Cellular senescence functions as a tumor suppressor that protects against cancer progression. α-Catulin, an α-catenin-related protein, is reported to have tumorigenic potential because it regulates the nuclear factor-κB (NF-κB) pathway, but little is known about its clinical relevance and the mechanism through which it regulates cancer progression. Here, we found that α-catulin mRNA levels were significantly upregulated in cancer cell lines and clinical oral squamous cell carcinomas, which positively correlated with tumor size (P=0.001) and American Joint Committee on Cancer (AJCC) stage (P=0.004). α-Catulin knockdown in the OC2 and A549 cancer cell lines dramatically decreased cell proliferation and contributed to cellular senescence, and inhibited OC2 xenograft growth. Mechanistic dissection showed that α-catulin depletion strongly induced the DNA-damage response (DDR) in both cell lines, via a p53/p21-dependent pathway in A549 cells, but a p53/p21-independent pathway in OC2 cells carrying mutant p53. Global gene expression analysis revealed that α-catulin knockdown altered cell-cycle regulation and DDR pathways at the presenescent stage as well as significantly downregulate several crucial genes related to mitotic chromosome condensation, DDR and DNA repair systems, which suggests that its depletion-induced cellular senescence might be caused by chromosome condensation failures, severe DNA damage and impaired DNA repair ability. Our study provides evidence that α-catulin promotes tumor growth by preventing cellular senescence and suggests that downregulating α-catulin may be a promising therapeutic approach for cancer treatment.
Cellular senescence is a state of irreversible growth arrest (Campisi and d’Adda di Fagagna, 2007) in which senescent cells remain metabolically active and display typical phenotype changes, including the absence of mitosis, an enlarged flattened morphology and high activity of senescence-associated β-galactosidase (SA-β-gal) (Dimri et al., 1995). It can be caused by the critical shortening of telomeres in a process known as replicative senescence or by various forms of stress such as oncogene activation and loss of tumor suppressors in a process known as premature senescence or stress-induced senescence. Moreover, evidence indicates that DNA damage is a crucial mediator of cellular senescence: it triggers the DNA-damage response (DDR) pathway, which enables cells to sense damaged DNA and to respond by repairing the damage or by arresting cell-cycle progression (Bartkova et al., 2006; Di Micco et al., 2006). Persistent DDR signaling is essential for both senescence initiation and maintenance (d’Adda di Fagagna, 2008). Genetic manipulations such as the aberrant expression of oncogenes abolishing the senescence response led to fully developed malignancy (Chen et al., 2005; Sarkisian et al., 2007). Cellular senescence is also a proven tumor-suppressive strategy that forces transformed cancer cells to (re-)enter irreversible cell-cycle arrest (Collado et al., 2007). Therefore, triggering the senescence of tumor cells may contribute to successful cancer therapy (Lleonart et al., 2009).
α-Catulin is an 82-kDa vinculin/α-catenin family protein encoded by the CTNNAL1 gene, which was originally identified as a downregulated transcript in sodium butyrate-treated pancreatic cancer cells that were undergoing differentiation and apoptosis (Zhang et al., 1998). α-Catulin has similarities to human vinculin and α-catenin, especially in the N-terminal region, which contains binding sites for β-catenin, talin and α-actinin, which suggests that α-catulin may be a cytoskeletal linker protein (Janssens et al., 1999). Despite the sequence similarity between α-catulin, α-catenin and vinculin, however, α-catulin did not inhibit the Wnt/β-catenin signaling pathway (Merdek et al., 2004). The α-CATULIN/CTNNAL1 gene localizes to chromosome 9q31-32 (Zhang et al., 1998; Park et al., 2002), a frequent site of allelic loss and a tumor suppressor marker that has been reported in many human cancers (Schultz et al., 1995; Miura et al., 1996), which implies that α-catulin may be a tumor suppressor. However, other studies (Park et al., 2002; Wiesner et al., 2008) report that α-catulin has tumorigenic potential because it binds directly to IKK-β and Lbc, a Rho-specific guanine nucleotide exchange factor, which promotes cell migration and increases resistance to apoptosis. The literature contains little evidence of the clinical significance and function of α-catulin in cancer progression. In this study, we analyzed the expression of α-catulin in clinical tumor specimens to clarify the correlation between α-catulin expression and tumor progression and to elucidate the mechanisms used by α-catulin in cellular senescence.
