Genotoxicity of 4-Hydroxy-2-Nonenal in Human Colon Tumor Cells Is Associated with Cellular Levels of Glutathione and the Modulation of Gluta
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《毒物学科学杂志》
Department of Nutritional Toxicology, Institute for Nutritional Sciences, Friedrich Schiller University, Dornburger Strae 25, D-07743 Jena, Germany
Department of Environmental and Occupational Health Sciences, University of Washington, 4225 Roosevelt Way NE, Suite 100, Seattle, Washington 98105
ABSTRACT
The cellular production of 4-hydroxy-2-nonenal (HNE), a product of endogenous lipid peroxidation, constitutes a genotoxic risk factor for carcinogenesis. Our previous studies have shown that human HT29 colon cells developed resistance toward HNE injury after treatment with butyrate, a diet-associated gut fermentation product. This resistance was attributed to the induction of certain glutathione S-transferases (hGSTP1-1, hGSTM2-2, and hGSTA1-1) and also for the tripeptide glutathione (GSH) synthesizing enzymes. In the present study, we have investigated in HT29 cells whether hGSTA4-4, which has a high substrate specificity for HNE, was also inducible by butyrate and, thus, could contribute to the previously observed chemoresistance. In addition, we investigated if cellular depletion of GSH by L-buthionine-S,R-sulfoximine (BSO) enhances chemosensitivity to HNE injury in HT29 cells. Incubation of HT29 cells with butyrate (2–4 mM) significantly elicited a 1.8 to 3-fold upregulation of steady state hGSTA4 mRNA over 8–24 h after treatment. Moreover, 4 mM butyrate tended to increase hGSTA4-4 protein concentrations. Incubation with 100 μM BSO decreased cellular GSH levels by 77% without significant changes in cell viability. Associated with this was a 2-fold higher level of HNE-induced DNA damage as measured by the comet assay. Collectively, the results of this study and our previous work indicate that the genotoxicity of HNE is highly dependent on cellular GSH status and those GSTs that contribute toward HNE conjugation, including hGSTA4-4. Since HNE contributes to colon carcinogenesis, the favorable modulation of the GSH/GST system by butyrate may contribute to chemoprevention and reduction of the risks.
Key Words: L-buthionine-(S,R)-sulfoximine (BSO); butyrate; DNA damage; glutathione (GSH); 4-hydroxy-2-nonenal (HNE); glutathione S-transferases; hGSTA4.
INTRODUCTION
The highly reactive aldehyde trans-4-hydroxy-2-nonenal (HNE) has a wide variety of biological activities. HNE is mutagenic and has strong effects on signal transduction pathways (Blanc et al., 1997; Chung et al., 1993; Esterbauer et al., 1990). It is an important endogenous factor of tissue injury risk, since it is the major electrophilic product of lipid peroxidation caused by oxidative stress (Chung et al., 1993). Formation of HNE is initiated by free radical-mediated degradation of -6-polyunsaturated fatty acids such as linoleic and arachidonic acid, which are relatively abundant fatty acids in membranes of human cells (Burcham, 1998; Esterbauer et al., 1990). Feng et al. (2004) described that conditions of oxidative stress potently enhanced cellular levels of HNE from 0.1–3 μM to 10 μM–5 mM. These types of oxidative stress, e.g., occur during chronic diseases like lymphoedemas (Siems et al., 2002) or during progression of colorectal cancer (Skrzydlewska et al., 2005). HNE can be further metabolized to an epoxide intermediate that interacts with DNA to form exocyclic etheno-guanine, -adenine, and -cytosine adducts (Chung et al., 1996). In addition, bulky exocyclic propane-type DNA adducts with guanine, and, in particular, 6-(1-hydroxyhexanyl)-8-hydroxy-1, N2-propano-2-deoxyguansine (4-HNE-dG) adducts may be formed in cellular reactions with HNE (Douki and Ames, 1994). We have recently shown that HNE is highly genotoxic to human colon tumor cells (Ebert et al., 2001). Additionally, damage to the tumor suppressor gene TP53 in adenoma cells and in primary colon cells has been shown using fluorescence in situ hybridization of the damaged gene (Schferhenrich et al., 2003a,b). Other investigators have recently reported that HNE-DNA adducts are preferentially formed at the third base of codons 249 and 174 in the TP53 gene (Hu et al., 2002). Collectively, these data emphasize the potentially important role of HNE in tumor initiation and progression (Cheeseman, 1993).
One of the primary pathways for detoxification of HNE in human cells is through conjugation with glutathione (GSH), a reaction, which is markedly enhanced by catalysis with certain glutathione S-transferases (GSTs) (Berhane et al., 1994). The alpha class GSTs, of which hGSTA1-1 is the most abundant form in human liver, are involved in HNE detoxification through GSH conjugation. In particular, hGSTA1-1 conjugates HNE with a Km = 50 μM and a kcat/Km = 0.058 s–1μM–1, whereas hGSTA4-4, a primarily mitochondrial GST, effectively conjugates HNE with an efficient Km = 49 μM and a very rapid kcat/Km of 2.7 s–1μM–1 (Cheng et al., 2001; Gardner and Gallagher, 2001). The human pi class isoform hGSTP1-1 can also catalyze the conjugation of HNE with GSH (Berhane et al., 1994). Comparatively, however, hGSTA4-4 exhibits the highest catalytic activity and turnover of HNE relative to the other GST isozymes. The specificity of GSTA4-4 is based on typical structural properties like an electrophilic substrate-binding pocket that consists of three structural modules. These form the binding site with a favorable positioning of Tyr212 to interact with the aldehyde group of the substrate and polarize it for reaction (Bruns et al., 1999). Because of this highly specific catalytic activity toward such aldehydes and related oxidants, it has been suggested that hGSTA4-4 and its rodent orthologues (mouse GSTA4-4 and rat GSTA4-4) have specifically evolved to protect against oxidative injury in vivo.
We have previously demonstrated that the gut fermentation product butyrate enhances the cellular levels of hGSTA1/2, hGSTM2-2, hGSTP1-1, but not hGSTM1-1 or hGSTT1-1 proteins in HT29 cells. Concomitant with the induction of these aforementioned GST isoforms were increases in GST catalytic activities, in hGSTP1 and hGSTM2 steady-state mRNA expression and also in cellular GSH levels (Ebert et al., 2003). However, what is not known is if butyrate exposure enhances the expression of hGSTA4-4, the primary HNE metabolizing GST (Hubatsch et al., 1998) and whether a downregulation of the GST/GSH system confers opposite effects, namely an enhanced chemosensitivity. In the present study we have investigated baseline expression levels of GSTA4-4 in the human colon cell line HT29 and tested the hypothesis that exposure to butyrate would result in induction of hGSTA4 mRNA and hGSTA4-4 protein expression. Once establishing the inducibility of hGSTA4-4, we further investigated if depletion of cellular GSH levels would increase the genotoxic response to HNE.
MATERIALS AND METHODS
Cells and reagents.
The human colon tumor cell line HT29 (ATCC, HTB-38) used in this study was established in 1964 and originated from human adenoma colon tissue (Fogh and Trempe, 1975). The HT29 cells were obtained from the American Tissue Culture Collection (ATCC, Rockville, MD). Sodium butyrate was purchased from Merck research laboratories (Schuchardt, Hhenbrunn, Germany). The Glutathione Assay-Kit and 4-hydroxynonenal (HNE) were purchased from Calbiochem-Novabiochem (Bad Soden, Germany). Positively charged nylon membranes, the DIG-RNA labeling kit, T7 RNA polymerase, anti-DIG-alkaline phosphatase antibody, and CDP-Star were supplied from Roche Diagnostics (Mannheim, Germany). Reverse transcriptase and oligo(dT) were purchased from SuperScript Invitrogen GmbH (Karlsruhe, Germany). The RNeasy mini kit and Hot Star Taq DNA Polymerase were purchased from Qiagen (Hilden, Germany). TRIzol-Reagent and cell culture materials, including Dulbecco's Modified Eagle Medium (DMEM), RPMI 1640 Minimal Essential Medium without L-glutamine (RPMI 1640), penicillin/streptomycin, and fetal calf serum (FCS) were purchased from Invitrogen GmbH (Karlsruhe, Germany). Goat anti-chicken IgY conjugated with horseradish peroxidase was purchased from CalBiotech (Santa Cruz). 4',6-Diamidino-2-phenylindole dihydrochloride (DAPI) and L-buthionine-(S,R)-sulfoximine (BSO) were purchased from Sigma-Aldrich (Deisenhofen, Germany). Enhanced Chemiluminescence (ECL)TM Western Blotting Detection Reagents were purchased from Amersham Biosciences Europe GmbH (Freiburg, Germany).
