR) and global genomic repair (GGR; ref. 1). TCR is required for the preferential removal of lesions from the transcribed strand of active genes, whereas GGR is responsible for the removal of DNA lesions from the bulk of the genome (1). Cell lines derived from individuals suffering from the disorders like xeroderma pigmentosum (XP) and Cockayne syndrome (CS) have defects in one or both of these NER subpathways. Specifically, XP-C cells have a defect in GGR but not TCR. In contrast, CS-A and CS-B cells are defective in TCR but not GGR (2), whereas XP-A cells have defects in both TCR and GGR (1).

TCR-deficient fibroblasts (e.g., CS-A and XP-A) are extremely sensitive to UV- and cisplatin-induced apoptosis (3–5), whereas XP-C cells with selective defect in GGR but with proficient TCR have normal resistance to cisplatin (6, 7). In addition, murine Xpc^−/− keratinocytes are characterized by a complete absence of apoptosis after UV exposure, in contrast to Xpa^−/− and Csb^−/− cells (8). These reports suggest that unrepaired DNA damage in the transcribed strand is an indispensable trigger for UV/cisplatin-induced apoptosis. However, increasing evidence has suggested that unrepaired DNA lesions in the nontranscribed strands or transcriptionally inactive genes can cause replication blockage, which also acts as an apoptosis-inducing signal (9). It has been reported that Xpa^−/− keratinocytes (TCR and GGR deficiency) showed greater apoptosis than Csb^−/− cells (TCR only deficiency) following UV irradiation (8). Given that both cells have deficient TCR, the additional GGR deficiency in Xpa^−/− cells must be a contributing factor in UV-induced apoptosis. Although transcription blockage is absent in XP-C cells due to proficient TCR, the replication blockage induced by deficient GGR is sufficient to trigger apoptosis in these cells. Therefore, the resistance of XP-C cells to UV/cisplatin-induced apoptosis could not be simply attributed to the proficient TCR. This prompts us to hypothesize that an important apoptosis element must be lost in XP-C cells.

XPC is a 940-amino acid protein and harbors domains that can bind to damaged DNA and repair factors (10–12). The major function of XPC is to recognize helix-distorting lesions located in a transcriptionally inactive genome or a nontranscribed strand of actively transcribed genes (13). Recent studies have suggested that beyond its role in DNA repair, the XPC protein is also involved in transcriptional process including both transcription activation and repression (14, 15). However, the relationship between XPC- and apoptosis-related gene transcription is yet to be established.

Caspases are a family of cysteine aspartate–specific proteases involved in the initiation or execution of apoptosis. Caspase-2 (casp-2) is the most conserved caspase across species and is one of the initiator caspases activated by various stimuli (reviewed in ref. 16). The casp-2 gene produces several alternative splicing isoforms. The inclusion of exon 9 leads to the inclusion of an in-frame stop codon in casp-2 short isoform (casp-2S) mRNA, thus producing a truncated protein that inhibits cell death, whereas the exclusion of exon 9 results in casp-2 long isoform (casp-2L) mRNA, whose protein product induces cell death (17, 18).

In this study, we have uncovered a novel function of XPC as a potent enhancer of apoptosis in the absence of any influence by p53. From a mechanistic standpoint, XPC protein downregulates antiapoptotic casp-2S through inhibition of its promoter activity and thus promotes DNA damage–induced activation of casp-9 and casp-6, which ultimately enhances cellular death.

SV40-transformed XP-C (GM15983) and fully corrected XP-C (GM16248) cells were purchased from NIGMS Human Genetic Cell Repository (Coriell Institute for Medical Research). HCT116(p53^−/−) and HCT116(p53^+/+) colorectal carcinoma cells were kindly provided by Dr. B. Vogelstein (Johns Hopkins University). 041-TR cells were provided by Dr. G. Stark (Cleveland Clinic Foundation). A2780-CP70 ovarian cancer cell line was provided by Dr. P. Modrich (Duke University), SKOV3 ovarian cancer cell line was provided by Dr. T. C. Hamilton (Fox Chase Cancer Center), H1299 and A549 cell lines were kindly provided by Dr. W. Duan (The Ohio State University). Except for the regular testing for mycoplasma contamination using a PCR-based assay, the cell lines were not specifically authenticated. The cultures were maintained as described in Supplementary Data, and the cells were treated with cisplatin or UV as described before (19).

pXPC3 plasmid containing XPC cDNA and its corresponding empty vector pEBS7 were kindly provided by Dr. R. Legerski (The University of Texas MD Anderson Cancer Center; ref. 20). XPC shRNA (short hairpin RNA)-containing plasmids (pLKO.1-puro) were purchased from Sigma. p3XFLAG-CMV-14-casp-2S construct was generated in our laboratory (Supplementary Data). Casp-2S promoter-luciferase construct Del4 has been described (21, 22). The plasmids were transfected into cells by either FuGENE6 (Roche Applied Science) or Lipofectamine LTX reagents (Invitrogen) according to manufacturer's instructions.

