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. Author manuscript; available in PMC: 2013 Aug 30.
Published in final edited form as: Toxicol Appl Pharmacol. 2012 Jun 28;263(2):218–224. doi: 10.1016/j.taap.2012.06.012

Suberoylanilide hydroxamic acid (SAHA) inhibits EGF-induced cell transformation via reduction of cyclin D1 mRNA stability

Jingjie Zhang a,b, Weiming Ouyang b, Jingxia Li b, Dongyun Zhang b, Yonghui Yu b, York Wang b, Xuejun Li a,*, Chuanshu Huang b,**
PMCID: PMC3758130  NIHMSID: NIHMS507092  PMID: 22749963

Abstract

Suberoylanilide hydroxamic acid (SAHA) inhibiting cancer cell growth has been associated with its down-regulation of cyclin D1 protein expression at transcription level or translation level. Here, we have demonstrated that SAHA inhibited EGF-induced Cl41 cell transformation via the decrease of cyclin D1 mRNA stability and induction of G0/G1 growth arrest. We found that SAHA treatment resulted in the dramatic inhibition of EGF-induced cell transformation, cyclin D1 protein expression and induction of G0/G1 growth arrest. Further studies showed that SAHA downregulation of cyclin D1 was only observed with endogenous cyclin D1, but not with reconstitutionally expressed cyclin D1 in the same cells, excluding the possibility of SAHA regulating cyclin D1 at level of protein degradation. Moreover, SAHA inhibited EGF-induced cyclin d1 mRNA level, whereas it did not show any inhibitory effect on cyclin D1 promoter-driven luciferase reporter activity under the same experimental conditions, suggesting that SAHA may decrease cyclin D1 mRNA stability. This notion was supported by the results that treatment of cells with SAHA decreased the half-life of cyclin D1 mRNA from 6.95 h to 2.57 h. Consistent with downregulation of cyclin D1 mRNA stability, SAHA treatment also attenuated HuR expression, which has been well-characterized as a positive regulator of cyclin D1 mRNA stability. Thus, our study identifies a novel mechanism responsible for SAHA inhibiting cell transformation via decreasing cyclin D1 mRNA stability and induction of G0/G1 growth arrest in Cl41 cells.

Keywords: SAHA, cyclin D1 mRNA stability, Cell transformation

Introduction

Histone deacetylase (HDAC) inhibitor is one of the most important classes of anticancer agents. Suberoylanilide hydroxamic (SAHA), one of the well-known HDAC inhibitors, has been shown with highly effective in inducing cell growth arrest and apoptosis, particularly in cancer cell lines in comparison to relative normal cells (Ungerstedt et al., 2005). Accumulated evidence indicates that SAHA exerts its inhibition of cancer cell growth through both histone-dependent and histone-independent mechanisms (Bolden et al., 2006). SAHA treatment increases p21 transcription via the accumulation of acetylated histones H3 and H4, by which SAHA mediates the cancer cell growth arrest (Gui et al., 2004). It has also been reported that SAHA treatment promotes cell apoptosis by induction of p53 protein acetylation and the protein stability in cancer cell lines (Kai et al., 2010). However, the potential chemopreventive effect of SAHA on skin carcinogenesis has not yet been reported. Thus, we evaluated the potential inhibitory effect of SAHA on EGF-induced cell transformation in Cl41 cells.

Cyclin D1 overexpression has been observed in many human cancer tissues, including breast cancer, lung cancer, pancreatic cancer, and non-melanoma skin cancer, particularly in keratoacanthomas and squamous cell carcinomas (SCCs) (Musgrove et al., 2011). In addition, cyclin D1 in carcinogenesis has been well demonstrated in previous studies (Donnellan and Chetty, 1998). We have also found that knockdown of cyclin D1 impairs cell transformation induced by environmental carcinogens in mouse skin epidermal Cl41 JB6 cells and human immortalized keratinocyte HaCaT cells (Ding et al., 2009; Ouyang et al., 2008; Zhang et al., 2009). Cyclin D1 plays a critical role in cell growth and proliferation through regulation of cell cycle progression (Resnitzky and Reed, 1995). Cyclin D1 forms complexes with CDK4 and CDK6, which is an essential in G1/S progression. Cyclin D1 also activates cyclin E/CDK2 and indirectly inactivates the function of tumor suppresser gene retinoblastoma protein (Fu et al., 2004). Previous studies indicate that SAHA treatment downregulates cyclin D1 via repressing cyclin D1 transcription and translation (Kawamata et al., 2007; Yamaguchi et al., 2005). Our current studies demonstrate that SAHA inhibits EGF-induced Cl41 cell transformation via decreasing cyclin D1 mRNA stability and induction of G0/G1 growth arrest.