Culture of HT29 cells.
The human colon tumor cell line HT29 was maintained in DMEM supplemented with 10% (v/v) FCS and penicillin (50 U/ml)/streptomycin (50 μg/ml) in a humidified 5% CO2 incubator at 37°C, where it grew strictly adherent. Under given laboratory conditions, the HT29 cells doubled their population number within 24 h (unpublished data). The experiments determining hGSTA4 mRNA and hGSTA4-4 protein expression were performed with passages from 30 to 53, and passages from 17 to 40 of HT29 cells were used for GSH-depletion trials and the comet assay studies. To retain standardized conditions, all experiments were performed using subconfluently grown cells, which 72 h after seeding had reached a degree of confluence of at least 80%.
Analysis of cell viability and growth.
The trypan blue exclusion test was used to determine membrane integrity reflecting cell viability after harvesting or incubating the cells in culture or in suspension. Studies on modulation of growth were performed to assess effective concentrations of BSO. For these experiments, the cells were seeded (10 x 103/well) into 96-well microtiter plates and allowed to attach. After 24 h, the cells were treated with different concentrations of BSO (0.001 mM) dissolved in culture medium. Survival was detected after 24 and 72 h by quantifying DNA with DAPI, a cell permanent stain that specifically binds double-stranded DNA. This exhibits a 20-fold higher fluorescence enhancement, which does not occur with single-stranded DNA (Tecan, 1997). By subsequent fluorometric analysis with ex/em 360/465 nm using a 96-well microplate Reader (Beyer-Sehlmeyer et al., 2003; Ebert et al., 2001) the total amount of DNA was measured and was used to represent the amount of surviving cells.
Cell treatment with butyrate.
Prior to treatment with sodium butyrate, 2 x 106 HT29 cells were allowed to attach and grow in DMEM for 48 h in 25-cm2 culture flasks. For experiments of mRNA and protein expression, the cells were treated with 2 and 4 mM sodium butyrate for 24 and 48 h. The investigated concentrations of butyrate were based on the EC50 and EC25 for cell survival after 48 h butyrate treatment as previously determined (Ebert et al., 2001). In addition, the treatment of the group receiving 2 mM butyrate was stopped after 1, 4, 8, 12, 16, or 24 h to characterize the time-dependent expression levels of hGSTA4 mRNA in culture.
Detection of hGSTA4 expression by Northern blotting and semiquantitative PCR.
Total RNA was isolated from HT29 cells using the TRIzol method (Chomczynski and Sacchi, 1987). After RNA isolation, 15 μg of total RNA was loaded onto a 1.2% denaturing agarose gel, separated for 3–4 h at 70 V, and blotted on a positively charged nylon membrane. Digoxygenin (DIG)-labeled probes for hGSTA4 and the glyceraldehyde-3-phosphate dehydrogenase (GAPDH) were prepared from total RNA by reverse transcriptase PCR followed by a gene-specific PCR to amplify the cDNAs and to introduce the T7 RNA polymerase promoter sequence for in vitro transcription. In vitro transcription and simultaneous DIG-labeling of hGSTA4 and GAPDH were performed using the DIG-RNA labeling kit and T7 RNA polymerase. The membrane was hybridized with 1 μg probe in 10 ml hybridization buffer overnight at 70°C. After washing with 2 and 0.3x saline sodium citrate (SSC) containing 0.1% sodium dodecyl sulphate (SDS), the membrane was incubated in blocking solution for 1 h at room temperature, followed by antibody incubation for further 30 min. Detection of the anti-DIG-alkaline phosphatase antibody was performed with CDP-Star on a HyperfilmTM ECL. The film was photographed for evaluation of the band intensities and the Quantity One 4.1 software (Bio-Rad) was used for quantitative evaluation. For the quantification of the expression levels, the peak densities of the bands were used. After the hGSTA4 mRNA levels were normalized with GAPDH, the peak densities of untreated cells were equated with 100%. The results represent the mean of the hGSTA4 mRNA expression values (±SEM) for the several treatments of six independent experiments.
Semiquantitative reverse transcriptase-PCR (RT-PCR) was used to quantitate and confirm the initial results from the Northern blotting experiments. For these experiments, total RNA was extracted with the RNeasy mini kit according to the manufacturer's instructions. A cDNA equivalent to 5 μg total RNA was prepared by first-strand synthesis using oligo(dT) primer (SuperScriptTM First-Strand Synthesis System). Semiquantitative RT-PCR was used to amplify GSTA4 mRNA levels and to compare its expression in butyrate-treated cells versus untreated cells. The expression of -actin served as an internal control. To avoid different cDNA concentrations between target gene (hGSTA4) and control gene (-actin), for each sample the template cDNA was added to the PCR reaction mix without primers. The mixture was aliquoted into two batches, one for GSTA4 and one for -actin. Subsequently, gene-specific PCR primers were added for hGSTA4 (Genbank NM_001512 5'-ACC TGG CAG CAA GAA GAA G-3', sense and 5'-CAG GAT AAG GAG AGC AGA AAG-3', antisense) and -actin (NM_001101 5'-GCT CGT CGT CGA CAA CGG CTC-3' sense, and 5'-TGG GTC ATC TTC TCG CGG TTG G-3', antisense). Based upon these published cDNA sequences, the predicted size of the hGSTA4 cDNA is 435 bp, and that of -actin is 337 bp. PCR reactions consisted of cDNA equivalent to 0.6 mg/25 ml of RNA, 1.25 U/25 ml of Hot Star Taq DNA Polymerase, either 0.25 mM of -actin primers or 0.5 mM of GSTA4 primers, and a final MgCl2 concentration of 1.5 mM. The cDNA was amplified by PCR, with 21 cycles for -actin and 28 cycles for GSTA4. After primary denaturation at 95°C for 5 min, the PCR cycles were run at 95°C for 15 s, 55°C for 30 s, and 72°C for 1 min. Densitometry evaluation of the ethidium bromide bands was performed with the Quantity One 4.1 Software (Bio-Rad). The results were expressed as ratio of hGSTA4 mRNA to -actin mRNA.
Relative quantification of GSTA4 mRNA transcript by real-time qPCR.
Expression of GSTA4 mRNAs was assessed by quantitative real-time PCR (SYBR Green I) system. One μl of cDNA (10 ng of total RNA equivalent) was used in a 25 μl PCR amplification reaction containing 2x iQ SYBR Green supermix (100 mM KCl, 40 mM Tris-HCl, pH 8.4, 0.4 mM each dNTP, 50 U/ml iTaq DNA polymerase, 6 mM MgCl2, SYBR Green I, 20 nM fluorescein, stabilizers), and 10 pmol each of target (GSTA4) and reference (-actin) gene specific primers. A region of GSTA4 and the -actin mRNA was amplified using these iGSTA4-F (sense 5'-CCGGATGGAGTCCGTGAGATGG-3'), iGSTA4-R (antisense 5'-CCATGGGCACTTGTTGGAACAGC-3'), i-actin_F (sense 5'-GCTCGTCGTCGACAACGGCTC-3'), and i-actin_R (antisense 5'-TCGCCCACATAGGAATCCTTCTG-3') primers. PCR cycles included one cycle of 95°C for 2 min followed by 40 cycles each of 94°C for 30 s, annealing temperature 60°C for 30 s, and 72°C for 40 s, and a final extension step of 72°C for 10 min in a iCycler iQ Real-Time PCR Detection System (Bio-Rad GmbH München, Germany). Product-specific amplification was confirmed by a melting curve analysis and agarose gel electrophoresis analysis. All experiments were performed in duplicates. The fluorescence threshold value (CT) was calculated using the iCycler iQ optical v3.0a system software. The relative quantification of GSTA4 mRNA expression was performed using the comparative CT (CT = CT control – CT reference) method. The normalization of the CT was obtained by calculating the average of the control value from the target CT subtracted by the reference gene, and the difference was determined by subtracting the value for control of the target (GSTA4) gene and the value for treatment of the target gene. The fold change was calculated according to the efficiency method (CT-method), where it is assumed that the PCR efficiency is 100% (E = 2; fold change = Edifference).
hGSTA4 protein expression with Western blotting.
