Differential effects of deoxycholic acid and taurodeoxycholic acid on NF-{kappa}B signal transduction and IL-8 gene expression in colonic epithelial cells

M. Mühlbauer,1 B. Allard,3 A. K. Bosserhoff,2 S. Kiessling,1 H. Herfarth,1 G. Rogler,1 J. Schölmerich,1 C. Jobin,3 and C. Hellerbrand1

1Department of Internal Medicine I and 2Institute of Pathology, University of Regensburg, D-93042 Regensburg, Germany; and 3University of North Carolina at Chapel Hill, North Carolina 27599

Submitted 7 August 2003 ; accepted in final form 8 January 2004


    ABSTRACT
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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Several effects of bile acids (BAs) on colonic epithelial cells (CECs) have been described, including induction of proliferation and apoptosis. Some of these effects are mediated through activation of the NF-{kappa}B transcriptional system. In this study, we investigated the molecular mechanisms underlying the BA-induced gene expression in CECs. The human CEC line HT-29 and primary human CECs were treated with dilutions of salts of deoxycholic acid (DCA) and taurodeoxycholic acid (TDCA). NF-{kappa}B binding activity was analyzed with EMSA, RelA translocation with immunofluorescence, and I{kappa}B{alpha}- and RelA-phosphorylation with Western blot analysis. IL-8 mRNA and protein expression were assessed by quantitative PCR and ELISA. Functional impact of NF-{kappa}B activation was determined by blocking the proteasome activity with MG132 or by preventing IKK activity with a dominant-negative IKK{beta} delivered by adenoviral dominant-negative (dn) IKK{beta} (Ad5dnIKK{beta}). DCA and TDCA induced IL-8 expression in a dose- and time-dependent manner. It is interesting that DCA but not TDCA induced I{kappa}B{alpha}-phophorylation, RelA translocation, and NF-{kappa}B binding activity. Accordingly, the proteasome inhibitor MG132 blocked DCA- but not TDCA-induced IL-8 gene expression. In contrast, TDCA-induced IL-8 gene expression correlated with enhanced RelA phosphorylation, which was blocked by Ad5dnIKK{beta}. Our data suggest that DCA-induced signal transduction mainly utilized the I{kappa}B degradation and RelA nuclear translocation pathway, whereas TDCA primarily induced IL-8 gene expression through RelA phosphorylation. These differences may have implications for the understanding of the pathophysiology of inflammation and carcinogenesis in the gut.

intestinal epithelial cells; colitis; inflammation; cancer


PRIMARY BILE ACIDS (BAs), predominantly cholic acid and chenodeoxycholic acid, are delivered into the intestinal tract as glycine or taurine conjugates. Most BAs are reabsorbed in the ileum as conjugates. However, during each enterohepatic cycle ~10% of the BAs escape into the colon in which they are exhaustively converted by the intestinal flora. Diets high in fat and low in fiber can raise the amount of taurine-conjugated BAs that passes daily through the colon >10-fold (24, 39). This is the consequence of a combined effect of increased BA production and increased taurine conjugation (12, 24). Only 1–3% of the BAs will eventually be excreted in the feces. Fecal BAs are almost completely deconjugated and have undergone various other reactions such as dehydroxylation, (de)hydrogenation, epimerization, etc. (23, 41, 43).

Intestinal epithelial cells are exposed to various concentrations and compositions of BAs throughout the fecal stream. Several effects of BAs on colonic epithelial cells (CECs) have been described, including induction of proliferation and apoptosis (30, 37). Furthermore, epidemiological and animal studies have demonstrated that BAs may be endogenous colon tumor promoters (2, 3, 42).

The mechanism by which BAs induce gene expression is not fully understood. BAs utilize multiple signaling cascades such as p53 (29), the mitogen-activated protein kinases ERK and p38 (28, 30), or PKC (27), phosphatidylinositol 3-kinase (PI3-kinase) (36), and the activator protein-1 (AP-1) transcription factor (28) to modify gene expression.