α-Catulin knockdown arrested cell proliferation and induced senescence in cancer cells
First, using quantitative reverse transcription (qRT)–PCR we measured the mRNA levels of α-catulin in two types of primary cultured healthy oral keratinocytes, the human lung cancer cell line A549 and several human oral cancer cell lines. α-Catulin mRNA levels were higher in cancer cells than in normal cells (Figure 1a). We generated two lentiviruses, designated shC01 and shB01, that produced specific short-hairpin RNA (shRNA) targeted to bases 324–344 and 1401–1421 of full-length α-catulin mRNA, respectively. To understand the role of α-catulin in tumorigenesis, we infected tumor cells with these lentiviruses to silence their α-catulin gene expression. After 1 week of infection and puromycin selection, we found that in OC2 and A549 cells α-catulin gene expression was silenced and cell proliferation dramatically reduced by the shB01 and shC01 viruses but not by the control shLuc viruses (Figures 1b–d). We also found that α-catulin knockdown in OC2 and A549 cells induced the accumulation of cell populations in S and G2/M cell-cycle phases (Figure 1e) as well as decreasing their specific markers cyclin A and cyclin B1, respectively, compared with shLuc-infected cells (Supplementary Figure 1). Less than 1.5% apoptotic cells were observed among shLuc- and α-catulin shRNA-infected cells (Figure 1e). Most of these cells had senescence-like morphology with an enlarged and flattened shape (Figure 1f). Notably, endogenous SA-β-gal activity and the oncogene-induced senescence marker DcR2 (Collado et al., 2005) were higher in α-catulin-silenced cancer cells 6 days after infection with α-catulin shRNA viruses (Figure 1g; Supplementary Figure 1). Taken together, these findings suggest that α-catulin knockdown induced cellular senescence as the major phenotype of cell death in OC2 and A549 cancer cells.
α-Catulin expression was frequent in oral squamous cell carcinoma tissues and positively correlated with tumor size
We then used qRT–PCR to examine α-catulin expression in oral cancer and adjacent non-tumor oral mucosa tissue samples from 63 oral cancer patients. α-Catulin levels were higher in 49 of the tumor tissue samples compared with adjacent non-tumor tissue samples (Figure 2a). All 63 patients were enrolled in the follow-up study, which lasted 77 months (median: 33 months). The total survival curve for the 63 oral squamous cell carcinoma (OSCC) patients is shown in Figure 2b. Although a Kaplan–Meier analysis showed no significant correlation between survival and α-catulin expression, we found that OSCC patients with higher α-catulin expression levels had a relatively lower cumulative survival rate. Furthermore, we investigated the clinical relevance of differential α-catulin expression in paired tumor and non-tumor tissues. α-Catulin overexpression in tumor tissue was not correlated with age and gender, cigarette smoking, betel nut chewing, alcohol drinking habits, tumor site or lymph node metastasis but was significantly correlated with tumor size (P=0.001) and AJCC staging (P=0.004) (Table 1). Although there was no significant correlation between the grade of tumor cell differentiation and α-catulin expression, α-catulin expression gradually increased with poorly differentiated grades of tumor (P=0.057).