HT29 cells were resuspended in ice-cold homogenizing buffer consisting of 250 mM sucrose, 20 mM Tris/HCl, 1 mM dithiothreitol, and 1 mM Pefabloc at pH 7.4 and homogenized using ultrasound. The homogenates were centrifuged (16,000 x g, 60 min, 4°C) and the supernatant divided into aliquots and frozen at –80°C until use. Total protein content was measured using the method by Bradford with bovine serum albumin (BSA) as standard protein (Bradford, 1976), and 100 μg total protein (1:1.5 dilution in loading buffer) was separated in a 15% polyacrylamide gel using the SDS-PAGE for 1 h at 170 V and transferred to a nitrocellulose membrane for 2 h at 300 V (Laemmli, 1970; Towbin et al., 1979). For Western blotting, the polyclonal anti-hGSTA4 peptide antibody was used as previously described, and the signal for hGSTA4-4 protein was detected using ECL. Normalization of hGSTA4-4 protein levels was performed on the basis of -actin (Gardner and Gallagher, 2001).
GSH depletion in HT29 cells by BSO and HNE.
HT29 cells (0.6–0.8 x 106) were seeded three-fold for each experiment in 6-well plates and cultured. After 48 h, the cells were transferred to fresh culture medium and cultured in the presence of BSO dissolved in DMEM for 18 h (Lu et al., 1999). For initial studies 1–1000 μM BSO were used, whereas in the main experiments concentrations of 1–100 μM BSO were tested. The HNE mediated depletion of GSH was investigated in subconfluently grown cells (2 x 106 per ml) that were suspended in serum-free RPMI 1640 and subsequently treated with HNE (100–250 μM or ethanol as the untreated control that was present at a maximum of 4% a concentration, which was neither cyto- nor genotoxic) for 30 min. Following treatment, intracellular GSH content was determined by preparing a 16,000 x g fraction of an acidified cell homogenate as previously described (Ebert et al., 2001; Pool-Zobel et al., 1998; Treptow-van Lishaut et al., 1999). The acidified supernatants were used for colorimetric measurement of total GSH content using the Calbiochem GSH assay-kit. Two parallel determinations per sample were performed for the GSH assay. All GSH values were normalized to the number of cells, with the results representing mean values of at least three independent experiments (Treptow-van Lishaut et al., 1999). For statistical analyzing, normalization of the GSH levels was performed on the basis of ethanol as solvent control for each single experiment.
Detection of DNA damage (Comet assay).
HT29 cells were seeded in 6-well plates (0.9 x 106) or in 25-cm2 culture flasks (3 x 106) and grown for 48 h before they were treated with BSO (0.1 mM) for 18 h to deplete GSH. The GSH-modulated cells (2 x 106/ml) were subsequently treated with 100–200 μM HNE (30 min, 37°C), and the viability was determined with trypan blue exclusion. The remaining cells were subjected to single cell gel electrophoresis (Ebert et al., 2001; Singh et al., 1988). Briefly, the cell pellets were mixed with 0.7% low-melting-point agarose, and 30 μl of the cell suspensions were distributed onto microscope slides precoated with 0.5% agarose. After the agarose solidified, the slides were immersed in a lysis solution (10 mM Tris–HCl, 100 mM Na2EDTA, 2.5 M NaCl, 10% DMSO, 1% Triton X-100, pH 10) for at least 60 min at 4°C. Slides were placed in an electrophoresis chamber containing alkaline buffer (1 mM Na2EDTA, 300 mM NaOH, pH 13) for DNA unwinding. After 20 min, the current was switched on, and electrophoresis was carried out at 1.25 V/cm and 300 mA for 20 min. The slides were removed from the electrophoresis chamber and washed three times for 5 min each with neutralization buffer (4.2 M Tris–HCl, 0.08 M Tris-base, pH 7.2). Slides were stained with ethidium bromide (Sigma, Deisenhofen, Germany), solved in aqua bidest at a concentration of 0.2 mg/ml. Microscopic evaluation of the comet-like images revealed the extent of damaged DNA (Singh et al., 1988). The degree of damage in each preparation was quantified using the image analysis system of Perceptive Instruments (Haverhill; Suffolk, UK). Fifty DNA spots were evaluated per slide. Mean values of tail intensity (the percentage of fluorescence in the comet tail) from three replicated slides per experiment were calculated and the means of at least three independently reproduced experiments are shown in the graphs.
Statistical analysis.
The analysis for statistically significant effects was performed with the Graph Pad Prism software 4.0 (GraphPad Software, Inc.). Differences were assessed using one-way ANOVA followed by post-hoc t-tests. Treatment-related effects were considered statistically significant at p < 0.05.
RESULTS
Effect of Butyrate Exposure on hGSTA4 mRNA and Protein Expression
Consistent with other studies of hGSTA4 mRNA expression in human tissues, baseline hGSTA4 expression was detected by Northern blotting in HT29 cells at relatively low levels (Fig. 1). Interestingly, the levels of hGSTA4 mRNA were significantly elevated 2-fold (p < 0.05) after butyrate treatment (2 and 4 mM) for 24 h. The mRNA levels of hGSTA4 and -actin (internal control) in HT29 cells were determined during a period of 24 h at 4-h intervals to find the time point at which the expression of hGSTA4 mRNA is most enhanced. These kinetics of hGSTA4 mRNA expression, determined using semiquantitative RT-PCR (Fig. 2), revealed that hGSTA4 mRNA levels began to increase after 8 h of butyrate (2 mM) treatment and reached the highest induction levels after 24 h, without however, reaching statistical significance. Further studies on the kinetics of expression levels were then performed with real-time qPCR (Fig. 3). Our findings show a time-dependent upregulation of GSTA4 mRNA expression that became significant (3-fold, p < 0.001) after 24 h of butyrate (2 mM) treatment. Thus, relative mRNA levels determined with real-time qPCR were more comparable to the results with Northern blot than with semiquantitative RT-PCR. Figure 4 shows that butyrate-mediated transcriptional activation also resulted in a higher hGSTA4-4 protein expression, which was especially apparent when regarding the representative sample blot. Even though the data of the pixel densitometry measurements were not statistically significant, probably due to the high levels of unavoidable experimental variation of this system, the results do appear to be biologically significant. Moreover, they are supportive of the mRNA data (Northern blot, real-time qPCR) and of the functional data (Comet assay with HNE) reported here.
Effects on GSH Levels of 4-Hydroxy-2-Nonenal
As discussed, HNE is inactivated by GSH conjugation, which can occur spontaneously or mediated enzymatically by GSTs. As shown in Figure 5, increasing concentrations of HNE resulted in a loss of GSH levels in HT29 cells. The absolute GSH content of the untreated cells (10.98 ± 2.14 nmol GSH/106 cells) was significantly reduced to 64% (6.89 ± 1.02 nmol GSH/106 cells, p < 0.01) after exposure to 100 μM HNE and to 49% (5.30 ± 1.17 nmol GSH/106 cells, p < 0.01) after treatment with 200 μM HNE, showing a dose-response relationship. The incubation of the HT29 cells with 250 μM HNE did not result in a higher loss of cellular GSH (5.46 ± 1.02 nmol GSH/106 cells, p < 0.01), but in a decreased cell survival (69.87 ± 7.62%, p < 0.05). This effect seems to be independent of the GSH level.
