AP-1 expression and DNA binding activity in human oral keratinocytes

Oral Oncology 38 (2002) 281–290 www.elsevier.com/locate/oraloncology

Acetaldehyde activates Jun/AP-1 expression and DNA binding activity in human oral keratinocytes Sherry R. Timmonsa, Joseph O. Nwankwob, Frederick E. Domannc,* a

Oral Sciences Graduate Program, B180ML, The University of Iowa, Iowa City, IA 52242, USA b Department of Biochemistry, B180ML, The University of Iowa, Iowa City, IA 52242, USA c Free Radical and Radiation Biology Graduate Program and Holden Comprehensive Cancer Center, B180ML, The University of Iowa, Iowa City, IA 52242, USA Received 9 April 2001; accepted 22 April 2001

Abstract Oral cancer is a significant health problem, particularly among individuals that ingest alcohol in combination with the use of tobacco products. The enhanced development of tobacco-initiated oral cancers by ethanol suggests that ethanol or one of its metabolites may act as a type of tumor promoter. Nevertheless, the mechanisms underlying the ability of ethanol to enhance oral carcinogenesis remain unclear. We hypothesize that acetaldehyde, the first metabolite of ethanol, may activate the expression and/ or activity of Jun/AP-1 in oral keratinocytes analogous to the phorbol ester TPA and other tumor promoters in epidermal keratinocytes. To test this hypothesis, we treated HPV immortalized, non-tumorigenic human oral keratinocytes with acetaldehyde at various concentrations and for various times and measured several parameters of Jun/AP-1expression and function. Our results indicated that c-Jun mRNA and protein levels increased in the acetaldehyde treated cells compared to untreated control cells. Moreover, Jun/AP-1 DNA binding activity was rapidly activated by acetaldehyde in a dose-dependent fashion. The increases in Jun protein and AP-1 DNA binding activity were accompanied by increased transactivation of an AP-1 responsive reporter construct as well as increased transcript levels of a candidate AP-1 responsive gene, stromelysin 3. The levels of acetaldehyde employed were minimally toxic to the cells as determined by MTT assays. Thus, acetaldehyde was found to activate the expression and activity of an oncogenic transcription factor in HPV-initiated cells. Taken together, these results suggest that acetaldehyde may participate, at least in part, in the promotion stage of oral carcinogenesis. # 2002 Elsevier Science Ltd. All rights reserved. Keywords: Alcohol; Oral cancer; Transcription factor; Tumor promotion; Carcinogenesis

1. Introduction Ethanol consumption is known to be a contributing risk factor for the development of oral squamous cell carcinoma. There is a correlation between history of ethanol exposure and the development of head and neck cancer, especially cancer of the oral cavity. In humans, the main source of ethanol exposure is through direct consumption [1]. However, ethanol itself is not mutagenic, thus the mechanisms underlying ethanolassociated oral carcinogenesis are not known. It has been suggested that ethanol might potentiate the effects of a carcinogen by facilitating the penetration of carcinogens across the oral mucosa. Studies examining * Corresponding author. Tel.: +1-319-335-8018; fax: +1-319-3358039. E-mail address: [email protected] (F.E. Domann).

the permeability constants of various oral tissues revealed an increase in the permeability to nitrosonornicotine (NNN) of the floor of mouth mucosa in the presence of 5% ethanol when compared to NNN alone [2]. Although this permeability effect cannot be entirely ruled out as a mechanism for ethanol’s effects on enhancing carcinogen-induced oral cancer, it seems more likely that a biologically derived metabolite of ethanol may act as a kind of tumor promoter, thus stimulating the proliferation and clonal expansion of the carcinogen-initiated cells. Many cells and tissues contain alcohol dehydrogenases (ADH) and cytochrome p450s (CYP) that metabolize ethanol to acetaldehyde. These include ADH3 and CYP2E1 [3–8]. In fact, an ADH enzyme has been isolated from human gingival and tongue homogenates [9]. In addition to generation of acetaldehyde by endogenous host enzymes, a growing body of evidence

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suggests that acetaldehyde production in the oral cavity is likely due to the indigenous microflora that inhabit the oral cavity [10–13]. It has been suggested that acetaldehyde, the first metabolite of ethanol, might be the causal factor in ethanol-associated oral cancers [14,15]. Acetaldehyde has been shown to cause DNA damage [16–20] and is known to be a mutagenic agent in various cell types. DNA damage can influence intracellular pathways, such as cell activation, growth, and proliferation. Acetaldehyde production after ethanol exposure has been shown to be higher in salivary flow of patients with head and neck cancer than in normal controls [12]. Moreover, acetaldehyde production in the oral cavity has been associated with poor dental status [21]. Microflora in the oral cavity, including bacteria and yeast, could lead to concentrations of acetaldehyde in the oral cavity sufficient to cause cellular effects. Nevertheless, the molecular mechanism(s) by which acetaldehyde exerts changes in cellular gene expression or phenotype is unknown. One possibility is that acetaldehyde poses a redox stress to the cell that leads to changes in gene expression through redox sensitive transcription factors such as NF-kB or AP-1. Activator protein 1 (AP-1) is a redox sensitive transcription factor complex that binds specific target DNA sequences in the promoter region of other genes and induces transcription of those genes. AP-1 is composed of members from the Jun and Fos protein families as well as members of the ATF transcription factor family. c-Jun is a member of the Jun family and is capable of forming homodimers or heterodimers with Fos and ATF family members. c-Jun is encoded by the c-jun gene, an immediate early response gene inducible by external stimuli such as the phorbol esters TPA, growth factors such as epidermal growth factor, cytokines, and carcinogens [22]. The jun family of genes have a proposed role in cell function during proliferation, differentiation, and apoptosis. For example, malignant epidermal keratinocytes exhibit higher constitutive expression of c-jun mRNA and oncoprotein and demonstrate enhanced AP-1 DNA binding activity and transactivating activity [23]. Moreover, interfering with Jun/AP-1 function with a dominant negative Jun protein caused reversion of the transformed phenotype and inhibited tumor formation in vivo [24]. Hybridization techniques to measure oncogene expression in total RNA preparations from head and neck squamous cell carcinomas as a means to evaluate chemotherapeutic response revealed an increase in c-jun expression in these types of cancers [25]. In addition, human epidermal keratinocytes exposed to ethanol displayed an increase in c-jun expression [26]. Tumor promoters such as TPA and okadaic acid lead to rapid and transient increases in c-jun expression and AP-1 DNA binding and transactivating activity [22,27– 31]. We hypothesized that acetaldehyde might contribute

to the oral carcinogenic process by altering c-jun protooncogene expression and AP-1 activity in oral keratinocytes and thus act in a manner analogous to that of a classical tumor promoting agent.