Whole-cell lysates were prepared by boiling cell pellets for 10 minutes in lysis buffer [2% SDS, 10% glycerol, 62 mmol/L Tris-HCl, at pH 6.8, and a complete mini-protease inhibitor cocktail (Roche Applied Science)]. For the detection of cytochrome c (cyto c) release, cells were harvested by trypsinization, and the cytosol was separated as described (23). Equal amounts of proteins were loaded, separated on a polyacrylamide gel, and transferred to a nitrocellulose membrane. Protein bands were immunodetected with appropriate antibodies (Supplementary Data).

Phosphatidylserine exposure on the outer leaflet of the plasma membrane was detected by the Annexin V–GFP apoptosis detection Kit II (BD Pharmingen) according to the manufacturer's instructions. Briefly, 1 × 10⁶ cells were pelleted following treatment and washed in PBS. Cells were then resuspended in 500 μL of binding buffer, mixed with Annexin V–GFP and propidium iodide and incubated at room temperature (22°C) for 5 to 10 minutes in the dark. The Annexin V–positive cells were analyzed by flow cytometry.

XP-C and XP-C+XPC cells growing on the coverslips were either treated with UV or untreated. The cells were further cultured for 24 hours and stained with MitoCapture reagents (BioVision) according to the manufacture's instruction. Cells were observed immediately under a Nikon Fluorescence Microscope E80i (Nikon) fitted with appropriate filters for FITC and Texas Red. The digital images were then captured with a cooled CCD camera and processed with the help of its SPOT software (Diagnostic Instruments).

siRNA directed against casp-2S (5′-GAAUACUACUGGUAAACUAUU-3′; 5′UTR), and a scramble nontargeting siRNA (5′-UUCUCCGAACGUGUCACGUdTdT-3′), were synthesized by Dharmacon Inc. siGENOME SMARTpool XPC siRNA was purchased from Dharmacon. siRNA (100 nmol/L) was transfected into cells with Lipofectamine 2000 transfection reagent (Invitrogen) according to the manufacture's instruction.

Real-time PCR was conducted as described previously (19). Special primers were designed to only amplify casp-2S or casp-2L, for example, the forward sequence of casp-2S primer covers the exon 9, whereas the forward sequence of casp-2L primer covers the junction of exon 8 and 10. The sequence of the primers is as follows: casp-2L: forward 5′-GCCTGCCGTGGAGATGAGACTGAT-3′; casp-2S: forward 5′-CACCGCCTCTCTTGCTCTAT-3′; casp-2L and casp-2S: reverse 5′- TTACCGGCATCACTCTCCTC-3′; GAPDH: forward 5′-GAAGGTGAAGGTCGGAGT -3′, reverse 5′-GAAGATGGTGATGGGATTTC-3′.

Cells were transiently transfected with casp-2S promoter-luciferase constructs (Del4). As an internal control, the pGL4.73 plasmid (Promega), which carries a Renilla luciferase gene, was also cotransfected into the cells to normalize for transfection efficiency. Luciferase assays were conducted with Dual-Luciferase Reporter Assay System (Promega) according to manufacturer's instruction.

HCT116(p53^−/−) cells were fixed with 1% formaldehyde for 10 minutes at room temperature. The chromatin immunoprecipitation (ChIP) assay was conducted as described previously (24) using 2 μg of either normal rabbit immunoglobulin G or rabbit anti-XPC antibodies. The casp-2S promoter–specific PCR reactions were carried out with 4 different pairs of primers (Supplementary Data).

