The initiative role of XPC protein in cisplatin DNA damaging treatment‐mediated cell cycle regulation (original) (raw)

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Gan Wang, Lynn Chuang, Xiaohong Zhang, Stephanie Colton, Alan Dombkowski, John Reiners, Amy Diakiw, Xiaoxin Susan Xu, The initiative role of XPC protein in cisplatin DNA damaging treatment‐mediated cell cycle regulation, Nucleic Acids Research, Volume 32, Issue 7, 1 April 2004, Pages 2231–2240, https://doi.org/10.1093/nar/gkh541
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Abstract

XPC is an important DNA damage recognition protein involved in DNA nucleotide excision repair. We have studied the role of the XPC protein in cisplatin treatment‐mediated cell cycle regulation. Through the comparison of microarray data obtained from human normal fibroblasts and two individual XPC‐defective cell lines, 486 genes were identified as XPC‐responsive genes in the cisplatin treatment (with a minimal 1.5‐fold change) and 297 of these genes were further mapped to biological pathways and gene ontologies. The cell cycle and cell proliferation‐related genes were the most affected genes by the XPC defect in the cisplatin treatment. Many other cellular function genes were also affected by the XPC defect in the treatment. Western blot hybridization results revealed that the XPC defect reduced the p53 responses to the cisplatin treatment. The ability to activate caspase‐3 was also attenuated in the XPC cells with the treatment. These results suggest that the XPC protein plays a critical role in initiating the cisplatin DNA damaging treatment‐mediated signal transduction process, resulting in activation of the p53 pathway and cell cycle arrest that allow DNA repair and apoptosis to take place. These results reveal an important role of the XPC protein in the cancer prevention.

Received October 14, 2003; Revised and Accepted March 23, 2004

INTRODUCTION

Cisplatin is an important antineoplastic drug that has been used in the treatment of many types of cancer (1). Cisplatin binds to the DNA template to form both intra‐ and inter‐strand cross‐links (ICL) (2–4). Recognition of this ICL damage can eventually lead to apoptotic cell death, which is an important mechanism for cisplatin in its anticancer action. Despite its great efficiency in the treatment of certain types of cancer, cisplatin and its analogs have been plagued by major problems of side effects and the presence of cancer cell resistance to these drugs. These factors have significantly undermined their curative potentials (5). To overcome these problems, considerable efforts have been directed at mechanistic studies of their anticancer activity (6–9). However, the mechanism of cisplatin in anticancer activity is still not fully understood.

XPC is an important DNA damage recognition protein involved in the DNA nucleotide excision repair (NER) process (10,11). The XPC protein recognizes a variety of bulky DNA damage, including UV irradiation‐generated cyclobutane pyrimidine dimers and cisplatin‐caused ICLs (12–21). Inside cells, the XPC protein binds tightly with an HR23B protein to form a stable XPC‐HR23B complex (22–25). The XPC protein also interacts with several other important protein components including the basal transcription factor TFIIH and the centrisomal protein CEN2 (23,24,26–28). Recent studies indicate that the XPC‐HR23B complex is the first protein component that recognizes and binds to the damaged sites (29–31). Importantly, XPC defects have been associated with many types of cancer (32–35). XPC gene knockout transgenic mice studies also reveal the predisposition of many types of cancer (36–38). It is possible that the XPC protein plays an important role in DNA damage‐mediated apoptotic cell death and cancer prevention. However, no studies have been done at this time to directly investigate the roles of the XPC protein in these aspects.

