Promoter CpG hypermethylation and downregulation of XAF1 expression in human urogenital malignancies: implication for attenuated p53 response to apoptotic stresses

Abstract

XIAP-associated factor 1 (XAF1) is a new candidate tumor suppressor, which has been known to exert proapoptotic effects by interfering with the caspase-inhibiting activity of XIAP. To explore the XAF1's candidacy for a suppressor in urogenital tumorigenesis, we investigated the XAF1 status in a series of cancer cell lines and primary tumors derived from the bladder, kidney and prostate. Expression of XAF1 transcript was undetectable or extremely low in 60% (3/5) of bladder, 66% (10/15) of kidney, and 100% (3/3) prostate cancer cell lines. Abnormal reduction of XAF1 was also found in 33% (18/55) of primary bladder and 40% (8/20) of primary kidney tumors, and showed a correlation with advanced stage and high grade of bladder tumor. Hypermethylation at 14 CpG sites in the 5′ proximal region of the XAF1 promoter was highly prevalent in cancers versus adjacent normal or benign tissues and tightly associated with reduced gene expression. XAF1 expression enhanced the apoptotic response of tumor cells to chemotherapeutic agents, such as etoposide or 5-FU. While XAF1 expression did not influence the subcellular distribution or expression of XIAP, it elevated the protein stability of p53 and its target gene expression. Moreover, the apoptosis-sensitizing and growth suppression function of XAF1 was markedly impeded by blockade of p53 function. Collectively, our study demonstrates that epigenetic alteration of XAF1 is frequent in human urogenital cancers and may contribute to the malignant progression of tumors by rendering tumor cells a survival advantage partially through the attenuated p53 response to apoptotic stresses.

Introduction

Apoptosis plays an essential role for elimination of defective or potentially dangerous cells and provides a defense against malignant transformation (Thompson, 1995). The inhibitor of apoptosis proteins (IAPs) are a family of intrinsic cellular antiapoptotic proteins (Deveraux and Reed, 1999; Sun et al., 1999). The human IAP family includes cIAP-1, cIAP-2, XIAP, NAIP, survivin, apollon, ILP2, and livin, and several members of human IAP family, including XIAP, c-IAP-1 and c-IAP-2, have been shown to prevent apoptosis by binding to caspase-3, -7 and -9 and thereby protecting them from cleavage activation (Deveraux et al, 1997; Roy et al., 1997; Green, 2000). Deregulation of IAPs plays a key role in the aberrantly increased cell viability and resistance to the anticancer therapy in human cancers whereas overexpression appears to suppress apoptosis against a large variety of triggers including tumor necrosis factor (TNF), Fas, staurosporin, etoposide and growth factor withdrawal (Duckett et al., 1996; Liston et al., 1996; Ambrosini et al., 1997; Li et al., 1998). Of the known human IAP proteins, XIAP is the most potent and versatile inhibitor of caspases and apoptosis. XIAP mRNA levels are relatively high in the majority of cancer cell lines and high levels of XIAP protein are generally found in human cancers including bladder and renal cell carcinomas (Fong et al., 2000; Tamm et al., 2001; Bilim et al., 2003). XIAP protects tumor cells from apoptosis induced by many triggers, including irradiation, serum withdrawal and chemotherapeutic compounds (Holcik et al., 2001). It has been shown that XIAP regulates Akt activity and caspase-3-dependent cleavage, and XIAP downregulation induces apoptosis in chemoresistant human ovarian cancer cells (Sasaki et al., 2000; Asselin et al., 2001).

The caspase-inhibiting activity of XIAP can be suppressed by two mitochondrial proteins, Smac/DIABLO and HtrA2, which are released from mitochondria into the cytoplasm during apoptosis, where they bind to the hydrophobic pocket in BIR3 of XIAP (Du et al., 2000; Verhagen et al., 2000; Suzuki et al., 2001). Recently, XIAP-associated factor 1 (XAF1), which is located at 17p13.2, was identified as a novel negative regulator of XIAP based on its ability to bind XIAP (Liston et al., 2001). XAF1 encodes 33.1 kDa protein with seven zinc fingers and its mRNA is ubiquitously expressed in all normal adult and fetal tissues, but absent or present at very low or undetectable levels in various cancer cell lines. XAF1 sensitizes tumor cells to the proapototic effects of etoposide and prevents the XIAP-mediated inhibition of caspase-3. Moreover, the antiapoptotic ability of XIAP in serum withdrawal or etoposide-induced apoptosis was restored in tumor cells by infection of adeno-XAF1 antisense. These observations thus suggest that deregulation of apoptosis through the loss of XAF1 expression might be important for malignant cell survival, and a high level of XIAP to XAF1 expression in cancer cells may provide a survival advantage through the relative increase of XIAP antiapoptotic function.

XAF1 was originally reported as a nuclear protein to exert its proapoptotic effect by directly interacting with XIAP and inducing XIAP sequestration in nuclear inclusions (Liston et al., 2001). A recent tissue microarray assay showed a significant reduction of XAF1 expression in 15 of 16 human melanoma cell lines and a large fraction of primary melanomas (Ng et al., 2004). While high level of XAF1 was detected in both nucleus and cytoplasm in normal melanocyte, the percentage of XAF1 positive cells and the staining intensity of XAF1 were significantly decreased in a majority of melanoma tissues compared to benign melanocytic nevi. Given the fact that XAF1 could trigger the relocalization of XIAP in the cytoplasm, it was proposed that loss of XAF1 expression in tumor may increase the functional pool of cytoplasmic XIAP, which in turn deregulate the normal apoptotic process and contribute to the malignant nature of tumors (Ng et al., 2004). Interestingly, it has been reported that the natural resistance of the adult motoneurons to axotomy is abolished when the ratio of XAF1 to XIAP is increased, and that XAF1 is increased and redistributed during reperfusion after focal cerebral ischemia in rats, which is accompanied by diffuse redistribution of XIAP and activation of caspase-3 (Perrelet et al., 2004; Siegelin et al., 2005). Therefore, the balance between XIAP and XAF1 may play an essential role for adult motoneuron protection and implicated in the pathophysiological mechanisms of reperfusion injury as well as tumorigenesis.

In the present study, we demonstrates that XAF1 expression is absent or significantly downregulated in a substantial fraction of urogenital malignancies by aberrant promoter CpG sites hypermethylation, and that abnormal reduction of XAF1 correlates with advanced stage and high grade of bladder tumor. Our data also show that XAF1 expression greatly increases tumor cell sensitivity to apoptosis triggered by chemotherapeutic drugs, suggesting that epigenetic inactivation of XAF1 might provide a selective advantage for tumor cell survival and thus contribute to the malignant progression of human urogenital cancers.

Results

Altered expression of XAF1 in human urogenital cancer cell lines and primary tumors

To investigate the candidacy of XAF1 as a suppressor in urogenital tumorigenesis, we initially characterized the expression status of XAF1 transcript in five bladder, 15 kidney and three prostate cancer cell lines. XAF1 expression was undetectable or very low in three of five (60%) bladder (T24, HT1197 and 253J/BV), 10 of 15 (66%) kidney (UOK107, UOK108, UOK109, UOK112, UOK115, UOK122, UOK123, UOK130, ACHN and WWC1), and all of three (100%) prostate (LNCaP, DU145 and PC-3) cancer cell lines (Figure 1a). By contrast, Smac/DIABLO and HtrA2, two other XIAP antagonists showed easily detectable expression in all cell lines tested. Expression levels (Target/GAPDH) of XAF1, Smac/DIABLO, and HtrA2 in cancer cell lines were observed within the range of 0.00–1.40 (mean; 0.68), 1.04–1.36 (mean; 1.18) and 0.82–1.22 (mean; 0.98), respectively (Figure 1b). XAF1 levels determined by semiquantitative RT–PCR were fairly consistent with the results from Northern blot assay (Figure 1a). It has been suggested that a high level of XIAP to XAF1 due to either elevated expression of XIAP or reduction of XAF1 may provide a survival advantage for tumor cells through the relative increase of XIAP antiapoptotic function (Liston et al., 2001). However, in contrast to XAF1, the majority of urogenital tumor cells we tested exhibited similar expression of XIAP at both mRNA and protein levels (Figure 1c).

