UV radiation-induced XPC translocation within chromatin is mediated by damaged-DNA binding protein, DDB2

Qi-En Wang1, Qianzheng Zhu1, Gulzar Wani1, Jianming Chen1 and Altaf A. Wani1,–4

1 Department of Radiology, 2 Department of Molecular and Cellular Biochemistry and 3 James Cancer Hospital and Solove Research Institute, The Ohio State University, Columbus, OH 43210, USA

4 To whom correspondence should be addressed at Department of Radiology, The Ohio State University, Columbus, OH 43210, USA Email: wani.2{at}osu.edu


    Abstract
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Supplementary material
 References
 
The tumor suppressor p53 protein has been established as an important factor in modulating the efficiency of global genomic repair. Our recent repair studies in human cells reported that p53 regulates the recruitment of XPC and TFIIH proteins to specific DNA damage sites. Here, we have examined the influence of p53 and damaged-DNA binding complex (DDB2) proteins on the distribution of XPC within damaged chromatin in vivo and the recruitment of XPC to DNA damage sites in situ. The results show that UV irradiation causes the translocation of XPC from a loosely bound form into a tight association with chromatin in vivo. The UV radiation-induced redistribution of XPC was equally compromised in p53-deficient, as well as DDB2-deficient, human cells. Similarly, rapid recruitment of XPC to DNA damage in situ was also impaired in both cell lines. Ectopic expression of DDB2 in p53-deficient cells overcame the requirement of p53 function for UV-induced translocation of XPC in vivo. Restoration of DDB2 function also enhanced the recruitment of XPC to DNA damage sites in situ and increased the global repair of cyclobutane pyrimidine dimer from the genome. These results indicate that DDB2 is a key downstream factor of p53 for regulating the movement of XPC to DNA damage in irradiated cells.

Abbreviations: CPD, cyclobutane pyrimidine dimer; DDB, damaged-DNA binding complex; GGR, global genome repair; LFS, Li-Fraumeni syndrome; NER, nucleotide excision repair; PBS, phosphate-buffered saline; 6–4PP, pyrimidine (6–4) pyrimidone photoproduct; TCR, transcription-coupled repair; XP, xeroderma pigmentosum


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Supplementary material
 References
 
Living organisms are continuously subjected to endogenous and exogenous agents that damage their genome. However, in order to maintain the integrity of the genetic material, cells have developed sophisticated systems for the repair of damaged DNA. Amongst these, the nucleotide excision repair (NER) pathway is a major DNA repair process that removes structurally unrelated DNA damage, such as UV light-induced DNA lesions and bulky DNA adducts from genomic DNA (1). NER includes two distinct subpathways, global genome repair (GGR) and transcription-coupled repair (TCR) (2). The GGR removes lesions from the entire genome, whereas, the TCR eliminates DNA damage from the transcribed strand of active genes (2). The biochemical steps of NER include damage recognition, dual incision and gap-filling DNA synthesis (3,4). Genetic and biochemical studies have identified the requirement of more than 20 different proteins to achieve complete NER (3,4). It is becoming clear that NER in vivo is mediated by the sequential assembly of these repair factors at the sites of the DNA lesions and XPC-hHR23B is believed to be the first recognition factor recruited in GGR (57). Following damage recognition events, TFIIH is recruited by XPC-hHR23B to open the DNA helix around the damage site (810). Other recognition factors, such as XPA and RPA, are believed to join the TFIIH-containing repair complex to verify the nature of DNA structural alteration (5). Once damage is validated, two endonucleases, XPF-ERCC1 and XPG, are recruited to the damage site for cleaving the damage-containing oligonucleotide of ~24–32 nt (10,11). Finally, the DNA structure is restored by subsequent DNA synthesis with the aid of auxiliary factors like PCNA.

It is now well recognized that the tumor suppressor p53 protein is intimately involved in NER. It contributes mainly to GGR, but not the TCR, of bulky DNA lesions. Moreover, p53 exhibits a certain degree of selectivity in its ability to influence NER, as the loss of p53 function affects the repair of cyclobutane pyrimidine dimers (CPD), but not of another major photoproduct, pyrimidine (6–4) pyrimidone photoproduct (6–4PP) (1216). Despite its demonstrated interaction with XPB and XPD components of TFIIH, however, p53 has failed to show an effect on NER in reconstituted cell-free repair systems (17,18). Recent observations indicate that the p53 protein is not physically associated with the sites of DNA damage in situ and, therefore, fail to support a direct role for p53 in NER (16,19). As a DNA damage responsive transcription factor, p53 activates a number of downstream genes, such as p21, GADD45 and DDB2 [a p48 subunit of damaged-DNA binding (DDB) complex], which are involved in cell cycle regulation, apoptosis and DNA repair (2022). Among these downstream targets of p53, DDB2 is most relevant to p53-dependent GGR. The transcription of DDB2 strongly depends on p53 function and the mRNA level of DDB2 increases after DNA damage in a p53-dependent manner (21). While the exact role of DDB2 in NER remains to be fully investigated, a number of studies implicate it as a factor in GGR (2325). For instance, DDB activity is absent in an inherited syndrome xeroderma pigmentosum group E (XP-E) cells due to a mutation of the DDB2 gene (26,27). This cell line has ~50% of the normal repair level as measured by unscheduled DNA synthesis after UV irradiation (2830). It has also been shown that XP-E cells (XP2RO) exhibit lesser GGR of CPD than the normal human cells (21).