2. Materials and methods 2.1. Cell culture HPV immortalized oral keratinocytes (HOK-16B) were cultured in keratinocyte basal medium (KBM) supplemented with growth factors (Clonetics Corp., San Diego, CA) at 37
C with 5% CO2 in air. Cells were trypsinized and reseeded at a 1:3 dilution every 5 days. A 1 mM stock solution of acetaldehyde (Fisher Scientific, Fair Lawn, NJ) in keratinocyte basal medium was prepared fresh before each use. The cell cultures were incubated with acetaldehyde at varying concentrations (0–100 mM) in basal keratinocyte medium. In some cases, cells were incubated in KBM supplemented with fetal bovine serum to measure serum inducible levels of c-jun and AP-1 activation. Incubation of cells with 100 ng/ml okadaic acid (Sigma-Aldrich, St. Louis, MO) served as a positive control for AP-1 DNA binding and transactivating activity. 2.2. MTT Assay for Cell Viability HOK-16B cells were seeded on 60 mm tissue culture plates (Corning Inc, Corning, NY) at 1.3106 cell density and incubated for 3 days. Cells were removed from serum supplemented medium and incubated in KBM for 18 h. Cells were treated with varying concentrations of acetaldehyde (0–100 mM) for 6 h at 37
C with 5% CO2 in air. Treatment medium was removed and cells were incubated in KBM containing MTT (3-(4,5dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide) for 2 h at 37
C with 5% CO2 in air. The medium was removed and the cells were gently rinsed twice with PBS (pH 7.4). Isopropanol (1 ml ) was added and absorbance read for 200 ml of the sample solution at 595 nm in a DU-64 Beckman spectrophotometer. 2.3. RNA isolation and northern blot analysis Total RNA was isolated from 90% confluent cells using the QIAGEN RNeasy kit (Chatsworth, CA). Quantitation was by absorbance measurement at 260 nm with the DU-64 spectrophotometer (Beckman Instruments Inc., Fullerton, CA). Ten micrograms of total RNA for each sample were electrophoresed in 1.2% agarose/2.2 M formaldehyde gels, transferred to nylon membranes (GeneScreen, New England Nuclear, Boston, MA) in 10saline/ sodium citrate and UV cross-linked (Stratalinker 1800

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auto-crosslink, Stratagene, La Jolla, CA). Blots were prehybridized, hybridized, and washed using standard techniques. The probe, a 0.63-kb partial cDNA of human c-jun (GenBank accession no. J04111; nt 16302255), was labeled with [a-32P] dCTP with the Prime-it II random priming kit (Stratagene) according to manufacturer’s instructions. The c-jun cDNA sequence was confirmed, plasmids digested with EcoRI and cDNA fragments purified from 1.5% agarose gels using QIAGEN Gel Extraction Kit before using as probes in Northern analysis. The membrane was washed under high-stringency conditions and exposed to X-ray film overnight. The membrane was stripped by immersing in a boiling water bath for 5 min and re-probed with a 1.6kb full length cDNA of human stromelysin 3 (GenBank accession no. XM009873) labeled with [a-32P] dCTP using the Prime-it II random priming kit (Stratagene) according to manufacturer’s instructions. The membrane was stripped again and re-probed with the labeled human glyceraldehyde-3-phosphate dehydrogenase (GAPDH) cDNA probe to enable a relative quantitation of c-jun transcripts.

The proteins were transferred to the membrane in transfer buffer at 30 V overnight at 4
C. The membrane was washed twice in TBST (10 mM Tris, pH 8.0, 150 mM sodium chloride, and 0.05% Tween-20 (Fisher Scientific Company, Fair Lawn, NJ), and blocked in 5% nonfat milk-TSBT for 1 h at RT with agitation. The membrane was washed twice for 5 min in TBST with agitation. Membrane was incubated with primary antibody (anti-c-Jun, SC-45, Santa Cruz Biotechnology, Santa Cruz, CA) diluted 1:200 in 5% nonfat milk-TBST for 18 h at 4
C with agitation. The membrane was washed four times for 10 min in TBST with agitation, then incubated with secondary antibody (anti-rabbit IgG conjugated with horseradish peroxidase, Boehringer Mannheim Corp., Indianapolis, IN) diluted 1:10,000 in 5% nonfat milk-TBST for 1 h at RT with agitation. The blot was washed for 10 min in TBST with agitation and washed twice for 20 min in TBST containing 0.5 M NaCl with agitation. An ECL detection kit (Amersham Corp., Arlington Heights, Il) was used according to manufacturer’s instructions to detect c-Jun protein bands. The blot was exposed to X-ray film for 1 min.

2.4. Preparation of protein extracts for Western analysis

2.6. Preparation of nuclear extracts and gel mobility shift assays

Tissue culture plates were washed twice with PBS. The plates were washed once with PB buffer (pH 7.8). Cells were scrape-harvested and the cell suspensions were placed in 1.5 ml eppendorf tubes. Suspensions were centrifuged for 5 min at 8160g and 4
C. The supernatant was removed. The pellet was resuspended in 50 ml PB buffer (pH 7.8) cell suspensions were sonicated for 15 s on ice. The cellular proteins were quantified using Biorad Protein Assay dye reagent concentrate (Bio-Rad Laboratories, Richmond, CA). The absorbance was measured at 595 nm on an ELISA plate reader and protein concentration determined. Twenty micrograms of protein were incubated with sample buffer (0.5 M Tris (pH 7), glycerol, 10% SDS, 2-mercaptoethanol (Sigma Chemical Company, St. Louis, MO), and 1% bromophenol blue dye (Fisher Scientific Company, Fair Lawn, NJ), heated for 5 min at 100
C, and cooled to room temperature. Proteins were separated by SDSPAGE with a 3% stacking gel and a 10% resolving gel. The gel was electrophoresed in an ice bath for 3 h at 100 V. 2.5. Western blotting for c-JUN protein The separated proteins were transferred to a nitrocellulose membrane (Schleicher and Schuell, Keene, NH) using the following method, the gel was soaked for 10 min in transfer buffer (3.03 g Tris, 14.45 g Glycine, 800 ml dd H2O, 200 ml Methanol; pH 8.3) and assembled into transfer apparatus (sponge–Whatman filter paper– gel–nitrocellulose membrane–Whatman filter paper).