In this work, we have studied the role of the XPC protein in cisplatin DNA damaging treatment‐mediated cellular response. Comparison of the microarray data obtained from human normal fibroblast (NF) cells and two individual XPC‐defective cell lines (XPC cells) revealed that 486 genes displayed altered transcription responses in both XPC cell lines with the cisplatin treatment (with P ≤ 0.01 and a minimal 1.5‐fold change). Among these genes, 297 genes were further mapped to biological pathways and gene ontologies. The genes related to cell cycle and cell proliferation were affected by the XPC defects in the cisplatin treatment to a greater extent than those of other cellular functions. To elucidate the results obtained from the microarray data, the cisplatin treatment‐mediated signal transduction process and the ability to activate caspase‐3, a key caspase in causing apoptosis, were further studied using western blot hybridization assays. The cisplatin treatment‐mediated p53 response was greatly attenuated in the XPC cells. The cisplatin treatment‐mediated activation of the caspase‐3 was also significantly reduced in the XPC cells. All these results suggest that the XPC protein plays a critical role in initiating the cisplatin DNA damaging treatment‐mediated signal transduction process by activating the p53 pathway and causing cell cycle arrest, which allows for DNA repair and apoptosis; and that the XPC defects cause a failure in DNA damage recognition, resulting in damaged cells escaping from the cell cycle arrest and an increase in the risk of mutation accumulation and cancer occurrence.

MATERIALS AND METHODS

Cell lines and plasmid

The human NF cells (GM00043 and GM00323) and xeroderma pigmentosum group C (XPC) cells (GM16684, GM02096, and GM00671) were obtained from the NIGMS Human Genetic Cell Repository (Corriel Institute, Camden, NJ). The GM16684 XPC cells carry a 2 bp deletion in exon 5 of the XPC gene (del AT 669–670), which results in a truncated XPC protein with the N‐terminal ∼220 amino acid residues. The GM02096 XPC cells carry a point mutation in the XPC gene that converts the proline at position 218 to histidine. The GM00671 XPC cells carry two mutations in the XPC gene: one results in the insertion of a valine residue after the valine at codon 580, while the other is a point mutation that created a nonconservative amino acid change near the C‐terminus of the protein. Both the NF and XPC cells were maintained in MEM medium supplemented with 20% FBS, 2× essential amino acids, nonessential amino acids, and vitamins.

A pDsRed‐XPC plasmid was constructed for this study. The pXPC3 plasmid, which carries the XPC gene cDNA (10), was digested with XhoI restriction enzyme to obtain a 3.2 kb DNA fragment containing the XPC gene cDNA. This 3.2 kb DNA fragment was inserted into the XhoI site of pDsRed2‐C1 (BE Clontech, Palo Alto, CA) with the correct orientation to obtain the pDsRed‐XPC plasmid.

Cisplatin treatment and RNA preparation

Both the NF and XPC cells were grown in cell growth medium at 37°C (with 5% CO2) to ∼70% confluence. Cisplatin (Sigma Corp., St Louis, MO) was freshly dissolved into dimethyl sulfoxide (DMSO**)** (20 mg/ml) and added to the cell culture medium to a final concentration of 10 µM. The cells were incubated at 37°C for 16 h and harvested from the flasks. Total RNA was isolated from the treated cells using a Qiagen RNeasy Purification System (Qiagen, Valencia, CA). RNA isolated from three individual treatments was mixed. As a control, total RNA was also isolated from NF and XPC cells that were treated with DMSO alone using a similar method. The quality of the isolated RNA was determined by an Agilent 2100 Bioanalyzer (Agilent Technologies, Wilmington, DE).

The microarray assay and data analysis

The RNA isolated from both untreated and cisplatin‐treated NF and XPC cells was individually labeled with a Cy‐5 fluorescence in a reverse transcription (RT) reaction (42°C for 1 h) to generate fluorescence‐labeled cDNA probes. A Universal Human RNA Reference Sample (Stratagene, La Jolla, CA) was labeled with a Cy‐3 fluorescence in a similar reaction. The Cy‐3 and Cy‐5 labeled cDNA probes were mixed together and cleaned using a Qiaquick PCR Purification Kit (Qiagen). The fluorescence‐labeled cDNA probes were added to the Agilent Human 1 cDNA Microarray slides, which contained 12 814 unique human cDNA clones (Agilent Technologies), for a competitive hybridization (65°C for 17 h). Each RNA sample was hybridized with three microarray slides. The slides were scanned using an Agilent two‐color laser detection scanner. The two‐color image was feature‐extracted using Agilent’s Feature Extraction software. The fluorescent intensities were normalized using low ESS intensity‐dependent normalization.

