Deoxycholic acid causes DNA damage in colonic cells with subsequent induction of caspases, COX-2 promoter activity and the transcription factors NF-kB and AP-1
B. Glinghammar,
H. Inoue1 and
J.J. Rafter,2
Department of Medical Nutrition, Karolinska Institutet, Novum, S-141 86 Huddinge, Sweden and
1 Department of Pharmacology, National Cardiovascular Center Research Institute, Osaka, Japan
 |
Abstract
|
---|
Evidence is accumulating that bile acids induce apoptosis in colonic cells. Therefore, it becomes important to study the underlying molecular mechanisms and the role of this phenomenon in tumor promotion. Minutes after exposure of HCT 116 and HT-29 cells to deoxycholate (DCA), DNA damage, measured using the COMET assay, was evident. Caspase-3 was rapidly activated in HCT 116 cells exposed to DCA, whereas in HT-29 cells, caspase-3 activation was delayed. Using transient transfections with reporter constructs, we showed that the transcription factors activator protein-1 (AP-1) and NF-kB were increased in HCT 116 cells, in a dose-dependent fashion, by DCA COX-2 promoter activity was also induced by DCA and using mutant COX-2 promoter plasmids, we showed that the ability of DCA to induce promoter activity was partly dependent upon a functional NF-kB and C/EBP site, and completely dependent on a functional c-AMP response element site. DNA damage thus appears to be the initiating event in DCA-induced apoptosis. In conclusion, the bile acid, DCA, has a major impact on apoptotic mechanisms in colonic cells and this may be contributing to its effect as a tumor promoter.
Abbreviations: APC, adenomatous polyposis coli; AP-1, activator protein-1; CRE, c-AMP response element; COX, cyclooxygenase; DCA, deoxycholic acid; DMEM, Dulbecco's modified Eagle's medium; FBS, fetal bovine serum; PARP, poly-ADP-ribose polymerase; PKC, protein kinase C; TRE, TPA response element.
 |
Introduction
|
---|
In colorectal cancer, genetic (1) and environmental factors contribute to the malignant transformation of colorectal epithelial cells. Epidemiological data implicate diet as a major environmental factor in colorectal carcinogenesis (2). A high fat consumption, seen in western societies, is associated with an increase in risk for colon cancer, as observed in many studies (3). One mechanism underlying this connection has been postulated to be increased levels of bile acids in the colon (4). Unconjugated deoxycholic acid (DCA) and chenodeoxycholic acid (CDCA) have been shown to be tumor promoters in animals (5,6) and higher levels of these bile acids have been reported in patients with adenomatous polyps and colon cancer (7,8). Earlier studies measured bile acids in total feces and attempted to correlate the levels to risk for colon cancer. However, more recently there has been a shift in focus towards the role of the aqueous phase of human feces (fecal water) in studies examining the mechanisms underlying the dietary etiology of colon cancer. The motivation for this is that components of this fecal fraction are more likely to be able to exert untoward effects on the cells of the colonic epithelium than components bound to food residues and the bacterial mass. The mechanism by which bile acids exert their tumor-promoting effect is poorly understood, although in recent years, many of the biological effects of bile acids at the molecular level are being clarified. Bile acids have, for example, been shown to activate protein kinase C (PKC) signaling pathways (9) involved in regulating cell proliferation, apoptosis and differentiation (10). DCA also induces COX-2 promoter activity, which leads to the formation of the cyclooxygenase (COX)-2 enzyme and consequent increases in prostaglandin production (11). In animal studies, blocking COX-2 chemically or by mutation causes a marked decrease in colon cancer development (12,13). COX-2 is therefore a potential target for therapeutics.
In this paper, we have further investigated some DCA effects upon cultured colonic cells and studied some of the mechanisms underlying the stress response it induces in the cell. This was done by studying the effect of DCA upon transcription factors, such as activator protein-1 (AP-1) and NF-kB and the inducible gene, COX-2. We demonstrate that DCA induces apoptosis in colonic cells, and that caspase-3 activation differs markedly between different colonic cells. This process can be inhibited by blocking p38 or by binding up calcium in the cell. Protecting the DNA with spermine also reduces the activation of caspase-3, induced by DCA. Cells exposed to DCA show severely damaged DNA. The DNA damage precedes the apoptosis in time and may be the reason why the cell enters apoptosis (14).
 |
Material and methods
|
---|
Chemicals
All chemicals were purchased from Calbiochem (Darmstadt, Germany) unless otherwise specified. DCA was prepared as stock solutions (0.1 mol/l) in water. SB 203580, BAPTA, PD98059, bisindolylmalemide, GO 6976, curcumin and apigenin were dissolved in dimethylsulfoxide. Spermine was dissolved in water.
Cells and culture
HT-29 and HCT 116 cell lines were purchased from ATCC (Rockville, MD) and used between passages 3 and 25 for all experiments. The HT-29 cell line is derived from a moderately well differentiated grade II human adenocarcinoma and is epithelial like. It harbors a mutated APC and expresses a truncated APC protein (15). HCT 116 cells are epithelial like and derived from a human carcinoma and harbor a normal APC gene (16). Both cells were maintained in Dulbecco's modified Eagle's medium (DMEM) with 10% fetal bovine serum (FBS), 2 mmol/l L-glutamine, 100 U/ml penicillin and 100 µg/ml streptomycin, in a humidified atmosphere of 95% air, 5% CO2 at 37°C. They were subcultivated every week and given fresh medium every other day. To obtain quiescent cells for the procedures outlined below, cultures were serum starved [0% FBS, 0.2% lacto albumin hydrolysate (LAH)] for 24 h.