We have demonstrated recently that the rapid recruitment of XPC and TFIIH to DNA damage requires normal p53 function. With regards to the role of DDB2 in GGR, our observations support the hypothesis that p53-dependent GGR is mediated by DDB2 (21). Thus, in this study, we examined the influence of p53 and DDB2 on the distribution of the GGR-specific recognition factor, XPC, before and after UV irradiation. By employing an extensive nuclear protein fractionation analysis, we have observed the translocation of XPC protein, from a loosely bound form to a tight in vivo chromatin association, following UV irradiation. Moreover, UV-induced redistribution of XPC is compromised in p53- or DDB2- deficient cells. More importantly, the data show that ectopic expression of the transfected DDB2 gene in p53-deficient cells, (i) compensates for the requirement of p53 function in UV-induced translocation of XPC in vivo, (ii) enhances the recruitment of XPC to DNA damage sites in situ and (iii) increases the removal of CPD from the genome. Lastly, histone hyperacetylation, induced by treatment of p53- or DDB2-deficient cells with sodium butyrate, did not affect the translocation of XPC to damaged chromatin in vivo or the recruitment of XPC to DNA damage sites in situ. The overall results suggest strongly that DDB2 is a downstream effector of p53 for regulating the recruitment of XPC to certain types of DNA damage.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Supplementary material
 References
 
Cell culture and treatment
The normal human skin fibroblasts (OSU-2) containing wild-type p53, were established and maintained in culture as described previously (31). Li-Fraumeni Syndrome (LFS) fibroblast 041 line (p53-Null, harboring a codon 184 frameshift mutation, resulting in premature termination of translation of p53 protein), was provided by Dr Michael Tainsky (Anderson Cancer Center, Austin, TX). Normal human fibroblast cell strain (WI38) and XP-E cell line (GM01389) were obtained from NIGMS Human Genetic Cell Repository (Coriell Institute for Medical Research, Camden, NJ). SV40-transformed WI38 cells (WI38/VA13) were provided by Dr Tsukasa Matsunaga (Kanazawa University, Japan). All the cell lines, except XP-E, were grown in DMEM supplemented with 10% fetal calf serum and antibiotics at 37°C in a humidified atmosphere of 5% CO2. XP-E cells were grown in MEM with 2x essential amino acid, 2x non-essential amino acid and 2x vitamin supplemented with 15% fetal calf serum and antibiotics. For overall UV exposure, the cells were washed with phosphate-buffered saline (PBS), irradiated with varying UV doses and incubated in suitable medium for the desired time period. The irradiation was performed with a germicidal lamp at a dose rate of 0.8 J/m2/s as measured by a Kettering model 65 radiometer (Cole Palmer Instrument Co., Vernon Hill, IL). For inhibition of histone deacetylases, the cells were grown to confluence and maintained in serum-free medium for 24 h before adding sodium butyrate to a final concentration of 5 mM. The cultures were kept in the inhibitor for another 24 h before UV irradiation.

Cell transfection
Exponentially growing 041 cells (3x105) were plated in 60-mm dishes 24 h prior to plasmid transfection. Cells were transfected with either pBJ5 empty vector or FLAG tagged p48 construct (provided by Dr Gilbert Chu, Stanford University, Stanford, CA) using FuGENE 6 transfection Reagent (Roche Molecular Biochemicals, Indianapolis, IN) according to manufacturer's instructions. After a 24 h post-transfection period, the cultures were shifted to serum-free medium for another 24 h and then subjected to UV treatment.

Whole cell extract preparation and fractionation of cellular proteins
For whole cell extract preparation, the cells were harvested by trypsinization and lysed by boiling for 10 min in a sample buffer (2% SDS, 10% glycerol, 10 mM DTT, 62 mM Tris–HCl, pH 6.8, and protease inhibitor cocktail of 10 µg/ml pepstatin, 1 µg/ml leupeptin and 1 mM PMSF). For fractionation of cellular proteins, the cells were harvested by trypsinization, washed twice with cold PBS, and subjected to sequential extraction with detergent and salt, according to the methods of Humphrey et al. (32) and Rapic-Otrin et al. (33), with some modifications (Figure 1A). Briefly, the cells were resuspended in a hypotonic buffer (10 mM HEPES, pH 7.9, 10 mM KCl, 1.5 mM MgCl2 and protease inhibitor cocktail) and lysed with 0.1% Triton X-100. The lysates were centrifuged and the supernatant, designated as S1, was further centrifuged at high speed to yield the clear supernatant S2 (cytoplasmic soluble proteins). The nuclear pellet was then washed twice with isotonic sucrose buffer (50 mM Tris–HCl, pH 7.4, 0.25 M sucrose, 5 mM MgCl2 and protease inhibitor cocktail) to yield STM fraction (cytoplasmic soluble proteins). Removal of the nuclear envelope by LS buffer (10 mM Tris–HCl, pH 7.4, 0.2 mM MgCl2 and protease inhibitor cocktail) containing 1% Triton X-100 yielded the Triton Wash fraction, TW (nucleoplasmic soluble proteins). The nuclear pellet was washed twice with LS buffer, yielding LS fraction (nucleoplasmic soluble proteins) and extracted consecutively with increasing concentrations (0.3, 0.5 and 2.0 M) of NaCl in LS buffer to result in supernatant fractions, respectively designated as 0.3, 0.5 and 2.0. The nuclear residue (NR) comprising of DNA and nuclear matrix was dissolved by sonication in LS buffer. In order to separate chromatin and nuclear matrix, the DNase I treatment and extraction method was carried out essentially as described (34,35). The nuclei were isolated as above, resuspended in digestion buffer with 1 U/µl DNase I (Invitrogen, Carlsbad, CA), and extracted with 0.25 M ammonium sulfate. The extract was then centrifuged and the supernatant contained the chromatin fraction. The pellet was further extracted with 2.0 M NaCl in LS buffer to yield a high salt (HS) wash fraction containing proteins bound to the nuclear matrix. The final pellet was resuspended in LS buffer and sonicated to release the nuclear matrix. Each protein fraction, corresponding to an equivalent cell number, was loaded for SDS–PAGE and analyzed by immunoblotting with indicated antibodies.