The image obtained from the two‐color scanner was analyzed by Significance Analysis of Microarray Data (SAM) software (developed at Stanford University) to identify the genes with significant alterations in their transcription levels. Within each cell line, genes differentially expressed in response to the treatment were identified using the replicate microarray data and a permutation‐based statistical test with an estimated false discovery rate of 5% (39). The genes that were affected by the XPC protein DNA damage recognition signal were further determined by the net fold change (ΔΔ) of the same gene between the XPC and the NF cells using an analysis of variance (ANOVA) with a standard parametric two‐way ANOVA with the _P_‐value ≤0.01 and the net fold change ≥1.5. The genes that were identified as statistically significant with the ANOVA were further analyzed using EASE software (GO Consortium) to categorize them into biological pathways and gene ontologies.

Real‐time PCR

The RT reaction was carried out using 2 µg of total RNA following the protocol for the Taqman Reverse Transcription Master Mix (Applied Biosystems, Foster City, CA). A primer optimization step is tested for each set of primers to determine the optimal primer concentrations. Once the optimal primer concentrations are determined, 10 ng of cDNA sample was used for the PCR and analyzed using the ABI Prism 7000 Sequence Detection System (Applied Biosystems). Sybr Green was used as a detector for monitoring the amplified double‐stranded DNA fragments. Cycle threshold (Ct) values were obtained from the ABI Prism 7000 software and the fold change was determined. As a control, the mRNA level of the GAPDH gene was also determined in the real‐time PCR assay for each RNA sample and used to minimize any experimental variations.

Western blot hybridization assay

The p53 (Pab1801) and p21 (c‐19) antibodies were purchased from Santa Cruz Biotechnologies (Santa Cruz, CA). The Phospho‐p53 Antibody Sampler Kit, which contains seven individual antibodies that recognize specific phosphorylation sites at the Ser6, Ser9, Ser15, Ser20, Ser37, Ser46 and Ser392 of the p53 protein, was purchased from Cell Signaling Technology, Inc. (Beverly, MA). The caspase‐3 antibody (H‐277) was purchased from Santa Cruz Biotechnologies. The NF and XPC cells were either treated with various concentrations of cisplatin (0, 10, 20 and 40 µM) for 24 h or treated with 20 µM cisplatin for various time points. The cells were harvested and lysed in RIPA cell lysis buffer (1× PBS, 1% NP‐40, 0.5% deoxycholic acid, 0.1% SDS). The cell lyses (30 µg of total protein) was mixed in 20 µl of solution containing 1× SDS gel loading buffer (60 mM Tris, pH 6.8, 1% SDS, 1% 2‐mercaptoethanol, 10% glycerol and 0.1% bromophenol blue) and boiled for 10 min. The cell lyses were analyzed by SDS–PAGE with a 12% gel. The proteins were transferred to a PVDF membrane and then hybridized with the desired primary and secondary antibodies. ECL reagent (Amersham, Arlington Heights, IL) was used to detect specific proteins in the membrane. The same membrane was then stripped in a stripping solution (62.5 mM Tris, pH 6.8, 2% SDS, 0.7% 2‐mercaptoethanol) and hybridized with β‐actin antibody to determine the level of β‐actin in each sample. The level of the desired proteins was calculated to the relative level of β‐actin in each sample to minimize experimental variations.

Transient transfection of the XPC cells with pDsRed‐XPC plasmid

The GM02096 XPC cells were seeded onto 100 mm tissue culture dishes and incubated in a tissue culture incubator to 60% confluence. The cells were then transfected with the pDsRed‐XPC plasmid using a cationic lipid (Superfect Transfection Reagent; Qiagen) (20 µg of plasmid DNA/50 µl of Superfect Reagent/100 mm dish) for 6 h. The medium was replaced with fresh cell growth medium and the cells were incubated at 37°C overnight. The cells were then treated with cisplatin at various concentrations (0–40 µM) for 24 h and then harvested and analyzed by western blot hybridization assay to determine the protein levels of p53, phosphorylated p53 and p21 in each sample.