Using 0.5 ml of cold Buffer A (10 mM HEPES, pH 7.9, 1.5 mM MgCl2, 10 mM KCL, and 0.5 mM dithiothreitol (DTT) the cells were scraped from the tissue culture plates. The cells were incubated on ice for 30 min. Cells were lysed using a Dounce homogenizer (Kontes Scientific Glassware, Vineland, NJ) and centrifuged for 30 s at 500g. The supernatant was removed and the cells recentrifuged for 30 s. The supernatant was removed and the pellets were resuspended in 15 ml cold Buffer C (20 mM HEPES, ph 7.9, 25% glycerol, 0.42 M NaCl, 1.5 mM MgCl2, 0.2 mM EDTA, 0.5 mM phenylmethylsulfonyl fluoride, and 0.5 mM DTT). The suspended nuclei were incubated on ice for 15 min and microcentrifuged for 5 min at 4
C and 16,000g. The supernatants were harvested and diluted 1:3 in cold Buffer D (20 mM HEPES, ph 7.9, 20% glycerol, 0.1 M KCl, 0.2 mM EDTA, 0.5 mM phenylmethylsulfonyl fluoride, and 0.5 mM DTT). The protein concentrations of the extracts were determined with BCA protein assay reagent (Pierce, Rockford, IL) according to manufacturer’s instructions. The oligonucleotide probe jun1URE, which contains the AP-1 binding site from nt 72 to63 of the c-jun promoter, 50 -agcttgTGACATCAccgtag-30 , was used to evaluate the formation of AP-1 DNA complexes after treatment with acetaldehyde. Probes were labeled with [a-32P] dCTP and Klenow DNA Polymerase. DNA binding assays were performed by incubating 2 mg of nuclear proteins with approximately 200,000 cpm of probe in the presence of 1 mg of poly (dIdC) (Pharmacia Inc.,

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Piscataway, NJ) and 1  gel shift buffer (10 mM Tris, pH 7.5, 50 mM NaCl, 1 mM MgCl2, 0.5 mM EDTA, 0.5 mM DTT, and 4% glycerol) at room temperature for 20 min. For the antibody clearance experiments, 1 ml of anti-c-Jun antibody (SC-45, Santa Cruz Biotechnology, Santa Cruz, CA) was incubated with the nuclear extracts for 1 h at room temperature before the probe was added. The binding reactions were loaded onto a 5% polyacrylamide gel and electrophoresed at 35 mA for about 1 h in 1 X TBE buffer. The gel was wrapped in plastic wrap and exposed to X-ray film overnight at 80
C.

activity, 10 ml complete scintillation fluid counting cocktail Budget-Solve (Research Products International Corp, Mt. Prospect, IL) was added to 200 ml of the upper xylene layer. The samples were counted using a Beckman liquid scintillation counter. For the b-galactosidase assay, the 40 ml of extract was incubated with 3 ml 100  Magnesium buffer (0.1 M MgCl2 and 4.5 M bmercaptoethanol), 66 ml 1  OPNG, and 191 ml 0.1 M sodium phosphate (1 M Na2HPO4 2H2O and 1 M NaH2PO4 2H2O). The samples were incubated at 37
C for 30 min and 500 ml of 1 M Na2CO3 was added to stop the reaction. The b-galactosidase activity was measured spectrophotometrically with absorbance read at 420 nm.

2.7. Plasmids, transfections and reporter assays The plasmid Col-CAT contains the collagenase promoter with an AP-1 binding site fused to the chloramphenicol acetyltransferase (CAT) reporter gene [22]. The plasmid CMV-b galactosidase was co-transfected to measure transfection efficiency (Clontech Laboratories, Inc., Palto Alto, CA). Cells were seeded at a density of 1106 cells per well on a 6-well plate (Corning Inc., Corning, NY). The cells were cultured in KBM supplemented with growth factors for 24 h. The medium was replaced with serum-free KBM. At 70% confluency, the cells were transfected with plasmid constructs using LipofectAMINE Plus reagent (Life Technologies Inc., Rockford, MD), according to manufacturer’s instructions. Each well contained 15 ml Lipofectamine Plus and 5 mg of either Col-CAT, CMV-bgal, or 1:4 ratio of ColCAT and CMV-bgal plasmid DNA, incubated at 37
C with 5% CO2 for 3.5 h. The transfection medium was removed, and cells were treated with serum-free KBM, with or without acetaldehyde (1 mM) or okadaic acid (100 ng/ml), incubated at 37
C with 5% CO2 for 24 h. The cells were harvested at the indicated times by scraping in 0.5 ml of 0.25 M Tris (pH 7.5). The cell suspensions were microcentrifuged for 1 min at room temperature and 500g, supernatant was removed, and the pellet was resuspended in 100 ml of 0.25 M Tris (pH 7.5). The cell suspensions were placed on ice and sonicated for 10 s. For the b-galactosidase assay, 50 ml of cell lysate was removed from the sonicant and stored in a separate tube. The remaining lysate was heated at 65
C for 10 min and cooled on ice. Lysates were centrifuged at 500g for 5 min at room temperature and the supernatant was used for the CAT assay. For the CAT assays, 50 ml of cellular protein was incubated with 10 ml n-butyryl-CoA (Sigma Chemical Co., Rockford, Il), 200,000 cpm [1,2-14C]chloramphenicol (ICN, Costa Mesa, CA), and 0.25 M Tris-HCL (pH 8.0) at 37
C for 20 h. The samples were briefly microcentrifuged at 500g and 300 ml mixed xylenes (EM Science, Gibbstown, NJ) were added to terminate the reaction. The samples were vortexed and microcentrifuged at 500g for 5 min at room temperature. To measure CAT