Assay for DNA damage
COMET assay
The method was basically that of Klaude et al. (17) with minor modifications. Clear microscope slides (Mentzel super frost, Kebo, Sweden) were pre-treated with 40 µl of 0.3% low melting point (LMP) agarose (type VII, Sigma, St Louis, MO) and allowed to air-dry. The cells were incubated with the test substance for 15 min at 37°C in a 5% CO2 atmosphere. Ten microliters of cell suspension (0.51x106 cells/ml) was mixed with 150 µl of LMP agarose (0.75% in PBS kept at 37°C). A Flexi-Strip spatula was used to distribute the mixture on the pre-coated slides, which were thereafter left to set on an ice tray. After solidification, the slides were treated as follows. Lysis was performed in darkness for 1 h with an ice-cold freshly prepared solution containing 2 M NaCl, 25 mM EDTA, 20 mM Tris and 0.5% Triton X-100 pH 10. The slides were then placed in an electrophoresis buffer (0.3 M NaOH and 1 mM EDTA) in darkness at room temperature for 45 min. Electrophoresis was performed at room temperature, in darkness, in a Bio-Rad (Munich, Germany) subcell GT unit containing the same buffer, for 30 min at 20 V (0.67 V/cm). After electrophoresis, the slides were neutralized in 0.4 M Tris pH 7.4, air-dried, fixed in methanol and stored in a dry and dust free box until analysis. The DNA was stained with ethidium bromide (10 µg/ml in TAE) for 5 min followed by destaining for 5 min in TAE. The comets were examined in a fluorescence microscope (Olympus BH2 with a 20x apochromatic oil immersion objective), using the program Comet Assay II (Perceptive Instruments, Liverpool, UK). Images of 50 randomly selected cells were analyzed from each sample and tail moment was determined as described previously (18). Cell viability was assessed before and after the incubation by trypan blue exclusion.
Damage against DNA in vitro
c-DNA from a PCR reaction was incubated (37°C, 30 min) with PBS only or with DCA at increasing concentrations. The products were loaded onto an agarose gel stained with ethidium bromide (10 µg/ml in TAE) and subjected to electrophoresis for 1 h. The fluorescence from the bands was quantified in an imager (FUJI LAS 1000, Fuji, Stockholm, Sweden).
Caspase-3 assay
HCT 116 and HT-29 cells were grown to near confluence in 10 cm Petri dishes. Cells were washed with PBS and trypsinized. The pellet was resuspended in DMEM medium (0.1% FBS) and DCA at different concentrations was added and incubated for indicated time periods. After incubation, cells were centrifuged and the pellet was lyzed in a cell lysis buffer (10 mM TrisHCl, 10 mM NaH2PO4/NaHPO4 pH 7.5, 130 mM NaCl, 1% Triton X-100, 10mM NaPPi). Protein concentration of the cell extract after centrifugation was measured with the Bradford method at 595 nm.
A 20 µl cell extract from each treatment was added to new tubes containing 250 µl HEPES buffer (40 mM HEPES pH 7.5, 20% glycerol, 4mM DTT) and 2.5 µl fluorogenic marker, DEVD-AMC [N-acetyl-Asp-Glu-Val-Asp-AMC (7-amino-4-methylcoumarin), Pharmingen, San Diego, CA]. After incubation for 1 h at 37°C, the mixture was excited at 380 nm in a spectrofluorometer and emission was detected at 400 nm.
Western blotting
HCT 116 and HT-29 cells were treated with DCA for indicated time periods. To study if poly-ADP-ribose polymerase (PARP) had been cleaved in the cells after DCA exposure, 20 µg whole cell extracts were subjected to SDSpolyacrylamide gel electrophoresis, which was performed under reducing conditions on 8% polyacrylamide as described by Laemmli (19). The resolved proteins were transferred to a nitrocellulose sheet as detailed by Towbin et al. (20) and subjected to Ponceaus staining. The nitrocellulose membrane was then incubated with mouse monoclonal antibodies against PARP (Pharmingen, San Diego, CA). The blots were probed with the corresponding secondary antibodies to IgG (Dakopatts, Stockholm, Sweden, 1:3000 dilution) conjugated to horseradish peroxidase. The ECL western blot detection system (Amersham, Buckinghamshire, UK) was used according to the manufacturer's instructions. The resulting bands were confirmed by comparing the size of the protein in the cell extract with known molecular markers (Bio-Rad Laboratories, Munchen, Germany).
Plasmid preparation
The COX-2 promoter constructs (1432/+59, 327/+59, 220/+59, 52/+59, CRM, KBM, ILM) were a kind gift from Drs Inoue and Tanabe (National Cardiovascular Center Research Institute, Osaka, Japan) (21,22). The reporter plasmids [TRE (TPA response element)]2tkLuc and (NF-kB)3 tkLuc were a gift from Dr Sam Okret (Karolinska Institute, Sweden) (23). The plasmid DNA was purified from bacteria cultures as described previously (24).
Transfection and luciferase assay
HCT 116 cells were grown to 50% confluence in a 75 cm2 culture dish. The medium was removed and the cells were washed with PBS. OPTIMEM (Life Technologies, Gibco, Scotland, UK), 2 µg/ml plasmid DNA and 10 µg/ml Lipofectin reagent (Life Technologies) were added to the culture dish. After 6 h of transfection, the transfection mixture was removed and the cells were washed with PBS and trypsinized. Full growth medium (DMEM 10%) was then added to the cells to inhibit the trypsin, and the cells were pelleted by centrifugation at 2000 r.p.m. Cells (30x103) were seeded out in a 24 well plate, and DMEM 10% was added to each well and incubated overnight. The cells were then starved in DMEM (0.2% LAH) for 24 h, before being incubated with DCA for an additional 15 h. The medium was removed and the cells were washed with PBS. Lysis buffer (100 µl, 25 mmol/l trisphosphate pH 7.8, 15% glycerol, 2% CHAPS, 1% lecithin, 1% bovine serum albumin, 0.1% EGTA pH 8.0, 8 mmol/l MgCl2, 1 mmol/l dithiothreitol and 0.4 mmol/l phenylmethylsulfonyl fluoride) was added to each well and the cells lysed during 30 min. A total of 50 µl of the cell lysates was transferred to a non-transparent 96 well plate and lucifierin mix (100 µl) (GenGlow, Bioorbit, Turku, Finland) was added per well. Assay for luciferase activity was performed in the automatic luminometer Lucy 1 (Anthos Labtec Instruments, Salzburg, Austria) according to the manufacturer's instructions. Luciferase activity was expressed per microgram of protein in the cell lysate. The luciferase activity of the untreated control cells in each experiment was set to 100% and the resulting activities for the test agents were calculated in relation (percent) to the control cells.