Figure 1
figure1

Expression status of XAF1, Smac/DIABLO, HtrA2 and XIAP in human bladder, kidney and prostate cancer cell lines. (a) Semiquantitative RT-PCR and Northern blot analysis of XAF1, Smac/DIABLO, and HtrA2 expression in cancer cell lines. (b) Expression levels of XAF1, Smac/DIABLO, and HtrA2 transcripts in cancer cell lines. Quantitation was achieved by densitometric scanning of RT-PCR products in ethidium bromide-stained gels and absolute area integrations of the curves representing each specimen were compared after adjustment for GAPDH. Data represent means of triplicate assays. (c) XIAP expression in urogenital cancer cell lines. Expression of XIAP mRNA was analysed by semiquantitative RT-PCR. For Western blot assay of XIAP protein expression, 30 μg of total protein was fractionated using 10% SDS–PAGE and XIAP was detected using an anti-XIAP monoclonal antibody and enhanced chemiluminescence.

Next, we evaluated XAF1 expression in 55 bladder transitional cell carcinoma (TCC) and 20 renal cell carcinoma (RCC) tissues. In both tumor types, a substantial fraction of tumors showed markedly reduced XAF1 expression (mean; 0.74 and 0.64 in TCC and RCC, respectively) compared to its noncancerous counterparts (mean; 1.20 and 1.16 in bladder and kidney, respectively) (P<0.005) (Figure 2a and b). In matched tissue sets examined, 50% (10 of 20) of TCC and 60% (12 of 20) of RCC cases revealed tumor-specific reduction of XAF1 (Figure 2c). We arbitrarily classified tumors with expression levels less than a half (<0.60 in TCC; <0.58 in RCC) of normal means as abnormally low expressors. Abnormal reduction of XAF1 was identified in 33% (18 of 55) of TCC and 35% (seven of 20) of RCC (Figure 2b). In bladder tumor, abnormal reduction of XAF1 was significantly higher in muscle-invasive tumors (T2–T4, 13 of 28 (46%)) compared to superficial tumors (Ta–T1, five of 27 (19%)) (P<0.001), and more frequent in high grade tumors (grade II–III, 14 of 37 (38%)) than low grade tumors (grade I, four of 18 (22%)) (P<0.01) (Figure 2d). However, XAF1 expression showed no correlation with morphologic patterns of tumors (papillary, 14 of 44 (32%); non-papillary, four of 11 (36%)), age (<40, two of six (33%); 41–60, five of 16 (31%); >61, 11 of 33 (33%)), sex (male, 15 of 47 (32%); female, three of eight (38%)), and smoking status (none, seven of 17 (41%); >10 years, 11 of 31 (36%)) of the bladder cancer patients. XAF1 association with clinicopathologic features of RCC was not defined in this study due to the small number of cases analyzed.

Figure 2
figure2

Expression status of XAF1 in human primary bladder and kidney carcinomas. (a) Semiquantitative RT–PCR analysis of XAF1, Smac/DIABLO, and HtrA2 expression in primary TCC of the bladder, primary RCC, and adjacent normal tissues. NB, normal bladder tissue; NK, normal kidney tissue. (b) Expression levels of XAF1, Smac/DIABLO, and HtrA2 transcripts in primary carcinomas. RT–PCR was repeated at least three times for each specimen and the means were obtained. Bar indicates the mean expression level of each specimen group (c) XAF1 expression in matched tissue sets. Expression level of XAF1 mRNA in cancer and adjacent noncancerous tissues were compared using matched sets obtained from the same cancer patients (P). N, normal tissue; T, tumor tissue. (d) Comparison of XAF1 expression levels between superficial (Ta–T1) and invasive (T2–T4) tumors and low- and high-grade tumors of the bladder.

Tumor-specific downregulation of XAF1 by promoter CpG sites hypermethylation

The XAF1 gene is located at the 17p13.2, approximately 3 cM to the p53 tumor suppressor, which undergoes frequent allelic losses in many human malignancies. To define whether altered expression of XAF1 is associated with gene deletion, we tested genomic level of XAF1 and p53 in the cell lines and tumor tissues. Five cancer cell lines (J82, T24, UOK122, UOK123 and PC-3), which have been known to carry allelic deletion of p53, exhibited low genomic level of p53, while none of the cell lines displayed detectable reduction of XAF1 gene level (Figure 3a). Similarly, all 75 primary tumors (55 TCCs and 20 RCCs) tested showed XAF1 gene level comparable to that of normal tissues while some tumors revealed low level of p53. To further define the genomic status of XAF1, loss of heterozygosity (LOH) of 17p13.1–13.2 region was evaluated using three polymorphic markers (D17S796, D17S1832 and D17S1828) (Figure 3b). Among 40 matched sets (20 TCCs and 20 RCCs) we tested, nine (23%) were informative for the centromeric marker (D17S796), which is located approximately 2.5 cM telomeric of the p53 locus, and LOH was detected from five (56%) cases. Intriguingly, four (80%) of the five LOH tumors showed a marked decrease in p53 gene level, while none of the tumors exhibited detectable reduction of XAF1 at both genomic and transcriptional levels. In addition, none of 10 (25%) cases informative for telomeric markers (D17S1832 and D17S1828) revealed LOH, suggesting that LOH at D17S796 might be associated with allelic loss of p53, and genomic deletion at this region including p53 locus might rarely extend into the XAF1 gene.

Figure 3
figure3

LOH and methylation analysis of XAF1 in cancer cell lines and primary tumors. (a) Genomic levels of XAF1 and p53 in cancer cell lines and tumor tissues. Exon 6 of XAF1 and exon 8 of p53 were amplified by intron-specific primers. Ten μl of the PCR products were resolved on a 2% agarose gel. GAPDH was used as an endogenous control. NB, normal bladder tissue; NK, normal kidney tissue. (b) No association of LOH at D17S796 with genomic status of XAF1. At centromeric marker D17S796, 5 tumor specimens exhibited LOH in the tumor DNA (T) compared with its corresponding normal DNA (N). Genomic level of XAF1 and p53 were evaluated using quantitative DNA-PCR. (c) Effect of 5-Aza-dC treatment on XAF1 expression in urogenital cancer cell lines. Cancer cell lines with no or low XAF1 expression were treated with 5-Aza-dC (5 μ M) for 5 days and XAF1 expression were evaluated by RT-PCR. C, control; T, treated. (d) Representative example of a dose-dependent reactivation of XAF1 mRNA expression by 5-Aza-dC in cancer cells. A kidney cancer cell line ACHN was treated with increasing doses of 5-Aza-dC (2.5–20 μ M) for 5days and expression level of XAF1 mRNA was determined by RT-PCR. (e) Methylation-specific PCR analysis of the XAF1 promoter in cancer cell lines and tissues. Fifty nanograms of bisulfite-modified genomic DNA was subjected to PCR amplification of the XAF1 promoter sequences using unmethylation (U)-specific and methylation (M)-specific primer sets, which cover nucleotides −354 to −13 (341 bp) and −687 to −458 (230 bp), respectively. NB, normal bladder tissue; NK, normal kidney tissue; CaP, carcinoma of the prostate.