GSH was depleted by BSO to determine whether this would enhance the sensitivity of the HT29 cells toward HNE. In preparation for this, we first investigated the effects of BSO on viability and GSH status of the HT29 cells. Table 1 summarizes the most important effects of BSO on viability and on GSH status. In contrast with a 24 h treatment, the exposure to 10 mM BSO for 72 h led to a marked reduction in viability to 67% of the level observed in control cells. However, doses of BSO as low as 100 μM elicited a dramatically loss of intracellular GSH in HT29 cells. Thus, concentrations up to 100 μM BSO, which did not cause a loss of viability, but resulted in a reduction of GSH levels from 13.23 ± 1.96 to 3.11 ± 0.6 nmol/106 cells (p < 0.001), were used for the further comet assay studies with HNE.
Comet Assay
As already mentioned, a second question of our study was to determine if an impaired GSH status of the HT29 cells would result in an enhancement of HNE-mediated genotoxicity. As observed in Figure 6, the cells that were treated with 0.1 mM BSO were markedly more sensitive toward DNA damage (p < 0.05) induction by HNE when compared to control cells. Specifically, a two-fold increase in HNE concentrations was needed to achieve an equivalent level of DNA damage in control cells, compared to the cells that were subjected to GSH depletion.
DISCUSSION
The GSTs constitute the major family of phase II detoxification enzymes involved in the cellular protection against electrophilic intermediates by catalyzing conjugation with GSH (Hayes and Pulford, 1995). Accordingly, elevation of GST and GSH expression by chemoprotective agents can afford cellular protection during the process of chemical carcinogenesis (Kensler and Helzlsouer, 1995; Wang et al., 1999). In this regard, several studies have provided evidence that GST isoenzymes and enzymes involved in the synthesis of GSH may be induced by nutritional factors in vivo and in vitro (Huber et al., 2002; Lampe and Peterson, 2002; Luceri et al., 2002; Nho and Jeffery, 2001; Treptow-van Lishaut et al., 1999). One of these inducing factors is butyrate, which is formed in the gut lumen as a metabolite of gut flora-mediated dietary fiber fermentation (Kobayashi and Fleming, 2001). We have previously demonstrated in human colon tumor HT29 cells that exposure to butyrate induces ERK1/2 phosphorylation, hGSTP1 mRNA, hGSTP1-1 protein, as well as total GST protein levels and catalytic activities, and maintains intracellular GSH concentrations (Ebert et al., 2001). These biochemical changes in GST/GSH levels by butyrate also decrease HNE-induced genotoxicity (Ebert et al., 2001). Furthermore, we have also shown that exposure to butyrate significantly enhances the expression of hGSTA1/2 and hGSTM2, concomitant with an elevation of GST catalytic activity (Ebert et al., 2003). Collectively, the aforementioned studies indicate a novel mechanism of chemoprotection by modulating GST expression. However, not known is if exposure to butyrate modulates the expression of hGSTA4-4, the primary HNE-metabolizing GST isoform and thus a contributor to chemoresistance to HNE. It is also not known if an opposite modulation of the GST/GSH system (e.g., by depletion of GSH) would instead increase HNE genotoxicity.
In the present study, we have provided new evidence that hGSTA4-4 protein levels and hGSTA4 mRNA expression are also inducible by butyrate. Since hGSTA4-4 is the GST isoenzyme with the highest affinity for HNE, these results provide an explanation for the acquired cellular resistance of HT29 cells cultured in the presence of butyrate against HNE genotoxicity. However, we cannot determine, at this point, what proportion of the previously reported acquired resistance is due to the induction of hGSTA4-4. For one, GSTA4-4 is clearly the GST isoform, which has the highest catalytic efficiency in HNE conjugation with GSH. On the other hand, according to our new data on expression analysis of GSTs with microarrays, it is expressed only in relatively low concentrations (1–2% of total GSTs) in HT29 that were upregulated after treatment with the maximal tolerated doses of butyrate of 4 mM for 48 h (Pool-Zobel et al., in press). The absolute amounts of those more abundant GST forms (e.g., hGSTA1, hGSTA2, hGSTP1) that also have catalytic activity toward HNE, albeit with lower catalytic efficiencies relative to hGSTA4-4, are likely contributors toward the butyrate-mediated chemoprotection against HNE.
In addition to GST, the present study provides additional evidence to indicate that the GSH status of the cells is a critical determinant of HT29 cell injury to HNE. GSH depletion by 0.1 mM BSO effectively enhanced HNE genotoxicity. Accordingly, dietary agents that modulate the expression level of GSH may also be modulators of HNE genotoxicity in human cells. Furthermore, we observed no cytotoxicity in HT29 cells when cellular GSH was reduced to nearly 50% with HNE concentrations increasing up to 200 μM. Only the highest concentration of HNE (250 μM) was significantly cytotoxic, which however seemed to be independent of the GSH level in HT29, since no further depletion of the GSH status was observed. Moreover, cytotoxic effects of GSH depletion have been reported to be tissue specific and only first occur when mitochondrial GSH is decreased to at least 20% of basic levels (Martensson et al., 1989a,b; Martensson and Meister, 1989). A reduction to the aforementioned very low levels of GSH implies that a longer incubation period of at least 24 h is needed to reach the GSH that is sequestered in mitochondria, evincing a slow turnover (Griffith and Meister, 1985).
Our results raise interesting questions regarding the mechanisms of butyrate-mediated GST induction in human HT29 cells. We know that the putative binding sites for transcription factors in the regulatory region of the hGSTA4 gene include AP1, STAT, GATA1, and NF-B (Desmots et al., 1998). In particular, AP1 and NF-B can be activated by butyrate in a reporter assay (Bhmer et al., personal communication). In addition, butyrate activates ERK of the MAP kinase pathway, an important component of induction of AP1-related transcription factors Jun and Fos (Ebert et al., 2001). In addition, posttranscriptional mechanisms can also play an important role in determining cell-specific expression of the hGSTP1 mRNA (Moffat et al., 1997). Global gene activation by modulation of histone acetylation probably plays an additional role in these activities, since butyrate and other short-chain fatty acids belong to the group of histone deacetylase inhibitors which activate a number of different target genes (Marks et al., 2001).
Interestingly, to our knowledge there are no other reports of hGSTA4 mRNA induction in human cell lines or tissues. One of our laboratories has utilized a human liver slice model system to test the potential of several known antioxidant inducers of rodent GST expression, including butylated hydroxyanisole (BHA) and tert-butylhydroquinone (Huisden et al., 2003). However, these results were, essentially negative with regard to effects of test agents on hGSTA4 mRNA expression. Given the presence, but relatively low expression, of hGSTA4 mRNA in most human tissues, others have hypothesized that hGSTA4 may function more as a housekeeping gene (Desmots et al., 1998). Accordingly, our present results indicate that this particular GST isoform may be an important target for chemoprevention studies, at least in models relevant to the human gastrointestinal tract. However, it must also be considered that our model (HT29 cells) are transformed cells and that the butyrate-mediated GST induction may potentially enhance the survival of transformed cells (Ebert et al., 2001; Henderson et al., 1998). Accordingly, additional work is needed on how the relative protein expression levels of GSTA4-4 in nontransformed human primary cells may be influenced by butyrate available from the gut lumen.
In conclusion, we have demonstrated that exposure to butyrate increases the expression of the primary HNE-metabolizing GST isoform hGSTA4 in HT29 cells. This increase in hGSTA4 probably contributes in part to the previously observed concomitant protection against HNE genotoxicity. Furthermore, the relative susceptibility of these cells is also codependent upon intracellular GSH status, since HNE genotoxicity was enhanced by decreasing GSH with BSO. Since HNE contributes to colon carcinogenesis by a number of different mechanisms involving genotoxicity, a favorable modulation of the GSH/GST system by dietary fiber, yielding butyrate, may ultimately be a modulator of risk to colon carcinogenesis. Future studies using nontransformed primary human cells and chemoprevention studies will shed light on the association between butyrate production and HNE genotoxicity.
ACKNOWLEDGMENTS
We thank the Bundesministerium für Landwirtschaft, Ernhrung und Forsten, Germany (BMELF No. 99 HS 039), the Deutsche Forschungsgemeinschaft (DFG PO 284/8–1) and the Bundesministerium für Forschung und Technologie, Germany (BMBF FKZ.01EA0103), for the financial support within the project. Additional support was provided by a grant from the National Institutes of Health (ES-R01ES09427).