Cytotoxicity assay
Cell proliferation/toxicity using the HT-29 and HCT 116 cells was measured using the Celltiter 96 proliferation kit (Promega, Madison,WI) as described previously (24).
TUNEL assay
Apoptosis was evaluated using the TUNEL [Tdt (terminal deoxynucleotidyl transferase)-mediated dUTP-x (x = biotin, fluorescein) nick end labeling] assay (Böerhinger-Manheim, Indianapolis, IN) and flow cytometry analysis. Apoptotic DNA cleavage may yield single-stranded as well as double-stranded DNA breaks (nicks). Both types of breaks can be detected by labeling the 3'-OH termini with modified nucleotides (for example, fluorescein-dUTP) in an enzymatic reaction. Thus, 0.5x106 freshly harvested cells (attached + floating) were washed with PBS and fixed in 2% formaldehyde in PBS (1 ml) for 15 min at room temperature. Cells were washed with PBS and incubated with 0.1% saponine in balanced salt solution (4 ml) for 3 min. Subsequently, the cells were incubated with the TUNEL reaction mixture, 50 µl of enzyme solution (Tdt) and 450 µl of label solution (fluorescein d-UTP) for 1 h at 37°C in the dark in a humidified atmosphere. During this incubation period, Tdt catalyzes the addition of fluorescein-dUTP to free 3'-OH groups in single and double-stranded DNA. Omission of Tdt from the staining protocol constituted the negative control. After washing the cells with PBS, the label incorporated into the damaged sites of DNA was measured using a FACScan flow cytometer (Becton Dickinson, Mountain View, CA). For every experiment 10 000 cells were analyzed.
Statistics
In order to analyze changes in induction of COX-2 promoter activity, AP-1- and NF-kB-dependent gene transcription, Student's t-test (two-tailed) was used. Pearson correlation was used to analyze correlations between variables. Descriptive and graphical methods were also used to characterize the data. All tests were performed with the software package STATISTICA 5.0 (Statsoft, Tulsa, OK) and the P-value is given for comparisons (*P < 0.05, **P < 0.01, ***P < 0.001).
 |
Results
|
---|
DCA induces DNA damage in intact cells but not on naked DNA
HCT 116 and HT-29 cells were exposed to DCA for 15 min at 37°C and analyzed for DNA damage using the single cell electrophoresis method. As seen in Figure 1A
, DCA caused a significant increase in DNA strand breaks which were 3-fold higher (P < 0.001) than untreated cells for both HCT 116 and HT-29 cells. There was no difference in the DNA damage response between HT-29 and HCT 116 cells treated with DCA. In Figure 1B
, DCA was incubated with naked DNA, for 30 min at 37°C, and the DNA products visualized on an agarose gel were quantified in an imager. DCA at 501000 µM did not have the capacity to break intact DNA in vitro.

View larger version (20K):
[in this window]
[in a new window]
|
Fig. 1. (A) Measurement of DNA strand breaks in HCT 116 and HT-29 cells after exposure to DCA 500 µM (15 min). Bars represent mean tail moment ± SD (n = 50). (B) Intact DNA exposed to DCA (01000 µM) for 30 min at 37°C and visualized on an agarose gel. L, Ladder; lane 1, untreated DNA; lane 2, DCA 50 µM; lane 3, DCA 250 µM; lane 4, DCA 500 µM; lane 5, DCA 1000 µM; lane 6, positive control, 0.4% ammonium persulfate; lane 7, positive control, 0.8% ammonium persulfate.
|
|
Caspase-3 activation in HCT 116 and HT-29 cells after exposure to DCA
We and others, have shown previously that DCA induces apoptosis in cultured colonic cells (25,26). In order to further investigate how fast the apoptotic process is initiated we compared the activation of caspase-3 in two different cell lines, HCT 116 and HT-29. The cells were exposed to DCA at 50500 µM from 0 to 75 min (Figure 2A
). HCT 116 cells exposed to 50 µM DCA did not show any significant caspase-3 activity over the time period studied. DCA (250 µM) induced a dose-dependent activation of caspase-3 after 15 min, reaching a maximum after 60 min (800% above untreated cells). DCA (500 µM) induced a higher doseresponse activation of caspase-3 and a maximum was reached after 45 min exposure (1800% above control). In HT-29 cells, no significant activation of caspase-3 was observed during this short-term exposure to DCA regardless of concentration used (50, 250 or 500 µM). In Figure 1A
, only DCA 500 µM is plotted for the HT-29 cells. In Figure 2B
, HCT 116 and HT-29 cells were treated with 250 µM DCA for 0.5, 1, 2, 4, 6, 8 and 24 h and activation of caspase-3 was studied. HCT 116 cells responded, as seen previously, with a rapid increase in caspase-3 activity, which reached a maximum after 60 min and then rapidly declined to basal levels after 24 h of treatment. In HT-29 cells, on the other hand, only a weak caspase-3 activity was observed after exposure to DCA for the earlier time points. However, after long-term exposure (24 h), the caspase-3 activity increased significantly (>300% above control, P < 0.001). The activation of caspase-3 by DCA in HT-29 cells was significantly delayed compared with HCT 116 cells, which may explain why HCT 116 cells are more sensitive to DCA than HT-29 cells [the numbers of surviving cells after 24 h exposure to DCA (250 µM) are 48 ± 9 and 96 ± 15% for HCT 116 and HT-29, respectively, absorbance of untreated cells was set to 100%]. We also measured total cell death (cytotoxicity assay) in HT-29 cells exposed to DCA for 72 h and determined to what extent this was due to apoptosis (TUNEL assay). There was a linear correlation between total cell death and apoptosis (r = 0.96, P < 0.001) and the majority of the cell death was due to apoptosis (data not shown).