Materials and methods

Cell culture and reagents

Mouse epidermal Cl41 cells and their transfectants were cultured in 5% Fetal bovine Serum (FBS) MEM containing with 1% penicillin/streptomycin and 2 mM l-glutamine (Life Technologies), and were maintained at 37 °C in 5% CO2 incubator. Human colon cancer cell lines HCT116 and HCT116 p21−/− were cultured in McCoy’s 5A supplemented with 10% FBS. The cultures were dissociated with trypsin and transferred to new 25 cm2 culture flasks twice a week. FBS was purchased from Life Technologies, Inc. (Gaithersburg, MD), MEM and McCoy’s 5A were from Calbiochem (San Diego, CA), Luciferase assay substrate, CellTiter-Glo® Luminescent Cell Viability Assay kit and EGF were from Promega (Madison, WI).

Plasmids and cell transfection

AP-1-luciferase reporter, NF-κB-luciferase reporter, and cyclin D1 promoter luciferase reporter were described in our previous studies (Ouyang et al., 2006, 2007a; Zhang et al., 2009). JB6 Cl41 cells stably transfected with cyclin D1 promoter-driven luciferase reporter plasmid, AP-1-luciferase reporter plasmid, and NF-κB-luciferase reporter plasmid were described in our previous publications (Ouyang et al., 2006; Zhang et al., 2009). The expression vector pEGFP-C3-cyclin D1 was constructed as follows: pEGFP-C3 was digested with EcoRI and BamHI. The human cyclin D1 cDNA fragment was amplified using specific primers (forward: 5′-GCGAATTCCCATGGAACACCAGCTCCTG-3′; reverse: 5′-GCGGATC CTCAGATGTCCACGTCCCGCA-3′). The fragment containing human cyclin D1 open reading frame, EcoRI site, and BamHI site, was inserted to the clone site of pEGFP-C3. The construct was sequenced by Genewiz (South Plainfield, NJ). Cl41 cells were transfected with either p-EGFP-C3 (GFP) or pEGFP-C3-cyclin D1 (GFP-cyclin D1) expression plasmids using PolyJet™ DNA in vitro transfection reagent (SignaGen Laboratories, Rockville, MD).

Anchorage-independent growth

Soft agar colony formation assay was performed as described previously (Ouyang et al., 2008; Zhang et al., 2009). Briefly, 2.5 ml of 0.5% agar in basal modified Eagle’s medium (BMEM) supplemented with 10% FBS and 20 ng/ml EGF was layered onto each well of 6-well tissue culture plates. 3×104 Cl41 cells or HCT116 cells were mixed with 1 ml of 0.5% agar BMEM supplemented with 10% FBS with or without 20 ng/ml EGF and layered on top of the 0.5% agar layer. The plates were incubated at 37 °C in 5% CO2 for three weeks. The colonies were then counted under microscopy and those with 32 cells were scored. The results were presented as colonies/104 cells.

Cell proliferation assay

2×103 Cl41 viable cells suspended in 100 μl medium containing 5% FBS were seeded into each well of 96-well plates and cultured till 70% confluent. The cells were treated with EGF (20 ng/ml) with or without SAHA at indicated doses for 24 h. The cell proliferation was determined using CellTiter-Glo® Luminescent Cell Viability Assay kit (Promega, Madison, WI) with a luminometer (Wallac 1420 Victor2 multilabel counter system). The results were expressed as proliferation index (relative luminescence signal to medium control).