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Department of Environmental and Occupational Health Sciences, University of Washington, 4225 Roosevelt Way NE, Suite 100, Seattle, Washington 98105
ABSTRACT
The cellular production of 4-hydroxy-2-nonenal (HNE), a product of endogenous lipid peroxidation, constitutes a genotoxic risk factor for carcinogenesis. Our previous studies have shown that human HT29 colon cells developed resistance toward HNE injury after treatment with butyrate, a diet-associated gut fermentation product. This resistance was attributed to the induction of certain glutathione S-transferases (hGSTP1-1, hGSTM2-2, and hGSTA1-1) and also for the tripeptide glutathione (GSH) synthesizing enzymes. In the present study, we have investigated in HT29 cells whether hGSTA4-4, which has a high substrate specificity for HNE, was also inducible by butyrate and, thus, could contribute to the previously observed chemoresistance. In addition, we investigated if cellular depletion of GSH by L-buthionine-S,R-sulfoximine (BSO) enhances chemosensitivity to HNE injury in HT29 cells. Incubation of HT29 cells with butyrate (2–4 mM) significantly elicited a 1.8 to 3-fold upregulation of steady state hGSTA4 mRNA over 8–24 h after treatment. Moreover, 4 mM butyrate tended to increase hGSTA4-4 protein concentrations. Incubation with 100 μM BSO decreased cellular GSH levels by 77% without significant changes in cell viability. Associated with this was a 2-fold higher level of HNE-induced DNA damage as measured by the comet assay. Collectively, the results of this study and our previous work indicate that the genotoxicity of HNE is highly dependent on cellular GSH status and those GSTs that contribute toward HNE conjugation, including hGSTA4-4. Since HNE contributes to colon carcinogenesis, the favorable modulation of the GSH/GST system by butyrate may contribute to chemoprevention and reduction of the risks.
Key Words: L-buthionine-(S,R)-sulfoximine (BSO); butyrate; DNA damage; glutathione (GSH); 4-hydroxy-2-nonenal (HNE); glutathione S-transferases; hGSTA4.
INTRODUCTION
The highly reactive aldehyde trans-4-hydroxy-2-nonenal (HNE) has a wide variety of biological activities. HNE is mutagenic and has strong effects on signal transduction pathways (Blanc et al., 1997; Chung et al., 1993; Esterbauer et al., 1990). It is an important endogenous factor of tissue injury risk, since it is the major electrophilic product of lipid peroxidation caused by oxidative stress (Chung et al., 1993). Formation of HNE is initiated by free radical-mediated degradation of -6-polyunsaturated fatty acids such as linoleic and arachidonic acid, which are relatively abundant fatty acids in membranes of human cells (Burcham, 1998; Esterbauer et al., 1990). Feng et al. (2004) described that conditions of oxidative stress potently enhanced cellular levels of HNE from 0.1–3 μM to 10 μM–5 mM. These types of oxidative stress, e.g., occur during chronic diseases like lymphoedemas (Siems et al., 2002) or during progression of colorectal cancer (Skrzydlewska et al., 2005). HNE can be further metabolized to an epoxide intermediate that interacts with DNA to form exocyclic etheno-guanine, -adenine, and -cytosine adducts (Chung et al., 1996). In addition, bulky exocyclic propane-type DNA adducts with guanine, and, in particular, 6-(1-hydroxyhexanyl)-8-hydroxy-1, N2-propano-2-deoxyguansine (4-HNE-dG) adducts may be formed in cellular reactions with HNE (Douki and Ames, 1994). We have recently shown that HNE is highly genotoxic to human colon tumor cells (Ebert et al., 2001). Additionally, damage to the tumor suppressor gene TP53 in adenoma cells and in primary colon cells has been shown using fluorescence in situ hybridization of the damaged gene (Schferhenrich et al., 2003a,b). Other investigators have recently reported that HNE-DNA adducts are preferentially formed at the third base of codons 249 and 174 in the TP53 gene (Hu et al., 2002). Collectively, these data emphasize the potentially important role of HNE in tumor initiation and progression (Cheeseman, 1993).
One of the primary pathways for detoxification of HNE in human cells is through conjugation with glutathione (GSH), a reaction, which is markedly enhanced by catalysis with certain glutathione S-transferases (GSTs) (Berhane et al., 1994). The alpha class GSTs, of which hGSTA1-1 is the most abundant form in human liver, are involved in HNE detoxification through GSH conjugation. In particular, hGSTA1-1 conjugates HNE with a Km = 50 μM and a kcat/Km = 0.058 s–1μM–1, whereas hGSTA4-4, a primarily mitochondrial GST, effectively conjugates HNE with an efficient Km = 49 μM and a very rapid kcat/Km of 2.7 s–1μM–1 (Cheng et al., 2001; Gardner and Gallagher, 2001). The human pi class isoform hGSTP1-1 can also catalyze the conjugation of HNE with GSH (Berhane et al., 1994). Comparatively, however, hGSTA4-4 exhibits the highest catalytic activity and turnover of HNE relative to the other GST isozymes. The specificity of GSTA4-4 is based on typical structural properties like an electrophilic substrate-binding pocket that consists of three structural modules. These form the binding site with a favorable positioning of Tyr212 to interact with the aldehyde group of the substrate and polarize it for reaction (Bruns et al., 1999). Because of this highly specific catalytic activity toward such aldehydes and related oxidants, it has been suggested that hGSTA4-4 and its rodent orthologues (mouse GSTA4-4 and rat GSTA4-4) have specifically evolved to protect against oxidative injury in vivo.
We have previously demonstrated that the gut fermentation product butyrate enhances the cellular levels of hGSTA1/2, hGSTM2-2, hGSTP1-1, but not hGSTM1-1 or hGSTT1-1 proteins in HT29 cells. Concomitant with the induction of these aforementioned GST isoforms were increases in GST catalytic activities, in hGSTP1 and hGSTM2 steady-state mRNA expression and also in cellular GSH levels (Ebert et al., 2003). However, what is not known is if butyrate exposure enhances the expression of hGSTA4-4, the primary HNE metabolizing GST (Hubatsch et al., 1998) and whether a downregulation of the GST/GSH system confers opposite effects, namely an enhanced chemosensitivity. In the present study we have investigated baseline expression levels of GSTA4-4 in the human colon cell line HT29 and tested the hypothesis that exposure to butyrate would result in induction of hGSTA4 mRNA and hGSTA4-4 protein expression. Once establishing the inducibility of hGSTA4-4, we further investigated if depletion of cellular GSH levels would increase the genotoxic response to HNE.
MATERIALS AND METHODS
Cells and reagents.
The human colon tumor cell line HT29 (ATCC, HTB-38) used in this study was established in 1964 and originated from human adenoma colon tissue (Fogh and Trempe, 1975). The HT29 cells were obtained from the American Tissue Culture Collection (ATCC, Rockville, MD). Sodium butyrate was purchased from Merck research laboratories (Schuchardt, Hhenbrunn, Germany). The Glutathione Assay-Kit and 4-hydroxynonenal (HNE) were purchased from Calbiochem-Novabiochem (Bad Soden, Germany). Positively charged nylon membranes, the DIG-RNA labeling kit, T7 RNA polymerase, anti-DIG-alkaline phosphatase antibody, and CDP-Star were supplied from Roche Diagnostics (Mannheim, Germany). Reverse transcriptase and oligo(dT) were purchased from SuperScript Invitrogen GmbH (Karlsruhe, Germany). The RNeasy mini kit and Hot Star Taq DNA Polymerase were purchased from Qiagen (Hilden, Germany). TRIzol-Reagent and cell culture materials, including Dulbecco's Modified Eagle Medium (DMEM), RPMI 1640 Minimal Essential Medium without L-glutamine (RPMI 1640), penicillin/streptomycin, and fetal calf serum (FCS) were purchased from Invitrogen GmbH (Karlsruhe, Germany). Goat anti-chicken IgY conjugated with horseradish peroxidase was purchased from CalBiotech (Santa Cruz). 4',6-Diamidino-2-phenylindole dihydrochloride (DAPI) and L-buthionine-(S,R)-sulfoximine (BSO) were purchased from Sigma-Aldrich (Deisenhofen, Germany). Enhanced Chemiluminescence (ECL)TM Western Blotting Detection Reagents were purchased from Amersham Biosciences Europe GmbH (Freiburg, Germany).