α-Catulin knockdown suppressed tumorigenicity in vivo
To investigate whether α-catulin depletion impairs tumorigenesis in vivo, we subcutaneously injected shLuc- or shC01-infected OC2 cells (2 × 106) into the posterior flank of non-obese diabetic (NOD)-severe combined immunodeficiency (SCID) mice and measured the tumor size on days 0, 6, 13, 20, 25, 27, 29, 31 and 33 post-inoculation. We found that the tumors became palpable and grew rapidly after day 25 in mice inoculated with shLuc-infected OC2 cells but grew slowly in mice inoculated with shC01-infected OC2 cells (Figure 3A). Tumors in the former group were significantly larger on day 33 than those in the latter group (tumor incidence: shLuc: 3 of 3; shC01: 3 of 4) (Figure 3A, inset). shLuc-induced tumors were three times heavier than shC01-induced tumors (Figure 3B), which indicated that inhibiting α-catulin expression in cancer cells delayed tumor formation. Moreover, α-catulin expression in shC01-infected OC2 cells was lower than in shLuc-infected OC2 cells (Figure 3C). Notably, immunohistochemical analysis revealed that the percentage of Ki-67-positive tumor cells was higher in shLuc infected than in shC01-infected OC2 cells (Figures 3D and E). In contrast, the percentage of DcR2-positive tumor cells was higher in shC01-infected OC2 cells and was morphologically enlarged and flattened compared with shLuc-infected cells (Figures 3D and F), strongly indicating tumor cell senescence in vivo driven by α-catulin knockdown. Taken together, these results suggest that α-catulin knockdown significantly suppressed tumor size by inducing senescence, which inhibited cancer cell proliferation in vivo.
Cellular senescence induced by α-catulin knockdown was p53 dependent in A549 cells but p53 independent in OC2 cells
p53/p21 is the key tumor suppressor pathway; it is trigged by DNA damage and has been implicated in senescence induction (d’Adda di Fagagna, 2008). Therefore, we also examined whether DNA damage and the p53/p21 pathway is involved in the senescence induced by α-catulin knockdown in OC2 and A549 cells. We found that α-catulin knockdown in OC2 and A549 cells resulted in increased phosphorylated form of histone H2AX (γ-H2AX, Ser139), which is essential for DDR signaling (Stucki et al., 2005), and p53 and p21 expression as well as nuclear accumulation (Figures 4a and b). These data suggest that the level of α-catulin expression is linked to changes in the p53/p21 pathway: a low level mediates cell-cycle arrest and causes cellular senescence.
Moreover, we clarified whether p53 activation was required for the senescence caused by α-catulin knockdown. We determined whether p53 knockdown rescues tumor cells from the deleterious effects of α-catulin knockdown and enables their continued proliferation. To do this, we co-infected A549 and OC2 cells with various combinations of lentiviruses expressing α-catulin shRNAs, p53 shRNAs, and several different negative control shRNAs. After puromycin selection, western blotting confirmed p53 knockdown in pooled clones (Figure 4f). We then determined that the effect of p53 depletion on SA-β-gal activity was that it abrogated α-catulin-knockdown-induced senescence and proliferative inhibition in A549 cells (Figures 4c–e, right). p53-mediated senescence induced by α-catulin knockdown was also confirmed in p53-siRNA-transfected A549 cells (Supplementary Figure 2). Moreover, p53 knockdown blocked the expression of p21 induced by α-catulin silencing in A549 cells but not the active expression of γ-H2AX, which suggested that DNA damage accumulated upstream of p53 (Figure 4f, right). p53 knockdown also inhibited DcR2 expression induced by α-catulin silencing, but increased the expression of cyclin A and cyclin B1 (Figure 4f, right), suggesting that p53 knockdown directed α-catulin-knockdown-induced senescence to cell proliferation. Unexpectedly, p53 knockdown in OC2 cells failed to abolish α-catulin-knockdown-induced cellular senescence and proliferative inhibition (Figures 4c–e, left). The cells still accumulated DcR2 and γ-H2AX, and cyclin A and cyclin B1 were not detected (Figure 4f, left). This suggests that α-catulin-knockdown-induced senescence in OC2 cells is not associated with the upregulation of the p53 pathway.