Next, to elucidate whether aberrant DNA methylation is associated with XAF1 downregulation, tumor cells with no or low expression (T24, HT1197, 253J, UOK123, ACHN, LNCaP, DU145 and PC-3) were treated with the demethylating agent 5-aza-2′-deoxycytidine (5-Aza-dC). XAF1 expression was reactivated or elevated in these cells following 5-Aza-dC treatment in a dose-dependent manner (Figure 3c and d). To determine the overall frequency of XAF1 hypermethylation in tumors, we carried out methylation-specific PCR (MSP) analysis of the XAF1 promoter sequences. Methylation-specific primers (MS and MAS) and unmethylation-specific primers (US and UAS) were designed to amplify nucleotides −687 to −458 and −354 to −13, respectively, based on our previous finding of a tight correlation of CpG sites hypermethylation in these regions with decreased XAF1 expression in gastric cancers (Byun et al., 2003). Methylation was detected from all 16 cell lines having no or abnormally low XAF1 expression, but not found from the seven cell lines with normal expression (Figure 3e). Both methylation and unmethylation signals were observed from T24 and HT1197 cells with low expression, and only methylation signal was identified in ACHN and PC-3 cells showing no expression. Likewise, methylation was detected in 16 of 18 (89%) TCC and six of seven (86%) RCC tissues, which show low expression, while all noncancerous tissues and tumors with normal XAF1 expression were unmethylated. In addition, 35% (seven of 20) of primary prostate carcinomas but none of 10 BPH specimens were identified to be methylated at these regions.

Aberrant CpG sites hypermethylation of the XAF1 promoter

To further define the relationship between aberrant hypermethylation and gene expression, we characterized the methylation status of 14 CpG sites located in the 5′ proximal region (nucleotides –20 to –695) of the XAF1 promoter using bisulfite DNA sequencing analysis (Figure 4a). Bisulfite-modified genomic DNA, which were extracted from nine cancer cell lines (two normal and seven abnormal expressors) and 14 primary carcinomas (six TCCs, four RCCs and four prostate tumors), were subjected to PCR amplification for the promoter region spanning the 14 CpG sites and 10 PCR clones from each specimens were sequenced to determine methylation frequency at individual CpG sites (complete methylation; 80–100%, partial methylation; 20–60% and unmethylation; 0%). The majority (12–14 sites; 86–100%) of the sites was completely or partially methylated in cells with no or extremely low expression (253J, LNCaP, ACHN, DU145 and PC-3) and 9–11 (64–79%) sites were partially methylated in low expressors (HT1197 and T24), whereas only 2–3 (14–21%) sites showed partial methylation in normal expressors (J82 and HT1376) (P<0.001) (Figure 4b). Likewise, six primary tumors (four TCCs and two RCCs) with abnormally low expression harbored complete or partial methylation at 10–12 (71–86%) CpG sites, whereas its adjacent noncancerous tissues as well as four tumors with normal expression carried partial methylation at less than four sites (P<0.001) (Figure 4c). Consistent with the result of MSP assay, two MSP-positive prostate tumors were found to have complete or partial methylation at 10–12 (71–86%) CpG sites, whereas two MSP-negative tumors and two BPH tissues were detected to have only partial methylation at 2 (14%) sites. Together, these results demonstrate that methylation of promoter CpG sites is tightly associated with downregulation of XAF1 expression in urogenital cancer cell lines and primary tumors (supplementary data; Table 1).

Figure 4
figure4

Bisulfite DNA sequencing analysis of the XAF1 promoter. (a) A map of the CpG sites of the 5′ upstream region of the XAF1 gene. Fourteen CpGs (nucleotide −23 to −692) analysed by bisulfite DNA sequencing are represented by vertical lines with circle and numbered 1–14. The first nucleotide of ATG start codon is indicated by an arrow at +1. (b) Methylation status of the 14 CpG sites in the XAF1 promoter region of cancer cell lines. The promoter region comprised of 14 CpGs was amplified by PCR using primers MS7 and MS2. The PCR products were cloned and 10 plasmid clones were sequenced for each cell line. Percent methylation was determined from the number of clones containing a methylated CpG at each position relative to the total number of clones analysed. Black, gray, and white circles represent complete methylation (>7 clones), partial methylation (1–6 clones), and unmethylation, respectively. (c) Methylation status of the 14 CpGs in primary tumor and its adjacent noncancerous tissues. NB, normal bladder tissue; NK, normal kidney tissue; CaP, carcinoma of the prostate.

Absence of somatic mutations of XAF1

To explore the presence of mutational alteration of XAF1, we performed RT–PCR–SSCP analysis for XAF1 transcripts expressed from 11 tumor cell lines (four bladder and seven kidney cell lines) and 75 primary tumor tissues (55 TCCs and 20 RCCs) and DNA-PCR-SSCP analysis of the XAF1 gene for 20 primary prostate tumors. The entire coding region of XAF1 transcript was amplified using seven sets of exon-specific primers for RT–PCR-SSCP and eight sets of intron-specific primers for DNA-PCR–SSCP. To improve mutation detection sensitivity, the same PCR products were digested using different restriction endonuclease(s) and SSCP was carried out under two different running conditions. However, we failed to detect any types of mutation leading to amino-acid substitutions or frameshifts, whereas all of the previously reported p53 mutations in the cell lines were detected under the same conditions.

Increased cellular response to apoptosis-inducing chemotherapeutic drugs by XAF1 expression

Frequent downregulation of XAF1 in tumors suggests that XAF1 may play an important role in tumorigenesis. To explore the growth-suppressive role for XAF1, we initially assessed XAF1 effect on cell proliferation. The XAF1-nonexpressing 253J cells were transfected with expression plasmids designed to produce XAF1 protein with a Flag tag at the N-terminus (Flag-XAF1), and XAF1-expressing HT1376 cells were transfected with siRNA against XAF1 to knockdown its endogenous XAF1 expression (Supplementary data; Figure s1). [3H]thymidine uptake analysis showed that both XAF1-restored 253J and XAF1-disrupted HT1376 have no detectable change in DNA replication rate compared to empty vector- or control siRNA-transfected cells (Figure 5a). Flow cytometric analysis also revealed that ectopic overexpression of XAF1 in 253J cells does not cause detectable change in the distribution of cell cycle phases, while it slightly increases basal level of apoptosis (Figure 5b). Restoration of XAF1 expression in 253J cells led to a slight increase of apoptosis in the absence of etoposide or 5-FU treatment but greatly activated apoptosis induction following treatment (Figure 5c). Likewise, siRNA-mediated XAF1 knockdown in HT1376 cells attenuated the apoptotic response of the cells to the drugs. These results indicate that expression of XAF1 strongly enhances the cellular sensitivity to apoptosis-inducing stimuli.

Figure 5
figure5

Effect of XAF1 expression on cell proliferation and apoptosis. (a) [3H]thymidine uptake assay of XAF1 effect on DNA synthesis. Cells were transfected with increasing doses of XAF1 expression plasmid or siRNA against XAF1. After 44 h transfection, cells were pulse-labeled for 4 h with 1 μCi/ml [3H]thymidine. The radioactivity incorporated into trichloroacetic acid-precipitable materials was counted by a liquid scintillation counter. Data represent means of triplicate assays (Bars, s.d.). (b) No effect of XAF1 expression on cell cycle progression. 253J cells were transfected with GFP-XAF1 expression plasmids, and the distribution of cell cycle phases of GFP-positive cells were analyzed using flow cytometry. (c) TUNEL assay for XAF1 effect on tumor cell apoptosis. Cells transfected with XAF1 expression plasmid or XAF1 siRNA were treated with etoposide (10 μ M) or 5-FU (10 μ M) for 36 h. Apoptotic cell death was measured by using TUNEL assay. Data represent means of triplicate assays (***P<0.001; **P<0.01; *P<0.05).