3. Results 3.1. Acetaldehyde caused c-jun mRNA accumulation in oral keratinocytes HOK-16B cells were treated with varying concentrations of acetaldehyde and c-jun mRNA was measured at various time points by Northern blot analysis. Cells were left untreated or treated with 1 mM or 5 mM acetaldehyde and total RNA was isolated from the cells after 2, 4 and 6 h of acetaldehyde exposure. c-jun mRNA levels were assessed using Northern blot hybridization with a c-jun specific cDNA probe. The results from this experiment are shown in Fig. 1. The cells treated with acetaldehyde at either dose showed a marked increase in c-jun mRNA within 2 h after treatment compared to untreated cells at the same time point (Fig. 1, lanes 3–5). When cells were exposed to acetaldehyde for 4 h, there was also a marked increase compared to untreated control cells, particularly at the 1 mM concentration (Fig. 1, lanes 6–8). After 6 h of exposure to 1 mM acetaldehyde, c-jun transcript levels were still above the basal level in untreated cells; however, at the 5 mM acetaldehyde dose, c-jun mRNA levels were similar to the basal expression in untreated control cells (Fig. 1, lanes 9–11). These results indicate that acetaldehyde induced an increase in steady state levels of c-jun mRNA in the HOK-16B cells and that the increase occurred at early time points, peaking as early as 2 h. While the higher acetaldehyde concentration (5 mM) did increase c-jun transcript levels above the basal levels in untreated cells, the higher concentration did not induce greater mRNA levels than those detected at the lower acetaldehyde concentration (1 mM). In fact, after 6 h of exposure to acetaldehyde, cjun mRNA levels in cells treated with 5 mM acetaldehyde were similar to those in untreated cells at the same time point. To normalize for equal sample loading and transfer efficiency, we stripped the membrane and re-probed it with a cDNA probe specific for glyceraldehyde-3-phosphate dehydrogenase (GAPDH).

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Fig. 1. Acetaldehyde induces c-jun mRNA accumulation in HPVimmortalized human oral keratinocytes. HPV immortalized HOK-16B cells were treated with acetaldehyde and then assayed at various time points for c-jun mRNA accumulation by Northern blot analysis with a human c-jun cDNA probe. Lane 1, HOK-16B cells treated with 100 ng/ml okadaic acid (OA) for 6 h (positive control); lane 2, HOK-16B cells in serum containing medium (SM) for 6 h. Lanes 3–11, HOK16B cells in serum free medium for 18 h followed by treatment with either 1 mM or 5 mM acetaldehyde for 2, 4, or 6 h as indicated on the figure. Typical results obtained from replicate experiments are shown.

Results from re-hybridizing the membrane with GAPDH verified equal loading and transfer of all the lanes (Fig. 1, lower panel). These results are consistent with the observation that c-jun is an early response gene whose expression is induced in response to external stimuli such as tumor promoters. 3.2. Acetaldehyde exposure increases c-Jun protein expression in oral keratinocytes Acetaldehyde increased c-Jun oncoprotein in a dose and time dependent manner. Cellular proteins were isolated from HOK-16B cells at 6 or 12 h after acetaldehyde addition, and c-Jun protein levels were assessed using Western blot analysis with anti- c-jun antibodies. Cells were treated with 1 or 5 mM acetaldehyde. At 6 h, cells had a low basal level of c-Jun protein and a marked increase in protein production after exposure to 5 mM acetaldehyde (Fig. 2, lanes 3 and 4). At 12 h in serum free medium (SFM), cells demonstrated a higher basal level of protein than at 6 h in SFM (Fig. 2, lane 5). Cells treated with 1 and 5 mM acetaldehyde for 12 h (lanes 6 and 7, respectively) expressed higher levels of c-Jun protein compared to cells treated with acetaldehyde for 6 h, however, no further increase in protein expression was observed in cells exposed to 1 mM versus cells

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Fig. 2. Acetaldehyde induces accumulation of c-Jun protein in HPVimmortalized human oral keratinocytes in a time and dose dependent manner. HOK-16B cells were exposed to either 1 mM or 5 mM acetaldehyde and then total cellular proteins were isolated and analyzed by western blot analysis using an antibody specific for c-Jun. Lane 1, purified recombinant AP-1 protein; lane 2, HepG2 cells were serum deprived for 18 h followed by treatment with 500 uM acetaldehyde for 6 h. Lanes 3 and 4, HOK-16B cells were incubated in serum deprived media for 18 h, then treated either with or without acetaldehyde as indicated on the figure. Lane 3, HOK-16B cells after an additional 6 h in serum-deprived medium (SFM); lane 4, treatment with 5 mM acetaldehyde for an additional 6 h. Lanes 5–7, HOK-16B cells were incubated in SFM for 18 h, then treated either with or without acetaldehyde as indicated on the figure. Lane 5, HOK-16B cells after an additional 12 h in serum-deprived medium (SFM); lane 6, treatment with 1 mM acetaldehyde for an additional 12 h; lane 7, HOK16B treatment with 5 mM acetaldehyde for an additional 12 h. N.S., nonspecific binding. Typical results are shown from replicate Western blots.

treated with 5 mM acetaldehyde for 12 h. Two bands were detected on Western blot analysis. The slower migrating band did not demonstrate any marked alteration whether or not acetaldehyde was present. This slower band likely represents non-specific binding by the antibody and thus serves as a useful internal control for equal amounts of protein loading and transfer in the Western blot. These findings indicate that acetaldehyde induced an increase in c-Jun protein in the HOK-16B cells. 3.3. Acetaldehyde enhances the DNA binding activity of AP-1 transcription factor Electrophoretic gel mobility shift assay was utilized to assess the effect of acetaldehyde on AP-1 DNA binding activity. Cells were treated with either okadaic acid (OA, 100 ng/ml) or acetaldehyde (Acet, 1 mM) in serum free medium. Alterations in AP-1 DNA binding activity were examined at 6, 12, and 24 h. There was a low basal level of AP-1 DNA binding activity with okadaic acid treatment at 6 h (Fig. 3a, lane 4) and this activity was similar to the serum induced DNA binding activity (Fig. 3a, lane 3). Acetaldehyde treatment for 6 h induced a marked increase in DNA binding activity (Fig. 3a, lane 5). Okadaic acid treatment at both 12 and

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24 h also demonstrated a marked increase in DNA binding activity (Fig. 3a, lanes 6 and 8, respectively) and these increases were greater than the induction noted with either OA or acetaldehyde at 6 h. Acetaldehyde treatment for 12 h induced an increase in binding activity, however, at 24 h the level of induction was similar to that seen at 6 h acetaldehyde exposure (Fig. 3a, lanes 7 and 9). These findings indicate that both okadaic acid and acetaldehyde were capable of inducing an increase in AP-1 DNA binding activity in HOK-16B cells, and though the induction by acetaldehyde was not as robust as that seen with okadaic acid, acetaldehyde induced an earlier and more persistent response than okadaic acid. To determine that the previously shifted complex specifically involved c-Jun/AP-1, electrophoretic gel mobility supershift assays were performed with an antic-jun antibody. Cells were treated with 1 mM acetaldehyde for 6 or 12 h. Nuclear extracts were isolated and incubated either with or without anti-c-jun antibody. Consistent with previous results, acetaldehyde induced DNA binding activity at both 6 and 12 h and the induction was slightly increased at 6 h (Fig. 3b, lanes 1 and 2, respectively). Incubating extracts with anti-c-jun antibody disrupted the formation of the complex and resulted in a decrease in the degree of AP-1DNA binding activity (Fig. 3b, lanes 3 and 4). These findings indicate that the protein binding the target DNA sequence is specifically c-Jun/AP-1. 3.4. Acetaldehyde increases AP-1 mediated transactivation through a cis-element in the collagenase promoter