RESULTS

Rationale and reliability of the microarray assay

To determine the role of the XPC protein in cisplatin DNA damaging treatment‐mediated cellular response, it is important to determine the level of alteration for all known genes after the treatment. The microarray assay provides a great opportunity for determining the alteration in transcription levels for thousands of genes in a single experiment. We have taken advantage of this assay to determine the genes that are affected by the XPC protein DNA damage recognition signal in the treatment. However, the microarray assay has some disadvantages. One of the most important concerns is its reliability. To minimize the experimental variations, the cells were treated in several independent experiments and the RNA isolated from these experiments were mixed and used as a template for the microarray assay. The quality of the isolated RNA samples was carefully determined using an Agilent 2100 Bioanalyzer before being applied to the microarray assay. To reduce the technical variations, each RNA sample was hybridized with three arrays and only data that was consistent in at least two of the three arrays was used in our data analysis. We also used a permutation‐based statistical test for data management and analysis to reduce the false discovery rate (39). With these carefully designed experiments, satisfactory results were obtained from our microarray experiments. Out of the total of 18 564 spots contained in the microarray, valuable data were extracted from 94 to 98% of the outliers in both the NF and XPC RNA samples.

Identification of the XPC‐responsive genes in the cisplatin treatment

Since the results obtained from our microarray assay revealed a high level of reproducibility, we further determined the genes that are affected by the XPC defect in the cisplatin treatment. The data obtained from the microarray assay were analyzed first by SAM to identify the genes with significant alteration in their transcription levels (Δ) in both the NF and XPC cells with an estimated 5% false discovery rate. The genes that are affected by the XPC defect were then determined in each XPC cell line by comparison of the altered genes between the NF and XPC cells with a minimal 1.5‐fold net change (ΔΔ) between the NF and the XPC cells with the treatment. The commonly affected genes were further determined between the XPC cell line GM02096 and GM16684. A total of 486 common genes were identified in both XPC cell lines with the cisplatin treatment. The correlation of the net fold change among these genes was 0.88, suggesting the high specificity of the genes affected by the individual XPC defects in the treatment (Fig. 1). These genes were further mapped to biological pathways and gene ontologies using the EASE analysis. Out of the total of 486 common genes identified in both XPC cell lines, 297 genes were mapped to known biological pathways and gene ontologies (Table 1) (40). These genes were designated as XPC‐responsive genes.

Determination of gene mRNA level by an RT–PCR based real‐time PCR assay

The results obtained from the microarray data analysis revealed the altered transcriptions of many important genes by the XPC defect in the cisplatin treatment. To validate the microarray data, we further determined the transcription levels in some of the chosen genes in both the NF (GM00043) and XPC (GM16684) cells using an RT–PCR based real‐time PCR assay (Table 2). The genes that were tested by the real‐time PCR assay included cdk1, cdk2, cycA2, cycD2, cycE1, cip1, IAP1, mlh1, msh2, PCNA, PLK, PPM1D and xpc. In the 14 genes tested by the real‐time PCR assay, 12 genes showed similar transcription alteration patterns as determined by the microarray assay, suggesting a very high reliability of the microarray data obtained in this study. Two of the tested genes, cycD2 and PLK, showed different transcription alterations between the real‐time PCR and the microarray assays. The difference observed in both assays might reflect the difference in DNA sequences detected by these two assays. Since the microarray assay uses cDNA as probes and relatively long sequences are targeted, homologous sequences and splicing variants of the same gene mRNA may be detected in this assay. In contrast, the real‐time PCR assay only uses short oligonucleotides and only the templates containing the oligonucleotide binding sequences will be detected by this assay.