No visible effect of XAF1 expression on subcellular localization of XIAP

To elucidate whether the proapoptotic action of XAF1 is accompanied with nuclear translocation of XIAP, the effect of XAF1 overexpression on the subcellular distribution of XIAP protein was investigated using fractionation assay. XAF1-transfected 253J cells were treated to etoposide for 24–72 h to induce apoptosis and equivalent amounts of protein from cytoplasmic and nuclear fractions were assessed for XIAP and XAF1 by Western blot analysis. In the absence of apoptotic stimuli, XAF1 protein was found in both the nucleus and the cytoplasm and endogenous XIAP protein was observed predominantly in the cytoplasm in both vector- and XAF1-transfected cells (Figure 6a). As predicted, etoposide treatment resulted in a significantly higher apoptosis in XAF1-transfected cells compared to vector-transfected cells (Figure 6b). However, XAF1-expressing 253J cells undergoing apoptosis exhibited no detectable nuclear translocation of XIAP up to 72 h after etoposide treatment, indicating that cytosolic XIAP proteins are not sequestered to the nucleus by XAF1 expression, which is sufficient to stimulate apoptosis induction. Similar results were obtained for the XIAP localization from the XAF1-restored LNCaP prostate cancer cells (data not shown). Additionally, etoposide-induced apoptosis was markedly activated by XAF1 transfection in a transfection dose-dependent manner, but mRNA and protein levels of XIAP in these cells with varying expression levels of XAF1 are comparable to those of vector-transfected or untransfected controls (Figures 5b and 6c).

Figure 6
figure6

Effect of XAF1 expression on cellular localization and expression of XIAP. (a) Fractionation assay for XAF1 effect on cellular distribution of XIAP protein in cells undergoing apoptosis. XAF1- or empty vector-transfected 253J cells were treated with etoposide. Nuclear and cytoplasmic extracts of the cells were prepared at the indicated time point and separated by 10% SDS–PAGE. Endogenous XIAP protein was detected by immunoblotting assay using an anti-XIAP antibody. U1 SnRNP 70 and β-tubulin were used as a control for purity of nuclear and cytoplasmic extracts, respectively (***P<0.001; **P<0.01). (b) Elevated tumor cell response to apoptotic stimuli by XAF1 expression. 253J cells transfected with Flag-XAF1 expression (3 μg) or empty vector (3 μg) were treated with etoposide (10 μ M) for 24–72 h. Number of apoptotic cell was determined using TUNEL assay. Data represent means of triplicate assays. Bars, s.d. (c) No visible effect of XAF1 on mRNA and protein expression level of XIAP during apoptosis. 253J cells transfected with increasing doses of XAF1 expression vectors were treated with etoposide (20 μ M) for 48 h and mRNA and protein levels of XIAP and p53 were determined by semiquantitative RT–PCR and Western blot assay.

Induction of p53 by XAF1 through the regulation of protein stability

We next investigated whether the apoptosis-sensitizing effect of XAF1 is associated with activation of p53, which plays a critical role in apoptosis induction triggered by a variety of stresses. Intriguingly, we found that p53 protein level is markedly elevated in XAF1-transfected 253J cells following etoposide treatment (Figure 7a and Figure s1). XAF1 expression led to a dose-associated induction of p53 protein and its target gene expression, including p21Waf1, PUMA and NOXA. Moreover, a higher and more sustained induction of p53 protein was detected in XAF1-versus vector-transfected cells, indicating that the presence of functional XAF1 causes a prolonged activation of p53 in response to apoptotic damage (Figure 7b). To define the role for XAF1 in p53 accumulation, the cells were exposed to the protein synthesis inhibitor cyclohexamide (CHX) and XAF1 effect on p53 stability was determined. As seen in Figure 7c, p53 accumulation in etoposide-treated cells was higher in XAF1-transfected cells compared to vector-transfected cells, supporting that XAF1 upregulation of p53 occurs at the post-translational level. To determine whether functional p53 is required for the proapoptotic action of XAF1, we utilized PC-3 prostate cancer cells, which express extremely low level of XAF1 and no functional p53 protein. While XAF1 transfection alone has a negligible effect on both basal and etoposide-induced apoptosis, co-transfection with wild-type p53 led to a dramatic elevation of apoptosis (Figure 7d). TUNEL assay demonstrated that after 36 h exposure to etoposide (20 μ M), approximately 69% of PC-3 cells display apoptotic death by co-transfection of XAF1 and p53, while only 18 and 29% of the cells undergo apoptosis by transfection of XAF1 and p53, respectively. Requirement of functional p53 for XAF1's proapoptotic effect was further characterized using 253J and its subline (253J/p53–175D) expressing a dominant negative mutant form of p53 (175D), which ablates apoptotic function of wild-type p53 (Ryan and Vousden, 1998). Flow cytometric analysis of the sub-G1 phase cells demonstrated that XAF1 expression strongly enhances apoptosis of parental cells, but its effect is markedly abrogated in the 253J/p53–175D cells. On this basis, we tested the growth suppression activity of XAF1 using colony formation assay. 253J cells were stably transfected with expression plasmids encoding sense or antisense XAF1, and XAF1 expression in the transfectants was verified by RT–PCR and immunobloting using anti-XAF1 antibody (data not shown). As seen in Figure 7e, introduction of sense XAF1 resulted in approximately threefold decrease in the number of neomycin-resistant colonies, and the XAF1-mediated inhibition of colony formation was significantly impeded by co-transfection of a dominant negative mutant form of p53 (175D).

Figure 7
figure7

XAF1 upregulation of p53 protein stability. (a) Induction of p53 and its target gene expression by XAF1. 253J bladder cancer cells were transfected with increasing does of Flag-XAF1 expression vectors and treated with etoposide (20 μ M) for 48 h. Protein level of p53 was analysed by Western blot assay and mRNA expression level of p53 target genes p21Waf1, PUMA and NOXA were determined by semi-quantitative RT–PCR. (b) Elevated and sustained induction of p53 by XAF1. XAF1- or vector-transfected 253J cells were pulse-treated with etoposide (50 μ M) for 6 h and p53 protein level was determined at the indicated times after treatment. (c) Enhancement of p53 protein stability by XAF1. The cells were pretreated with CHX (10 mM) for 1 h before etoposide treatment (20 μ M) and p53 protein level was examined 12 h after etoposide treatment. (d) Requirement of functional p53 for the apoptotic action of XAF1. PC-3 (p53-deficient), 253J (wild-type p53), or 253J/p53–175D cells were transfected separately or simultaneously with expression vectors encoding XAF1 or wild-type p53 and its effect on apoptosis was determined using TUNEL assay (PC-3) or flow cytometric analysis of sub-G1 fraction (253J). Data represent means of triplicate assays (Bars, s.d.; ***P<0.001; **P<0.01; *P<0.05). (e) XAF1 inhibition of tumor cell growth and its dependency on functional p53. 253J cells (5 × 104) transfected the indicated transgene were seeded in soft agar and maintained in the presence of neomycin for 4 weeks. Neomycin-resistant colonies were stained with crystal violet. Assays were performed in triplicate and average number of colonies and s.d. were calculated (***P<0.001).