Flow cytometry assay

Cl41 cells were cultured in 6-well plates until 70%–80% confluent. Cell culture medium was replaced with 0.1% FBS medium for 36 h. The cells were then treated with EGF (20 ng/ml) with or without SAHA at indicated concentrations in the medium containing 1% FBS. Cells were fixed in ice-cold 70% ethanol and stained with PI buffer (0.1% Triton X-100, 0.2 mg/ml RNase A, and 0.05 mg/ml PI) for 15 min. The samples were subjected to flow cytometry (Beckman) for cell cycle analysis.

Western blottings

Cl41 cells and their transfectants (24 h after transfection) were cultured in each well of 6-well plates with normal medium until 70%–80% confluence. Cell culture medium was replaced by medium with 0.1% FBS for 36 h. Following that the culture medium was changed to MEM with 1% FBS and cells were treated with SAHA for 0.5 h followed by treatment with SAHA and/or EGF for the indicated concentrations and time periods. After exposure to EGF and SAHA, cells were washed with ice-cold PBS, and then extracted with cell lysis buffer (10 mM Tris–HCl, pH 7.4, 1% SDS, 1 mM Na3VO4, and proteasome inhibitor). The cell extracts were separated on polyacrylamide-SDS gels, transferred and probed with each of the antibodies against GAPDH (Cell Signaling, Beverly, MA), GFP, cyclin D1, VHL, HuR (Santa Cruz Biotechnology, Santa Cruz, CA), Nucleolin and β-Actin (Sigma, St. Louis, MO). The protein bands specifically bound to the primary antibodies were detected using alkaline phosphatase-linked secondary antibody and ECF (enhanced chemifluorescence) western blotting analysis system (Amersham Pharmacia Biotech, Piscataway, NJ) as previously described (Zhang et al., 2009).

Reverse transcription polymerase chain reaction (RT-PCR)

Cl41 cells and their transfectants (24 h after transfection) were cultured in 6-well plates until 70%–80% confluence. Cell culture medium was changed to 0.1% FBS medium for 36 h and then changed to 1% FBS medium and cells were exposed to SAHA with or without EGF and Actinomycin D (Act D), in the same manner as the cells treated for western blotting assay. After treatment for indicated time periods, total RNAs were extracted from cells using Trizol reagent (Invitrogen, Carlsbad, California). Total cDNAs were synthesized using oligdT(20) primer by SuperScript™ First-Strand Synthesis system (Invitrogen, Carlsbad, California). cyclin D1, GFP-cyclin D1 and β-actin mRNA amounts presenting in the cells were determined by semiquantitative RT-PCR assay. Mouse cyclin D1 (forward 5′-TCCCTTGACTGCCGAGAAG-3′, reverse 5′-AGACCAGCCTCTTCCTCCAC-3′) and β-actin (forward: 5′-CCTGTGGCATCCATGAAACT-3′, reverse: 5′-GTGCTAGGAGCCAGAGCA GT-3′) primers (Invitrogen) were used to determine the mRNA amount of endogenous cyclin D1 and β-actin, respectively. Human cyclin D1 (forward: 5′-GAGGTCTGCGAGGAACAGAAGTG-3′, reverse: 5′-GAGGGCGGATTGGAAATGAACTTC-3′) primer (Invitrogen) was used to detect the mRNA amount of reconstitutive GFP-cyclin D1 in Cl41 cells. The PCR products were separated on 3% agarose gels, stained with EB, and scanned the images from a UV light, as described previously (Zhang et al., 2009).

Luciferase assay

Cl41 and its stable transfectants transfected with AP-1-luciferase reporter plasmid, NF-κB-luciferase reporter plasmid, and cyclin D1 promoter luciferase reporter plasmid, respectively, were seeded into 96-well plates and starved by replacing culturing medium with 0.1% FBS MEM prior to exposure to EGF and SAHA, at indicated time of periods and concentration. Following that, cells were lysed for luciferase assay using luciferase substrate as previously described (Ouyang et al., 2007b).