Fig. 3. Acetaldehyde causes a pronounced increase in DNA binding activity of the Jun/AP-1 complex in HPV-immortalized human oral keratinocytes as determined by gel mobility shift assay. (a) Time course and dose response for acetaldehyde mediated activation of AP1 DNA binding activity. Lane 1, probe only; lane 2, HOK-16B cells in serum containing medium; lane 3, HOK-16B cells after 24 h in serum free medium. Lanes 4–9, HOK-16B cells were serum deprived for 18 h followed by treatment with either okadaic acid (100 ng/ml) or acetaldehyde (1 mM). Lane 4, okadaic acid for an additional 6 h; lane 5, acetaldehyde for an additional 6 h; lane 6, okadaic acid for an additional 12 h, lane 7, acetaldehyde for an additional 12 h; lane 8, okadaic acid for an additional 24 h; lane 9, acetaldehyde for an additional 24 h. OA, okadaic acid. Acet, acetaldehyde. (b) Acetaldehyde induced formation of an AP-1 specific DNA binding complex. Antibody specific for c-jun competes for c-Jun binding in formation of the Jun/AP-1DNA complex in HPV-immortalized oral keratinocytes as determined by gel supershift mobility assay. HOK-16B cells were serum-deprived for 18 h followed by treatment with 1 mM acetaldehyde for varying times of exposure. Lane 1, HOK-16B cells treated with acetaldehyde for an additional 6 h; lane 2, HOK-16B cells treated with acetaldehyde for an additional 12 h; lane 3, HOK-16B cells treated with acetaldehyde for an additional 6 h and nuclear extracts were incubated with anti-c-Jun antibody; lane 4, HOK-16B cells treated with acetaldehyde for an additional 12 h and nuclear extracts were incubated with anti-c-Jun antibody. Typical results obtained from replicate experiments are shown.

To determine the effects of acetaldehyde on inducing an AP-1 responsive gene we performed transient transfections using a collagenase-CAT promoter reporter construct [22]. The collagenase promoter sequence utilized in the transfection assays contains an AP-1 response element as previously described [22]. Cells were transiently transfected using LipofectAMINE reagent in serum free medium followed by treatment with okadaic acid (100 ng/ml) or acetaldehyde (1 mM) for 24 h. Cells were lysed and CAT activities were determined. Untreated cells demonstrated a low basal level of CAT activity while cells treated with okadaic acid or acetaldehyde demonstrated higher CAT activity. Cells treated with okadaic acid demonstrated a tenfold increase in CAT activity compared to untreated cells (Fig. 4). Cells treated with acetaldehyde demonstrated a twofold increase in CAT activity compared to untreated cells (Fig. 4). Although the transactivation response to acetaldehyde was not as great as to OA, the AP-1 cis element was clearly transactivated by acetaldehyde. These results indicated that acetaldehyde activated an AP-1 responsive cis-element in human oral keratinocytes, and suggested that acetaldehyde may function as a

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Fig. 4. Acetaldehyde caused transactivation of an AP-1 responsive ciselement in the human collagenase promoter in HPV-immortalized human oral keratinocytes. Acetaldehyde induced transactivation of the collagenase promoter was measured in HOK-16B cells transiently transfected with a chloramphenicol acetyl transferase (CAT) reporter fused to an AP-1 responsive cis-element from the human collagenase promoter. Cells were serum-deprived for 18 h and transiently transfected using LipofectAMINE reagent for 3.5 h. Lane 1, HOK-16B cells were transfected with 5 mg pCMV-bgal vector; lane 2, HOK16B cells were co-transfected with 4 mg Col-CAT and 1 mg pCMV-bgal vectors; lane 3, HOK-16B cells were co-transfected with 4 mg Col-CAT and 1 mg pCMV-bgal vectors followed by treatment with 100 ng/ml okadaic acid for 24 h; lane 4, HOK-16B cells were co-transfected with 4 mg Col-CAT and 1 mg pCMV-bgal vectors followed by treatment with 1 mM acetaldehyde for 24 h (n=3).

tumor-promoting stimulus in part by activating AP-1 responsive genes in preneoplastic cells. 3.5. Acetaldehyde caused accumulation of mRNA encoding stromelysin-3 (MMP-11), a candidate AP-1 responsive gene, in oral keratinocytes HOK-16B cells were treated with varying concentrations of acetaldehyde and stromelysin-3 (MMP-11) mRNA was measured at various time points by northern blot analysis. Cells were left untreated or treated with 1 mM or 5 mM acetaldehyde and total RNA was isolated from the cells after 2, 4 and 6 h of acetaldehyde exposure. Stromelysin-3 mRNA levels were assessed using northern blot hybridization with a stromelysin-3 specific cDNA probe. The results from this experiment are shown in Fig. 5. The cells treated with 1 mM acetaldehyde for either 4 or 6 h showed a marked increase in stromelysin-3 mRNA compared to untreated control cells at the same time points (Fig. 5, lanes 6–7 and 9–10, respectively). After 4 or 6 h exposure to 5 mM acetaldehyde, stromelysin-3 transcript levels were similar to the basal expression in untreated control cells (Fig. 5, lanes 8 and 11). These results indicate that acetaldehyde induced an increase in steady state levels of stromelysin3 mRNA in the HOK-16B cells and that the increase occurred at early time points, peaking as early as 4 h. However, the time course for stomelysin-3 mRNA accumulation was delayed relative to that of c-jun mRNA accumulation as would be expected if increases

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Fig. 5. Acetaldehyde induces stromelysin-3 mRNA accumulation in HPV-immortalized human oral keratinocytes. HPV immortalized HOK-16B cells were treated with acetaldehyde and then assayed at various time points for stromelysin- mRNA accumulation by Northern blot analysis with a human stromelysin-3 cDNA probe. Lane 1, HOK-16B cells treated with 100 ng/ml okadaic acid (OA) for 6 h (positive control); lane 2, HOK-16B cells in serum containing medium (SM) for 6 h. Lanes 3–11, HOK-16B cells in serum free medium for 18 h followed by treatment with either 0, 1, or 5 mM acetaldehyde for 2, 4, or 6 h as indicated on the figure.