Because p53 depletion did not affect α-catulin-knockdown-induced senescence in OC2 cells, we sequenced the complete coding region of TP53, which was PCR amplified from A549 and OC2 cell cDNA. This analysis confirmed the wild-type TP53 cDNA sequence for A549 and revealed a point mutation of A to G at position 394 in OC2 cells—Lys132 mutated to Glu132 within the DNA-binding domain of p53 (Supplementary Figure 3). Moreover, the p16/Rb pathway is crucial for senescence induction, but our data indicated that p16/Rb was not activated by α-catulin silencing in OC2 cells (Supplementary Figure 4). To further confirm the p53-dependent and p53-independent functions of α-catulin-knockdown-induced senescence, we performed experiments in two further cell lines, MCF-7 human breast cancer cells and CL1-0 human lung adenocarcinoma cells carrying wild-type and mutant p53, respectively. Our results also showed that p53 knockdown blocked α-catulin-depletion-induced senescence and proliferative inhibition in MCF-7 cells (Supplementary Figures 5a–c, right), which was consistent with the results observed in A549 cells. However, p53 knockdown in CL1-0 cells did not affect the cellular senescence and proliferative inhibition driven by α-catulin depletion (Supplementary Figures 5a–c, left), which was also consistent with the observations in OC2 cells. Therefore, these results suggest that α-catulin-knockdown-induced senescence is p53 dependent in p53 wild-type cells but p53 independent in mutant p53-expressing cells.
α-Catulin knockdown significantly altered cell-cycle regulation and DDR-related pathways
To further investigate the mechanism by which α-catulin-knockdown induces senescence in OC2 cells, we used cDNA microarray analysis to examine the genes regulated as early as the presenescent stage (on day 2 and day 4), during which the expression of SA-β-gal activity and DcR2 was undetectable (Figure 5a). A MetaCore software analysis of global genes revealed that cell-cycle regulation and DDR were the two main pathways that significantly altered OC2 cells at the presenescent stage (Figure 5b). Notably, the most prominently altered cell-cycle regulation pathway in α-catulin-knockdown cells was the chromosome condensation in prometaphase (CCP), which includes the CCNA, CCNB, CDK1, INCENP, AURKA, NCAPD2, NCAPG2, SMC2, SMC4, TOP2A, H3F3B and H1FX genes, all of which were markedly downregulated (Figure 5c). An analysis of the time course of each prominently altered gene expression among these pathways was performed using quantitative real-time PCR (Figure 6a). We found that two DNA-damage-induced genes—growth arrest and DNA-damage-inducible protein GADD45A and 14-3-3 protein sigma SFN—were markedly upregulated at the presenescent stage. These genes are expressed upon cell-cycle arrest and are involved in DNA repair. On the other hand, the genes related to CCP, including SMC2, SMC4, NCAPG2, TOP2A and AURKA, and the DNA repair-related genes, including MSH2, MSH6, RAD50 and BRCA1, were strongly downregulated at 2 days and with a moderate decrease by 4 days. In addition, the active expression of γ-H2AX was also initially induced at the presenescent stage (Figure 6b). Indeed, there was a higher percentage of 4′,6′ diamino-2-phenylindole·2HCl (DAPI)-stained nuclei in α-catulin shRNAs-infected OC2 cells with a giant nucleus at the presenescent stage, than in shLuc-infected OC2 cells (Figure 6c), suggesting that α-catulin depletion-induced chromatin relaxation at presenescence may be primarily due to the loss of the genes involved in chromosome condensation. Taken together, these results suggest that the loss of CCP and the impairment of DDR and DNA repair ability during presenescence are the major causative triggers accelerating the onset of cellular senescence (on day 6, Figure 5a) by α-catulin knockdown in OC2 cells.