Statistical analysis

The student’s t-test was used to determine the significance between treated and untreated group. The results are expressed as mean±SD.

Results

SAHA inhibited anchorage-independent colony formation induced by EGF in Cl41 cells

SAHA possesses significant inhibitory activity on cell proliferation in various human cancer cell lines (Musgrove et al., 2011). However, the potential inhibitory effect of SAHA on tumor promoter EGF-induced cell transformation has not been explored yet. Cells are capable of yielding anchorage-independent growth when neoplastic transformation occurs upon exposure to carcinogens or tumor promoters, such as TPA, EGF, and TNF-α. Soft agar has been considered to be the most accurate experimental method to detect cell capability of anchorage-independent growth (Anderson et al., 2007). EGF has been reported as a tumor promoter in many experimental systems, such as mouse epidermal Cl41 cells (DiGiovanni et al., 1994; Huang et al., 1999). Therefore, we employed the EGF-induced Cl41 cell transformation model to evaluate the inhibitory effect of SAHA. As shown in Fig. 1A, EGF-induced cell transformation was dramatically attenuated, in a dose-dependent manner, by co-incubation of cells with SAHA. Consistent with SAHA inhibition of EGF-induced cell transformation, SAHA treatment also led to a dramatic inhibition of Cl41 cell proliferation and induced a significant G0/G1 growth arrest without mediating cell apoptosis in a dose-dependent manner (Figs. 1B–D). These results indicate that SAHA treatment inhibits EGF-induced cell transformation and such inhibition of cell transformation might be associated with SAHA induction of G0/G1 growth arrest.

Fig. 1.

Fig. 1

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SAHA inhibited EGF-induced cell transformation, cell proliferation, and induced G0/G1 growth arrest in Cl41 cells. (A) Cl41 cells were exposed to indicated concentrations of SAHA in combination with EGF for cell transformation assay in soft agar as described in our published studies (Huang et al., 1999). After being cultured in a 37 °C 5% CO2 for three weeks, the colony formation was observed under microscope and the number of colonies was scored and presented as colonies per 104 seeded cells. The symbol (*) indicates a significant increase in comparison to that of medium control (p<0.05). The symbol (#) indicates a significant decrease in comparison to that with EGF treatment (p<0.05). (B) Cl41 cells were treated with SAHA and EGF as indicated for cell proliferation assay, and the results are expressed as cell proliferation index as described in Materials and methods. Each bar indicates the mean and SD of triplicate assays. The symbol (*) indicates a significant increase in comparison to that of medium control (p<0.05). The symbol (#) indicates a significant decrease in comparison to that with EGF treatment (p<0.05). (C and D) Cl41 cells were seeded into each well of 6-well plates. After cells were treated for 24 h with EGF and SAHA at indicated concentrations, the cell morphology were photographed under microscope (C); and the cell cycle was analyzed by PI staining with flow cytometry (D).

p21 induction by SAHA did not play a role in SAHA inhibition of anchorage-independent growth

p21 is a well-known cyclin-dependent kinase inhibitor and functions as a negative regulator of cell cycle at both G1 and S phase (Richon et al., 2000). Since SAHA has been reported to induce p21 protein expression (Gui et al., 2004; Richon et al., 2000), we determined whether SAHA treatment was able to upregulate p21 expression in Cl41 cells. As shown in Fig. 2A, SAHA treatment resulted in p21 induction in Cl41 cells in a dose-dependent manner. To evaluate the potential role of p21 induction in SAHA inhibition of anchorage-independent growth, we compared the SAHA inhibition of anchorage-independent growth in p21-defecient cells (HCT116/p21−/−) with its parental HCT116 cells with wild-type p21 expression. The identification of the p21 induction in HCT116 cells and deficient of p21 expression in HCT116/p21−/− cells was indicated in Fig. 2B. Our results revealed that SAHA treatment did show a similar inhibition of anchorage-independent cell growth between HCT116 cells and HCT116/p21−/− cells (Fig. 2C). These results strongly suggest that SAHA inhibition of anchorage-independent growth was through p21-independent mechanism.