in c-Jun/AP-1 were responsible for the downstream upregulation of stromelysin-3. While the higher acetaldehyde concentration (5 mM) did not induce greater mRNA levels than those detected at basal level, the lower acetaldehyde concentration (1 mM) did induce a marked increase in transcript levels. In fact, even after 6 h of exposure to acetaldehyde, stromelysin-3 mRNA levels in cells treated with 5 mM acetaldehyde were similar to those in untreated cells at the same time point. The membrane was re-probed with a [a-32P]-labeled, 1.4 kb full length human collagenase-1 cDNA probe and there was no alteration in collagenase-1 mRNA levels after exposure to varying concentrations of acetaldehyde for various time points (data not shown). To normalize for equal sample loading and transfer efficiency, we stripped the membrane and re-probed it with a cDNA probe specific for glyceraldehydes-3-phosphate dehydrogenase (GAPDH). Results from re-hybridizing the membrane with GAPDH verified equal loading and transfer of all the lanes (Fig. 5, lower panel). These results suggest that stromelysin-3, a candidate AP-1 responsive gene, is induced in response to acetaldehyde. 3.6. Acetaldehyde caused a minimal decrease in cell survival at the concentrations employed to induce accumulation of c-jun mRNA, protein, and DNA binding activity To determine cell survival in response to acetaldehyde we performed the MTT assay. Cells were treated for 6 h with serum-deprived medium or varying concentrations of acetaldehyde in SFM. Cells treated with 10 mM or less acetaldehyde demonstrated a high percentage (87– 100%) of cell survival (Fig. 6). Concentrations of acetaldehyde greater than 10 mM resulted in a marked decrease in cell survival. Cells treated with 25, 50, or 100

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Fig. 6. Acetaldehyde had limited toxicity to HPV-immortalized oral keratinocytes at doses sufficient to activate c-Jun expression and activity. Cell viability was determined by the MTT assay. Cells were serum-deprived for 18 h followed by treatment of varying concentration of acetaldehyde for 6 h. Cells were treated with 0, 1, 10, 25, 50, and 100 mM acetaldehyde in serum free medium (n=4).

mM acetaldehyde resulted in a 50–70% decrease in cell survival in comparison to untreated cells (59, 50.8, and 33.3%, respectively). These results suggested that acetaldehyde was toxic to this cell line only at extremely high concentrations and the dosages utilized throughout our study, 1 or 5 mM, were non-toxic to the HOK-16B cells over the time period studied.

4. Discussion Oral squamous cell carcinoma is a malignant lesion that causes a significant amount of morbidity and mortality. Oral cancers are often not detected until advanced stages of disease when treatment options are limited. Epidemiological evidence has implicated two major contributing factors to increased incidence of human oral cancer as tobacco and ethanol. Although ingestion of ethanol is known to be a contributing risk factor for the development of oral squamous cell carcinoma, the mechanism by which this agent enhances oral carcinogenesis is not known. Ethanol is metabolized to acetaldehyde, an agent known to cause DNA damage and oxidative stress. Our study tested the hypothesis that acetaldehyde, the first metabolite of ethanol, may be a contributing factor for human oral carcinogenesis by causing increased expression of the c-jun protooncogene and increased Jun/AP-1 DNA binding and transactivating activity in a manner similar to that of known tumor promoters such as TPA and OA. It is widely accepted that epithelial carcinogenesis is a multi-step process involving initiation, promotion, and progression. Previous studies have evaluated the effect of human papilloma virus (HPV) infection and carcinogenic compounds in oral keratinocytes [32–34]. Cells infected with HPV and exposed to benzo(a)pyrene or

tobacco specific compounds demonstrated malignant phenotypic changes, while cells infected by HPV alone remained immortal but not tumorigenic. Our studies were performed under the premise that acetaldehyde, and thus ethanol indirectly, would also act as a tumor promoter on cells previously initiated by human papilloma virus (HPV) infection. There are several potential sources of acetaldehyde as a result of ethanol metabolism in mammals. Ethanol can be metabolized through both oxidative and nonoxidative mechanisms as well as through both enzymatic and non-enzymatic pathways. Some of the enzymatic pathways involve tissue alcohol dehydrogenases (ADHs), the microsomal system involving enzymes such as CYP2E1, and the peroxisomal system. The non-enzymatic mechanisms involve metal chelation in the presence of hydroxyl radicals and non-oxidative mechanisms involve the formation of fatty acid ethyl esters from a fatty acid synthetase. For example, in the oral cavity the enzymes involved in ethanol metabolism, alcohol and acetaldehyde dehydrogenase (ADH and ALDH, respectively), have been isolated from tongue and attached gingiva [9]. It was determined that high Km, enzymatically active ADHs were present but low Km ALDHs lacked enzymatic activity in these tissues. Different isoforms of alcohol dehydrogenase have been linked with increased susceptibility to cancer development [4–7]. Also, CYP2E1, cytochrome P4502E1, can metabolize ethanol to form reactive oxygen species. In HepG2 cells stably transfected with CYP2E1, exposure to ethanol was toxic and reduced cell viability, increased LDH leakage, and altered cellular morphology when compared to cells transfected with a control vector [35]. Besides tissue sources, microflora indigenous to the oral cavity are also capable of producing acetaldehyde from ethanol. These microflora can utilize ethanol/acetaldehyde as an energy source under anaerobic conditions through alcohol dehydrogenase. Studies evaluating the role of microbial flora in acetaldehyde production have shown that pre-treatment with the antiseptic chlorohexidine decreased the amount of acetaldehyde produced from ethanol metabolism [10,12]. Interestingly, a previous study evaluating the mouthwashings of patients with oral/pharyngeal cancers found that patients with cancer produced more acetaldehyde from ethanol than patients without cancer, suggesting a link between cancer development and the capacity to metabolize ethanol [12]. Thus, microbial production of acetaldehyde may play a role in the exposure of oral keratinocytes to acetaldehyde. Previous studies have examined the effect of tumor promoting agents such as TPA and okadaic acid on Jun/AP-1 expression in other cells lines [23,31]. Our studies demonstrated that exposing HPV immortalized oral keratinocytes to okadaic acid induced an increase in c-jun mRNA and Jun/AP-1 transactivating activity.