The role of the XPC protein in the cisplatin DNA damaging treatment‐mediated cell cycle regulation

The results obtained from the EASE data analysis revealed that the genes involved in the cell cycle and cell proliferation‐related cellular functions scored at the top of the list (Table 1), suggesting a very important role of the XPC protein in the cisplatin DNA damaging treatment‐mediated cell cycle regulation and apoptosis. To further study the role of the XPC protein in these processes, we analyzed the gene expression profile of the cell cycle‐related genes (Table 3 and Fig. 2). The transcription levels of many cell cycle‐related genes were significantly altered by the XPC defects in the treatment (Table 3). Several cell cycle‐related genes, including cdc7, cip1 (p21), culL3, G0S2, GADD45, mcm7, psen1, sesn1, stat1 and tsc1, were up‐regulated in the cisplatin‐treated NF cells; the degree of this up‐regulation was attenuated in both XPC cell lines (GM16684 and GM02096) with the treatment. Several other genes, including cdc2, fancg, myc, ppp5C, stk6 and ttk, were down‐regulated in the cisplatin‐treated NF cells; the degrees of down‐regulation of these genes, however, were reduced in both of the cisplatin‐treated XPC cells. These results suggest that the XPC protein plays a very important role in the cisplatin DNA damaging treatment‐mediated cell cycle regulation and that the XPC defects result in a failure of the cell cycle regulation in damaged cells.

The genes involved in the apoptotic cell death process were also analyzed with the microarray data (Table 4). Indeed, the transcription levels for several important apoptosis‐related genes were significantly affected by the XPC defect in the cisplatin treatment. In the NF cells, the cisplatin treatment led to a significant increase in the transcription levels for several important apoptosis‐related genes including cip1 (p21), casp3, casp4, psen1 and stat1. In contrast, the transcription levels of these genes in the cisplatin‐treated XPC cells either remained unchanged or displayed only low levels of increase. These results suggest that the XPC protein DNA damage recognition signal is also involved in the cisplatin DNA damaging treatment‐mediated apoptosis process.

The role of the XPC protein in the cisplatin DNA damaging treatment‐mediated DNA repair

Because of the important function of the XPC protein DNA damage recognition in initiation of the DNA repair process, we also determined the DNA repair genes that were affected by the XPC defects in the cisplatin treatment (Table 5). The transcription levels of many important DNA repair‐related genes were affected by the XPC defects in the cisplatin treatment. For example, the transcription levels of the DNA repair genes ADPRTL1, GADD45A and ube2B were significantly increased when the NF cells were treated with cisplatin. In contrast, the transcription levels of these genes in the cisplatin‐treated XPC cells either remained unchanged or displayed only very low levels of increase. Many other important DNA repair genes, including fancg, lig1, mlh1, pol δ_1_, rad51 and rpa3, displayed significant decreases in their transcription levels when the NF cells were treated with cisplatin. In comparison, the transcription levels of these genes either remained unchanged or showed only small decreases in the cisplatin‐treated XPC cells. These results suggest that the XPC protein DNA damage recognition signal plays a very important role in the cisplatin treatment‐mediated DNA repair process.

The XPC defects correlate with attenuated p53 and p21 responses to the cisplatin treatment

Although the microarray results suggest that the XPC protein plays an important role in the cisplatin DNA damaging treatment‐mediated signaling process, the signal transduction pathway utilized in this process is unclear. The altered cip1 (p21) transcription response from the XPC defect in the cisplatin treatment suggests that the p53 signal transduction pathway is likely to be involved in the process. To test this possibility, a western blot hybridization assay was performed to determine the levels of both the p53 and p21 proteins in NF and XPC cells that were treated with various concentrations of cisplatin (Fig. 3A and B, top). The cisplatin treatment led to an increase in the levels of p53 and p21 protein in the NF cells. In contrast, the p53 and p21 responses in the cisplatin‐treated XPC cells were attenuated and the changes were substantially less (Fig. 3A and B, top). When pDsRed‐XPC, an expression vector that carries a CMV promoter‐driven XPC gene cDNA, was transiently transfected into a XPC cell line, however, the cisplatin treatment‐mediated p53 and p21 responses were re‐established (Fig. 3A and B, top). As an internal control, the level of β‐actin remained unchanged in both the NF and XPC cells with the treatment (Fig. 3A and B, bottom).