Here, we provide evidence that the expression of α-catulin is upregulated in most human cancer cells and clinical oral cancer tissues and positively correlated with tumor size and cancer stage of OSCC. Knockdown of α-catulin in cancer cells bearing either wild-type or mutant p53 is sufficient to trigger DDR and eventually induce cellular senescence in vitro as well as suppress tumor formation in vivo. These results suggest that α-catulin functions as an oncoprotein, sustaining proliferation by preventing cellular senescence and promoting tumor progression.
Ample evidence suggests that p53/p21 is crucial for mediating DDR (d’Adda di Fagagna, 2008) and induces growth arrest to allow DNA repair, or promotes cellular senescence. Our results show that p53/p21 activation mediates α-catulin-knockdown-induced cellular senescence in p53 wild-type cells (A549 and MCF-7). Moreover, it has been reported that p53 mutations impair the DDR pathway (Bartkova et al., 2005; Gorgoulis et al., 2005) because mutant p53 is often more stable than wild-type p53 in the nucleus and is a dominant-negative inhibitor against wild-type p53 (de Vries et al., 2002), which allows cancer cells to develop by bypassing senescence. However, this is not consistent with our finding that OC2 and CL1-0 cells expressing mutant p53 also show α-catulin-knockdown-induced senescence, which suggests that pathways other than p53/p21 lead to senescence.
Our cDNA microarray data revealed that α-catulin-knockdown-induced senescence in OC2 cells may be caused primarily by the downregulation of many cell-cycle regulation-related genes, especially by CCP. Moreover, an analysis of the time course found that several CCP-related genes, including SMC2, SMC4, NCAPG2, TOP2A and AURKA, were strongly downregulated as early as 2 days after α-catulin silencing. Loss of CCP genes has been reported to block centrosome separation, spindle assembly and chromosome segregation during mitosis. SMC2, SMC4 and NCAPG2 are three of five components of the condensing II complex, a complex that establishes the mitotic chromosome architecture and has a central role in chromosome assembly and segregation (Hudson et al., 2009). The loss of SMC4 stops the resolution of sister chromatids and cause the formation of anaphase chromatin bridges and DNA breakages (Coelho et al., 2003). Loss of TOP2A has been demonstrated to drastically disrupt chromatin condensation by preventing the proper formation of the kinetochore (Mikhailov et al., 2002). AURKA has a critical role in controlling centrosome separation, spindle assembly and chromosome segregation during mitosis (Carmena and Earnshaw, 2003). A recent study reported that the inhibition of ARUKA induces cancer cell senescence in vitro and in vivo (Huck et al., 2010). Although the association between chromatin relaxation and cellular senescence remains unclear, importantly, chromatin relaxation has been hypothesized to be a response to DNA damage and to be related to chromosomal instability (Murr et al., 2006; Ziv et al., 2006). Moreover, chromosomal instability induces aneuploidy, which eventually leads to cellular senescence (Suzuki et al., 2002; Walen, 2007). These findings strongly suggest that the inhibition of α-catulin expression may lead to the absence of chromosome condensation, caused by the inhibition of these genes, result in chromosomal instability and cause senescence.
DNA repair mechanisms maintain genomic fidelity (Pastink et al., 2001). The loss of these repair genes usually contributes to genetic defects and eventually causes genomic instability (Pastink et al., 2001; van Gent et al., 2001), which predisposes the cell to senescence (Suzuki et al., 2002; Walen, 2007). Notably, our study showed that several DNA repair-related genes were markedly suppressed in presenescent OC2 cells induced by α-catulin depletion. MSH2, MSH6 and PCNA are reported to have key roles in mismatch repair (Kunkel and Erie, 2005); ATM, BRCA1 and RAD50 are required in homologous recombination repair and non-homologous end-joining repair (Shrivastav et al., 2008); and XPC is involved in nucleotide excision repair (Volker et al., 2001; Wakasugi et al., 2002). When the expression of these DNA repair genes was inhibited in α-catulin-knockdown cells, this was powerfully reflected in the high incidence of genomic instability. On the other hand, we found that DNA damage upregulated the expression of the GADD45A and SFN (14-3-3 sigma) genes. GADD45A is known to have a central role as a cellular stress sensor (Li et al., 2009), and its expression increased following stressful growth arrest conditions and treatment with DNA-damaging agents (Rosemary Siafakas and Richardson, 2009). Moreover, upregulated SFN gene expression inhibits Akt-mediated cell growth, transformation and tumorigenesis, all of which cause DNA damage (Yang et al., 2006). Taken together with these findings, our study strongly suggests that one major driving force behind α-catulin-knockdown-induced senescence in OC2 cells is insufficiency of DDR and the impairment of DNA repair ability, which causes severe DNA damage and genomic instability, both of which eventually result in irreversible senescence.