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Fig. 1. Effect of UV irradiation on the distribution of NER proteins in normal human cells. (A) Outline of the scheme for sequential salt extraction of cellular proteins. Cells were processed according to the described steps to yield distinct cytoplasmic and nuclear protein fractions for analysis of individual NER factors. (B) Repair-proficient human fibroblast, OSU-2, were growth arrested for 24 h before UV irradiation at 20 J/m2 and allowed to repair for 30 min. The protein fractions were prepared from control unirradiated and irradiated cells according to the outlined scheme in (A). Individual protein fractions, corresponding to equivalent cell numbers, were subjected to 8% SDS–PAGE and analyzed by immunoblotting with anti-XPC, anti-XPB, anti-XPA, anti-actin and anti-lamin B antibodies. (C) Nuclei isolated from unirradiated and UV-irradiated OSU-2 cells were subjected to DNase I treatment and proteins were extracted with 0.25 M ammonium sulfate to separate the chromatin and nuclear matrix. The nuclear matrix was washed and further extracted with 2.0 M NaCl to release the matrix-associated proteins. Each protein fraction was separated by SDS–PAGE and analyzed by western blotting with anti-XPC antibody.

 
Western blot analysis of proteins
The proteins were quantified, separated by SDS–PAGE and the immunoblot analysis was performed as described earlier by using chemiluminescent detection (12). The following antibodies and dilution factors were used: rabbit anti-XPC (kindly provided by F.Hanaoka, Osaka University, Japan; 1:10 000), rabbit anti-p21 (Santa Cruz Biotechnology, Santa Cruz, CA; 1:200), rabbit anti-XPA (Santa Cruz Biotechnology; 1:1000), rabbit anti-XPB (Santa Cruz Biotechnology; 1:1000), monoclonal anti-p53(DO-1) (Santa Cruz Biotechnology; 1:200), monoclonal anti-actin (Neomarkers, Fremont, CA; 1:500) and anti-lamin B (Santa Cruz Biotechnology; 1:1000).

Localized micropore UV irradiation and immunofluorescent staining
To perform micropore UV irradiation, the cells grown on glass coverslips were washed with pre-warmed PBS and UV irradiated as described (16). Briefly, an isopore polycarbonate filter (Millipore, Bedford, MA), containing pores of a 5 mm diameter, was placed on top of the cell monolayer. The filter-covered coverslips were irradiated from above with desired doses of UV-C (254 nm) at a dose rate of 0.8 J/m2/s. The filter was then gently removed and the cells were either processed immediately, or maintained in serum-free medium for the desired period before fixation and processing. For immunofluorescent staining, cells growing on coverslips were washed twice with cold PBS and subsequently fixed and permeabilized in buffer containing freshly made 2% paraformaldehyde and 0.5% Triton X-100 in PBS at 4°C for 30 min. The cells were blocked with normal goat serum (NGS), incubated with primary antibody against XPC (rabbit anti-XPC at 1:5000) in PBS with 1.5% NGS, and stained with secondary antibody (FITC-conjugated goat anti-rabbit IgG at 1:200 dilution). After first step staining, the cells were treated with 2 M HCl at 37°C for 10 min to denature the DNA, followed by a PBS rinse to remove HCl. The slides were incubated with 1:1000 dilution of mouse anti-CPD antibody (TDM2, generously provided by Dr T.Matsunaga, Kanazawa University, Japan) and secondary antibody (Texas Red conjugated goat anti-mouse IgG at 1:400 dilution) in PBS with 1.5% NGS. The cells, after washing with PBS containing 0.1% Tween-20, were mounted in an antifade containing medium with 0.75 µg/ml of 4',6'-diamidino-2-phenylindole (DAPI, Vector Laboratories, Burlingame, CA) as a DNA counter stain. Fluorescence images were obtained with a Nikon Fluorescence Microscope E800 (Nikon, Tokyo, Japan) fitted with appropriate filters for DAPI, 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, Sterling Heights, MI).

Quantification of CPD by immuno-slot blot assay
The amount of initial dimer formation and the damage remaining in DNA after cellular repair were quantified using non-competitive immuno-slot blot assay, essentially as described earlier (31,36). Briefly, after UV exposure and desired incubation periods, cells were recovered by trypsinization and immediately lysed for DNA isolation. For damage estimation at each point, several increasing concentrations of unirradiated, irradiated and repaired DNA samples were evaluated by standard immunoassay using dimer-specific polyclonal antibody, as described earlier (37). The damage levels were calculated by comparing the band intensities of the samples with UV-irradiated DNA standard samples run in parallel with all the blots.