View larger version (16K):
[in this window]
[in a new window]
|
Fig. 2. (A) Dose-dependent effects of DCA on caspase-3 activity in HCT 116 and HT-29 cells after short-term treatment. HCT 116 cells: DCA 50 µM (triangle), DCA 250 µM (circle), DCA 500 µM (square); HT-29 cells: DCA 500 µM (cross). (B) Dose-dependent effects of DCA on caspase-3 activity in HCT 116 and HT-29 cells after long-term treatment. HCT 116: DCA 250 µM (squares), HT-29: DCA 250 µM (triangle).
|
|
PARP cleavage in HCT 116 and HT-29 cells after DCA exposure
In order to study if protein substrates for caspase-3 in the cell are cleaved as rapidly as was observed in the in vitro system above, western blots on the same cell extracts (HCT 116, DCA 250 µM, 075 min) were performed and the membrane was probed with an antibody directed against the PARP protein. The result (Figure 3
) shows PARP cleavage in the cell extracts (116 and 85 kDa) which correlates with cleavage of the fluorogenic marker, DEVD-AMC in vitro seen in Figure 2A
. The protein fragment of PARP (85 kDa) was visible in cells which were treated with DCA (250 µM) for 45 min. In HT-29 cells, no cleavage of PARP was seen after these early time periods (075 min, data not shown). However, long-term exposure (up to 72 h) of HT-29 cells to DCA results in cleavage of PARP, which we have demonstrated previously (26).

View larger version (35K):
[in this window]
[in a new window]
|
Fig. 3. Western blot, showing PARP protein (116 kDa) from cell extracts (HCT 116 cells) treated with DCA (250 µM) for 1575 min. Lane 1, untreated 75 min; lane 2, 15 min treatment; lane 3, 30 min; lane 4, 45 min treatment; lane 5, 60 min treatment; lane 6, 75 min, treatment. The lower band appearing in lanes 46 represent cleaved PARP (85 kDa).
|
|
Blocking of caspase-3 activation by inhibitors
Inhibitors were used in order to test if caspase-3 activity induced by DCA (500 µM) in HCT 116 cells could be prevented and to give some information about the signal transduction pathways involved. HCT 116 cells were used in these and subsequent experiments because of the larger and more rapid caspase response to DCA in these cells and the greater practical difficulties in transfecting the HT-29 cells. All inhibitors were pre-incubated for 15 min, before exposure to DCA. In Figure 4A
, a dose-dependent inhibition of caspase-3 activity was observed when the cells had been pre-incubated with the antioxidant curcumin (1050 µM). In a similar manner, the internal calcium chelator, BAPTA, was able to prevent caspase-3 from being activated. The PKC blocking agent (GO 6976, 50 nM) had no significant inhibitory effect on caspase-3 activity (Figure 4B
). However, bisindolylmalemide (100 nM) had a partial blocking effect on caspase-3 activation by DCA. Interestingly, by blocking the stress-related map kinase, p38 (SB 203580, 30 µM), a complete suppression of caspase-3 activity was observed. Blocking MEK1/2 using PD98059 (10 µM) had, however, no effect on caspase-3 activation. The DNA protective amine, spermine (1 mM) could partially block induction of caspase-3 activation. Fifty percent of the actual apoptosis induced by DCA (500 µM) in HT-29 cells could be blocked by cell-permeable caspase-3 (Z-DEVD-FMK) and caspase 6 (Z-VEID-FMK) inhibitors at 100 µM (A.Haza, Department of Medical Nutrition, Karolinska Institutet, personal communication).

View larger version (34K):
[in this window]
[in a new window]
|
Fig. 4. (A) Caspase-3 activity in HCT 116 cells, treated with DCA (500 µM) for 30 min, with and without pre-treatment of inhibitors. CUR 10CUR 50, curcumin 1050 µM; BP 1030, BAPTA 1030 µM. (B) BIS, bisindolylmalemide 100 nM; GO, GO 6976 50 nM; PD, PD 98059 (10 µM); SB, SB 203580 (30 µM); SPER, spermine (1 mM). Bars represent mean ± SD (n = 2).
|
|
Effects on NF-kB-dependent gene transcription by DCA
It has been reported previously that NF-kB is induced under cellular stress and NF-kB inhibitory effects on caspases have been described (27). It therefore became important to study if DCA induced NF-kB in HCT 116 cells.
HCT 116 cells were transiently transfected with a NF-kB-tk-luc plasmid and exposed to DCA at different concentrations for 15 h. Luciferase activity (Figure 5
) was quantified and showed a dose-dependent increase in NF-kB-dependent gene transcription from 180% (P < 0.05) at DCA (100 µM) up to 300% induction (P < 0.05) at DCA (300 µM). DCA concentrations <100 µM did not induce the NF-kB-dependent gene transcription. TNF-
(400 U) was used as a positive control for induction of NF-kB-dependent gene transcription (Figure 5
).