Fig. 2.

Fig. 2

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SAHA inhibited EGF-induced cell transformation and cyclin D1 expression in Cl41 cells. (A and B) Cl41 and HCT116 cells were treated with SAHA in the indicated concentrations for 12 h. Western blotting assay was carried out to determine p21 expression. (C) The capacity of anchorage-independent colony formation between HCT 116 and HCT 116 p21−/− was performed. The symbol (*) indicates a significant decrease as compared with medium control (p<0.05). Each bar indicates the mean and SD of triplicate assays. (D) Cl41 cells were pretreated with SAHA for 0.5 h and then co-treated with EGF and SAHA for 12 h and the expression of cyclin D1 was determined by western blot.

SAHA treatment inhibited expression of endogenous cyclin D1, but not exogenous cyclin D1

HDAC inhibitor has shown to suppress cyclin D1 at the protein degradation and translation level (Alao et al., 2006; Kawamata et al., 2007). To determine whether cyclin D1 expression is downregulated upon SAHA treatment, Cl41 cells were treated with SAHA for various time periods. As shown in Fig. 3A, SAHA treatment significantly inhibited both the basal level and the EGF-induced level of cyclin D1 protein expression in a time-dependent manner. To elucidate the molecular mechanisms for SAHA inhibition of cyclin D1 protein expression, SAHA was applied to cells transfected with GFP-cyclin D1. The results showed that SAHA treatment did not show any observable inhibitory effect on exogenous reconstitutional expression of GFP-cyclin D1, whereas it did attenuate endogenous cyclin D1 expression in same cells (Fig. 3B). Consistent with SAHA inhibition of cyclin D1 protein expression, these results demonstrated from RT-PCR assay, indicated that SAHA inhibited endogenous cyclin D1 mRNA expression (Figs. 3C and D), but did not affect reconstitutional exogenous cyclin D1 mRNA expression (Fig. 3D). Our results strongly suggest that SAHA is able to inhibit cyclin D1 mRNA expression in Cl41 cells through independent of protein degradation level.

Fig. 3.

Fig. 3

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SAHA attenuated expression of endogenous cyclin D1, but not of constitutive expressed cyclin D1. (A) Cl41 cells were pretreated with SAHA for 0.5 h, and then co-treated with EGF and SAHA at the indicated time periods. The cyclin D1 expression levels were determined by western blot. (B and D) Cl41 cells transfected with GFP-tagged cyclin D1 and its vector control transfectant were exposed to SAHA for the indicated time periods, and cyclin D1 protein (B) and mRNA (D) were determined by western blot and RT-PCR, respectively. (C) Cl41 cells were treated with EGF and SAHA for the indicated time periods and RT-PCR was performed to determine cyclin D1 mRNA expression.

SAHA treatment reduced cyclin D1 mRNA stability, but did not affect cyclin D1 transcription

The above results suggested that SAHA was able to inhibit cyclin D1 mRNA expression in Cl41 cells, revealing that cyclin D1 down-regulation may occur at transcriptional level. AP-1 and NF-κB have been considered to be two of the major transcription factors that are responsible for the regulation of cyclin D1 expression (Barre and Perkins, 2007; Shaulian and Karin, 2001; Witzel et al., 2010). To test the potential inhibitory effect of SAHA on EGF-induced AP-1 and NF-κB activity, Cl41 stable transfectants with either AP-1-luciferase reporter (Cl41 AP-1) or NFκB-luciferase reporter (Cl41 NF-κB) were employed. As shown in Figs. 4A and B, SAHA co-treatment significantly enhanced EGF-induced both AP-1 activity and NF-κB activity. Furthermore, SAHA also promoted cyclin D1 promoter-driven luciferase reporter activity following EGF incubation (Fig. 4C). Our results demonstrated that transcriptional regulation is not involved in SAHA downregulating cyclin D1 mRNA expression. Thus, we anticipated that SAHA treatment might decrease cyclin D1 mRNA stability. This notion was greatly supported by the results showing that co-treatment of cells with SAHA and actinomycin D increased cyclin D1 mRNA degradation rate in comparison to that observed in cells treated with actinomycin D alone (Fig. 4D). We further determined the effect of SAHA on the half-life of cyclin D1 mRNA. The results showed that the cyclin D1 mRNA half-life in Cl41 cells was about 6.95 h, whereas SAHA treatment reduced the cyclin D1 mRNA half-life to 2.57 h (Figs. 4E and F).