    Results
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 Supplementary material
 References
 
UV irradiation causes translocation of XPC from a loosely bound form into a tight association with chromatin
NER factors, e.g. XPC and XPB (TFIIH), are sequentially recruited to the DNA damage sites immediately after UV irradiation (7,16). It has been suggested that such recruitment is due to the movement of the NER proteins (7,16). To examine the NER factor recruitment events in the context of their association with chromatin, we first analyzed the cellular and subnuclear distribution of XPC, XPB and XPA proteins in normal human fibroblast cells. The proteins were fractionated into eight components, with different concentrations of detergent and/or salt, using a published procedure (32,33) with appropriate modifications (Figure 1A). The presence of XPC, XPB and XPA in each fraction was examined by western blotting analysis and the results are shown in Figure 1B. In unirradiated normal human cells, XPC was initially detected in 0.3, 0.5 and 2.0 M salt fractions (Figure 1B), indicating that XPC resides within the nucleus and the majority of the protein is associated with chromatin. Thirty minutes after UV irradiation of intact cells, XPC was completely depleted from the 0.3 M fraction, indicating its tighter association with chromatin upon UV irradiation. Most XPB protein was detectable in 0.3, 0.5 and 2.0 M salt, and a small amount in NR fractions. This indicates that the majority of XPB protein is initially bound to chromatin and a smaller HS-resistant portion is tightly bound to the nuclear matrix. Interestingly, incubation of cells for 0.5 h following UV irradiation did not show any discernible change in the distribution pattern of XPB (Figure 1B). Unlike the distribution of XPC and XPB proteins, most of the XPA protein was detectable in TW and LS fraction with a small amount accompanying 0.3 M salt (Figure 1B). This indicates that the majority of XPA protein resides in a soluble form within the nucleoplasm. UV irradiation of cells did not change the association of XPA protein. It should be noted that in all the protein analysis experiments, cytoskeleton structural protein, actin and nuclear matrix protein, lamin B, were used as the internal references for reproducible fractionation. As expected, actin was readily detected in all the fractions, whereas lamin B was only present in the NR fraction (Figure 1B).

Another scheme using treatment with DNase I was applied to differentiate between the association of XPC protein with chromatin or nuclear matrix. Soluble proteins in nucleoplasm were first extracted with 1% Triton X-100, and chromatin-associated proteins were recovered upon DNase I digestion and extraction with 0.25 M ammonium sulfate. This treatment releases ~95% of the histone proteins (35). Proteins bound to the nuclear matrix were released with 2 M NaCl to separate them from structural nuclear matrix proteins. About 60% of XPC protein was detected in the chromatin fraction (Figure 1C). However, the XPC protein distribution did not change upon UV irradiation of cells. These results confirm that observed translocation of XPC upon UV irradiation occurs exclusively within the chromatin milieu.

Lack of p53 compromises the UV radiation-induced translocation of XPC within chromatin
We have demonstrated that the recruitment of XPC and TFIIH to DNA damage sites is largely dependent upon the functional status of p53 (16). To investigate the factor association in a chromatin context, XPC protein was fractionated from p53-deficient LFS cells (Figure 2A). Like that of normal p53-wt cells, XPC protein was detected in 0.3, 0.5 and 2.0 M salt fractions, indicating an initial p53-independent cellular distribution of XPC. Unlike normal cells, the XPC subnuclear distribution in p53-deficient 041 cells was not affected by UV irradiation at 20 J/m2. The distribution pattern was clearly different from that of p53-proficient cells in which the same dose of UV irradiation causes a distinct shift of XPC protein from the lower 0.3 M salt fraction. However, irradiation of 041 cells at 50 J/m2 and higher doses resulted in the XPC translocation patterns similar to that of OSU-2 cells (data not shown). CPD and 6–4PP, induced in genomic DNA at a 3:1 ratio, are the main lesions of cellular UV irradiation. The translocation patterns in these two cell types can be explained by the well-known fact that the repair of CPD is dependent, whereas that of 6–4PP is independent of the cellular p53 status. The data suggest that the number of CPD lesions induced by 20 J/m2 are sufficient to cause a demonstrable XPC translocation in p53-proficient OSU-2 cells and not in p53-deficient 041 cells. Despite their prompt repair, however, the low levels of 6–4PP induced by doses below 50 J/m2 preclude the detection of XPC translocation in p53-deficient 041 cells. To confirm the observed patterns of XPC translocation, similar experiments were also conducted with isogenic normal human fibroblast WI38 (Figure 2B) and SV40-transformed WI38VA13 cell lines (Figure 2C). UV radiation-induced translocation of XPC protein was again remarkable in WI38 but not in WI38VA13 cell lines. Since p53 function is disrupted due to the presence of SV40 large T antigen in WI38VA13 cells, it can be concluded that UV induced tighter association of XPC with chromatin is dependent upon the p53 function. In essence, the loss or disruption of p53 function compromises the UV-induced XPC translocation in irradiated cells.



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Fig. 2. Influence of p53 status on the UV-radiation induced redistribution of XPC protein. Various normal or p53-deficient cells were grown to confluence, growth arrested for 24 h before UV irradiation at 20 J/m2. The cells were incubated for 30 min for repair to commence and the protein fractions isolated according to the standard scheme in Figure 1A. Individual protein fractions, corresponding to equivalent cell numbers, were subjected to 8% SDS–PAGE and analyzed by immunoblotting with anti-XPC, anti-actin and anti-lamin B antibodies. Only representative actin and lamin B blots are shown for reference. (A) Human LFS 041 fibroblast, (B) normal human fibroblasts WI38 and (C) SV40-transformed human WI38VA13 fibroblast.