View larger version (31K):
[in this window]
[in a new window]
|
Fig. 5. Effects of DCA on NF-kB-dependent gene transcription in HCT 116 cells exposed to increasing concentrations of DCA for 15 h, before assay of luciferase activity. TNF- , positive control. Bars represent mean ± SD (n = 4).
|
|
Inhibition of AP-1-dependent gene transcription induced by DCA
HCT 116 cells were transfected with a TRE-plasmid and exposed to DCA (250 µM) with and without inhibitors. The AP-1-dependent gene transcription was increased in HCT 116 cells in a dose-dependent fashion by DCA (Figure 6
). Pre-treatment with GO 6976 (50 nM) had no significant effect upon AP-1-dependent gene transcription induced by DCA (250 µM) and neither had the MEK1/2 inhibitor, PD98059 (10 µM) or apigenin (20 µM). However, blocking the stress-related map kinase, p38, using SB 203580 (30 µM) totally abolished the AP-1-dependent gene transcription induced by DCA. Binding up calcium in the cell, using BAPTA (30 µM), also prevented AP-1 induction. Blocking PKC with bisindolylmalemide (100 nM) could partly prevent induction of AP-1. Curcumin (40 µM) completely abolished AP-1 induction induced by DCA.

View larger version (47K):
[in this window]
[in a new window]
|
Fig. 6. Effects of DCA (250 µM) with and without inhibitors on AP-1-dependent gene transcription in HCT 116 cells. GO, GO 6976 50 nM; PD, PD 98059 10 µM; SB, SB 203580 30 µM; BP, BAPTA 30 µM; BIS, bisindolylmalemide 10 nM; API, apigenin 20 µM; CUR, curcumin 40 µM. Bars represent mean ± SD (n = 4).
|
|
Response elements in COX-2 promoter involved in transactivation by DCA
We have shown previously that DCA induces the COX-2 promoter in HCT 116 cells (28). The purpose of the present experiment was to understand what response elements in the COX-2 promoter are involved in DCA induced transactivation. Different lengths of the COX-2 promoter plasmid were transiently transfected into HCT 116 cells and the cells were exposed to DCA 250 mM for 15 h. In Figure 7A
, the full length COX-2 promoter construct (1432/+59), containing sites for SP1, GRE, GATA, NF-kB, PEA3, SP1, NF-kB, AP-2, C/EBP and c-AMP response element (CRE) (29) induced a luciferase activity of 533 ± 33% above control (100%). The response of the deleted construct (327/+59), containing sites for SP1, NF-kB, AP-2, C/EBP, CRE was of the same magnitude, 588 ± 32%. A clear reduction in response was observed with the (220/+59) construct (AP-2, C/EBP, CRE), which gave an induction of 284 ± 80%. The shortest construct (52/+59), lacking all response elements gave no induction compared with untreated cells (control).

View larger version (17K):
[in this window]
[in a new window]
|
Fig. 7. (A) Deletion mutants of the COX-2 promoter, transiently transfected into HCT 116 cells and exposed to DCA 250 µM for 15 h. 1432/+59 represents the COX-2 plasmid, with 1432 bases upstream from the transcription start site of the COX-2 promoter. (B) Mutants of the COX-2 promoter plasmid, transiently transfected into HCT 116 cells and exposed to DCA 250 µM. KBM, represents the 327/+59 COX-2 plasmid, with a mutation in the NF-kB site. ILM, represents the 327/+59 COX-2 plasmid, with a mutation in the C/EBP site. CRM represents the 327/+59 COX-2 plasmid, with a mutation in the CRE site. Bars represent mean ± SD (n = 8).
|
|
In Figure 7B
, the wild-type (327/+59) plasmid, and mutated forms of the (327/+59) plasmid were transfected into HCT 116 cells and exposed to DCA 250 µM. Using the KBM plasmid (327/+59, NF-kB site mutated), resulted in a significant reduction in luciferase response compared with wild-type (327/+59), 167 ± 17 and 588 ± 32%, respectively. Using the ILM plasmid (C/EBP mutation), a 50% reduction (269 ± 40%) was observed compared with the wild-type (327/+59) construct. Comparing the CRM plasmid (CRE mutation) with the wild-type plasmid, a complete suppression of promoter activity was observed, 75 ± 58%.
 |
Discussion
|
---|
The precise mechanisms by which bile acids act as tumor promoters are not fully understood. Much work has focused upon the ability of bile acids to induce apoptosis in colonic cells, as a possible mechanism to explain its tumor-promoting effect in the colon (30). In this paper, we have studied the molecular effects resulting from exposure to DCA in the colonic cell lines HCT 116 and HT-29. The concentrations of DCA employed in the present study may seem to be high, but such concentrations have been reported to occur in the fecal water of risk groups for colon cancer (31,32). Shortly after exposure of these cells to DCA, when no cell death is evident, DNA damage can be detected. The mechanism behind damage of DNA after DCA exposure is not known. The fact that caspase-3 is rapidly induced in association with DNA damage and could be inhibited by DNA protective agents like spermine, supports the hypothesis that DNA damage may be inititating the apoptosis program. The antioxidant curcumin also reduced caspase-3 activity, which may indicate that DCA is provoking a free radical process in the cells that may lead to damage of the DNA. Stultz-Washo et al. (33) have shown that reactive nitrogen species are formed (peroxynitrite, coming from nitric oxide and superoxide) in HT-29 cells after DCA exposure (500 µM). The work by Craven et al. (34) has earlier shown that bile salt increases reactive oxygen production in vivo. Our observation that DCA did not have the capacity to induce damage to naked DNA in vitro also suggests that it is not DCA per se which is responsible for the damage, but rather a DCA-induced cellular process such as those mentioned above.