To determine if p53 phosphorylation is involved in the XPC protein DNA damage recognition‐mediated signal transduction process, we further determined the level of the phosphorylated p53 protein in the cisplatin‐treated NF and XPC cells using pooled phospho‐p53 antibodies that recognize seven known serine phosphorylation sites in the p53 protein (Ser6, Ser9, Ser15, Ser20, Ser37, Ser46 and Ser392) (Fig. 3C). Indeed, the cisplatin treatment led to a dose‐dependent increase in the phosphorylated p53 protein in the NF cells. In contrast, the level of the phosphorylated p53 protein was attenuated in the cisplatin‐treated XPC cells. Again, once the pDsRed‐XPC plasmid was transiently transfected in a XPC cell line (GM00671), the p53 phosphorylation response was re‐established. Further studies revealed that both the Ser15 and Ser392 of the p53 protein were phosphorylated while the other tested sites (Ser6, Ser9, Ser20, Ser37 and Ser46) remained unphosphorylated in the cisplatin‐treated NF cells (data not shown).

A time course experiment was also performed to study the kinetics of p53 and p21 response to the cisplatin treatment in both the NF and XPC cells (Fig. 4). When the NF cells were treated with cisplatin for up to 8 h, the level of p53 protein remained unchanged (Fig. 4A). When the NF cells were treated with cisplatin for 16 h or longer, the level of p53 protein was significantly increased (Fig. 4A). Similar patterns were also seen in the phosphorylated p53 protein and the p21 protein in the cisplatin‐treated NF cells (Fig. 4B and C). In contrast, the levels of increase for both the p53 and the phosphorylated p53 proteins were attenuated in the cisplatin‐treated XPC cells (Fig. 4A–C). As an internal control, the level of β‐actin remained unchanged during the treatment (Fig. 4D).

The XPC defect correlates with attenuated caspase‐3 activation in the cisplatin treatment

The results obtained from the microarray data strongly suggest that the XPC protein plays an important role in initiating the cisplatin DNA damage‐mediated signal transduction process, resulting in DNA repair and apoptosis. To further validate the microarray data, we also determined the level of active caspase‐3 (19 and 17 kDa), a key caspase that directly cleaves other proteins and causes apoptosis, in the cisplatin‐treated NF and XPC cells (Fig. 5). Activation of caspase‐3 was observed when the NF cells were treated with cisplatin at 40 µM; in contrast, a much lower level of the active caspase‐3 was seen when the XPC cells were treated with cisplatin at the same concentration. As an internal control, the level of β‐actin remains unchanged in the cisplatin treatment. These results suggest that a functional XPC protein is required for the cisplatin treatment‐mediated apoptotic cell death.

DISCUSSION

In this study, we have determined the role of the XPC protein in the cisplatin DNA damaging treatment‐mediated cellular response. Comparison of the microarray data between the NF and the two individual XPC cell lines identified 485 common genes with significant alterations in their transcription levels in both of the XPC cell lines with the cisplatin treatment (with P ≤0.01 and a net fold change ≥1.5). The EASE analysis mapped 297 of these genes to known biological pathways and gene ontologies. To validate the microarray data, we further determined the transcription levels of some chosen genes using a real‐time PCR assay. The results obtained from the real‐time PCR assay are consistent with the microarray data in most of the tested genes (12 of 14 genes). These results suggested a high reliability of the microarray data obtained in our studies.

The EASE data analysis suggests that the genes related to cell cycle and cell proliferation were affected to a greater degree by the XPC defect than genes related to other cellular functions in the cisplatin treatment, suggesting the important role of the XPC protein in the cisplatin DNA damaging treatment‐mediated cell cycle arrest and apoptosis process. A further study was conducted to validate this observation. The result obtained from the western blot hybridization assay indicated that the XPC defect attenuated the cisplatin treatment‐mediated caspase‐3 activation, suggesting a critical role of the XPC protein in initiating the cisplatin DNA damaging treatment‐mediated signal transduction process and the subsequent apoptotic cell death.