 
DDB2 mutation affects the UV radiation-induced translocation of XPC within chromatin as well as the recruitment of XPC to DNA damage sites
Since DDB2 is involved in the initial CPD recognition steps of GGR, and its expression is strongly dependent on p53 (21), we investigated whether the observed action of p53 in UV-induced XPC translocation is actually mediated through DDB2. Cellular distribution of XPC protein was examined in a DDB2-deficient XP-E cell line. In this cell line, compound heterozygous mutations in DDB2 lead to a L350P point mutation from one allele and an Asn-349 deletion from the other (38). The protein levels of p53 and its downstream target gene p21, were first quantified to confirm the p53 functional status, as well as its UV response in the XP-E cell line. Western blot analysis showed that the basal p53 protein level was very low in this cell line. However, after UV irradiation, the time-dependent induction of p53 responded as expected from the normal p53-wt cells. The p21 protein was also induced by 24 h (Figure 3A). A recent publication indicates that while regulation of p53 was impaired in several kinds of XP-E cells, the levels of p53 and p21 were very well inducible upon UV irradiation (39). We then fractionated XPC protein in unirradiated and UV irradiated XP-E cells according to the standard scheme. As shown in Figure 3B, most of XPC protein was detected in 0.3, 0.5 and 2.0 M salt fractions and this distribution of XPC is similar to that of normal cells. More importantly, the distribution of XPC remained unaltered after UV irradiation. This XPC distribution is clearly similar to that of p53-deficient 041 and SV40 transformed WI38VA13 cells. Thus, these results indicate that per se DDB2, and not p53, is causally related to the UV radiation-induced XPC translocation within chromatin.



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Fig. 3. Involvement of DDB2 in UV radiation-induced XPC translocation within chromatin. (A) p53 and p21 induction response upon UV irradiation are normal in XP-E cell line (GM01389). Confluent XP-E cells were UV irradiated at 20 J/m2 and incubated for indicated time. Whole cell extracts were prepared, proteins separated by SDS–PAGE, blotted on a nitrocellulose filter and examined for the presence of p53 and p21 using anti-p53 and anti-p21 antibodies, respectively. The amount of actin, as a standard control for sample loading, was determined with anti-actin antibody. (B) XPC protein fails to translocate in chromatin following UV irradiation in XP-E cells. XP-E cells were UV or mock irradiated at 20 J/m2, allowed to repair for 30 min and then subjected to sequential salt extraction according to standard scheme of Figure 1A. Individual protein fractions, corresponding to equivalent cell numbers, were subjected to 8% SDS–PAGE and analyzed by immunoblotting with anti-XPC. (C) XPC protein fails to recruit to UV-induced damage sites in DDB2-deficient XP-E cells. Normal OSU-2 cells and XP-E cells grown on the coverslips were UV irradiated at 40 J/m2 through a micropore filter (5 µm pore diameter). The cells were allowed to repair for 30 min and in situ analysis of damage and protein translocation determined upon double immunostaining with rabbit anti-XPC and mouse anti-CPD antibodies as described in the Materials and methods. A color version is available as supplementary material online.

 
We next examined whether DDB2 is required in UV-induced XPC recruitment to local DNA damage sites by micropore UV irradiation. Normal and DDB2-deficient XP-E cells were UV irradiated through a 5 µm pore filter and allowed to repair the damage for 30 min. XPC protein and CPD damage were specifically visualized through their cognate antibodies. As shown in Figure 3C, the XPC-specific signal was seen to intensify within subnuclear spots and co-localized with the CPD signal in normal cells at 30 min following local UV irradiation. This confirmed the rapid recruitment of XPC to damage sites within cells (7,16). On the contrary, in both unirradiated and UV-irradiated XP-E cells, characteristic spatial redistribution of XPC was not observed. For instance, the XPC-specific signal was uniformly present through the entire nuclear area, despite a very clear CPD signal within the subnuclear spots (Figure 3C). These results indicate that for GGR to commence effectively, DDB2 is required for the rapid initial recruitment of XPC to damage sites. In view of the fact that p53 status in the XP-E cell line is normal, these results further suggest that DDB2 is a downstream factor to that of p53 in regulating the rapid and efficient recruitment of XPC in damage processing by GGR.

Ectopic expression of DDB2 in p53-null cells compensates for the defective early events of GGR
If p53 exerts its NER regulatory function through transcriptional activation of DDB2, it should be possible to restore the DNA repair function of p53 by enforced expression of DDB2. To test this possibility, we transfected a FLAG tagged p48 gene into p53-deficient 041 cell line, and assessed the UV radiation-induced XPC translocation, damage site recruitment, as well as DNA repair capacity in FLAG-p48 and control vector transfected cells. Once the expression of transfected FLAG-p48 was confirmed, and the level of transfected FLAG-p48 compared among different samples, the distribution of FLAG-tagged p48 was examined by the standard fractionation scheme. As shown in Figure 4A, most of p48 was detected in 0.3 M salt fraction, some in 0.5 M salt and NR fractions, and a minor amount in the S2, STW and 2.0 M salt fractions. After UV irradiation, the amount of p48 protein decreased considerably in the 0.3 M salt fraction and increased significantly in 2.0 M salt fraction. Moreover, p48 was completely depleted from S2 and STW cytoplasmic fractions upon UV irradiation. Consistent with earlier reports (33), these results indicate that UV radiation results in a tight association of DDB2 with damaged chromatin.