A critical signaling cascade controlling inducible gene expression is the NF-{kappa}B transcriptional system. Many inducers such as cytokines, growth factors, and bacteria activate the NF-{kappa}B signaling cascade that then triggers transcription of a wide array of proinflammatory and immune system-modulating genes. The main integrator of signal-induced NF-{kappa}B activation is the I{kappa}B kinase (IKK) complex. The subunit IKK{beta} is responsible for I{kappa}B{alpha} serine residues 32 and 36 phosphorylation, a process that marks the protein for ubiquitination and subsequent proteolytic degradation through the proteasome pathway (20). Destruction of the NF-{kappa}B inhibitor releases the transcription factor that migrates to the nucleus, binds to {kappa}B-dependent gene promoters, and induces transcription.

Numerous studies have linked the NF-{kappa}B signaling cascade to important biological processes in the gut, such as carcinogenesis, as well as innate and immune responses. For example, systemic or local delivery of RelA oligonucleotide antisense has been shown to prevent and reverse experimental colitis without apparent damage to the intestinal barrier (25). Also, selective deletion of IKK{beta} in intestinal epithelial cells prevented ischemia-reperfusion-induced TNF secretion and lung injury (6). Therefore, an inducer of NF-{kappa}B signal transduction may impact the state of intestinal homeostasis. Because BAs induce various proinflammatory gene expressions in different cell types, it is important to resolve the molecular mechanism responsible for this effect.

In this study, we characterized molecular mechanisms underlying the BA-induced gene expression in CECs. We report that BAs induce IL-8 gene expression through activation of the transcription factor NF-{kappa}B. It is interesting that deoxycholic acid (DCA) signaled to NF-{kappa}B through proteasome-mediated I{kappa}B degradation, whereas taurodeoxycholic acid (TDCA) mainly enhanced RelA phosphorylation.


    MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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Reagents

DCA and TDCA were obtained as sodium salts (Sigma, St. Louis, MO). Stock solutions of BAs (10 mM) were prepared in water and stored at 4°C. Before starting stimulation experiments, they were incubated in a water bath at 37°C and brought to the final working solution with medium without FCS plus penicillin/streptomycin.

TNF served as control as a strong inducer of NF-{kappa}B activation and was obtained from R&D Systems (Minneapolis, MN). Pharmacological blockade of the proteasome was performed by preincubation with MG132 (Peptides International, Minoh-Shi Osaka, Japan) at a concentration of 10 µg/ml.

Cells and Cell Culture

Isolation of CECs. CECs were isolated as previously described (34). Briefly, intestinal mucosa was stripped from the submucosa within 30 min after bowel resection and rinsed several times with PBS at room temperature. The mucus was removed by rotating the resection two times in 1 mM DTT (Sigma) for 15 min at 37°C. After being washed with PBS to remove DTT, the mucosa was rotated in 2 mM EDTA in Hanks' balanced salt solution without calcium and magnesium (PAA, Linz, Austria) for 10 min at 37°C. The resulting supernatant-containing debris and mainly villus cells was discarded. The remaining mucosa was vortexed in PBS, and the supernatant containing complete crypts and some single cells was collected into a 15-ml tube. Vortexing was repeated until the supernatant was almost clear. To separate crypts from single cells (partially nonepithelial cells), the suspension was allowed to settle down for 3–5 min. These steps were all carried out at room temperature. The sedimented crypts were collected and washed with PBS, and cell viability was assessed by 0.1% trypan blue exclusion and flow cytometry.