Because our results (Supplementary Data S1A and B and S2A–D) and others (6, 7) have shown that XPC-deficient cells have lower UV and cisplatin sensitivity than other NER-deficient cells, we wanted to determine whether XPC-deficient cells are also resistant to DNA damage–induced apoptosis. A SV40-transformed human XP-C cell line and its stably XPC-restored counterpart were used to investigate the role of XPC protein in various DNA-damaging agent–induced apoptosis. As shown in Fig. 1A–D, following treatment with diverse DNA-damaging agents, XPC-restored cells exhibited enhanced apoptosis than parental XPC-deficient cells, as reflected by the characteristic increase of PARP cleavage in these cells. Enhanced UV-induced apoptosis in XPC-restored cells was also confirmed by Annexin V staining (Fig. 1E) and sub-G₁ cell analysis (Supplementary Data S3). Furthermore, we transiently transfected XPC cDNA-containing plasmids into XP-C cells and again found the increased apoptosis upon UV irradiation compared with the empty vector–transfected XP-C cells (Supplementary Data S4A and B). To further confirm the independent role of XPC in apoptosis, without the potential influence via its canonical function in NER, we knocked down the expression of XPC in NER-deficient XP-A cells and analyzed their apoptosis upon UV irradiation. Our findings indicated that XPC-deficiency compromised UV-induced apoptosis in XP-A cells which are completely deficient in both GGR and TCR (Fig. 1F and G). These observations showed that XPC protein can facilitate DNA damage–induced apoptosis independent of its known function in DNA repair.

The above-mentioned data were obtained from SV40-transformed XP-C and XP-A cells, in which the transcriptional properties of p53 are suppressed. To further investigate the role of an involvement of p53 in the XPC-mediated apoptosis, we analyzed the effect of XPC deficiency on UV-induced cellular apoptosis in the presence or absence of p53. XPC shRNA was stably transfected into 041-TR cells carrying tetracycline (Tet)-regulated p53 and 2 new cell lines with downregulated XPC were established (Fig. 2A). These cell lines were irradiated with UV either in the presence or absence of Tet, and the apoptosis was analyzed by PARP cleavage and Annexin V staining. As shown in Fig. 2B–E, XPC knockdown compromised UV-induced apoptosis when p53 is eliminated by Tet, while it had no significant influence on apoptosis when p53 was present and responded normally to induction by DNA damage. This finding was also confirmed in HCT116 cells with differing status of p53 (Supplementary Data S5A–D). These data indicated that XPC-mediated apoptosis is independent of functional p53, and this novel effect can be obscured by the regular p53-mediated apoptosis.

For insights into the mechanism by which XPC protein enhances apoptosis, we focused on the contribution of XPC to the intrinsic apoptotic pathway. We first analyzed cyto c release from mitochondria following UV irradiation of XPC-deficient and XPC-proficient cells. As shown in Fig. 3A, despite the higher level of apoptosis in XPC-proficient cells as reflected by increased PARP cleavage, UV irradiation caused a comparable release of cyto c into cytosol fraction from both cell lines. Furthermore, we analyzed the disruption of the mitochondrial transmembrane potential and found that both XPC-deficient and XPC-proficient cells displayed the similar diffused green fluorescence upon UV irradiation (Fig. 3B), further supporting that XPC protein does not influence DNA damage–induced mitochondrial permeabilization, and indicated that XPC could be playing its role at steps downstream of cyto c release during apoptotic cascade. We then determined the activation of various caspases functioning in the downstream of cyto c release in both XP-C and XP-C+XPC cells following UV irradiation. As shown in Fig. 3C and D, casp-2L, casp-6, and casp-9 activation prominently coincided with the enhanced PARP cleavage of XPC-proficient cells, whereas casp-3 and casp-7 activation was comparable in XP-C and XP-C+XPC cells. This suggested that XPC-mediated apoptosis might be occurring through the enhanced activation of casp-2L, casp-9, and casp-6. Furthermore, we also found that pan-caspase inhibitor z-VAD blocked, whereas casp-3 inhibitor z-DEVD had no influence on UV-induced apoptosis in both XP-C and XP-C+XPC cells, although casp-3 activity was higher in XP-C+XPC cells than XP-C cells upon UV irradiation (Supplementary Data S6A–D). This indicates that the functional role of casp-3 in UV-induced apoptosis in XP-C cells is apparently being substituted by other executor caspases.

Casp-2 exists as 2 distinct isoforms, proapoptotic casp-2L and antiapoptotic casp-2S (17, 18). So, we determined the effects of XPC on the expression of both casp-2L and casp-2S in cultured cells. As shown in Fig. 4A, in contrast to casp-2L, casp-2S protein was upregulated in XPC-deficient cells, indicating that XPC protein inhibits the expression of casp-2S. Real-time quantitative reverse transcriptase (RT)-PCR analysis further indicated that XPC repressed the transcription of casp-2S without an influence on casp-2L transcription (Fig. 4B and C). To further confirm the inhibitory effect of XPC on the expression of casp-2S, we transiently transfected various amounts of XPC cDNA-containing constructs into XP-C cells, or down-regulated XPC expression by transfecting XPC siRNA into HCT116(p53^−/−) cells, and determined the protein levels of casp-2S and casp-2L. As expected, with the increase of XPC expression, casp-2S showed decreased level whereas casp-2L level did not change. Similarly, with the decrease of XPC expression, casp-2S exhibited increased level and again casp-2L level kept constant (Fig. 4D and E).