We have studied the XPC protein DNA damage recognition‐mediated signal transduction process. The results obtained from our western blot hybridization assays suggest that the p53 signal transduction pathway is involved in the XPC protein DNA damage recognition‐mediated signal transduction process. This provides an important understanding for the mechanism of bulky DNA damage‐mediated cell cycle regulation. Although many DNA damaging reagents, including many commonly used anticancer drugs, can generate bulky DNA damage and cause cell death, the proteins that recognize the bulky DNA damage and initiate the signaling process have not been identified. The results obtained from this study suggest that the XPC protein DNA damage recognition plays a critical role in initiation of the DNA damage‐mediated signal transduction process. It is possible that the DNA damage recognition and binding by the XPC protein affects the interactions of XPC protein with other protein components such as the complex formation of XPC‐TFIIH, which results in enhanced interaction of the TFIIH with the p53 protein (41) and causes phosphorylation of the p53 protein by CDK7 (42,43), leading to subsequent activation of the p53 signal transduction pathway. In addition, a recent study published by Zou and Elledge suggests that the single‐stranded DNA is the activating signal for DNA damage‐induced signaling process (44). Another possibility is that the XPC protein initiates the DNA damage‐mediated signaling process by recognizing the DNA damage and recruiting other DNA repair proteins, including the TFIIH, to the damaged site. Once the XPC‐TFIIH complex forms, the helicase activities carried in the TFIIH cause the DNA strands to unwind, generating the single‐stranded DNA regions around the damaged site and resulting in activation of the DNA damage‐mediated signal transduction process. Further studies are needed to determine how the XPC protein DNA damage recognition signal leads to activation of the p53 signaling pathway.

The work published by others demonstrates the regulation of the XPC gene by the p53 protein in the DNA damaging treatment (21,45–47). The results obtained in this study suggest the involvement of the XPC protein in initiating the cisplatin DNA damaging treatment‐mediated signal transduction process. It is possible that both the XPC and p53 proteins can regulate each other in response to the DNA damage treatment. Since the XPC‐HR23B complex is the first protein component that recognizes and binds to the damaged sites, it is likely that the DNA damage recognition signal caused by the XPC protein will first activate the p53 protein. In response, the active p53 protein then regulates the transcription of many DNA damage‐responsive genes, including the XPC gene, resulting in cell cycle arrest and subsequent DNA repair and apoptosis.

XPC defects have been associated with many types of cancer (32–35). XPC gene knockout mice studies also revealed the predisposition of many types of cancer (36–38). The results obtained from our recent mutagenesis study demonstrated that the XPC defect caused an increase in mutation accumulation with DNA damaging treatment (19). The results of this study suggest that the XPC protein is involved in the cisplatin DNA damage‐mediated signaling process. All these results suggest that the XPC protein plays an important role in eliminating damaged cells and in preventing cancer occurrence and that XPC defects can lead to a high risk of cancer occurrence. Therefore, the results obtained from this study provide an important understanding for the molecular mechanism of cancer occurrence and the role of the XPC protein in cancer prevention. In addition, these results provide important insights into the mechanism of cancer cell drug resistance. Therefore, the XPC protein may provide an important biomarker for cancer prevention, diagnosis and treatment.

SUPPLEMENTARY MATERIAL

Supplementary Material is available at NAR Online.

ACKNOWLEDGEMENTS

We thank Dr R. Legerski for providing us the pXPC‐3 plasmid, which was used to construct the pDsRed‐XPC plasmid in our complementation study. We thank Drs R. Novak and P. Stemmer for their helpful discussions and critical reading of this manuscript and R. Paxton for his critical reading of this manuscript. Performance of this work was facilitated by the Microarray and Bioinformatic Core, the Cell Culture Core, and the Imaging and Flow Cytometry Core of the Environmental Health Sciences Center in Molecular and Cellular Toxicology with Human Applications at Wayne State University (P30 ES06639). This work is supported by grant R01ES09699 from the National Institute of Environmental Health Sciences, NIH to G.W. and a pilot project from the Environmental Health Sciences Center at Wayne State University (P30 ES06639).