Culture of CECs. The 5 x 105 cells were resuspended in 400 µl MEM supplemented with Earle's salts, 5% FCS, 2 mM glutamine, 100 U/ml penicillin, 100 µg/ml streptomycin, 100 µg/ml gentamycin, and 2.5 µg/ml fungizone (JRH Bioscience, Lenexa, KS). Media were purchased from Biochrom (Berlin, Germany), and supplements were obtained from Sigma. Approximately 1 x 106 CECs were seeded into Millicell-CM culture plate inserts (Millipore, Eschborn, Germany) suitable for six-well plates, coated with 0.5 mg/ml collagen A (Biochrom, Berlin, Germany). The medium containing the isolated cells was placed inside the filter inserts. The cells were rapidly forced to contact the collagen coating of the membranes as the medium passed the filters. Just enough medium was added to cover the cells with a liquid film. The cells were incubated at 37°C in 10% CO2.

HT-29 cells. Human colon carcinoma HT-29 cells were obtained from American Type Culture Collection (Manassas, VA). These cell cultures were maintained in Dulbecco's high glucose medium supplemented with 10% heat-inactivated FCS and 5% penicillin/streptomycin at 37°C in a humidified atmosphere of 5% CO2 in air.

Cell stimulation. Before stimulation in individual experiments, cells were starved for 24 h to avoid the confounding variable of serum-induced signaling. TNF at a concentration of 10 ng/ml served as control.

RNA Isolation and IL-8 mRNA Quantification

RNA was isolated by using the RNeasy Mini Kit including a RNase-free DNase step following the manufacturer's instructions (Qiagen, Hilden, Germany). RNA amounts were analyzed by using a fluorescence microplate reader following the instructions of the RiboGreen RNA Quantitation Reagent and kit (MoBiTec, Göttingen, Germany). Integrity of the RNA was verified by agarose gel electrophoresis, and ribosomal bands were visualized with ethidium bromide staining. First-strand cDNA was synthesized by using 1 µg of total RNA and the AMV-reverse transcription reaction (Promega, Madison, WI) in a total volume of 25 µl utilizing oligo(dT) primers.

To quantify the expression of cDNAs, the real-time PCR LightCycler system (Roche, Mannheim, Germany) was used. For PCR, 1–3 µl cDNA preparation, 0.5–2.4 µl 25 mM MgCl2, 0.5 µM forward and reverse primer, and 2 µl of SYBRgreen LightCycler mix in a total volume of 20 µl were applied.

The following sets of primers were used: {beta}-actin forward, 5'-cta cgt cgc cct gga ctt cga gc-3'; {beta}-actin reverse, 5'-gat gga gcc gcc gat cca cac g-3'; IL-8 forward, 5'-tct gca gct ctg tgt gaa ggt gca gtt-3'; and IL-8 reverse, 5'-aac cct ctg cac cca gtt ttc ct-3'. The following PCR program was performed: 600 s at 95°C (initial denaturation); 20°C/s temperature transition rate up to 95°C for 15 s, 10 s at 58–68°C, 22 s at 72°C, and 10 s at 82°C acquisition mode single, repeated 40 times (amplification). MgCl2 concentration and annealing temperature were optimized. The PCR reaction was evaluated by melting curve analysis following the manufacturer's instructions and checking the PCR products on 1.8% agarose gels. Each quantitative PCR was performed at least in duplicate for two sets of RNA preparations.

IL-8 ELISA

Cells were plated in 24-well plates and serum starved for 24 h before stimulation. Subsequently, supernatants were collected and centrifuged to remove cellular debris, and IL-8 concentration was analyzed by a sandwich ELISA following the instructions (Biosource, Camarillo, CA).

LDH Release

The release of cytosolic lactate dehydrogenase (LDH) was measured to investigate significant membrane disruption using the cytotoxicity detection kit (LDH) (Boehringer-Mannheim, Mannheim, Germany) following the manufacturer's instructions. LDH release is given as the percentage of possible maximum release by induction of total cell lysis with Triton X 2%.

Additionally, cytotoxicity was analyzed morphologically and by trypan blue exclusion dye test, using a phase contrast microscope (Leitz, Wetzlar, Germany)

Nuclear Extracts

Nuclear extracts were prepared as previously described (4). Protease inhibitors in the form of the complete minitablets (Roche, Mannheim, Germany) were used.