Discussion

The 17p13 region, where the XAF1 gene is located, undergoes frequent allelic losses in a variety of human malignancies including bladder cancer, and the tumor-specific loss or downregulation of XAF1 expression suggests a possible role for XAF1 in the suppression of malignancy (Fong et al., 2000; Liston et al., 2001; Steidl et al., 2002). However, the mechanism by which XAF1 is downregulated in human cancers has been poorly characterized. Allelic loss of the XAF1 gene has been initially predicted as a possible mechanism for XAF1 inactivation, based on the microsatellite analysis of the NCI 60 cell line panel showing a decreased heterozygosity within the XAF1 region at 17p13.2 (Fong et al., 2000). XAF1 is located at approximately 3 cM telomeric of the p53 tumor suppressor, which is frequently associated with LOH, underscoring the importance of resolving the basis for the XAF1 downregulation in cancer cells. However, our previous study showed that LOH at the centromeric region of the XAF1 locus is rarely extends into the XAF1 gene in human gastric cancers (Byun et al., 2003). Moreover, tumors with LOH at telomeric markers of the p53 gene exhibited low genomic levels of p53 while none of these tumors show detectable reduction of XAF1 gene level, supporting that LOH at the 17p13 region is associated with allelic deletion of p53 but not of XAF1. Consistent with this, all urogenital cancer cell lines and primary tumors tested in this study have normal level of the XAF1 gene, while a substantial fraction of the same set of tumor specimens shows significantly reduced genomic level of p53, indicating that genomic deletion of XAF1 might be infrequent in human urogenital cancers.

It has been well documented that hypermethylation in CpG-rich promoter or transcription regulatory region is strongly associated with transcriptional silencing and a critical event leading to the epigenetic inactivation of tumor suppressor genes in human cancers. In this study, we found that XAF1 expression is reactivated or upregulated in no or low expressor tumor cells following 5-Aza-dC treatment and aberrant hypermethylation at 14 CpG sites located within the 5′ proximal region (nucleotides −23 to −692) of the promoter is tightly associated with downregulation of XAF1 expression in both cancer cell lines and primary tumors. Thus, this suggests that hypermethylation of the 5′ proximal region might be critical for the transcriptional silencing of XAF1 in urogenital cancers and possibly other human malignancies.

Very recently, XAF1 was identified as a novel interferon (IFN)-stimulated gene that contributes to IFN-dependent sensitization of cells to TRAIL-induced apoptosis (Leaman et al., 2002). XAF1 mRNA was upregulated by IFN-α2 and IFN-β and high levels of XAF1 protein were induced predominantly in cell lines sensitive to the proapoptotic effects of IFN-β. It was also found that XAF1-expressing melanoma cells are sensitive to TRAIL-induced apoptosis compared with nonexpressing cells and the degree of sensitization is correlated with the level of XAF1 expressed. Intriguingly, we found that epigenetic inactivation of XAF1 is significantly high in muscle-invasive bladder tumors compared with superficial tumors and more frequent in high-grade tumors than low-grade tumors, suggesting its contribution to the malignant progression of human bladder tumors. Instillation of Bacillus Calmette-Guerin (BCG) is one of the most successful immunotherapies for superficial bladder cancer and carcinoma in situ. BCG seems to induce tumor regression by producing a proinflammatory Th1 cytokine response, including IFN-γ, TNF-α and TRAIL (Jackson et al., 1995). IFNs induce apoptosis and/or growth arrest of bladder cancer cells and combination of IFN with BCG results in an additive or synergistic effect against clinical bladder cancers (Luo et al., 1999; Benedict et al., 2004). However, some bladder cancer cells are resistant against to BCG- or IFN-induced tumor regression, leading to the conjecture that defective response to growth suppression effect of BCG and IFNs may give the tumors a selective advantage and abet escape from T-cell antitumor response. Recently, IFN-α was found to stimulate marked increases in TRAIL expression and induce early activation of caspase-8 (Papageorgiou et al., 2004). IFN-α-induced apoptosis was significantly blocked by anti-TRAIL antibody whereas sensitivity to IFN-α in IFN-resistant bladder cancer cells was increased by treatment with a TRAIL-sensitizing agent such as Bortezomib. It was also demonstrated that patients who responded to BCG therapy have significantly higher urine TRAIL levels compared with nonresponders and TRAIL plays a crucial role in BCG-induced antitumor effects (Ludwig et al., 2004). These findings thus suggest that IFN-induced apoptosis in bladder cancer cells involves autocrine TRAIL production and overcoming TRAIL resistance may be very effective in restoring IFN sensitivity in human bladder tumors (Papageorgiou et al., 2004). In this context, it could be suspected that bladder cancers with XAF1 inactivation might be more resistant to BCG or IFN combined immunotherapy than cancers with normal XAF1 expression, and restoration of functional XAF1 could be effective in overcoming TRAIL resistance by sensitization of tumor cells to TRAIL-induced apoptosis. Therefore, it will be valuable to examine that expression status of XAF1 could be a clinically useful marker for cancer treatment including IFN and BCG therapy.

XAF1 was originally identified as a nuclear protein that could inactivate anticaspase function of XIAP via sequestering XIAP protein to the nucleus (Liston et al., 2001). However, we could not detect the nuclear translocation of XIAP protein in tumor cells undergoing apoptosis by XAF1 overexpression. XIAP protein was predominantly observed in the cytoplasm and its level was decreased in apoptosis-undergoing cells after etoposide treatment. Additionally, restoration of XAF1 expression in XAF1-deficient cells markedly accelerated apoptosis induction by etoposide, but no visible effect on XIAP protein level was identified in the cells. This observation raises the possibility that the apoptosis-sensitizing function of XAF1 could be exerted, in part, through the XIAP-independent pathway. In this context, our finding that XAF1 activates p53 protein accumulation lends support to the notion that XAF1's proapoptotic effect may not be solely dependent on its XIAP-interfering activity. Furthermore, we found that XAF1 enhances the protein stability of p53, thus leading to a prolonged activation of p53 and its target gene expression under stress condition, and the proapoptotic effect of XAF1 was dramatically increased in the presence of functional p53. It is therefore conceivable that inactivation of XAF1 may decrease the apoptotic sensitivity of tumor cells by the attenuated p53 response to various apoptotic stimuli. This is in line with our result that XAF1-mediated suppression of colony-forming ability of cancer cells is substantially impeded by blockade of p53 function. Thus, functional p53 might be critical for the apoptosis-sensitizing action of XAF1 although the p53-independent proapoptotic role for XAF1 cannot be excluded. In fact, the existence of p53-independent function of XAF1 was suggested by our data showing that the apoptotic sensitivity of p53-deficient cells is reduced by knockdown of XAF1 expression using siRNA. Taken together, these findings support that XAF1 might be a novel effector of p53 in signaling apoptosis and tumor suppression function of XAF1 might stem partially from its capability to enhance the p53 apoptosis-signaling pathway.

In conclusion, the data presented here clearly demonstrate that XAF1 undergoes epigenetic silencing in a considerable proportion of human urogenital malignancies. An association of abnormal XAF1 expression with advanced tumor stage and higher grade of bladder tumor further supports its implication in the malignant progression of human tumors. Additionally, our study raises the possibility that loss or abnormal dowwnregulation of XAF1 expression might render tumor cells a survival advantage by attenuating the apoptotic response of p53 to various stress conditions. It will be therefore valuable to explore the possible application of XAF1 as a molecular marker for detection and treatment of human urogenital malignancies.

Materials and methods

Primary tumor tissues and cancer cell lines

A total of 95 primary tumor specimens, including 55 TCC and corresponding normal bladder mucosa, 20 RCC and adjacent noncancerous kidney tissues, and 20 prostate carcinomas were obtained by surgical resection in the Kyung Hee University Medical Center (Seoul, Korea). Ten benign prostatic hyperplasia tissues were also obtained from patients with no evidence of cancer. Tissue specimens were snap-frozen immediately in liquid N2 and stored at −80°C until used. Signed informed consent was obtained from each patient. Bits of primary tumors and adjacent portions of each tumor were fixed and used for hematoxylin and eosin staining for histopathological evaluation. Tumor specimens composed of at least 70% carcinoma cells and adjacent tissues found not to contain tumor cells were chosen for molecular analysis. Six human bladder cancer cell lines (T24, J82, HT1197, HT1376, 253J and 253J-BV) and three human prostate cancer cell lines (LNCaP, DU145 and PC-3) were obtained from American Type Culture Collection (Rockville, MD, USA) or Korea Cell Line Bank (Seoul National University, Seoul, Korea). Total cellular RNA and genomic DNA specimens extracted from 15 human RCC cell lines (UOK101, UOK105, UOK107, UOK108, UOK109, UOK110, UOK112, UOK115, UOK121, UOK122, UOK123, UOK124, UOK130, ACHN and WWC1) were provided by Dr deVere White (University of California at Davis, CA, USA).