Culture of HT29 cells.
The human colon tumor cell line HT29 was maintained in DMEM supplemented with 10% (v/v) FCS and penicillin (50 U/ml)/streptomycin (50 μg/ml) in a humidified 5% CO2 incubator at 37°C, where it grew strictly adherent. Under given laboratory conditions, the HT29 cells doubled their population number within 24 h (unpublished data). The experiments determining hGSTA4 mRNA and hGSTA4-4 protein expression were performed with passages from 30 to 53, and passages from 17 to 40 of HT29 cells were used for GSH-depletion trials and the comet assay studies. To retain standardized conditions, all experiments were performed using subconfluently grown cells, which 72 h after seeding had reached a degree of confluence of at least 80%.
Analysis of cell viability and growth.
The trypan blue exclusion test was used to determine membrane integrity reflecting cell viability after harvesting or incubating the cells in culture or in suspension. Studies on modulation of growth were performed to assess effective concentrations of BSO. For these experiments, the cells were seeded (10 x 103/well) into 96-well microtiter plates and allowed to attach. After 24 h, the cells were treated with different concentrations of BSO (0.001 mM) dissolved in culture medium. Survival was detected after 24 and 72 h by quantifying DNA with DAPI, a cell permanent stain that specifically binds double-stranded DNA. This exhibits a 20-fold higher fluorescence enhancement, which does not occur with single-stranded DNA (Tecan, 1997). By subsequent fluorometric analysis with ex/em 360/465 nm using a 96-well microplate Reader (Beyer-Sehlmeyer et al., 2003; Ebert et al., 2001) the total amount of DNA was measured and was used to represent the amount of surviving cells.
Cell treatment with butyrate.
Prior to treatment with sodium butyrate, 2 x 106 HT29 cells were allowed to attach and grow in DMEM for 48 h in 25-cm2 culture flasks. For experiments of mRNA and protein expression, the cells were treated with 2 and 4 mM sodium butyrate for 24 and 48 h. The investigated concentrations of butyrate were based on the EC50 and EC25 for cell survival after 48 h butyrate treatment as previously determined (Ebert et al., 2001). In addition, the treatment of the group receiving 2 mM butyrate was stopped after 1, 4, 8, 12, 16, or 24 h to characterize the time-dependent expression levels of hGSTA4 mRNA in culture.
Detection of hGSTA4 expression by Northern blotting and semiquantitative PCR.
Total RNA was isolated from HT29 cells using the TRIzol method (Chomczynski and Sacchi, 1987). After RNA isolation, 15 μg of total RNA was loaded onto a 1.2% denaturing agarose gel, separated for 3–4 h at 70 V, and blotted on a positively charged nylon membrane. Digoxygenin (DIG)-labeled probes for hGSTA4 and the glyceraldehyde-3-phosphate dehydrogenase (GAPDH) were prepared from total RNA by reverse transcriptase PCR followed by a gene-specific PCR to amplify the cDNAs and to introduce the T7 RNA polymerase promoter sequence for in vitro transcription. In vitro transcription and simultaneous DIG-labeling of hGSTA4 and GAPDH were performed using the DIG-RNA labeling kit and T7 RNA polymerase. The membrane was hybridized with 1 μg probe in 10 ml hybridization buffer overnight at 70°C. After washing with 2 and 0.3x saline sodium citrate (SSC) containing 0.1% sodium dodecyl sulphate (SDS), the membrane was incubated in blocking solution for 1 h at room temperature, followed by antibody incubation for further 30 min. Detection of the anti-DIG-alkaline phosphatase antibody was performed with CDP-Star on a HyperfilmTM ECL. The film was photographed for evaluation of the band intensities and the Quantity One 4.1 software (Bio-Rad) was used for quantitative evaluation. For the quantification of the expression levels, the peak densities of the bands were used. After the hGSTA4 mRNA levels were normalized with GAPDH, the peak densities of untreated cells were equated with 100%. The results represent the mean of the hGSTA4 mRNA expression values (±SEM) for the several treatments of six independent experiments.
Semiquantitative reverse transcriptase-PCR (RT-PCR) was used to quantitate and confirm the initial results from the Northern blotting experiments. For these experiments, total RNA was extracted with the RNeasy mini kit according to the manufacturer's instructions. A cDNA equivalent to 5 μg total RNA was prepared by first-strand synthesis using oligo(dT) primer (SuperScriptTM First-Strand Synthesis System). Semiquantitative RT-PCR was used to amplify GSTA4 mRNA levels and to compare its expression in butyrate-treated cells versus untreated cells. The expression of -actin served as an internal control. To avoid different cDNA concentrations between target gene (hGSTA4) and control gene (-actin), for each sample the template cDNA was added to the PCR reaction mix without primers. The mixture was aliquoted into two batches, one for GSTA4 and one for -actin. Subsequently, gene-specific PCR primers were added for hGSTA4 (Genbank NM_001512 5'-ACC TGG CAG CAA GAA GAA G-3', sense and 5'-CAG GAT AAG GAG AGC AGA AAG-3', antisense) and -actin (NM_001101 5'-GCT CGT CGT CGA CAA CGG CTC-3' sense, and 5'-TGG GTC ATC TTC TCG CGG TTG G-3', antisense). Based upon these published cDNA sequences, the predicted size of the hGSTA4 cDNA is 435 bp, and that of -actin is 337 bp. PCR reactions consisted of cDNA equivalent to 0.6 mg/25 ml of RNA, 1.25 U/25 ml of Hot Star Taq DNA Polymerase, either 0.25 mM of -actin primers or 0.5 mM of GSTA4 primers, and a final MgCl2 concentration of 1.5 mM. The cDNA was amplified by PCR, with 21 cycles for -actin and 28 cycles for GSTA4. After primary denaturation at 95°C for 5 min, the PCR cycles were run at 95°C for 15 s, 55°C for 30 s, and 72°C for 1 min. Densitometry evaluation of the ethidium bromide bands was performed with the Quantity One 4.1 Software (Bio-Rad). The results were expressed as ratio of hGSTA4 mRNA to -actin mRNA.
Relative quantification of GSTA4 mRNA transcript by real-time qPCR.
Expression of GSTA4 mRNAs was assessed by quantitative real-time PCR (SYBR Green I) system. One μl of cDNA (10 ng of total RNA equivalent) was used in a 25 μl PCR amplification reaction containing 2x iQ SYBR Green supermix (100 mM KCl, 40 mM Tris-HCl, pH 8.4, 0.4 mM each dNTP, 50 U/ml iTaq DNA polymerase, 6 mM MgCl2, SYBR Green I, 20 nM fluorescein, stabilizers), and 10 pmol each of target (GSTA4) and reference (-actin) gene specific primers. A region of GSTA4 and the -actin mRNA was amplified using these iGSTA4-F (sense 5'-CCGGATGGAGTCCGTGAGATGG-3'), iGSTA4-R (antisense 5'-CCATGGGCACTTGTTGGAACAGC-3'), i-actin_F (sense 5'-GCTCGTCGTCGACAACGGCTC-3'), and i-actin_R (antisense 5'-TCGCCCACATAGGAATCCTTCTG-3') primers. PCR cycles included one cycle of 95°C for 2 min followed by 40 cycles each of 94°C for 30 s, annealing temperature 60°C for 30 s, and 72°C for 40 s, and a final extension step of 72°C for 10 min in a iCycler iQ Real-Time PCR Detection System (Bio-Rad GmbH München, Germany). Product-specific amplification was confirmed by a melting curve analysis and agarose gel electrophoresis analysis. All experiments were performed in duplicates. The fluorescence threshold value (CT) was calculated using the iCycler iQ optical v3.0a system software. The relative quantification of GSTA4 mRNA expression was performed using the comparative CT (CT = CT control – CT reference) method. The normalization of the CT was obtained by calculating the average of the control value from the target CT subtracted by the reference gene, and the difference was determined by subtracting the value for control of the target (GSTA4) gene and the value for treatment of the target gene. The fold change was calculated according to the efficiency method (CT-method), where it is assumed that the PCR efficiency is 100% (E = 2; fold change = Edifference).
hGSTA4 protein expression with Western blotting.