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Studies have demonstrated an increase in c-jun expression and AP-1 DNA binding activity when HepG2 cells were exposed to acetaldehyde [36]. Similarly, when we exposed HPV immortalized oral keratinocytes to acetaldehyde we detected an increase in c-jun mRNA, c-Jun oncoprotein, Jun/AP-1 DNA binding activity, and AP-1 transactivating activity. AP-1 is a transcription factor capable of turning on the transcription of target genes. AP-1 responsive genes include matrix metalloproteinases, such as stromelysin and collagenase [23,37,38]. These enzymes function to degrade the extracellular matrix and allow cells to violate the basement membrane barrier, a hallmark of malignancy. Our promoterreporter studies utilized a fragment of the collagenase promoter containing an AP-1 response element and we demonstrated that acetaldehyde could induce AP-1 to stimulate transcription from this promoter element. Moreover, treatment of HOK16B cells with acetaldehyde led to an increase in steady state mRNA levels of the extra-cellular matrix degrading enzyme stromelysin3 (MMP-11). Together with collagenase-1, this gene has previously been implicated as a candidate AP-1 target gene [29]. In addition, stromelysin-3 is associated with increased local invasiveness in head and neck squamous cell carcinomas [39]. Taken together, these findings suggest a possible mechanism for acetaldehyde, an ethanol metabolite, in promoting oral carcinogenesis. Further studies are needed to fully elucidate the relationship between acetaldehyde, AP-1, and oral cancer. Studies transfecting a dominant negative AP-1 (TAM67) into oral keratinocytes would be valuable in further examining a relationship between acetaldehyde and AP-1 [24]. These studies would further support the contention that acetaldehyde may exert some if not all of its cellular alterations through AP-1. Studies examining the tumorigenic potential in nude mice by HPV immortalized oral keratinocytes exposed to acetaldehyde would help establish a relationship of acetaldehyde as a tumor promoting agent. In our in vitro studies, HPV-immortalized, non-tumorigenic oral keratinocytes demonstrated an increase in c-jun mRNA, c-Jun oncoprotein, and Jun/AP-1 DNA binding and transactivating activity with exposure to acetaldehyde. Taken together, these data suggest that acetaldehyde induction of c-jun expression may play a role in oral keratinocyte carcinogenesis by acting as a promoting agent on previously initiated cells.

Acknowledgements We would like to thank Dr. No Hee Park for generously donating the HPV-immortalized oral keratinocytes utilized in this research project. This work was supported in part by Public Health Service grant CA-73612 to F.E.D. from the National Cancer Institute. S.R.T.

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received salary support from Dental Scientist Training Grant K16-DE-00175 from the National Institutes of Health. References [1] Riveros-Rosas H, Julian-Sanchez A, Pina E. Enzymology of ethanol and acetaldehyde metabolism in mammals. Archives of Medical Research 1997;28(4):453–71. [2] Squier CA, Cox P, Hall BK. Enhanced penetration of nitrosonornicotine across oral mucosa in the presence of ethanol. Journal of Oral Pathology 1986;15(5):276–9. [3] Bakker BM, Bro C, Kotter P, Luttik MA, van Dijken DP, Pronk JT. The mitochondrial alcohol dehydrogenase Adh3p is involved in a redox shuttle in Saccharomyces cerevisiae. Journal of Bacteriology 2000;182(17):4730–7. [4] Borras E, Coutelle C, Rosell A, Fernandez-Muixi F, Broch M, Crosas B. Genetic polymorphism of alcohol dehydrogenase in europeans: the ADH2*2 allele decreases the risk for alcoholism and is associated with ADH3*1. Hepatology 2000;31(4):984–9. [5] Bosron WF, Lumeng L, Li TK. Genetic polymorphism of enzymes of alcohol metabolism and susceptibility to alcoholic liver disease. Molecular Aspects of Medicine 1988;10(2):147–58. [6] Bouchardy C, Hirvonen A, Coutell C, Ward PJ, Dayer P, Benhamou S. Role of alcohol dehydrogenase 3 and cytochrome P-4502E1 genotypes in susceptibility to cancers of the upper aerodigestive tract. International Journal of Cancer 2000;87(5): 734–40. [7] Coutelle C, Ward PJ, Fleury B, Quattrocchi P, Chambrin H, Iron A, Couzigou P, Cassaigne A. Laryngeal and oropharyngeal cancer, and alcohol dehydrogenase 3 and glutathione S-transferase M1 polymorphisms. Human Genetics 1997;99(3):319–25. [8] Kunitoh S, Imaoka S, Hiroi T, Yabusaki Y, Monna T, Funae Y. Acetaldehyde as well as ethanol is metabolized by human CYP2E1. Journal of Pharmacology and Experimental Therapeutics 1997;280(2):527–32. [9] Dong YJ, Peng TK, Yin SJ. Expression and activities of class IV alcohol dehydrogenase and class III aldehyde dehydrogenase in human mouth. Alcohol 1996;13(3):257–62. [10] Homann N, Jousimies-Somer H, Jokelainen K, Heine R, Salaspuro M. High acetaldehyde levels in saliva after ethanol consumption: methodological aspects and pathogenetic implications. Carcinogenesis 1997;18(9):1739–43. [11] Homann N, Tillonen J, Meurman JH, Rintamaki H, Lindqvist C, Rautio M, Jousimies-Somer H, Salaspuro M. Increased salivary acetaldehyde levels in heavy drinkers and smokers: a microbiological approach to oral cavity cancer. Carcinogenesis 2000;21(4):663–8. [12] Jokelainen K, Heikkonen E, Roine R, Lehtonen H, Salaspuro M. Increased acetaldehyde production by mouthwashings from patients with oral cavity, laryngeal, or pharyngeal cancer. Alcoholism: Clinical and Experimental Research 1996;20(7):1206–10. [13] Jones AW. Measuring and reporting the concentration of acetaldehyde in human breath. Alcohol and Alcoholism 1995;30(3): 271–85. [14] Harty LC, Caporaso NE, Hayes RB, Winn DM, Bravo-Otero E, Blot WJ, et al. Alcohol dehydrogenase 3 genotype and risk of oral cavity and pharyngeal cancers [see comments]. Journal of the National Cancer Institute 1997;89(22):1698–705. [15] Tillonen J, Homann N, Rautio M, Jousimies-Somer H, Salaspuro M. Role of yeasts in the salivary acetaldehyde production from ethanol among risk groups for ethanol-associated oral cavity cancer. Alcoholism: Clinical and Experimental Research 1999;23(8):1409–15. [16] Dellarco VL. A mutagenicity assessment of acetaldehyde. Mutation Research 1988;195(1):1–20.