Semi-quantitative RT- and genomic PCR analysis

The strategy for our semi-quantitative RT–PCR was previously described (Chi et al., 1994; Tricoli et al., 1996). Briefly, 1 μg of RNA was converted to cDNA by reverse transcription using random hexamer primers and MoMuLV reverse transcriptase (Life Technologies Inc., Gaithersburg, MD, USA). For quantitative evaluation of XAF1 expression levels by RT–PCR, we initially performed the PCR reaction over a range of cycles (20–40 cycles) using serially diluted cDNA, and 1:4 diluted cDNA (12.5 ng/50 μl PCR reaction) undergoing 24–36 cycles was observed to be within the logarithmic phase of amplification and yielded reproducible results with primers all primers used for XAF1, Smac/DIABLO, HtrA2, XIAP, p53, p21Waf1, PUMA, NOXA and an endogenous expression standard gene GAPDH (Chi et al., 1998; Byun et al., 2003). PCR was performed for 30–34 cycles at 95°C (1 min), 58–62°C (0.5 min) and 72°C (1 min) in 1.5 mM MgCl2-containing reaction buffer (PCR buffer II, Perkin Elmer). For quantitative genomic PCR, genomic DNA was extracted from the same cells of the tissues or cell lines from the DNA phase after RNA was extracted. Two-hundred ng of genomic DNA were used for amplification of the exon 6 region of XAF1, exon 8 of p53, and intron 5 of GAPDH using intron-specific primers as previously described (Chi et al., 1999). RT- and genomic PCR products (10 μl) were resolved on 2% agarose gels. Quantitation was achieved by densitometric scanning of the ethidium bromide-stained gels. Absolute area integrations of the curves representing each specimen were then compared after adjustment for GAPDH. Integration and analysis was performed using Molecular Analyst software program (Bio-Rad, Hercules, CA, USA). Quantitative PCR was repeated at least three times for each specimen and the mean was obtained. Gene expression data was analysed by χ2 test and Student's t-test. P<0.05 was considered statistically significant.

Immunoblotting assay

Cells were lysed in a lysis buffer containing 20 mM Tris (pH 7.4), 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1% Triton, 2.5 mM sodium phosphate, 1 mM β-glycerolphosphate, 1 mM Na3VO4, 1 μg/ml leupeptin, and 1 mM PMSF. The cell lysate was clarified by centrifugation and 20 μg of total protein were supplemented with Laemmli buffer and loaded on a 10% sodium dodecyl suflate–polyacrylamide gel (SDS–PAGE) for electrophoresis. Western analyses were performed using antibodies specific for XIAP (BD Biosciences, San Diego, CA, USA), p53 (Santa Cruz Biotechnology, CA, USA), β-tubulin (Sigma, St Louis, MO, USA), and U1 SnRNP 70 (Santa Cruz Biotechnology, CA, USA). Antibody binding was detected by enhanced chemiluminescence (Amersham Biosciences, Piscataway, NJ, USA) using a secondary antibody conjugated to horseradish peroxidase. For stripping, the blots were incubated in a stripping buffer (0.2 M glycine (pH 2.2), 0.1% SDS, 1% Tween-20) at room temperature for 60 min.

Loss of heterozygosity analysis

LOH was determined using three polymorphic CA markers (D17S796, D17S1832 and D17S1828) localized at chromosome 17p13.1–13.2. PCR amplification was performed on each tumor and normal DNA sample pair obtained from 40 patients using primers as previously described (Byun et al., 2003). PCR products (10 μl) were electrophoresed on nondenaturing 6% polyacrylamide gels. Signal intensity of fragments and the relative ratio of both tumor and normal allele intensities were determined by scanning desitometry. As certain numbers of noncancerous cells might be present in tumor tissues, LOH was assigned when the intensity ratio of the two tumor alleles differed by at least 50% from that observed on its corresponding normal DNA. The same PCR products were also subjected to nonisotopic SSCP analysis for verification of LOH.

Nonisotopic RT–PCR–SSCP analysis

To detect possible sequence alterations in XAF1, we performed nonisotopic RT–PCR–SSCP analysis. The XAF1 transcript was amplified with seven sets of primers that were designed to cover the entire coding region of the gene. Sequences of the primers used will be obtained upon request. The PCR products of over 300 bp in lengths were digested with endonuclease(s) to increase the sensitivity of SSCP analysis. RT–PCR–SSCP analysis of p53 transcripts was performed as described previously (Chi et al., 1994). PCR products (10 μl) mixed with 10 μl of 0.5 N NaOH, 10 mM EDTA, and 15 μl of denaturing loading buffer (95% formamide, 20 mM EDTA, 0.05% bromophenol blue and 0.05% xylene cyanol). After heating at 95°C for 5 min, samples were loaded in wells precooled to 4°C and run using 8% nondenaturating acrylamide gels containing 10% glycerol at 4–8°C and 18–22°C.

5-Aza-dC treatment

To assess reactivation of XAF1 expression, cell lines showing no or low levels of XAF1 transcript were plated in six-well tissue plates 48 h before treatment. 5-Aza-dC (Sigma, St Louis, MO, USA) was added to the fresh medium at concentration of 2.5–20 μ M in duplicate and cells were harvested after 5 days of treatment (Lee et al., 2001). Using cDNA obtained from cancer cell lines, mRNA transcript of XAF1 before and after 5-Aza-dC treatment were analysed by RT–PCR.

Methylation-specific PCR analysis

Genomic DNA (1 μg) in a volume of 50 μl was denatured by NaOH (final concentration 0.3 M). Thirty micorliters of 10 mM hydroquinone and 520 μl of 3 M sodium bisulfite (pH 5.0) were added and incubated at 55°C for 16–20 h. DNA samples were purified using Wizard DNA clean-up system (Promega Corp., Madison, WI, USA), again treated with NaOH at 37°C for 15 min, precipitated with ethanol, and resuspended in distilled water. Fifty nanograms of bisulfite-modified or unmodified DNA were subjected to PCR amplification. PCR was performed separately with a methylation-specififc primer set, MS (sense; 5′-IndexTermTTTATTTTATTGGTAGACGTTACG-3′) and MAS (antisense; 5′-IndexTermATAACTCCTAAACTTCCAAACG-3′) and a nonmethylation-specific primer set, US (sense; 5′-IndexTermTTTGGAAAAGGGATGGAGATTTAGATG-3′) and UAS (antisense; 5′-IndexTermACAAACTTTCAATTAAATTTCA-3′) for 38 cycles at 95°C for 1 min, at 60–63°C for 1 min, and 72°C for 1 min. Ten micoliters of PCR product were visualized on a 2% agarose gel.

Bisulfite DNA sequencing analysis for the XAF1 promoter

Fifty nanograms of bisulfite-modified DNA were subjected to PCR amplification of the XAF1 promoter region using primers MS7 (sense; 5′-IndexTermAATTTTGTTATTTTTTTTAGAG-3′) and MS2 (antisense; 5′-IndexTermCATATTCTACTCTCTACAAAC-3′) for nucleotides −700 to −20. The PCR products were cloned into pCRII vectors (Invitrogen, Carlsbad, CA, USA) and 10 clones of each specimen were sequenced by automated fluorescence-based DNA sequencing to determine the methylation status.