EMSA. Nuclear extracts (5 µg) were incubated with a radiolabeled, double-stranded oligonucleotide-containing class I myosin heavy chain {kappa}B binding site (GGCTGGGATTCCCCATCT), separated by electrophoresis, and analyzed by autoradiography as described previously (13). Specificity of the probe was evaluated by incubating the nuclear extracts with an excess (100x) of unlabeled oligonucleotide.

Quantification of activated nuclear NF-{kappa}B concentration. Activated NF-{kappa}B was quantified in nuclear extracts with the ELISA-based kit TransAm from Active Motif (Rixensart, Belgium) according to the manufacturer's instructions as described previously (33). ELISA plates were coated with oligonucleotide (5'-GGGACTTTCC-3') coding for an NF-{kappa}B consensus site. Plates were preincubated with a binding buffer containing DTT and herring sperm DNA. Twenty micrograms of nuclear extracts, solved in 20 µl lysis buffer containing DTT and protease inhibitors, were added per well and incubated at room temperature for 1 h. After completion of a washing step, an anti-RelA antibody was added and incubated for another hour at room temperature. After an additional washing step, a horseradish peroxidase-conjugated secondary antibody was added, followed by an additional hour of incubation at room temperature. After a last washing step, developing solution was added, and the absorption was measured at 450 nm.

RelA Immunofluorescence

Cellular RelA distribution was analyzed by immunofluorescence as described previously (13). Briefly, HT-29 cells grown on chamber slides (Nunc, Naperville, IL) or primary CECs were stimulated for various time intervals (30 min to 3 h) with DCA, TDCA, or TNF-{alpha} (10 ng/ml), briefly washed with PBS, and then fixed with ice-cold 100% methanol for 10 min. After being blocked for 30 min with 10% nonimmune goat serum (NGS; Sigma), cells were incubated with rabbit anti-RelA antibody (Rockland, Gilbertsville, PA) diluted 1:200 in 10% NGS/PBS. Slides were washed three times with PBS before 30-min incubation with rhodamine isothiocyanate-conjugated goat anti-rabbit IgG antibody (Jackson ImmunoResearch, West Grove, PA) diluted 1:100 in 10% NGS/PBS. The RelA localization was visualized with a fluorescence light microscope (Leica, Bensheim, Germany). Double staining with Hoechst dye for detecting nuclear DNA was performed as described previously (35).

Adenoviruses

HT-29 cells were infected overnight with adenoviral dominant-negative (dn) IKK{beta} (Ad5dnIKK{beta}) or Ad5I{kappa}B{alpha}AA in serum-free media (Opti-MEM; GIBCO, Grand Island, NY) at 50 multiplicity of infection. The dnIKK{beta} construct cloned in adenoviral vector consisted of a point mutation in the kinase domain (K44A) as described previously (35) and contained an extra 24-bp DNA nucleotides coding for the FLAG peptide (DYLDDDDL). The Ad5I{kappa}BAA virus has been characterized and described previously (19). Ad5GFP containing green fluorescent protein was used as negative control. After a washing step, fresh medium containing serum without antibiotics was added, and cells were used for subsequent experiments.

Western Blot Analysis

HT-29 cells were stimulated for various times (0–3 h) with DCA or TDCA. The cells were lysed in 1x Laemmli buffer, and 20 µg of protein was subjected to electrophoresis on 10% SDS-polyacryl-amide gels. Anti-phosphoserine I{kappa}B{alpha} (Cell Signaling, Beverly, MA) and anti-phosphoserine RelA (S536; Cell Signaling) antibodies were used to detect immunoreactive phospho-I{kappa}B{alpha} and phospho-RelA, respectively, using enhanced chemiluminescence light-detecting kit (Amersham, Arlinghton Heights, IL) as previously described (11). To proof equal loading, the blots were additionally analyzed for actin expression using anti-actin antibodies from Santa Cruz Biotechnology (Santa Cruz, CA). Furthermore, anti-flag antibodies (Kodak Eastman) were applied to demonstrate efficient gene expression of the FLAG-tagged dnIKK{beta} in HT-29 cells after adenoviral transfection with Ad5dnIKK{beta}.