HT29 cells were resuspended in ice-cold homogenizing buffer consisting of 250 mM sucrose, 20 mM Tris/HCl, 1 mM dithiothreitol, and 1 mM Pefabloc at pH 7.4 and homogenized using ultrasound. The homogenates were centrifuged (16,000 x g, 60 min, 4°C) and the supernatant divided into aliquots and frozen at –80°C until use. Total protein content was measured using the method by Bradford with bovine serum albumin (BSA) as standard protein (Bradford, 1976), and 100 μg total protein (1:1.5 dilution in loading buffer) was separated in a 15% polyacrylamide gel using the SDS-PAGE for 1 h at 170 V and transferred to a nitrocellulose membrane for 2 h at 300 V (Laemmli, 1970; Towbin et al., 1979). For Western blotting, the polyclonal anti-hGSTA4 peptide antibody was used as previously described, and the signal for hGSTA4-4 protein was detected using ECL. Normalization of hGSTA4-4 protein levels was performed on the basis of -actin (Gardner and Gallagher, 2001).
GSH depletion in HT29 cells by BSO and HNE.
HT29 cells (0.6–0.8 x 106) were seeded three-fold for each experiment in 6-well plates and cultured. After 48 h, the cells were transferred to fresh culture medium and cultured in the presence of BSO dissolved in DMEM for 18 h (Lu et al., 1999). For initial studies 1–1000 μM BSO were used, whereas in the main experiments concentrations of 1–100 μM BSO were tested. The HNE mediated depletion of GSH was investigated in subconfluently grown cells (2 x 106 per ml) that were suspended in serum-free RPMI 1640 and subsequently treated with HNE (100–250 μM or ethanol as the untreated control that was present at a maximum of 4% a concentration, which was neither cyto- nor genotoxic) for 30 min. Following treatment, intracellular GSH content was determined by preparing a 16,000 x g fraction of an acidified cell homogenate as previously described (Ebert et al., 2001; Pool-Zobel et al., 1998; Treptow-van Lishaut et al., 1999). The acidified supernatants were used for colorimetric measurement of total GSH content using the Calbiochem GSH assay-kit. Two parallel determinations per sample were performed for the GSH assay. All GSH values were normalized to the number of cells, with the results representing mean values of at least three independent experiments (Treptow-van Lishaut et al., 1999). For statistical analyzing, normalization of the GSH levels was performed on the basis of ethanol as solvent control for each single experiment.
Detection of DNA damage (Comet assay).
HT29 cells were seeded in 6-well plates (0.9 x 106) or in 25-cm2 culture flasks (3 x 106) and grown for 48 h before they were treated with BSO (0.1 mM) for 18 h to deplete GSH. The GSH-modulated cells (2 x 106/ml) were subsequently treated with 100–200 μM HNE (30 min, 37°C), and the viability was determined with trypan blue exclusion. The remaining cells were subjected to single cell gel electrophoresis (Ebert et al., 2001; Singh et al., 1988). Briefly, the cell pellets were mixed with 0.7% low-melting-point agarose, and 30 μl of the cell suspensions were distributed onto microscope slides precoated with 0.5% agarose. After the agarose solidified, the slides were immersed in a lysis solution (10 mM Tris–HCl, 100 mM Na2EDTA, 2.5 M NaCl, 10% DMSO, 1% Triton X-100, pH 10) for at least 60 min at 4°C. Slides were placed in an electrophoresis chamber containing alkaline buffer (1 mM Na2EDTA, 300 mM NaOH, pH 13) for DNA unwinding. After 20 min, the current was switched on, and electrophoresis was carried out at 1.25 V/cm and 300 mA for 20 min. The slides were removed from the electrophoresis chamber and washed three times for 5 min each with neutralization buffer (4.2 M Tris–HCl, 0.08 M Tris-base, pH 7.2). Slides were stained with ethidium bromide (Sigma, Deisenhofen, Germany), solved in aqua bidest at a concentration of 0.2 mg/ml. Microscopic evaluation of the comet-like images revealed the extent of damaged DNA (Singh et al., 1988). The degree of damage in each preparation was quantified using the image analysis system of Perceptive Instruments (Haverhill; Suffolk, UK). Fifty DNA spots were evaluated per slide. Mean values of tail intensity (the percentage of fluorescence in the comet tail) from three replicated slides per experiment were calculated and the means of at least three independently reproduced experiments are shown in the graphs.
Statistical analysis.
The analysis for statistically significant effects was performed with the Graph Pad Prism software 4.0 (GraphPad Software, Inc.). Differences were assessed using one-way ANOVA followed by post-hoc t-tests. Treatment-related effects were considered statistically significant at p < 0.05.
RESULTS
Effect of Butyrate Exposure on hGSTA4 mRNA and Protein Expression
Consistent with other studies of hGSTA4 mRNA expression in human tissues, baseline hGSTA4 expression was detected by Northern blotting in HT29 cells at relatively low levels (Fig. 1). Interestingly, the levels of hGSTA4 mRNA were significantly elevated 2-fold (p < 0.05) after butyrate treatment (2 and 4 mM) for 24 h. The mRNA levels of hGSTA4 and -actin (internal control) in HT29 cells were determined during a period of 24 h at 4-h intervals to find the time point at which the expression of hGSTA4 mRNA is most enhanced. These kinetics of hGSTA4 mRNA expression, determined using semiquantitative RT-PCR (Fig. 2), revealed that hGSTA4 mRNA levels began to increase after 8 h of butyrate (2 mM) treatment and reached the highest induction levels after 24 h, without however, reaching statistical significance. Further studies on the kinetics of expression levels were then performed with real-time qPCR (Fig. 3). Our findings show a time-dependent upregulation of GSTA4 mRNA expression that became significant (3-fold, p < 0.001) after 24 h of butyrate (2 mM) treatment. Thus, relative mRNA levels determined with real-time qPCR were more comparable to the results with Northern blot than with semiquantitative RT-PCR. Figure 4 shows that butyrate-mediated transcriptional activation also resulted in a higher hGSTA4-4 protein expression, which was especially apparent when regarding the representative sample blot. Even though the data of the pixel densitometry measurements were not statistically significant, probably due to the high levels of unavoidable experimental variation of this system, the results do appear to be biologically significant. Moreover, they are supportive of the mRNA data (Northern blot, real-time qPCR) and of the functional data (Comet assay with HNE) reported here.
Effects on GSH Levels of 4-Hydroxy-2-Nonenal
As discussed, HNE is inactivated by GSH conjugation, which can occur spontaneously or mediated enzymatically by GSTs. As shown in Figure 5, increasing concentrations of HNE resulted in a loss of GSH levels in HT29 cells. The absolute GSH content of the untreated cells (10.98 ± 2.14 nmol GSH/106 cells) was significantly reduced to 64% (6.89 ± 1.02 nmol GSH/106 cells, p < 0.01) after exposure to 100 μM HNE and to 49% (5.30 ± 1.17 nmol GSH/106 cells, p < 0.01) after treatment with 200 μM HNE, showing a dose-response relationship. The incubation of the HT29 cells with 250 μM HNE did not result in a higher loss of cellular GSH (5.46 ± 1.02 nmol GSH/106 cells, p < 0.01), but in a decreased cell survival (69.87 ± 7.62%, p < 0.05). This effect seems to be independent of the GSH level.