It has been reported that casp-2S can be transcribed from an alternate promoter within intron 1 of the casp-2 gene (21, 22). Thus, we used the casp-2S promoter-luciferase reporter construct, described earlier (21, 22), to investigate the repressive effect of XPC on the casp-2S promoter activity. Our data showed that XPC expression led to a substantial decrease in the reporter activity from the construct containing casp-2S promoter (Del4; Fig. 4F), indicating that XPC may reduce the synthesis of casp-2S pre-mRNA. We further detected the influence of downregulation of XPC on the activity of casp-2S promoter in HCT116 cells. As shown in Fig. 4G, knockdown of XPC increased the casp-2S promoter activity in HCT116 cells. To gain insight into the mechanism through which XPC regulates casp-2S promoter activity, we conducted ChIP assay to investigate whether XPC protein directly binds to the putative casp-2S promoter (–3,006–1,952) site (21). As shown in Fig. 4H, the strong casp-2S promoter (Seq 1–4), but not casp-2 exon-10 PCR product, in anti-XPC antibody-recruited DNA fragment indicated the enrichment of XPC to casp-2S promoter. Taken together, these data showed that XPC protein specifically regulates the expression of antiapoptotic casp-2S by directly binding to casp-2S promoter region and inhibiting its activity.

Earlier it has been reported that casp-2 is required for stress-induced apoptosis (25). However, that study could not have separated the roles of casp-2L and casp-2S because the siRNA used in their experiments actually targeted both casp-2L and casp-2S (25). By the siRNA targeting of casp-2L and casp-2S (Supplementary Data S7A and B) and casp-2 inhibition (Supplementary Data S7C), we failed to find any influence of casp-2 downregulation on UV-induced apoptosis in both XP-C and XP-C+XPC cells observed in this study.

To specifically investigate the role of casp-2S in XPC-mediated apoptosis, we designed an siRNA that only targeted casp-2S (Fig. 5A) and analyzed the UV-induced apoptosis in both XPC-deficient and XPC-proficient cells before and after casp-2S siRNA knockdown. The results indicated that downregulation of casp-2S significantly enhanced the apoptosis in the deficient XP-C cells but not the restored XP-C+XPC cells following UV irradiation (Fig. 5B–D). Overexpression of casp-2S in casp-2S siRNA–transfected XP-C cells reduced UV-induced apoptosis to the original level (Fig. 5E and F). Furthermore, downregulation of casp-2S simultaneously enhanced the activation of casp-9 and casp-6 in XPC-deficient cells, but not in XPC-proficient cells (Fig. 5C and D). Notably, we were unable to observe any influence of casp-2S knockdown on the casp-2L activation. The above results showed that XPC enhances apoptosis through downregulation of antiapoptotic casp-2 short isoform, which is an inhibitor of casp-9 and casp-6 activation.

To determine whether the level of XPC affects chemotherapeutic agent–induced apoptosis in human cancer cells, we overexpressed XPC in various human cancer cell lines of different p53 status and analyzed cisplatin-induced apoptosis. As shown in Fig. 6, XPC overexpression enhanced the cisplatin-induced apoptosis in p53-deficient human ovarian carcinoma cells SKOV3 and human non–small cell lung carcinoma cells H1299, as well as p53 heterozygous ovarian carcinoma cells A2780/CP70. However, XPC overexpression did not exhibit augmented apoptosis in p53-proficient A549 cells upon cisplatin treatment. These data indicated that elevation of XPC level in p53-deficient cancer cells can overcome their resistance to cisplatin.

XPC deficiency has been associated with cancer predisposition in both human and animal models (26–30). Deficient NER is established as the main contributor to the increasing cancer rates. Here, we have provided evidence showing that XPC protein can also enhance DNA damage–induced apoptosis. Thus, the loss of XPC-mediated apoptotic pathway could also be a contributory factor in the cancer susceptibility of XPC-deficient individuals.