Construction of expression plasmids, siRNA and transfection

Expression vectors encoding sense or antisense XAF1 were constructed by a PCR based approach using primers XAF1-FLAG-14 (sense; 5′-IndexTermATGGATTACAAGGATGACGACGATAAGATGGAAGGAGACTTCTCGGT-3′) and XAF1–15 (antisense; 5′-IndexTermGTTAAAAGTGAAATCTTTTGAATT-3′). The PCR products were first ligated to the pCR2.1-TOPO vector (Invitrogen, Carlsbad, CA, USA) and then subcloned into the pcDNA3.1-FLAG vector (Invitrogen, Carlsbad, CA, USA). Approximately, 1 × 105 cells were plated per six-well plate in media containing 10% fetal bovine serum to give 50–60% confluence, and the transfection of constructs was performed using FuGene6 (Roche, Mannheim, Germany) according to the manufacturer's protocol. The transfection efficiency was monitored using a fluorescence microscopy for FLAG or CAT assay (Roche, Mannheim, Germany). Expression vector encoding human mutant p53 (R175D), which give rise to a protein with impaired apoptotic and cell cycle arrest functions, was kindly provided by Dr KH Vousden (Beatson Institute for Cancer Research, Glagsow, UK). Small interfering RNA (siRNA) duplex against XAF1 (5′-IndexTermATGTTGTCCAGACTCAGAG-3′) and control siRNA duplex served as negative control were synthesized by Dharmacon Research (Lafayette, CO, USA). For transfection, 1 × 105 cells were plated on 60-mm-diameter dishes 24 h before transfection. The cells were incubated with a siRNA-Oligofectamine mixture at 37°C for 4 h, followed by addition of fresh medium containing 10% fetal bovine serum.

Cell proliferation and DNA synthesis assays

Cells were seeded in six-well plate at the density of 2 × 104 cells per cell in duplicate and were maintained in the presence of 10% FBS. The following day, cells were transfected with expression vector or siRNA as described above. Cell numbers were counted using a hemocytometer for 4 days at 24-h intervals. DNA synthesis was measured by determining the incorporation of [3H]thymidine. After 44 h transfection, cells were pulse-labeled for 4 h with 1 μCi/ml of [3H]thymidine (Amersham Pharmacia Biotech, NJ, USA) and the radioactivity incorporated into trichloroacetic acid-precipitable materials was counted by a liquid scintillation counter. For flow cytometry analysis, cells were harvested 48 h after transfection and fixed with 70% ethanol and resuspended in 1 ml of PBS containing 50 mg/ml RNase and 50 mg/ml propidium iodide (Sigma, St Louis, MO, USA). The assay was performed on a FACScan flow cytometer (Becton Dickinson, San Jose, CA, USA), and the cell cycle profile was analysed using MultiCycle software (Phoenix Flow Systems, San Diego, CA, USA).

TUNEL assay

TUNEL assay was performed to evaluate apoptotic effect of XAF1. Briefly, 253J and HT1376 cells were transfected with XAF1-expressing vector and siRNA for XAF1, respectively, and the cells were treated with etoposide (25 μ M) or 5-FU (25 μ M) for 24–72 h. The cells were then fixed with 4% paraformaldehyde in PBS, and the buffer containing 3% bovine serum albumin and 0.1% Triton X-100 was added and incubated for 15 min at 4°C. The cells were labeled by the terminal deoxynucleotidyltransferase-mediated dUTP-biotin nick end labeling (TUNEL) reaction mixture using the In Situ Cell Death Detection kit (Roche, Mannheim, Germany). TUNEL signals of cells were visualized directly under microscopy.

Preparation of cytosolic and nuclear extracts

Nuclear and cytosolic extracts from XAF1-or vector-transfected cells were prepared for immunoblotting assay for XAF1 and XIAP at the indicated time points. Cells were harvested in 1 ml of phosphate-buffered saline with a rubber policeman. Samples were centrifuged for 1 min at 6000 g (4°C), and the resulting cell pellets were resuspended in 100 μl of low salt buffer (20 mM HEPES, pH 7.9, 10 mM KCl, 0.1 mM NaVO4, 1 mM EDTA, 1 mM EGTA, 0.2% Nonidet P-40, 10% glycerol, supplemented with a set of proteinase inhibitors, Complete™). After 10 min of incubation on ice, the samples were centrifuged at 13 000 g for 2 min (4°C), and the supernatants (cytosolic extracts) were immediately frozen in a dry ice/ethanol bath. Pelleted nuclei were resuspended in 60 μl of high salt buffer (20 mM HEPES, pH 7.9, 420 mM NaCl, 10 mM KCl, 0.1 mM NaVO4, 1 mM EDTA, 1 mM EGTA, 20% glycerol, supplemented with Complete™), and nuclear proteins were extracted by shaking on ice for 30 min. Samples were centrifuged at 13 000 g for 10 min (4°C), and the supernatants were taken as nuclear extracts.

Colony formation assay

253J human bladder carcinoma cells were transfected with expression vectors encoding sense XAF1, antisense XAF1, wild-type 53 or mutant p53 (R175D) using FuGene6 (Roche, Mannheim, Germany). Transfected cells (5 × 104) were maintained grown in soft agar in the presence of G418 for 4 weeks. Colonies were stained with 0.5% crystal violet in 20% ethanol.

Statistical analysis

The results of DNA replication, apoptosis, colony forming assays were expressed as mean±s.d. A Student's t-test was used to determine the statistical significance of the difference. The χ2 test was used to determine the statistical significance of expression and methylation levels between tumor and normal tissues. A P-value of <0.05 was considered significant.

References

  1. Ambrosini G, Adida C, Altieri DC . (1997). A novel anti-apoptosis gene, survivin, expressed in cancer and lymphoma. Nature Med 3: 917–921.

  2. Asselin E, Mills GB, Tsang BK . (2001). XIAP regulates Akt activity and caspase-3-dependent cleavage during cisplatin-induced apoptosis in human ovarian epithelial cancer cells. Cancer Res 61: 1862–1868.

  3. Benedict WF, Tao Z, Kim CS, Zhang X, Zhou JH, Adam L et al. (2004). Intravesical Ad-IFNα causes marked regression of human bladder cancer growing orthotopically in nude mice and overcomes resistance to IFN-α protein. Mol Ther 10: 525–532.

  4. Bilim V, Kasahara T, Hara N, Takahashi K, Tomita Y . (2003). Role of XIAP in the malignant phenotype of transitional cell cancer (TCC) and therapeutic activity of XIAP antisense oligonucleotides against multidrug-resistant TCC in vitro. Int J Cancer 103: 29–37.

  5. Byun DS, Cho K, Ryu BK, Lee MG, Kang MJ, Kim HR et al. (2003). Hypermethylation of XIAP-associated factor 1, a putative tumor suppressor gene from the 17p13.2 locus, in human gastric adenocarcinomas. Cancer Res 63: 7068–7075.

  6. Chi SG, Chang SG, Lee SJ, Lee CH, Kim JI, Park JH . (1999). Elevated and biallelic expression of p73 is associated with progression of human bladder cancer. Cancer Res 59: 2791–2793.

  7. Chi SG, deVere White RW, Meyers FJ, Siders D, Lee F, Gumerlock PH . (1994). p53 in prostate: frequent expressed transition mutations. J Natl Cancer Inst 86: 926–933.

  8. Chi SG, Kim HJ, Park BJ, Min HJ, Park JH, Kim YW et al. (1998). Mutational abrogation of the PTEN/MMAC1 gene in gastrointestinal polyps in patients with Cowden disease. Gastroenterology 115: 1084–1089.