The activation of caspase-3 in HCT 116 and HT-29 cells differed markedly between the two cells after DCA exposure. There was a rapid and transient induction of caspase-3 activation upon DCA treatment (250 and 500 µM) in HCT 116 cells, and a delayed response (24 h) in HT-29 cells. This may be one explanation for our observation that HT-29 cells were more resistant than HCT 116 cells to cell death after 24 h exposure to DCA, despite equal damage to DNA induced by the bile acid. It is interesting to note that HCT 116 cells harbor normal APC and p53 genes (15,35), both of which are mutated and non-functional in HT-29 cells (16,35). Both these genes are important for the initiation of the apoptosis program, and may contribute to explaining the difference observed between the two cell lines in the apoptosis response to DCA treatment (14,36).
DCA is inducing stress in the cells, and the stress-related map kinase, p38 is phosphorylated as shown previously (37). Interestingly, in our system blocking p38 could prevent caspase-3 activation induced by DCA. The activation of caspase-3 after DCA exposure seems to be calcium dependent, as the activation of caspase-3 could also be prevented by pre-incubation with BAPTA, an internal calcium chelator.
The results with the PKC inhibitors indicate that the induction of caspase-3 activity by bile acid is mediated, at least in part, by specific isoforms of PKC. Interestingly, blocking MEK1/2 had no effect on induction of caspase activity.
We measured the total cell death in HT-29 cells exposed to DCA at increasing concentrations, and in addition we quantified the number of apoptotic cells using the TUNEL staining. The result showed that there was a good correlation between the total cell death and apoptosis. At higher concentrations of DCA (>250 µM), the cell death was due to both necrosis and apoptosis; however, the majority of the cells were undergoing apoptosis. Evidence that DCA induces apoptosis in vivo under normal physiological conditions does exist. Perfusion studies in the rat colon have demonstrated that DCA has the capacity to induce the cell death referred to above (38).
Transfection of a NF-kB-tk-luc into HCT 116 cells and exposure (15 h) to increasing concentrations of DCA resulted in a dose-dependent increase of reporter gene activity, indicating that exposure to bile acid led to NF-kB activation. It is interesting, that the caspase-3 activity in the HCT 116 cells already after 15 h exposure to DCA had returned to basal levels. This becomes interesting, as it has been reported previously that NF-kB can inhibit caspase-3 activity via IAP activation (27). This may be interpreted as a survival mechanism, and is part of the cells response to the stress provoked by DCA. In transient transfection studies with a TRE-tk-luc plasmid in HCT 116 cells, DCA exposure resulted in a dose-dependent increase in AP-1-dependent gene transcription. This activation of AP-1 could be prevented by blocking p38, binding up calcium in the cells, as well as by the antioxidant curcumin. Blocking MEK1/2 had no effect, and results with PKC inhibitors indicated that the AP-1 activation was mediated, at least in part, by specific PKC isoforms. There is a good correlation regarding the inhibitors that are needed to block activation of caspase-3- and AP-1-dependent gene transcription, which indicates that they are both part of the same stress response.
COX-2 is also involved in the stress response, and activity of COX-2 results in production of prostaglandins, which can make the cells more resistant to apoptosis (39). In order to elucidate the main transactivation domain in the COX-2 promoter, we transfected the HCT 116 cells with different lengths of the COX-2 promoter (luciferase reporter constructs) and exposed them to DCA. We observed that for the longer (1432/+59) and the shorter (327/+59) plasmid construct, there was no difference in reporter gene activity induced by DCA. However, between the (327/+59) and (220/+59) plasmid construct, there was a significant decrease in reporter gene activity. The difference between these two promoter constructs is that the (220/+59) form lacks the NF-kB response element. Use of the shortest plasmid construct, (52/+59), which only contains a TATA box, resulted in no induction by DCA compared with untreated cells. Using mutant (327/+59) plasmids, we could demonstrate that a mutation in the NF-kB binding site significantly reduced the reporter gene activity compared with the response using wild-type (327/+59). Mutation of the C/EBP binding site resulted in a 50% decrease in reporter gene activity. Finally, mutation of the CRE (c-AMP response element) totally abolished the reporter gene activity induced by DCA. In summary, the results indicate that DCA induction of the COX-2 gene is complex, and involves multiple transcription factors binding to NF-kB, C/EBP and CRE elements. It is interesting to note that the CRE in the COX-2 promoter is similar to a TRE element, and may bind AP-1 transcription factors as well as CREB factors/ATF (40). In addition, it has been demonstrated that C/EBP factors can also dimerize with AP-1 transcription factors (41). Our results are in agreement with those of Zhang et al. (11) who showed that DCA treatment of esophageal adenocarcinoma cells resulted in activation of the COX-2 promoter. However, our results hopefully extend our understanding of bile acid induced signaling by mapping the response elements in the COX-2 promoter that are required for full activation of gene transcription in colonic cells.
In conclusion, DCA induces apoptosis in HCT 116 and HT-29 cells, and cell death is delayed in HT-29 cells, which correlates with delayed activation of caspase-3. The activation of caspase-3 is dependent upon active p38 and normal calcium signaling. Cells undergoing DCA-induced apoptosis, but which are still viable have reduced caspase-3 levels and NF-kB and AP-1 are induced. In these cells, COX-2 promoter activity is also induced, and mapping of the main transactivation domain, indicates that full activation of the promoter needs functional NF-kB, C/EBP and CRE elements located in the 327 bases up from start of transcription. Which transcription factors bind to these elements of the COX-2 promoter when the cells are exposed to DCA is not fully elucidated and requires further investigation. Finally, the bile acid, DCA, has a major impact on the apoptotic machinery in colonic cells but the role of this effect in tumor promotion requires further work.
 |
Notes
|
---|
2 To whom correspondence should be addressed Email: joseph.rafter{at}mednut.ki.se 
 |
Acknowledgments
|
---|
This work was supported by the Swedish Cancer Society. B.G. is grateful to Robert Lundbergs foundation for financial support.