Fig. 4.

Fig. 4

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SAHA promoted cyclin D1 mRNA decay. (A–C) Cl41 cells stably transfected with AP-1 luciferase reporter plasmid (A), NF-κB luciferase reporter plasmid (B), and cyclin D1 promoter luciferase reporter plasmid (C) were pretreated with SAHA in the indicated concentration for 0.5 h followed by co-treatments with SAHA and EGF for 12 h. The luciferase activity was measured as described in Materials and methods. The results were presented as relative AP-1 activity (A), relative NF-κB activity (B) and relative cyclin D1 promoter activity (C). The symbol (*) indicates a significant increase as compared with that of medium control (p<0.05). The symbol (#) indicates a significant increase as compared with those treated with EGF alone (p<0.05). (D) Cl41 cells were treated with actinomycin D (20 μM) with or without SAHA (2 μM) for 6 h. RNA was then isolated and RT-PCR was performed to determine cyclin D1 mRNA levels. (E and F) Cl41 cells were seeded into 6-well plates and cultured till 70–80% confluent. The medium was replaced with 0.1% FBS MEM for 36 h for synchronization of cell cycle. The cells were then cultured in 2% FBS MEM for 8 h. The cyclin D1 mRNA half life was assessed by treatment of actinomycin D (20 μM) in the presence or absence of SAHA (2 μM) for 2 and 4 h. The result was a representative one from three independent experiments (E), and cyclin D1 mRNA half-life was graphically displayed (F). (G) Cl41 cells were cultured in 6-well plate till 70–80% confluent. The cells were treated with different concentrations of SAHA for 0.5 h followed by co-treatment with EGF and SAHA for additional time periods as indicated. The cell extracts were then subjected to western blot assay.

HuR downregulation by SAHA was associated with its reduction of cyclin D1 mRNA degradation

Several RNA-binding proteins, such as HuR, Nucleolin, and VHL, have been reported to bind their target mRNA and increase mRNA stability (Abdelmohsen and Gorospe, 2010; Audic and Hartley, 2004; Ginisty et al., 1999). Thus, we tested whether those RNA-binding proteins were involved in the SAHA downregulation of cyclin D1 mRNA stability. The results showed that although SAHA treatment led to dramatically reduction of HuR protein expression, it did not show any observable inhibitory effect on the expression of either VHL or Nucleolin (Fig. 4G). Taking into consideration the fact that HuR has particularly been well-documented to up-regulate cyclin D1 mRNA stability through binding with its AU rich domain of 3′-UTR (Abdelmohsen and Gorospe, 2010; Audic and Hartley, 2004), we anticipated that HuR downregulation by SAHA would be responsible for its biological effect on regulating cyclin D1 mRNA stability.

Discussion

The histone deacetylase inhibitor SAHA has shown strong inhibition of cancer cell growth in various human cancers via inducing cell apoptosis and causing cell cycle arrest (Musgrove et al., 2011). However, the preventive effect and mechanism of SAHA on skin tumor promotion upon EGF treatment has not been identified. In the current study, we have demonstrated that SAHA inhibited EGF-induced cell proliferation and cell transformation in mouse epidermal JB6 Cl41 cells, which suggested the chemopreventive effect of SAHA on skin carcinogenesis. In addition, our studies showed that SAHA inhibited the expression of both basic level and EGF-induced cyclin D1 expression with induction of G0/G1 cell growth arrest. Further study showed that SAHA regulated cyclin D1 mRNA stability, rather than either protein degradation or transcription of cyclin D1. Additional experiments indicated that SAHA might inhibit cyclin D1 mRNA stability via down-regulation of HuR expression. Since published studies from various groups, including ours, had reported that cyclin D1 induction was responsible for cell transformation in various experimental systems (Bianchi et al., 1993; Ouyang et al., 2008; Yamamoto et al., 2002), we anticipated that SAHA downregulation of cyclin D1 expression was responsible for its inhibition of EGF-induced Cl41 cell transformation via mediating G0/G1 growth arrest.