  9. Deveraux QL, Reed JC . (1999). IAP family proteins-suppressors of apoptosis. Genes Dev 13: 239–252.

  10. Deveraux QL, Takahashi R, Salvesen GS, Reed JC . (1997). X-linked IAP is a direct inhibitor of cell-death proteases. Nature 388: 300–304.

  11. Du C, Fang M, Li Y, Li L, Wang X . (2000). Smac, a mitochondrial protein that promotes cytochrome c-dependent caspase activation by eliminating IAP inhibition. Cell 102: 33–42.

  12. Duckett CS, Nava VE, Gedrich RW, Clem RJ, Van Dongen JL, Gilfillan MC et al. (1996). A conserved family of cellular genes related to the baculovirus iap gene and encoding apoptosis inhibitors. EMBO J 15: 2685–2689.

  13. Fong WG, Liston P, Rajcan-Separovic E, St Jean M, Craig C, Korneluk RG . (2000). Expression and genetic analysis of XIAP-associated factor 1 (XAF1) in cancer cell lines. Genomics 70: 113–122.

  14. Green DR . (2000). Apoptotic pathways: paper wraps stone blunts scissors. Cell 102: 1–4.

  15. Holcik M, Gibson H, Korneluk RG . (2001). Apoptotic brake and promising therapeutic target. Apoptosis 6: 253–261.

  16. Jackson AM, Alexandroff AB, Kelly RW, Skibinska A, Esuvaranathan K, Prescott S et al. (1995). Changes in urinary cytokines and soluble intercellular adhesion molecule-1 (ICAM-1) in bladder cancer patients after bacillus Calmette-Guerin (BCG) immunotherapy. Clin Exp Immunol 99: 369–375.

  17. Leaman DW, Chawla-Sarkar M, Vyas K, Reheman M, Tamai K, Toji S et al. (2002). Identification of X-linked inhibitor of apoptosis-associated factor-1 as an interferon-stimulated gene that augments TRAIL Apo2L-induced apoptosis. J Biol Chem 277: 28504–28511.

  18. Lee MG, Kim HY, Byun DS, Lee SJ, Lee CH, Kim JI et al. (2001). Frequent epigenetic inactivation of RASSF1A in human bladder carcinoma. Cancer Res 61: 6688–6692.

  19. Li J, Kim JM, Liston P, Li M, Miyazaki T, Mackenzie AE et al. (1998). Expression of inhibitor of apoptosis proteins (IAPs) in rat granulosa cells during ovarian follicular development and atresia. Endocrinology 139: 1321–1328.

  20. Liston P, Fong WG, Kelly NL, Toji S, Miyazaki T, Conte D et al. (2001). Identification of XAF1 as an antagonist of XIAP anti-Caspase activity. Nat Cell Biol 3: 128–133.

  21. Liston P, Roy N, Tamai K, Lefebvre C, Baird S, Cherton-Horvat G et al. (1996). Suppression of apoptosis in mammalian cells by NAIP and a related family of IAP genes. Nature 379: 349–353.

  22. Ludwig AT, Moore JM, Luo Y, Chen X, Saltsgaver NA, O’Donnell MA et al. (2004). Tumor necrosis factor-related apoptosis-inducing ligand: a novel mechanism for Bacillus Calmette-Guerin-induced antitumor activity. Cancer Res 64: 3386–3390.

  23. Luo Y, Chen X, Downs TM, DeWolf WC, O’Donnell MA . (1999). IFN-α2B enhances Th1 cytokine responses in bladder cancer patients receiving Mycobacterium bovis bacillus Calmette-Guerin immunotherapy. J Immunol 162: 2399–2405.

  24. Ng KCP, Campos EI, Martinka M, Li G . (2004). XAF1 expression is significantly reduced in human melanoma. J Invest Dermatol 123: 1127–1134.

  25. Papageorgiou A, Lashinger L, Millikan R, Grossman HB, Benedict W, Dinney CP et al. (2004). Role of tumor necrosis factor-related apoptosis-inducing ligand in interferon-induced apoptosis in human bladder cancer cells. Cancer Res 64: 8973–8979.

  26. Perrelet D, Perrin FE, Liston P, Korneluk RG, MacKenzie A, Ferrer-Alcon M et al. (2004). Motoneuron resistance to apoptotic cell death in vivo correlates with the ratio between X-linked inhibitor of apoptosis proteins (XIAPs) and its inhibitor, XIAP-associated factor 1. J Neurosci 24: 3777–3785.

  27. Roy N, Deveraux QL, Takahashi R, Salvesen GS, Reed JC . (1997). The c-IAP-1 and c-IAP-2 proteins are direct inhibitors of specific caspases. EMBO J 16: 6914–6925.

  28. Ryan KM, Vousden KH . (1998). Characterization of structural p53 mutants which show selective defects in apoptosis but not cell cycle arrest. Mol Cell Biol 18: 3692–3698.

  29. Sasaki H, Sheng Y, Kotsuji F, Tsang BK . (2000). Down-regulation of X-linked inhibitor of apoptosis protein induces apoptosis in chemoresistant human ovarian cancer cells. Cancer Res 60: 5659–5666.

  30. Siegelin M, Touzani O, Toutain J, Liston P, Rami A . (2005). Induction and redistribution of XAF1, a new antagonist of XIAP in the rat brain after transient focal ischemia. Neurobiol Dis 20: 509–518.

  31. Steidl C, Simon R, Burger H, Brinkschmidt C, Hertle L, Bocker W et al. (2002). Patterns of chromosomal aberrations in urinary bladder tumours and adjacent urothelium. J Pathol 198: 115–120.

  32. Sun C, Cai M, Gunasekera AH, Meadows RP, Wang H, Chen J et al. (1999). NMR structure and mutagenesis of the inhibitor-of-apoptosis protein XIAP. Nature 401: 818–822.

  33. Suzuki Y, Imai Y, Nakayama H, Takahashi K, Takio K, Takahashi R . (2001). A serine protease, HtrA2, is released from the mitochondria and interacts with XIAP, inducing cell death. Mol Cell 8: 613–621.

  34. Tamm I, Kornblau SM, Segall H, Krajewski S, Welsh K, Kitada S et al. (2001). Expression and prognostic significance of IAP-family genes in human cancers and myeloid leukemias. Clin Cancer Res 6: 1796–1803.

  35. Thompson CB . (1995). Apoptosis in the pathogenesis and treatment of disease. Science 267: 1456–1462.

  36. Tricoli JV, Gumerlock PH, Yao JL, Chi SG, D’Souza SA, Nestok BR et al. (1996). Alterations of the retinoblastoma gene in human prostate adenocarcinoma. Genes Chromosomes & Cancer 15: 108–114.

  37. Verhagen AM, Ekert PG, Pakusch M, Silke J, Connolly LM, Reid GE et al. (2000). Identification of DIABLO, a mammalian protein that promotes apoptosis by binding to and antagonizing IAP proteins. Cell 102: 43–53.

Download references

Acknowledgements

This work was supported in part by grants from Korea Science and Engineering Foundation (R02-2003-000-10031-0), Korea Research Foundation (2003-070-C00031), and the National Cancer Center (0420230), Korea.

Author information

Correspondence to S-G Chi.

Additional information

Supplementary Information accompanies the paper on the Oncogene website (http://www.nature.com/onc).

Supplementary information

Supplementary Figure S1 (GIF 64 kb)

Supplementary data (DOC 28 kb)

Rights and permissions

Reprints and Permissions

About this article

Keywords

  • XAF1
  • promoter hypermethylation
  • CpG site
  • urogenital cancer
  • apoptosis
  • p53

Further reading