 |
References
|
---|
-
Fearon,E.R. and Vogelstein,B. (1990) A genetic model for colorectal tumorigenesis. Cell, 61, 759767.[ISI][Medline]
-
Sandler,R.S., Lyles,C.M., Peipins,L.A., McAuliffe,C.A., Woosley,J.T. and Kupper,L.L. (1993) Diet and risk of colorectal adenomas: macronutrients, cholesterol and fiber. J. Natl Cancer Inst., 85, 884891.[Abstract]
-
Jenkins,D.J., Jenkins,A.L., Rao,A.V. and Thompson,L.U. (1986) Cancer risk: possible protective role of high carbohydrate high fiber diets. Am. J. Gastroenterol., 81, 931935.[ISI][Medline]
-
Weisburger,J.H., Reddy,B.S., Barnes,W.S. and Wynder,E.L. (1983) Bile acids, but not neutral sterols, are tumor promoters in the colon in man and in rodents. Environ. Health Perspect., 50, 101107.[ISI][Medline]
-
Reddy,B.S., Watanabe,K., Weisburger,J.H. and Wynder,E.L. (1977) Promoting effect of bile acids in colon carcinogenesis in germ-free and conventional F344 rats. Cancer Res., 37, 32383242.[ISI][Medline]
-
Mahmoud,N.N., Dannenberg,A.J., Bilinski,R.T., Mestre,J.R., Chadburn,A., Churchill,M., Martucci,C. and Bertagnolli,M.M. (1999) Administration of an unconjugated bile acid increases duodenal tumors in a murine model of familial adenomatous polyposis. Carcinogenesis, 20, 299303.[Abstract/Free Full Text]
-
Imray,C.H., Radley,S., Davis,A., Barker,G., Hendrickse,C.W., Donovan,I.A., Lawson,A.M., Baker,P.R. and Neoptolemos,J.P. (1992) Faecal unconjugated bile acids in patients with colorectal cancer or polyps. Gut, 33, 12391245.[Abstract]
-
Reddy,B.S., Mastromarino,A., Gustafson,C., Lipkin,M. and Wynder,E.L. (1976) Fecal bile acids and neutral sterols in patients with familial polyposis. Cancer, 38, 16941698.[ISI][Medline]
-
Huang,X.P., Fan,X.T., Desjeux,J.F. and Castagna,M. (1992) Bile acids, non-phorbol-ester-type tumor promoters, stimulate the phosphorylation of protein kinase C substrates in human platelets and colon cell line HT29. Int. J. Cancer., 52, 444450.[ISI][Medline]
-
Clemens,M.J., Trayner,I. and Menaya,J. (1992) The role of protein kinase C isoenzymes in the regulation of cell proliferation and differentiation. J. Cell Sci., 103, 881887.[Free Full Text]
-
Zhang,F., Subbaramaiah,K., Altorki,N. and Dannenberg,A.J. (1998) Dihydroxy bile acids activate the transcription of cyclooxygenase-2. J. Biol. Chem., 273, 24242428.[Abstract/Free Full Text]
-
Oshima,M., Dinchuk,J.E., Kargman,S.L., Oshima,H., Hancock,B., Kwong,E., Trzaskos,J.M., Evans,J.F. and Taketo,M.M. (1996) Suppression of intestinal polyposis in Apc delta716 knockout mice by inhibition of cyclooxygenase 2 (COX-2). Cell, 87, 803809.[ISI][Medline]
-
Jacoby,R.F., Seibert,K., Cole,C.E., Kelloff,G. and Lubet,R.A. (2000) The cyclooxygenase-2 inhibitor celecoxib is a potent preventive and therapeutic agent in the min mouse model of adenomatous polyposis. Cancer Res., 60, 50405044.[Abstract/Free Full Text]
-
Rich,T., Allen,R.L. and Wyllie,A.H. (2000) Defying death after DNA damage. Nature, 407, 777783.[ISI][Medline]
-
Hsi,L.C., Angerman-Stewart,J. and Eling,T.E. (1999) Introduction of full-length APC modulates cyclooxygenase-2 expression in HT-29 human colorectal carcinoma cells at the translational level. Carcinogenesis, 20, 20452049.[Abstract/Free Full Text]
-
Kutchera,W., Jones,D.A., Matsunami,N., Groden,J., McIntyre,T.M., Zimmerman,G.A., White,R.L. and Prescott,S.M. (1996) Prostaglandin H synthase 2 is expressed abnormally in human colon cancer: evidence for a transcriptional effect. Proc. Natl Acad. Sci. USA, 93, 48164820.[Abstract/Free Full Text]
-
Klaude,M., Eriksson,S., Nygren,J. and Ahnstrom,G. (1996) The comet assay: mechanisms and technical considerations. Mutat. Res., 363, 8996.[ISI][Medline]
-
Venturi,M., Hambly,R.J., Glinghammar,B., Rafter,J.J. and Rowland,I.R. (1997) Genotoxic activity in human faecal water and the role of bile acids: a study using the alkaline comet assay. Carcinogenesis, 18, 23532359.[Abstract]
-
Laemmli,U.K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature, 227, 680685.[ISI][Medline]
-
Towbin,H., Staehelin,T. and Gordon,J. (1979) Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure and some applications. Proc. Natl Acad. Sci. USA, 76, 43504354.[Abstract]
-
Inoue,H., Yokoyama,C., Hara,S., Tone,Y. and Tanabe,T. (1995) Transcriptional regulation of human prostaglandin-endoperoxide synthase-2 gene by lipopolysaccharide and phorbol ester in vascular endothelial cells. Involvement of both nuclear factor for interleukin-6 expression site and cAMP response element. J. Biol. Chem., 270, 2496524871.[Abstract/Free Full Text]
-