Cellular senescence is thought to be important for preventing unregulated growth and malignant transformation (Campisi and d’Adda di Fagagna, 2007). Cellular senescence induced by loss of function of oncogenes such as Cdk4 (Zou et al., 2002) and CIP2A (Li et al., 2008) has been considered as a therapeutic approach and has been verified by previous studies (Lleonart et al., 2009). Knockdown of α-catulin is crucial for ensuring the irreversibility of the senescence arrest even in p53 mutant cells. Our findings not only offer new perspectives in the modulation of senescence by α-catulin but also suggest a novel therapeutic target for cancer treatment.
Materials and methods
OC2 and OECM1 human oral cancer cell lines established from Taiwanese men with a history of betel quid chewing were obtained from Dr RC Chang (Veterans General Hospital, Taipei, Taiwan) and Dr SY Liu (Chi-Mei Medical Center, Tainan, Taiwan), respectively. An HSC3 cell line derived from human tongue carcinoma with lymph node metastasis was from the JCRB cell bank of Japan. OC2, OECM1 and HSC3 cells were maintained in RPMI 1640 medium, Dulbecco's modified Eagle's medium. A549 human lung cancer cells (American Type Culture Collection, Washington, DC, USA) were maintained in Dulbecco's modified Eagle's medium. Medium was supplemented with 10% fetal bovine serum and 100 units/ml of penicillin and streptomycin (Invitrogen, Carlsbad, CA, USA). Human normal oral keratinocytes (NHOK), taken with informed consent, were isolated from the gingival tissue of healthy patients in the Department of Dentistry, National Cheng Kung University Hospital. The NHOK were cultured in keratinocyte-serum-free medium supplied with L-glutamine, epidermal growth factor (EGF) and bovine pituitary extract (BPE) (Invitrogen) and then incubated at 37°C in a humidified 5% CO2 atmosphere.
RNA extraction and qRT–PCR
Total RNA was isolated using a reagent (Trizol; Invitrogen) according to the manufacturer's protocol. First-strand cDNA was synthesized from 1 μg of total RNA using reverse transcription (ImProm-II Reverse Transcription System; Promega, Madison, WI, USA). qRT–PCR was done (LightCycler 480 Instrument; Roche Applied Sciences, Indianapolis, IN, USA) using SYBR Green I dye. Each 10 μl reaction contained 4 mmol/l of MgCl2, 200 nmol/l each of forward and reverse primers, 1 μl of dye (LightCycler Fast Start DNA Master SYBR Green I; Roche) and 2 μl of 10-fold diluted cDNA. Primer sequences are listed in Supplementary Table 1. Gene expression levels of α-catulin and a housekeeping gene (glyceraldehyde 3-phosphate dehydrogenase (GAPDH) or β2-microglobulin) were determined for each sample. Relative quantities of α-catulin mRNA were calculated using the comparative Ct method, subtracting Ct (α-catulin) from Ct (housekeeping gene) to derive ΔCt for each sample. Analyses were done in triplicate to confirm the data.