Aberrant expression of cyclin D1 has been observed in many human cancers (Knudsen et al., 2006; Motokura et al., 1991). For example, amplification of cyclin D1 mRNA has been observed in about 45% of breast carcinomas (Buckley et al., 1993). Liang et al. reported that in a total of 307 patients with nonmelanocytic skin cancer, more than 50% cases showed cyclin D1 overexpression, whereas few cases were observed in normal skin tissue (Liang et al., 2000). Clinic studies also suggested that incidents of cancer patients with elevated cyclin D1 expression were related to poor prognosis (Bae et al., 2001). Further investigation showed that cyclin D1 played the critical role in cancer development, especially in skin carcinogenesis (Bianchi et al., 1993; Yamamoto et al., 2002). Conditional cyclin D1 overexpression in transgenic mice was more sensitive to dimethylbenz [a]anthracene-induced skin tumor (Yamamoto et al., 2002). Our previous studies also showed that knockdown of cyclin D1 attenuated arsenite-induced cell transformation in Cl41 and HaCat cells (Ouyang et al., 2008; Zhang et al., 2009). The molecular mechanisms responsible for cyclin D1-mediated cancer cell growth have been investigated in many experimental systems. For example, early studies indicated that cyclin D1 overexpression promoted cancer cell proliferation through the regulation of cell cycle progression (Ouyang et al., 2005, 2008). Recently published work has shown that cyclin D1 expression increased DNA damage repair, thereby protecting cancer cells from radiation-induced cell apoptosis (Biliran et al., 2005; Jirawatnotai et al., 2011). As the critical protein for cancer cell growth and proliferation, cyclin D1 had been used as a target for cancer therapy. SAHA had been identified as processing significant anticancer activity via inducing cell growth arrest and apoptosis (Marks, 2007). However, there were no reports investigating the potential utilization of SAHA as a chemopreventive agent for prevention of skin carcinogenesis. In current studies, the EGF-induced Cl41 cell transformation model was employed to elucidate this possibility. Our results indicated that SAHA incubation suppressed EGF-induced Cl41 cell proliferation and transformation with induction of cell G0/G1 growth arrest in a dose-dependent manner, suggesting that SAHA might be an agent that could be used for the prevention of skin cancer development at the tumor promotion stage. In addition, our results showed that SAHA pre-treatment also attenuated EGF-induced cyclin D1 protein expression. The results obtained from ChIP (ChIP, chromatin immunoprecipitation) assay by Yamaguchi et al. demonstrated that SAHA (25 μM) could suppress PMA-induced cyclin D1 mRNA transcription through the inhibition of activated c-Jun binding to the cyclin d1 promoter region in esophageal squamous cell carcinoma cells (Yamaguchi et al., 2005). Another study suggested that SAHA treatment decreased cyclin D1 protein translation in mantle cell lymphoma cells, whereas mRNA and protein degradation of cyclin D1 was minimally affected (Kawamata et al., 2007). Moreover, eukaryotic translation initiation factor 4E binding protein (eIF4E-BP) was downregulated and the cap site binding activity of eIF4E was inhibited in SAHA-treated cells (Kawamata et al., 2007). Thus, it was thought that SAHA could downregulate cyclin D1 expression in cancer cells at transcriptional and translational levels. However, here we found that SAHA treatment specifically inhibited endogenous cyclin D1 expression at both protein and mRNA levels, whereas it didn’t affect constitutively expressed cyclin D1. These results excluded the possibility of protein degradation involvement in the inhibition of cyclin D1 expression in EGF-treated Cl41 cells by SAHA. Furthermore, we found that although cyclin D1 mRNA was significantly downregulated, cyclin D1 promoter activity was slightly increased upon SAHA treatment, indicating that SAHA might regulate cyclin D1 expression via the modulating its mRNA stability in EGF-treated Cl41 cells. This notion was greatly supported by the results showing that SAHA treatment markedly increased cyclin D1 mRNA degradation and decreased cyclin D1 mRNA half-life in the cells co-treated with actinomycin D. Taken together, we demonstrated that SAHA inhibited EGF-induced cyclin D1 expression by mediating cyclin D1 mRNA degradation. This mechanism was distinctly different from the results that have been reported in cancer cells.