Inoue,H., Nanayama,T., Hara,S., Yokoyama,C. and Tanabe,T. (1994) The cyclic AMP response element plays an essential role in the expression of the human prostaglandin-endoperoxide synthase 2 gene in differentiated U937 monocytic cells. FEBS Lett., 350, 5154.[ISI][Medline]
-
Caldenhoven,E., Liden,J., Wissink,S., Van de Stolpe,A., Raaijmakers,J., Koenderman,L., Okret,S., Gustafsson,J.A. and Van der Saag,P.T. (1995) Negative cross-talk between RelA and the glucocorticoid receptor: a possible mechanism for the antiinflammatory action of glucocorticoids. Mol. Endocrinol., 9, 401412.[Abstract]
-
Glinghammar,B., Holmberg,K. and Rafter,J. (1999) Effects of colonic lumenal components on AP-1-dependent gene transcription in cultured human colon carcinoma cells. Carcinogenesis, 20, 969976.[Abstract/Free Full Text]
-
Hague,A., Elder,D.J., Hicks,D.J. and Paraskeva,C. (1995) Apoptosis in colorectal tumour cells: induction by the short chain fatty acids butyrate, propionate and acetate and by the bile salt deoxycholate. Int. J. Cancer, 60, 400406.[ISI][Medline]
-
Haza,A.I., Glinghammar,B., Grandien,A. and Rafter,J. (2000) Effect of colonic luminal components on induction of apoptosis in human colonic cell lines. Nutr. Cancer, 36, 7989.[ISI][Medline]
-
LaCasse,E.C., Baird,S., Korneluk,R.G. and MacKenzie,A.E. (1998) The inhibitors of apoptosis (IAPs) and their emerging role in cancer. Oncogene, 17, 32473259.[ISI][Medline]
-
Glinghammar,B. and Rafter,J. (2001) Colonic luminal contents induce cyclooxygenase 2 transcription in human colon carcinoma cells. Gastroenterology, 120, 401410.[ISI][Medline]
-
Tazawa,R., Xu,X.M., Wu,K.K. and Wang,L.H. (1994) Characterization of the genomic structure, chromosomal location and promoter of human prostaglandin H synthase-2 gene. Biochem. Biophys. Res. Commun., 203, 190199.[ISI][Medline]
-
Payne,C.M., Bernstein,H., Bernstein,C. and Garewal,H. (1995) Role of apoptosis in biology and pathology: resistance to apoptosis in colon carcinogenesis. Ultrastruct. Pathol., 19, 221248.[ISI][Medline]
-
Ejderhamn,J., Rafter,J.J. and Strandvik,B. (1991) Faecal bile acid excretion in children with inflammatory bowel disease. Gut, 32, 13461351.[Abstract]
-
de Kok,T.M., van Faassen,A., Glinghammar,B., Pachen,D.M., Eng,M., Rafter,J.J., Baeten,C.G., Engels,L.G. and Kleinjans,J.C. (1999) Bile acid concentrations, cytotoxicity and pH of fecal water from patients with colorectal adenomas. Dig. Dis. Sci., 44, 22182225.[ISI][Medline]
-
Washo-Stultz,D., Hoglen,N., Bernstein,H., Bernstein,C. and Payne,C.M. (1999) Role of nitric oxide and peroxynitrite in bile salt-induced apoptosis: relevance to colon carcinogenesis. Nutr. Cancer, 35, 180188.[ISI][Medline]
-
Craven,P.A., Pfanstiel,J. and DeRubertis,F.R. (1986) Role of reactive oxygen in bile salt stimulation of colonic epithelial proliferation. J. Clin. Invest., 77, 850859.[ISI][Medline]
-
Chendil,D., Oakes,R., Alcock,R.A., Patel,N., Mayhew,C., Mohiuddin,M., Gallicchio,V.S. and Ahmed,M.M. (2000) Low dose fractionated radiation enhances the radiosensitization effect of paclitaxel in colorectal tumor cells with mutant p53. Cancer, 89, 18931900.[ISI][Medline]
-
Browne,S.J., Williams,A.C., Hague,A., Butt,A.J. and Paraskeva,C. (1994) Loss of APC protein expressed by human colonic epithelial cells and the appearance of a specific low-molecular-weight form is associated with apoptosis in vitro. Int. J. Cancer, 59, 5664.[ISI][Medline]
-
Qiao,D., Chen,W., Stratagoules,E.D. and Martinez,J.D. (2000) Bile acid-induced activation of activator protein-1 requires both extracellular signal-regulated kinase and protein kinase C signaling. J. Biol. Chem., 275, 1509015098.[Abstract/Free Full Text]
-
Rafter,J.J., Eng,W.W., Furrer,R., Medline,A. and Bruce,W.R. (1986) Effects of calcium and pH on the mucosal damage produced by deoxycholic acid in the rat colon. Gut, 27, 13201329.[Abstract]
-
Tsujii,M. and DuBois,R.N. (1995) Alterations in cellular adhesion and apoptosis in epithelial cells overexpressing prostaglandin endoperoxide synthase 2. Cell, 83, 493501.[ISI][Medline]
-
Nomura,N., Zu,Y.L., Maekawa,T., Tabata,S., Akiyama,T. and Ishii,S. (1993) Isolation and characterization of a novel member of the gene family encoding the cAMP response element-binding protein CRE-BP1. J. Biol. Chem., 268, 42594266.[Abstract/Free Full Text]
-
Hsu,W., Kerppola,T.K., Chen,P.L., Curran,T. and Chen-Kiang,S. (1994) Fos and Jun repress transcription activation by NF-IL6 through association at the basic zipper region. Mol. Cell. Biol., 14, 268276[Abstract]
Received June 22, 2001;
accepted January 14, 2002.