Lentivirus production and infection
shRNA targeting α-catulin, p53 and luciferase were expressed from the pLKO.1 hairpin vector, which harbors an expression cassette for a puromycin resistance gene driven by the human phosphoglycerate kinase promoter. The shRNA sequences used in this study are shown in Supplementary Table S1. shRNA lentiviruses were generated from HEK293T packaging cells. Cell culture supernatant containing lentivirus was harvested every 24 h until 72 h after transfection. Viral titers were determined by transmuting HEK293 T cells using diluted culture supernatants and tested by counting the number of viable cells after 2 days of culture in the presence or absence of puromycin (Sigma-Aldrich, St Louis, MO, USA). Viral supernatants were stored at −80°C. Cells were infected with shRNA or control lentiviruses in the presence of 8 μg/ml of polybrene (Sigma-Aldrich). After 48 h of infection, the cells were treated with 1 μg/ml of puromycin for selection. Puromycin-resistant cells were pooled for subsequent analysis.
Colony formation assay
Approximately 105 cells were seeded per 10 cm dish and cultured in growth medium with puromycin (1 μg/ml) for 14 days. Clonogenic survival was determined by staining the colonies with crystal violet and visualizing them with a digital camera.
MTS proliferation assay
Cell proliferation was determined by using CellTiter 96 AQueous One Solution Cell Proliferation Assay (MTS) kit (Promega, Madison, WI, USA). Approximately 103 cells were seeded in 96-well plate and cultured in growth medium with puromycin (1 μg/ml). Following phosphate-buffered saline washing and addition of MTS reagent, absorbance at 490 nm was recorded.
For the analysis of cell-cycle distribution, both floating and attached cells were collected and centrifuged before washed with cold phosphate-buffered saline, and then fixed in 70% cold ethanol overnight at −20°C. Propidium iodide staining was performed after incubation of the cells with 50 μg/ml propidium iodide and 20 μg/ml RNase A in the dark at room temperature for 30 min, which were then analyzed by flow cytometry (BD Biosciences, San Jose, CA, USA).
SA-β-gal activity was measured using a kit (Senescence β-Galactosidase Staining Kit; Cell Signaling Technology, Beverly, MA, USA) according to the manufacturer's instructions. Light microscopy was used to identify and count senescent (blue stained) cells.
Cells seeded on a coverslip were fixed in 4% paraformaldehyde for 15 min. After they had been permeabilized with 0.25% Triton X-100, the cells were blocked in 5% bovine serum albumin (BSA) for 30 min at room temperature and then incubated with antibodies against γ-H2AX (Ser 139) (#2577; Cell Signaling Technology), p53 (sc-126) or p21 (sc-817) (Santa Cruz Biotechnology, Santa Cruz, CA, USA). Rhodamine phalloidin (Alexa Fluor 633-conjugated phalloidin; Invitrogen) and DAPI were used for F-actin and nuclear staining.
Whole-cell lysates were resolved by sodium dodecyl sulfate polyacrylamide gel electrophoresis. Proteins were transferred onto a polyvinylidene difluoride membrane (Millipore, Billerica, MA, USA) and incubated with the primary antibody, and then incubated with horseradish peroxidase-conjugated secondary antibodies. Specific proteins were detected using enhanced enhanced chemiluminescence (ECL) chemiluminescence reagent (Amersham Biosciences, Piscataway, NJ, USA). The primary antibodies used for western blotting were α-catulin (B01P; Abnova, Taipei, Taiwan), γ-H2AX (Ser 139, #2577; Cell Signaling Technology), p53 (sc-126) and p21 (sc-817) (Santa Cruz Biotechnology); DcR2 (ab2019; Abcam, Cambridge, UK), the Cyclin Antibody Sampler kit (#9869; Cell Signaling Technology) and β-actin (Sigma-Aldrich).
Patients and tissue samples
The study was approved by the institutional review boards of Chi-Mei Hospital and National Cheng Kung University Hospital, Tainan, Taiwan. Sixty-three paired primary OSCC samples were included in this study. Immediately after surgery, cancerous and adjacent non-cancerous tissues were stored in liquid nitrogen. Pathologists determined the histological grade of each specimen. The OSCC samples were staged according to the American Joint Committee on Cancer (5th AJCC) 1997 cancer staging guidelines.