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Fig. 4. Effect of ectopic DDB2 expression in p53-deficient human cells. (A) Human LFS 041 cells were transiently transfected with either empty pBJ5 vector or FLAG-p48 construct 48 h before UV irradiation at 20 J/m2. After 30 min incubation, the cells were subjected to sequential salt extraction according to standard scheme of Figure 1A. The p48 expression and distribution was determined by western blotting with an anti-FLAG antibody. The presence of XPC protein was also examined upon western blotting with an anti-XPC antibody in protein fractions from empty vector transfected (B) and FLAG-p48 transfected (C) 041 cells. (D) Spatiotemporal analysis of damage and NER factor translocation was determined in control (pBJ5) and DDB2 expressing (p48) 041 cells irradiated through micropore (5 µm pore diameter) filters at 40 J/m2. The cells on coverslips were incubated for 30 min and the in situ detection of damage and XPC protein performed as described in the Materials and methods. (E) The ratio of XPC to CPD foci was determined upon scoring >300 individual nuclei for merged fluorescent signals of XPC and CPD foci. (F) 041 cells, transiently transfected with either empty (pBJ5) vector or FLAG-p48 construct for 24 h were incubated in serum-free medium for another 24 h to stop cell growth. The growth-arrested cells were UV irradiated at 20 J/m2 and allowed to repair for desired periods. Total DNA was purified and the amounts of CPD remaining were quantified by immuno-slot analysis with anti-CPD antibody as described in the Materials and methods. A color version is available as supplementary material online.

 
We next investigated the effect of DDB2 expression on the degree of UV radiation-induced shift in the association of XPC with chromatin (Figure 4B and C). Transient transfection of the control vector did not change the distribution of XPC in 041 cells under both unirradiated and UV irradiated conditions (Figure 4B). However, expression of DDB2 provoked a typical redistribution of XPC upon UV irradiation. For example, ectopic expression of p48 had no effect on the original XPC distribution in unirradiated cells and was comparable with control vector-transfected cells. Whereas in p48 transfected cells, UV irradiation caused a decrease in XPC proteins released by 0.3 M salt wash, a transition signified by normal cell response (Figure 4C). This observation suggests that the presence of DDB2 protein in cells, irrespective of their p53 status, increases the association of XPC with chromatin following UV treatment.

The influence of p48 expression on XPC recruitment to damaged sites in situ was further determined by spatio-temporal analysis following micropore UV irradiation (Figure 4D and E). First, we compared the transfection efficiencies for each sample by counting the FLAG-p48 expressing cells, and found that ~70% of cells have been transiently transfected by FLAG-p48 construct. Cells transfected with p48 expression construct exhibited ~37, 21 and 5% XPC and CPD co-localizing foci at 0.5, 1.0 and 2.0 h post-UV irradiation, respectively. On the other hand, cells transfected with the control vector demonstrated that the ratio of XPC and CPD co-localizing foci to total CPD foci was only 20, 10 and 2% within the same post-irradiation period. These data indicate that p48 expression alone, in a p53-null background, enhances XPC recruitment to DNA damage sites. Recent reports also found that enforced expression of p48 can activate the recruitment of ectopic XPC to CPD in SV40-transformed XP-A cell line (19,40). Taken together, the transfection experiments suggest that DDB2 plays a key regulatory role in the damage recognition by XPC.

Finally, we also determined the effect of p48 expression on the GGR of CPD (Figure 4F). p53-deficient 041 cells transfected with the control vector exhibited the typical slow kinetics of CPD repair. Unrepaired CPD remaining in the genome were 98, 91, 80 and 69% at 2, 4, 8 and 24 h post-UV irradiation time, respectively. In contrast, in cells transfected with p48, the unrepaired CPD decreased to 91, 83, 63 and 47% at the same times following UV irradiation. Transient expression of p48 seemed to increase GGR efficiency at all time points but was more pronounced at the late time points. These results are fully consistent with reports that indicate that ectopic expression of DDB2 enhances GGR in p53-deficient cells (41).

Histone acetylation does not affect XPC translocation or recruitment but increases GGR of CPD
DDB2 has been proposed as a chromatin accessibility factor for DNA repair as it has the demonstrated ability to bind p300 and STAGA (42,43). To explore the relationship between DDB2-regulated XPC translocation, XPC recruitment and chromatin modification, p53-deficient 041 and DDB2-deficient XP-E cells were pre-treated with a well-established histone deacetylase inhibitor, sodium butyrate (44,47). The treated cells were then UV irradiated and the distribution of XPC as well as histone acetylation was examined. As expected, the level of acetylated histone H3 increased significantly in all cell lines tested after 1 h treatment with sodium butyrate (data not shown). However, for both 041 and XP-E cells, the treatment of sodium butyrate did not affect the distribution of XPC, as comparable levels were observed in various cellular fractions from treated and untreated cells (Figure 5A and B). Interestingly, UV irradiation also failed to show the UV radiation-induced translocation of XPC in sodium butyrate treated 041 and XP-E cells (Figure 5A and B). Similarly, in p53-deficient 041 cells, treatment of sodium butyrate also did not affect the XPC recruitment (Figure 5C). The ratio of XPC to CPD foci, showing co-localization of XPC to DNA damage sites, was comparable in both treated and untreated cells (Figure 5D). Moreover, no XPC accumulation was found either in sodium butyrate-treated or untreated XP-E cells after micropore UV irradiation (data not shown). These results indicate that global histone hyperacetylation induced by sodium butyrate treatment cannot substitute for the function of DDB2 in regulating UV-induced XPC translocation. The data suggest that DDB2 plays a direct role in the recognition of UV radiation-induced CPD by XPC protein.