GSH was depleted by BSO to determine whether this would enhance the sensitivity of the HT29 cells toward HNE. In preparation for this, we first investigated the effects of BSO on viability and GSH status of the HT29 cells. Table 1 summarizes the most important effects of BSO on viability and on GSH status. In contrast with a 24 h treatment, the exposure to 10 mM BSO for 72 h led to a marked reduction in viability to 67% of the level observed in control cells. However, doses of BSO as low as 100 μM elicited a dramatically loss of intracellular GSH in HT29 cells. Thus, concentrations up to 100 μM BSO, which did not cause a loss of viability, but resulted in a reduction of GSH levels from 13.23 ± 1.96 to 3.11 ± 0.6 nmol/106 cells (p < 0.001), were used for the further comet assay studies with HNE.
Comet Assay
As already mentioned, a second question of our study was to determine if an impaired GSH status of the HT29 cells would result in an enhancement of HNE-mediated genotoxicity. As observed in Figure 6, the cells that were treated with 0.1 mM BSO were markedly more sensitive toward DNA damage (p < 0.05) induction by HNE when compared to control cells. Specifically, a two-fold increase in HNE concentrations was needed to achieve an equivalent level of DNA damage in control cells, compared to the cells that were subjected to GSH depletion.
DISCUSSION
The GSTs constitute the major family of phase II detoxification enzymes involved in the cellular protection against electrophilic intermediates by catalyzing conjugation with GSH (Hayes and Pulford, 1995). Accordingly, elevation of GST and GSH expression by chemoprotective agents can afford cellular protection during the process of chemical carcinogenesis (Kensler and Helzlsouer, 1995; Wang et al., 1999). In this regard, several studies have provided evidence that GST isoenzymes and enzymes involved in the synthesis of GSH may be induced by nutritional factors in vivo and in vitro (Huber et al., 2002; Lampe and Peterson, 2002; Luceri et al., 2002; Nho and Jeffery, 2001; Treptow-van Lishaut et al., 1999). One of these inducing factors is butyrate, which is formed in the gut lumen as a metabolite of gut flora-mediated dietary fiber fermentation (Kobayashi and Fleming, 2001). We have previously demonstrated in human colon tumor HT29 cells that exposure to butyrate induces ERK1/2 phosphorylation, hGSTP1 mRNA, hGSTP1-1 protein, as well as total GST protein levels and catalytic activities, and maintains intracellular GSH concentrations (Ebert et al., 2001). These biochemical changes in GST/GSH levels by butyrate also decrease HNE-induced genotoxicity (Ebert et al., 2001). Furthermore, we have also shown that exposure to butyrate significantly enhances the expression of hGSTA1/2 and hGSTM2, concomitant with an elevation of GST catalytic activity (Ebert et al., 2003). Collectively, the aforementioned studies indicate a novel mechanism of chemoprotection by modulating GST expression. However, not known is if exposure to butyrate modulates the expression of hGSTA4-4, the primary HNE-metabolizing GST isoform and thus a contributor to chemoresistance to HNE. It is also not known if an opposite modulation of the GST/GSH system (e.g., by depletion of GSH) would instead increase HNE genotoxicity.
In the present study, we have provided new evidence that hGSTA4-4 protein levels and hGSTA4 mRNA expression are also inducible by butyrate. Since hGSTA4-4 is the GST isoenzyme with the highest affinity for HNE, these results provide an explanation for the acquired cellular resistance of HT29 cells cultured in the presence of butyrate against HNE genotoxicity. However, we cannot determine, at this point, what proportion of the previously reported acquired resistance is due to the induction of hGSTA4-4. For one, GSTA4-4 is clearly the GST isoform, which has the highest catalytic efficiency in HNE conjugation with GSH. On the other hand, according to our new data on expression analysis of GSTs with microarrays, it is expressed only in relatively low concentrations (1–2% of total GSTs) in HT29 that were upregulated after treatment with the maximal tolerated doses of butyrate of 4 mM for 48 h (Pool-Zobel et al., in press). The absolute amounts of those more abundant GST forms (e.g., hGSTA1, hGSTA2, hGSTP1) that also have catalytic activity toward HNE, albeit with lower catalytic efficiencies relative to hGSTA4-4, are likely contributors toward the butyrate-mediated chemoprotection against HNE.
In addition to GST, the present study provides additional evidence to indicate that the GSH status of the cells is a critical determinant of HT29 cell injury to HNE. GSH depletion by 0.1 mM BSO effectively enhanced HNE genotoxicity. Accordingly, dietary agents that modulate the expression level of GSH may also be modulators of HNE genotoxicity in human cells. Furthermore, we observed no cytotoxicity in HT29 cells when cellular GSH was reduced to nearly 50% with HNE concentrations increasing up to 200 μM. Only the highest concentration of HNE (250 μM) was significantly cytotoxic, which however seemed to be independent of the GSH level in HT29, since no further depletion of the GSH status was observed. Moreover, cytotoxic effects of GSH depletion have been reported to be tissue specific and only first occur when mitochondrial GSH is decreased to at least 20% of basic levels (Martensson et al., 1989a,b; Martensson and Meister, 1989). A reduction to the aforementioned very low levels of GSH implies that a longer incubation period of at least 24 h is needed to reach the GSH that is sequestered in mitochondria, evincing a slow turnover (Griffith and Meister, 1985).
Our results raise interesting questions regarding the mechanisms of butyrate-mediated GST induction in human HT29 cells. We know that the putative binding sites for transcription factors in the regulatory region of the hGSTA4 gene include AP1, STAT, GATA1, and NF-B (Desmots et al., 1998). In particular, AP1 and NF-B can be activated by butyrate in a reporter assay (Bhmer et al., personal communication). In addition, butyrate activates ERK of the MAP kinase pathway, an important component of induction of AP1-related transcription factors Jun and Fos (Ebert et al., 2001). In addition, posttranscriptional mechanisms can also play an important role in determining cell-specific expression of the hGSTP1 mRNA (Moffat et al., 1997). Global gene activation by modulation of histone acetylation probably plays an additional role in these activities, since butyrate and other short-chain fatty acids belong to the group of histone deacetylase inhibitors which activate a number of different target genes (Marks et al., 2001).
Interestingly, to our knowledge there are no other reports of hGSTA4 mRNA induction in human cell lines or tissues. One of our laboratories has utilized a human liver slice model system to test the potential of several known antioxidant inducers of rodent GST expression, including butylated hydroxyanisole (BHA) and tert-butylhydroquinone (Huisden et al., 2003). However, these results were, essentially negative with regard to effects of test agents on hGSTA4 mRNA expression. Given the presence, but relatively low expression, of hGSTA4 mRNA in most human tissues, others have hypothesized that hGSTA4 may function more as a housekeeping gene (Desmots et al., 1998). Accordingly, our present results indicate that this particular GST isoform may be an important target for chemoprevention studies, at least in models relevant to the human gastrointestinal tract. However, it must also be considered that our model (HT29 cells) are transformed cells and that the butyrate-mediated GST induction may potentially enhance the survival of transformed cells (Ebert et al., 2001; Henderson et al., 1998). Accordingly, additional work is needed on how the relative protein expression levels of GSTA4-4 in nontransformed human primary cells may be influenced by butyrate available from the gut lumen.
In conclusion, we have demonstrated that exposure to butyrate increases the expression of the primary HNE-metabolizing GST isoform hGSTA4 in HT29 cells. This increase in hGSTA4 probably contributes in part to the previously observed concomitant protection against HNE genotoxicity. Furthermore, the relative susceptibility of these cells is also codependent upon intracellular GSH status, since HNE genotoxicity was enhanced by decreasing GSH with BSO. Since HNE contributes to colon carcinogenesis by a number of different mechanisms involving genotoxicity, a favorable modulation of the GSH/GST system by dietary fiber, yielding butyrate, may ultimately be a modulator of risk to colon carcinogenesis. Future studies using nontransformed primary human cells and chemoprevention studies will shed light on the association between butyrate production and HNE genotoxicity.
ACKNOWLEDGMENTS
We thank the Bundesministerium für Landwirtschaft, Ernhrung und Forsten, Germany (BMELF No. 99 HS 039), the Deutsche Forschungsgemeinschaft (DFG PO 284/8–1) and the Bundesministerium für Forschung und Technologie, Germany (BMBF FKZ.01EA0103), for the financial support within the project. Additional support was provided by a grant from the National Institutes of Health (ES-R01ES09427).
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