Survival and statistical analysis
A χ2 test was used to analyze the clinicopathological variables of qRT–PCR results. Survival rates were calculated using the Kaplan–Meier method, and statistical significance was determined using the log-rank test. Statistical significance was set at P<0.05. All statistical analyses were performed using SPSS 13.0 for Windows (SPSS Inc., Chicago, IL, USA).
Tumorigenicity assays in NOD/SCID mice
After lentiviral infection and puromycin selection (1 μg/ml), shLuc- or shC01-infected OC2 cells (2 × 106) in 50 μl of phosphate-buffered saline were subcutaneously injected into the posterior flank of 6-week-old NOD–SCID mice. Tumor size (mm3) was measured for each mouse on days 0, 6, 13, 20, 25, 27, 29, 31 and 33 post-inoculation. On day 33, primary tumors were excised and weighed. Tumor size was monitored by measuring the length (L), width (W) and height (H), and calculated with the formula (L × W × H).
Histopathology and immunohistochemistry
Specimens fixed in 10% buffered formalin were embedded in paraffin and then sectioned and stained with hematoxylin and eosin. Immunohistochemical analysis of the paraffin sections was done using primary antibody for Ki-67 (550609; BD Pharmingen, San Diego, CA, USA) and DcR2 (ab2019; Abcam); the sections were then incubated with anti-mouse/rabbit immunoglobulin G-horseradish peroxidase-conjugated secondary antibody. The signal was detected using a kit (Aminoethyl Carbazole Substrate Kit; Zymed Laboratories Inc., San Francisco, CA, USA). The sections were counterstained with hematoxylin. Image analysis of Ki-67- and DcR2-positive cell was further quantified by using Histoquest software (Tissue Gnostics, Vienna, Austria).
Microarray analysis and pathway analysis
Microarray hybridization was done at the Phalanx Biotech Service Center (Hsinchu, Taiwan). RNA integrity and quality were assessed using a kit (RNA 6000 Nano Assay; Agilent Technologies, Inc., Santa Clara, CA, USA), a spectrophotometer (NanoDrop ND-1000; Thermo Fisher Scientific Inc., Wilmington, DE, USA) and agarose gel electrophoresis. RNA was labeled with Cy5 using aminoallyl RNA labeling and then hybridized in duplicate on full human genome arrays (Whole Genome Human OneArray HOA 4.3; Phalanx Biotech) containing 30 968 human genome probes and 1082 experimental control probes in a one-block array format, and with each probe a 60 mer oligonucleotide designed in the sense direction. Filtered data were log2 transformed and corrected using quantile normalization before their average ratios and significance values were calculated. Genes that showed a significant (P<0.01) expression difference between α-Catulin silenced (shB01 and shC01 infected) and non-silenced (shLuc) were analyzed for pathways (GeneGo Map Folders of MetaCore software suite; GeneGo, St Joseph, MI, USA). Significance probability was calculated using the hypergeometrical distribution based on gene ontology terms. Because one gene is frequently involved in multiple pathways, all pathways corresponding to the genes with significance probability were listed.
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This work was supported by grants NSC 96-2311-B-006-005-MY3, NSC 97-2314-B-384-003-MY3, NSC 99-3112-B-006-011 and NSC 99-2627-B-006-003 from the National Science Council, and DOH99-TD-C-111-003 from the Department of Health, Taiwan. RNAi reagents were obtained from the National RNAi Core Facility located at the Institute of Molecular Biology/Genomic Research Center, Academia Sinica, supported by the National Research Program for Genomic Medicine Grants of National Science Council, Taiwan (NSC 97-3112-B-001-016).
The authors declare no conflict of interest.
Supplementary Information) accompanies the paper on the Oncogene website
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