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Fig. 5. The effects of histone deacetylase inhibition on UV radiation-induced XPC translocation within chromatin. Growth-arrested 041 (A) and XP-E (B) cells were pre-treated with 5 mM sodium butyrate (NaBu) or mock treated for 24 h and UV irradiated at 20 J/m2. The cells were incubated in serum-free medium with or without 5 mM sodium butyrate for another 30 min. The protein fractions were prepared according to standard scheme and individual fractions were immunoanalyzed for XPC protein as in Figure 1. (C) 041 cells grown on coverslips were pre-treated with 5 mM sodium butyrate or mock-treated for 24 h before UV irradiation at 40 J/m2 through a micropore filter (5 µm pore diameter). After 30 min incubation in the medium with or without 5 mM sodium butyrate, in situ analysis of CPD and XPC were conducted as described in the Materials and methods. (D) The ratio of XPC to CPD foci was determined upon scoring >300 individual nuclei for merged fluorescent signals of XPC and CPD foci. (E) Growth-arrested OSU-2, LFS 041 and XP-E cells were pre-treated with 5 mM sodium butyrate or mock-treated for 24 h. The cells were UV irradiated at 20 J/m2 and incubated in serum-free medium with or without 5 mM sodium butyrate for various post-treatment times. Total DNA was purified and CPD remaining was quantified by immuno-slot analysis with anti-CPD antibody as described in the Materials and methods. A color version is available as supplementary material online.

 
To examine the efficiency of GGR upon treatment of cells with butyrate, CPD removal was analyzed in normal, p53- and DDB2-deficient cells pre-treated with 5 mM sodium butyrate for 24 h before UV irradiation. As shown in Figure 5E, CPD remaining in genomic DNA of untreated normal cells was 86, 79, 63 and 41% at 2, 4, 8 and 24 h post-irradiation times, respectively. During the same post-irradiation times, however, CPD remaining was 66, 61, 52 and 42% in butyrate-treated cells. Apparently, butyrate treatment increased the initial removal of CPD up to 8 h post-irradiation time. Nevertheless, butyrate treatment increased the GGR of CPD in both p53-deficient 041 and DDB2-deficient XP-E cells at all time points (Figure 5E). Sodium butyrate treatment led to an additional 18 and 26% increase in CPD removal, respectively, at 8 and 24 h in p53-deficient 041 cells. Similarly, CPD removal was increased by 38 and 34% at corresponding 8 and 24 h in DDB2-deficient XP-E cells. Thus, butyrate-induced hyperacetylation of histones seems to have a profound influence on the slow rate of damage processing in p53-deficient 041 as well as DDB2-deficient XP-E cells.


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Association of XPC with chromatin following UV irradiation
We have examined the distribution of NER factors, XPA, XPB and XPC, and demonstrated a UV-radiation dependent alteration in the chromatin association of XPC protein distribution in cells of varying DNA repair background. Protein fractionation experiments revealed that UV irradiation induces a shift of XPC protein from LS (loose association) to HS (tight association) fractions. However, most of the XPA protein was found in simple detergent washes (mainly nucleoplasm) and only a small amount in 0.3 M salt fraction. The distribution of XPA and XPB was not influenced by UV irradiation. Since XPA, XPB and XPC are all recruited to DNA-damage sites (7,16,19), it is inferred that the UV-induced tight association of XPC with chromatin reflects its high affinity to DNA lesions. It is well known that XPA, XPB and XPC all bind to damaged DNA. However, the molar affinities for DNA, as well as the preference of binding damaged over non-damaged DNA, are different for these proteins. For example, XPB does not exhibit any preferential binding to damaged DNA (48); XPA has a nominal ability to discriminate damaged DNA (49,50); whereas XPC-hHR23B, like that of DDB2, has the highest affinity for damaged DNA (51). Thus, in terms of damaged DNA discrimination potential, the proteins are ranked as XPC-hHR23B {cong} UV-DDB > XPA (51). However, some studies show a different binding preference and a dramatic redistribution of XPA rather than XPC following UV irradiation (52,53). The difference could be attributed to varying processing methodologies for generating different protein fractions. For example, in one study, DNase I was used to treat nuclei for solubilizing chromatin before extraction with LS buffer to yield soluble bulky chromatin (52,53). In this procedure, some of the chromatin-bound proteins would be lost with soluble chromatin. However, the scheme used in the present study does not disrupt the chromatin structure and the proteins bound to chromatin were released by extraction with an incremental concentration of salt. This fractionation scheme allows an orderly detection of chromatin-bound proteins. For instance, besides XPC, we also detect a clear UV-induced tight association of DDB2 with chromatin and these results are consistent with recent reports using the similar protein fractionation procedure (33). Both of the data sets are consistent with the movement of DDB2 to DNA damage sites following UV irradiation (19,54).

p53 effect in NER is mediated by DDB2
It has been shown that the basal level of DDB2 transcription, irrespective of DNA damage in cells, is dependent on the presence of p53 protein. In p53-deficient, i.e. 041 cells, the basal level of DDB2 mRNA is greatly reduced and fails to induce after treatment with UV or ionizing radiation (21). Based on these observations, DDB2 has been suggested as the p53 downstream factor for the regulation of GGR. In our previous in situ study, it has been shown that the deficiency of p53 compromises the recruitment of XPC to DNA damage sites following UV irradiation (16). Since p53 itself is not present at the sites of DNA damage (16,19), it is easy to speculate that p53-regulated DDB2 plays the actual role of directing the recruitment of XPC to DNA damage sites. The present study tested this hypothesis through a coordinated inter-related series of experiments on initial XPC-chromatin association, co-localization of XPC to locally induced DNA damage and NER efficiency regulated by ectopic expression of DDB2. The combined data support the conclusion that DDB2 is indeed the key downstream factor of p53-mediated regulation of GGR. However, the results do not exclude the possibility of participation by additional intermediary factors and of more complex mechanisms by which p53 regulates the expression and function of the p48 protein. While significant accumulation of p53 occurs 2–4 h after UV irradiation, the regulatory influence of p53 in modulating the expression of p48 can only be seen after 24 h of UV irradiation (33). Interestingly, it has been shown that p48 is degraded and DDB activity lost at earlier post-UV irradiation times (33). It appears that the UV-induced p48 degradation occurs at DNA damage sites where p48 and other NER factors are recruited immediately after introducing damage in cellular DNA. One plausible explanation for this conundrum could be that p53 maintains critical basal levels of p48 protein for prompt usage in the initial steps of NER. It should be noted, however, that the regulation of DDB2 expression is not the only mechanism by which p53 is involved in NER. It is reported that UV irradiation causes chromatin relaxation via chromatin decondensation and this requires only p53 but not DDB2 function (55).