Figure 1. Correlation of the common genes affected by the XPC defects in the cisplatin treatment determined by microarray assay. The common genes with a net fold change ≥1.5 in both the GM16684 and GM02096 XPC cells are compared. The _x_‐axis shows the net fold change of the genes in GM16684 XPC cells and the _y_‐axis shows the net fold change of the genes in GM02096 XPC cells. Each dot represents one common gene. The correlation of the net fold change among these genes was 0.88 between these two XPC cell lines.

Figure 2. The effect of XPC defects on transcription of the cell cycle‐related genes with the cisplatin treatment. The common genes identified in both the GM16684 and GM02096 XPC cell lines with a net fold change ≥1.5 are color‐mapped. If the ΔΔ > 0, the gene is counted as up‐regulated and colored red in the map. If the ΔΔ < 0, the gene is counted as down‐regulated and colored green in the map. If the ΔΔ < 1.5, no data are available from the microarray analysis, or the net fold change ≥1.5 is presented in only one of the two XPC cell lines, the gene is uncolored in the map.

Figure 3. The cisplatin treatment‐mediated p53 and p21 responses in both NF and XPC cells. The NF (GM00043), XPC (GM16684 and GM00671) and XPC gene cDNA transiently transfected GM00671 XPC cells were treated with cisplatin at various concentrations for 24 h. The cell lyses were analyzed by western blot hybridization assay. (A) The level of p53 protein in the cisplatin‐treated NF and XPC cells. (B) The level of p21 protein in the cisplatin‐treated NF and XPC cells. (C) The level of p53 protein phosphorylation in the cisplatin‐treated NF and XPC cells.

Figure 4. Detection of the p53 and p21 proteins in both the NF and XPC cells with the cisplatin treatment. Both the NF (GM00043) and XPC (GM16684) cells were treated with cisplatin (20 µM) for various lengths of times. The cell lyses were analyzed by western blot hybridization assay for the levels of p53, phosphorylated p53, p21 and β‐actin proteins. (A) The level of p53 protein during the cisplatin treatment. (B) The level of the phosphorylated p53 protein during the cisplatin treatment. (C) The level of the p21 protein during the cisplatin treatment. (D) The level of the β‐actin protein during the cisplatin treatment. All the protein levels were detected with the same western blot membrane.

Figure 5. Detection of caspase‐3 activation in both the NF and XPC cells with the cisplatin treatment. Both the GM00043 NF cell and GM16684 XPC cells were treated with 0, 10, 20, and 40 µM cisplatin for 24 h. The cell lyses were analyzed by western blot hybridization assay for detection of the active caspase‐3 (19 and 17 kDa) (top). Some unspecific protein bands were cross‐reacted with the caspase‐3 polyclonal antibody. The bands specific for the caspase‐3 are indicated by arrows. The same membrane was also used for detection of the level of β‐actin (bottom).

Table 1.

Characterization of some of the common genes affected by the XPC defects in the cisplatin treatmenta

aCharacterization of whole list of the common genes are provided as Supplementary Material.

Table 1.

Characterization of some of the common genes affected by the XPC defects in the cisplatin treatmenta

aCharacterization of whole list of the common genes are provided as Supplementary Material.

Table 2.

Determination of the gene transcription alteration in both the NF and XPC cells by the real‐time PCR assay

Table 2.

Determination of the gene transcription alteration in both the NF and XPC cells by the real‐time PCR assay

Table 3.

The common cell cycle‐related genes that were affected by the XPC defects in the cisplatin treatment

Table 3.

The common cell cycle‐related genes that were affected by the XPC defects in the cisplatin treatment

Table 4.

The common apoptosis‐related genes that were affected by the XPC defects in the cisplatin treatment

Table 4.

The common apoptosis‐related genes that were affected by the XPC defects in the cisplatin treatment

Table 5.

The common DNA repair‐related genes that were affected by the XPC defects in the cisplatin treatment

Table 5.

The common DNA repair‐related genes that were affected by the XPC defects in the cisplatin treatment

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