290

S.R. Timmons et al. / Oral Oncology 38 (2002) 281–290

[17] Feron VJ, Kruysse A, Woutersen RA. Respiratory tract tumours in hamsters exposed to acetaldehyde vapour alone or simultaneously to benzo(a)pyrene or diethylnitrosamine. European Journal of Cancer & Clinical Oncology 1982;18(1):13–31. [18] Helander A, Lindahl-Kiessling K. Increased frequency of acetaldehyde-induced sister-chromatid exchanges in human lymphocytes treated with an aldehyde dehydrogenase inhibitor. Mutation Research 1991;264(3):103–7. [19] Nakamura K, Iwahashi K, Itoh M, Ameno K, Ijiri I, Takeuchi Y, Suwaki H. Immunohistochemical study on acetaldehyde adducts in alcohol-fed mice. Alcoholism: Clinical and Experimental Research 2000;24(4 Suppl):93S–96S. [20] Ristow H, Seyfarth A, Lochmann ER. Chromosomal damages by ethanol and acetaldehyde in Saccharomyces cerevisiae as studied by pulsed field gel electrophoresis. Mutation Research 1995;326(2):165–70. [21] Homann N, Tillonen J, Rintamaki H, Salaspuro M, Lindquist C, Meurman JH. Poor dental status increase acetaldehyde production from ethanol in saliva: a possible link to increased oral cancer risk among heavy drinkers. Oral Oncology 2001;37:153–8. [22] Angel P, Imagawa M, Chiu R, Stein B, Imbra RJ, Rahmsdorf HJ, Jonat C, Herrlich P, Karin M. Phorbol ester-inducible genes contain a common cis element recognized by a TPA-modulated trans-acting factor. Cell 1987;49(6):729–39. [23] Domann FE, Levy JP, Finch JS, Bowden GT. Constitutive AP-1 DNA binding and transactivating ability of malignant but not benign mouse epidermal cells. Molecular Carcinogenesis 1994;9(2):61–6. [24] Domann FE, Levy JP, Birrer MJ, Bowden GT. Stable expression of a c-Jun deletion mutant in two malignant mouse epidermal cell lines blocks tumor formation in nude mice. Cell Growth and Differentiation 1994;5(1): 9–16. [25] Riva C, Lavieille JP, Reyt E, Brambilla E, Lunardi J, Brambilla C. Differential c-myc, c-jun, c-raf and p53 expression in squamous cell carcinoma of the head and neck: implication in drug and radioresistance. European Journal of Cancer Part B, Oral Oncology 1995;31B(6):384–91. [26] Kharbanda S, Nakamura T, Kufe D. Induction of the c-jun proto-oncogene by a protein kinase C-dependent mechanism during exposure of human epidermal keratinocytes to ethanol. Biochemical Pharmacology 1993;45(3):675–81. [27] Kim SJ, Lafyatis R, Kim KY, Angel P, Fujiki H, Karin M, Sporn MB, Roberts AB. Regulation of collagenase gene expression by okadaic acid, an inhibitor of protein phosphatases. Cell Regulation 1990;1(3):269–78. [28] Spandidos DA. Mechanism of carcinogenesis: the role of onco-

[29]

[30]

[31]

[32]

[33]

[34]

[35]

[36]

[37]

[38]

[39]

genes, transcriptional enhancers and growth factors. Anticancer Research 1985;5(5):485–98. Young MR, Li JJ, Rincon M, Flavell RA, Sathyanarayana BK, Hunziker R, Colburn N. Transgenic mice demonstrate AP-1 (activator protein-1) transactivation is required for tumor promotion. Proceedings of the National Academy of Sciences of the United States of America 1996;96(17):9827–32. Zhou H, Lin A, Gu Z, Chen S, Park NH, Chiu R. 12-O-tetradecanoylphorbol-13-acetate (TPA)-induced c-Jun N-terminal kinase (JNK) phosphatase renders immortalized or transformed epithelial cells refractory to TPA-inducible JNK activity. Journal of Biological Chemistry 2000;275(30):22868–75. Peng J, Bowden GT, Domann FE. Activation of AP-1 by okadaic acid in mouse keratinocytes associated with hyperphosphorylation of c-jun. Molecular Carcinogenesis 1997;18(1):37–43. Li SL, Kim MS, Cherrick HM, Doniger J, Park NH. Sequential combined tumorigenic effect of HPV-16 and chemical carcinogens. Carcinogenesis 1992;13(11):1981–7. Kim MS, Shin KH, Baek JH, Cherrick HM, Park NH. HPV-16, tobacco-specific N-nitrosamine, and N-methyl-N0 -nitro-N-nitrosoguanidine in oral carcinogenesis. Cancer Research 1993; 53(20):4811–6. Park NH, Gujuluva CN, Baek JH, Cherrick HM, Shin KH, Min BM. Combined oral carcinogenicity of HPV-16 and benzo(a)pyrene: an in vitro multistep carcinogenesis model. Oncogene 1995;10(11): 2145–53. Cederbaum AI, Wu D. Ethanol cytotoxicity to a transfected HepG2 cell line expressing human cytochrome P4502E1. Journal of Biological Chemistry 1996;271(39):23914–9. Roman J, Gimenez A, Lluis JM, Gasso M, Rubio M, Caballeria J, Pares A, Rodes J, Fernandez-Checa JC. Enhanced DNA binding and activation of transcription factors NF-kappa B and AP-1 by acetaldehyde in HEPG2 cells. Journal of Biological Chemistry 2000;275(19): 14684–90. Casini A, Ceni E, Salzano R, Milani S, Schuppan D, Surrenti C. Acetaldehyde regulates the gene expression of matrix-metalloproteinase-1 and-2 in human fat-storing cells. Life Sciences 1994;55(17):1311–6. Angel P, Karin M. Specific members of the Jun protein family regulate collagenase expression in response to various extracellular stimuli. Matrix Supplement 1992;1:156–64. Muller D, Wolf C, Abecassis J, Millon X, Engelmann A, Bronner G, Rouyer N, Rio MC, Eber M, Methlin G. Increased stromelysin 3 gene expression is associated with increased local invasiveness in head and neck squamous cell carcinomas. Cancer Research 1993;53(1):165–9.