The role of DDB2 in GGR within chromatin
Repair of DNA damage within the densely packed chromosomal structure requires a mandatory disassembly of chromatin structure for the sake of allowing access to repair machinery. Accordingly, DDB2, while not required for NER in vitro (56), stimulates in vivo repair rates when microinjected into XP-E cells (57,58). This coheres with the proposed role for DDB2 in NER within the chromatin context (58). Accumulating evidence now seems to support the idea that DDB is involved in chromatin remodeling through interaction with CBP/p300 and STAGA proteins (42,43). However, several questions must be answered before such a role of DDB2 in NER is unequivocally established. For example, are the chromatin remodeling HATs actually recruited to DNA damage sites? Does damage recognition by DDB2 also accompany histone acetylation and chromatin remodeling? What is the nature of association, if any, of DDB2 and XPC, when engaged in recognition of the same damage sites?

The presented data show that sodium butyrate treatment stimulates the initial rate of NER in normal cells and increases GGR of CPD in both p53-deficient 041 and DDB2-deficient XP-E cells at all the time points. If butyrate-induced histone acetylation alters NER, it would seem that histone acetylation is a factor in the efficient repair of CPD in chromatin. However, the treatment with sodium butyrate does not affect XPC-chromatin association or XPC recruitment to DNA damage. Thus, treatment of sodium butyrate can increase CPD removal without increasing XPC translocation to damage sites. This result is consistent with the published reports that show a 2-fold increase of UV-induced unscheduled DNA synthesis in sodium butyrate treated XP-C cells (59). It is possible that the increased removal of CPD is due to TCR, which does not require XPC protein. Alternatively, histone hyperacetylation could be making the DNA around lesion sites sufficiently distorted, that it mimics the structure formed during TCR and for which XPC is no longer required to initiate repair. This suggestion is supported by studies which show that several structurally distorted artificial DNA substrates can be repaired in vitro without the requirement of either the transcription or the XPC complex (60,61). XPC-independent incision still releases the lesion as part of a 24–32 nt oligonucleotide, showing that the NER process otherwise operates normally (60,61). It may be noted that histones are not the only substrates of histone deactylases as high-mobility group proteins are also acetylated by these enzymes. So their action has a wide range of effects on the function of the histones and high-mobility group proteins in remodeling chromatin structure as well as regulating gene expression (6264). However, at present we cannot rule out the possibility that the increased NER is due to sodium butyrate-mediated induction or repression of gene expression.

The above-mentioned observations also indicate that the action of DDB2 in NER is more involved than envisaged from the analysis of NER within chromatin. First, histone acetylation and relaxed chromatin is not by itself sufficient to allow the recruitment of XPC to damage sites, implying that DDB2 is acting through other mechanisms. Secondly, the lesions recognized by DDB2 are passed on to XPC for downstream processing steps of NER. Thirdly, DDB2 is supposedly degraded, while XPC and other NER factors are loaded onto damaged DNA. Thus, the requirement of DDB2 in the recruitment of XPC to DNA damage suggests a direct but accessory role for the ultimate damage recognition by XPC. This suggestion is supported by a study showing that DDB can greatly stimulate the excision of CPD in repair assays reconstituted in vitro with XPA, RPA, XPC-hHR23B, TFIIH, XPF-ERCCI and XPG (54).

In summary, we have demonstrated a p53-regulated, UV-induced tight association of XPC with chromatin as a very early event of damage recognition. The XPC-chromatin association is due to the physical binding of XPC to damaged DNA as the XPC protein can be detected in localized DNA damage sites generated through micropore UV irradiation (7,16). The effect of ectopic expression of DDB2 on the proficiency of NER in p53-deficient cells provides the needed evidence for DDB2 acting as a downstream factor of p53 in regulating both the UV-induced association of XPC with chromatin and the recruitment of XPC to DNA damage.


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Supplementary material can be found at: http://www.carcin.oupjournals.org


    Acknowledgments
 
We thank Dr Michael Tainsky (Anderson Cancer Center, Austin TX) for providing the LFS 041 cells, Dr Tsukasa Matsunaga (Kanazawa University, Japan) for WI38VA13 cells and TDM-2 antibody, Dr Fumio Hanaoka (Osaka University, Japan) for anti-XPC antibody, Dr Gilbert Chu for FLAG tagged p48 construct (Stanford University, Stanford, CA) and Dr Gustavo Leone (OSU Comprehensive Cancer Center) for the use of Nikon E800 microscope. This work was supported by NIH grants CA93413 and ES6074.


    References
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 Abstract
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 Materials and methods
 Results
 Discussion
 Supplementary material
 References
 

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Received September 8, 2003; revised January 6, 2004; accepted January 10, 2004.