Statistical Analysis

Statistical significance was evaluated by the two-tailed Student's t-test for paired data. P < 0.05 was considered statistically significant.


    RESULTS
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 MATERIALS AND METHODS
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Effects of DCA and TDCA on IL-8 Secretion

HT-29 cells were stimulated with different concentrations of DCA and TDCA, and IL-8 secretion was analyzed in the supernatant 24 h after stimulation. Both BA induced a dose-dependant increase of IL-8 secretion with maximum induction varying from fivefold for DCA to eightfold for TDCA, respectively (Fig. 1, A and B). Optimal doses inducing IL-8 secretion varied considerably between DCA and TDCA. DCA (300 µM) compared with 2 mM TDCA resulted in maximal IL-8 secretion. However, also at lower concentrations of individual BA, as occurring in the aqueous phase of human feces, effects on IL-8 secretion of HT-29 cells were observed.



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Fig. 1. Effects of deoxycholic acid (DCA) and taurodeoxycholic acid (TDCA) on IL-8 secretion and LDH release. HT-29 cells were starved for 24 h and stimulated with the indicated concentrations of DCA (A and C) and TDCA (B and D) for 16 h, and IL-8 secretion (A and B) and LDH release (C and D) were analyzed in the supernatant. Data represent the means ± SD from 3 separate experiments. *P < 0.05 compared with untreated control. IL-8 secretion is depicted as x-fold stimulation. LDH release is set in relation to levels of 100% cell death induced by Triton X (%).

 
In parallel to IL-8 secretion, LDH release of HT-29 cells into the supernatant was determined to analyze cytotoxic effects of the BA concentrations used (Fig. 1, C and D). Significantly increased IL-8 secretion was observed already at BA concentrations revealing no or only minimal LDH release. It is interesting that a significant increase of IL-8 secretion was found even at BA concentrations inducing a LDH release corresponding to 100% cell death. However, this LDH release did not occur immediately after BA stimulation, but started to increase 4 h after exposure to the BA and reached a maximum after 16 h of stimulation (data not shown).

Time Course of DCA- and TDCA-Induced IL-8 mRNA Expression and IL-8 Secretion

DCA- and TDCA-induced IL-8 secretion was completely blocked by preincubation of HT-29 cells with actinomycin D, indicating that the IL-8 induction was dependent on transcription (data not shown). This was further confirmed by analyzing the effects of DCA and TDCA on IL-8 mRNA expression. HT-29 cells were stimulated with DCA and TDCA for different time intervals, and IL-8 mRNA accumulation was quantified by using real-time PCR. Both BA induced IL-8 mRNA after 1 h of stimulation that peaked at 6 h (Fig. 2, A and B). This induction differed significantly from TNF-induced IL-8 mRNA induction that started earlier, peaked at 3 h, and sharply dropped at 6 h (Fig. 2C).



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Fig. 2. Time course of DCA- and TDCA-induced IL-8 mRNA-expression and IL-8 secretion. HT-29 cells were stimulated with DCA or TDCA. TNF served as control. At the indicated time intervals after stimulation 1) RNA was isolated, and IL-8 mRNA expression was quantified in relation to {beta}-actin expression by real time PCR (AC) and 2) IL-8 secretion was determined compared with unstimulated cells (DF). Data represent the means ± SD from 3 separate experiments. *P < 0.05 compared with untreated control.

 
We then determined the profile of IL-8 secretion in BA-stimulated HT-29 cells. BA induced low amounts of IL-8 secretion after 2 and 4 h but a strong increase (10-fold) after 24 h (Fig. 2, D and E). Again, the time course of TNF-induced IL-8 secretion showed significant differences, revealing a rapid increase of secretion beginning already 2 h after stimulation and reaching a plateau after 8–12 h (Fig. 2F).