The resistance of XPC-deficient cells to apoptosis was initially attributed to the proficient TCR in these cells (6, 7). The intact TCR pathway removes UV- or cisplatin-induced DNA lesions from the transcribed strand of the active genes, thus eliminates the apoptosis triggers. However, our data show that XPC-deficient cells are not only resistant to apoptosis induced by UV and cisplatin treatment, which induce DNA lesions repaired by NER, but also resistant to apoptosis induced by ionizing radiation (IR) and etoposide, which induce double strand breaks (DSB). Given the putative role of XPC in the repair of DSBs (31), XPC-deficient cells should have more DNA lesions, or in other words apoptosis triggers, than XPC-proficient cell. Therefore, the compromised apoptosis, in response to IR and etoposide in XPC-deficient cells, could not be attributed to diminished apoptosis signals. This indicates that XPC protein may participate in apoptosis process through a novel function beyond its canonical role in DNA repair. Furthermore, our finding that downregulation of XPC in XP-A cells compromised UV-induced apoptosis further reinforced this notion. Because XP-A cell line is deficient in both GGR and TCR, the knockdown of XPC expression does not further reduce its NER efficiency, and the DNA lesions in the entire genome including transcribed and nontranscribed strands are comparable between XPC-knockdown and XPC-proficient XP-A cells. Thus, the reduced apoptosis in XP-A cells transfected with XPC siRNA can only be attributed to the loss of XPC protein and indicates that XPC participates in DNA damage–induced apoptosis independent of NER.

It was previously reported that XPC protein plays an essential role in cisplatin-induced apoptosis, perhaps through the activation of p53- and p73-dependent apoptotic pathways (32). Here, we have identified a new pathway in which XPC is shown to enhance DNA damage–induced apoptosis in the absence of functional p53. More interestingly, XPC seems to enhance apoptosis only when p53 function is deficient, indicating that XPC is redundant in cisplatin- or UV-induced apoptosis in the presence of functional p53, probably because the dominant p53-driven apoptosis obscures the underlying contribution of the XPC protein. Therefore, the role of this XPC-casp-2S pathway, similar to the role of p73 pathway (33), is not critical to apoptosis in the cells with wild-type p53. However, considering the high p53 mutation prevalence among human tumors, induction of XPC-mediated apoptosis pathway offers a unique strategy for sensitizing p53-deficient tumors to cancer therapy.

Although we were unable to alter UV-induced apoptosis by casp-2 siRNA that targets both casp-2L and casp-2S, we clearly showed that specific knockdown of casp-2S overcomes the resistance of XPC-deficient cells to apoptosis. As indicated above, casp-2 exists as 2 distinct isoforms with opposing functions: casp-2L induces while casp-2S inhibits cell death upon overexpression (17, 18). Because both isoforms were knocked out or downregulated simultaneously in casp-2 knockout mice studies and the RNA interference cell culture studies, the biological function of these 2 casp-2 isoforms, especially the antiapoptotic function of casp-2S was not clarified (34). In addition, the results about the antiapoptotic function of casp-2S obtained by overexpression experiments are also controversial (18, 21, 35). By a specific casp-2S siRNA designed in this study, we have unambiguously revealed that downregulation of casp-2S can enhance UV-induced apoptosis in XPC-deficient cells, but not XPC-proficient cells. This indicates that the cellular endogenous XPC level plays a critical role in investigations of the function of casp-2S by manipulation of cellular casp-2S levels.

The expression of casp-2S can be regulated by alternative promoter activation (21). It has been reported that p73 is able to specifically transactivate the expression of casp-2S through directly binding to a Sp-1 binding site–containing region in the promoter of casp-2S. In contrast, our data clearly showed that the binding of XPC protein to casp-2S promoter inhibits casp-2S expression. Given XPC is able to regulate gene transcription in 2 opposite ways with different mechanisms (14, 15), it is possible that XPC can function as a direct transcriptional repressor of casp-2S.

In summary, XPC protein is proposed to enhance DNA damage–induced apoptosis through intrinsic apoptotic pathway (Fig. 7). XPC protein downregulates the expression of antiapoptotic casp-2S through inhibiting its promoter activity. The downregulation of casp-2S would attenuate its inhibitory effect on the activation of casp-9 and casp-6, resulting in augmented apoptosis.

No potential conflicts of interest were disclosed.

The authors greatly acknowledge Drs. George Stark for 041-TR cell line, Bert Vogelstein for HCT116 cell line, Paul Modrich for A2780/CP70 cell line, Thomas Hamilton for SKOV3 cell line, and Wenrui Duan for H1299 and A549 cell lines. The authors also thank Dr. Randy Legerski for providing pXPC3 plasmids and Yi-Wen Huang for assisting real-time PCR analysis.

This study was supported by NIH grants CA151248 to Q.-E. Wang and ES2388, ES12991, and CA93413 to A.A. Wani.

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