Regulation of mRNA stabilization is one of the major mechanisms responsible for cells controlling protein expression and is regulated by multiple proteins (Lal et al., 2004; Marderosian et al., 2006). Microarray analyses suggested that approximately 40%–50% of changes in gene expression in response to extracellular treatment occurred due to altered mRNA stability (Benjamin and Moroni, 2007; Garneau et al., 2007). The defect in regulation of mRNA stability might lead to complicated disorders, including cancers (Benjamin and Moroni, 2007). The mRNA of some oncogenes, such as c-fos and c-myc, are stabilized and overexpressed in many cancer cells, which mediates cancer cell proliferation (Aghib et al., 1990; Blancato et al., 2004; Meijlink et al., 1985). It has also been reported that Ras-induced oncogenic transformation involved in abnormal of gadd153 mRNA degradation (Rong et al., 2005) and the alteration of cyclin D1 mRNA stability has been implicated in cancer cell proliferation and cell transformation (Nguyen-Chi and Morello, 2008). For example, cyclin D1 mRNA with 3′ truncation was identified in human breast cells. This truncation could result in increasing cyclin D1 mRNA stability and overexpression in human breast cancer cells (Lebwohl et al., 1994). In mantle cell lymphoma, point mutation and genomic deletion led to cyclin D1 mRNA lack of degradation elements in 3′-UTR, which increased cyclin D1 mRNA stability and promoted a high proliferation of lymphoma cells. Clinical studies indicate that the patients who express full-length cyclin D1 mRNA live much longer than those that express the truncated cyclin D1 mRNA (Wiestner et al., 2007). The results obtained from in vitro experiments also demonstrate that cyclin D1 mRNA stability could be regulated via AU-rich elements (ARE) located in the 3′-untranslated region (3′-UTR) (Audic and Hartley, 2004). Several RNA binding proteins, including HuR, AUF1 and Tristetraprolin (TTP), have been reported to bind to 3′-UTR of cyclin D1 mRNA and regulate its turnover (Lal et al., 2004; Marderosian et al., 2006). HuR, containing three RNA binding motifs, was able to stabilize mRNA via recognization and interaction with AU-rich domain of target genes (Abdelmohsen and Gorospe, 2010). Knockdown of HuR significantly promoted cyclin D1 mRNA decay and inhibited cell colony formation in oral cancer cells, indicating the essential role of HuR in mediating the oncogenic effect on cancer cells (Kakuguchi et al., 2010; Lal et al., 2004). Here we found that SAHA downregulated HuR expression, accompanied by a reduction of cyclin D1 mRNA stability, inhibition of EGF-induced Cl41 cell transformation, as well as induction of G0/G1 growth arrest. Our results strongly suggest that HuR downregulation might be responsible for SAHA inhibition of cyclin D1 expression and cell transformation via induction of G0/G1 cell growth arrest.

In summary, our studies demonstrate the chemopreventive effect of SAHA on EGF-induced cell proliferation and transformation in mouse epidermal Cl41 cells, is associated with its induction of cell G0/G1 growth arrest by downregulation of HuR expression and reduction of cyclin D1 mRNA stability. These results highlight a novel mechanism underlying SAHA regulation of cyclin D1 expression in Cl41 cells, which is distinctly different from the findings that have been reported in cancer cells.

Acknowledgments

This work was supported in part by grants from NIH/NCI CA112557, NIH/NIEHS ES000260 and ES010344, NSFC30928023 and NSFC 30971516.

Footnotes

Conflict of interest

There’s no conflict of interest.

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