Effects of BAs on NF-{kappa}B Signal Transduction

To dissect the molecular mechanism of BA-induced IL-8 gene expression in CECs, we investigated the effect of BA on the activation of the transcription factor NF-{kappa}B. The first step of NF-{kappa}B activation is signal-induced I{kappa}B{alpha} phosphorylation/ degradation and nuclear translocation of the transcription factor. It is interesting that DCA induced I{kappa}B{alpha} serine 32 phosphorylation in HT-29 cells, whereas TDCA revealed only minimal effects (Fig. 3).



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Fig. 3. Effects of DCA and TDCA on the phosphorylation of I{kappa}B{alpha}. HT-29 cells were stimulated for the indicated times with DCA (250 µM) or TDCA (1,500 µM). TNF (5 ng/ml) served as control. Cells were lysed, and immunoreactive phospho-I{kappa}B{alpha} (P-I{kappa}B{alpha}) was detected by Western blot analysis. Actin was analyzed on the same blot to demonstrate equal loading (bottom).

 
This was followed by nuclear RelA translocation as shown by immunofluorescence (Fig. 4). Stimulation with DCA resulted in a RelA translocation beginning at 1–2 h and was most evident after 3 h (Fig. 4C). However, compared with TNF-induced RelA translocation, which served as a positive control (Fig. 4D), nuclear staining occurred later and was less prominent. In contrast, only minimal RelA translocation was observed in TDCA-stimulated cells (data not shown).



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Fig. 4. Effects of DCA on nuclear translocation of RelA. HT-29 cells were stimulated with DCA (250 µM) for 30 min (B) and 3 h (C), and RelA translocation was analyzed by immunofluorescence. As a positive control, cells were stimulated with TNF (10 ng/ml) for 30 min (D). Unstimulated cells served as negative control (A). Double staining with Hoechst dye was performed to reveal nuclear DNA (EH).

 
Evidence of nuclear RelA translocation was confirmed by analyzing the concentration of activated NF-{kappa}B in nuclear extracts using a new ELISA-based technique (Fig. 5, A and B). These experiments revealed a time- and dose-dependent increase of the nuclear concentration of activated NF-{kappa}B in HT-29 cells after stimulation with DCA. NF-{kappa}B activity reached a plateau 3 h after stimulation with DCA and remained elevated for up to 12 h (Fig. 5A). Furthermore, stimulation with DCA resulted in a clear dose-dependent increase of activated nuclear NF-{kappa}B (Fig. 5B) that paralleled IL-8 secretion (Fig. 1).



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Fig. 5. Effects of bile acids on NF-{kappa}B activation. HT-29 cells were stimulated with 250 µM DCA for different time intervals (A) or with different DCA concentrations for a defined time interval (3 h) (B), respectively. Nuclear concentration of activated NF-{kappa}B were analyzed by ELISA. C: HT-29 cells were stimulated with DCA (250 µM) or TDCA (1,500 µM) for different time intervals or with TNF (10 ng/ml) for 30 min, respectively. Nuclear extracts were prepared, and NF-{kappa}B and activator protein-1 (AP-1) binding activities were analyzed by EMSA.

 
In contrast, TDCA failed to significantly enhance NF-{kappa}B activity in HT-29 cells over a wide range of doses (500–5,000 µM) and stimulation intervals (1–12 h), respectively (data not shown).

To further confirm interaction of NF-{kappa}B with gene promoter elements, we analyzed nuclear extracts derived from HT-29 cells stimulated with either DCA or TDCA with EMSA. Weak induction of NF-{kappa}B binding activity was observed 1 h after stimulation with DCA. However, binding activity increased with time, reaching a maximum 3 h after stimulation with DCA in accordance with the time course of RelA translocation. In contrast, NF-{kappa}B DNA binding activity was not significantly induced in TDCA-stimulated cells (Fig. 5C, top). On the other hand, TDCA strongly induced AP-1 DNA binding activity (Fig. 5C, bottom), suggesting a selective effect of this BA on different signal transduction pathways in HT-29 cells.

Functional Role of NF-{kappa}B in BA-Induced IL-8 Gene Expression

We have previously shown that cytokine-induced IL-8 gene expression is mediated by NF-{kappa}B in intestinal epithelial cells (18, 19). To verify the role of NF-{kappa}B in BA-induced IL-8 gene expression, we used MG132, a pharmacological inhibitor of the proteasome. Whereas MG132 completely blocked DCA-induced IL-8 mRNA expression and secretion, only moderate effects were seen with TDCA-stimulated cells (Fig. 6).



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Fig. 6. Effect of MG132 on DCA- and TDCA-induced IL-8 secretion. HT-29 cells were preincubated with the proteasome inhibitor MG132 (10 µg/ml) for 30 min before stimulation with DCA or TDCA. IL-8 secretion into the supernatant was determined by ELISA, 16 h after BA stimulation. Data represent the means ± SD from 3 separate experiments. *P < 0.05 compared with untreated control; #P < 0.05 compared with DCA-stimulated cells.

 
Because TDCA induced IL-8 gene expression without inducing the classical NF-{kappa}B pathway, we then sought to explore the mechanism of action of this BA. We (10, 11) have previously shown that signal-induced RelA phosphorylation plays a critical role in bacteria-induced gene expression in CECs. It is interesting that TDCA induced RelA serine 536 phosphorylation at 30 and 60 min, whereas DCA revealed only minimal effects, and TDCA-induced RelA phosphorylation was blocked by adenoviral delivery of dnIKK{beta} (Fig. 7A). In addition, TDCA-induced IL-8 mRNA expression was blocked in Ad5dnIKK{beta}-infected cells (Fig. 7B) but not in control-infected cells (data not shown). These data suggest that TDCA primarily induced IL-8 gene expression through RelA phosphorylation, whereas DCA utilized mainly signal-induced I{kappa}B degradation and RelA nuclear translocation.



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Fig. 7. TDCA-induced RelA phosphorylation and IL-8 mRNA expression is inhibited by Ad5dnIKK{beta}. HT-29 cells were infected with Ad5dnIKK{beta} or control virus Ad5GFP before stimulation with TDCA (1,500 µM) and DCA (250 µM). A: cells were lysed, and immunoreactive phospho-RelA was detected by Western blot analysis. B: mRNA was isolated, and IL-8 expression was analyzed by PCR.

 
Effects of BAs on primary CECs

To further analyze the physiological impact of BA on proinflammatory gene expression, we investigated primary human colonic CECs ex vivo in a cell-culture model recently established in our laboratory (34). DCA and TDCA induced IL-8 secretion in primary CECs, although induction was lower than seen in HT-29 cells (Fig. 8). Similarly, TNF-induced IL-8 secretion was lower in CECs than in HT-29 cells (twofold compared with tenfold, respectively).



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Fig. 8. Effects of DCA and TDCA on IL-8 secretion of primary colonic epithelial cells (CECs). Primary human CECs were seeded on collagen-coated membranes with 10% serum in culture medium as described in MATERIALS AND METHODS and subsequently stimulated with DCA (250 µM) or TDCA (1,500 µM) for 16 h. IL-8 secretion was determined by ELISA. TNF (10 ng/ml) served as control. Data represent the means ± SD from three separate experiments. *P < 0.05 compared with untreated control.

 
Moreover, DCA induced RelA nuclear translocation in CECs. However, in accordance with the data obtained with the HT-29 cell line, TDCA revealed no effects (Fig. 9).



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Fig. 9. Effects of DCA and TDCA on RelA translocation in primary CECs. Primary CECs were seeded on collagen-coated membranes with 10% serum in culture medium as described in MATERIALS AND METHODS. Subsequently, cells were stimulated with DCA (250 µM) (B) or TDCA (1,500 µM) (C) for 3 h. Immediately after stimulation, cells were quickly trypsinized and attached to glass slides by cytospin, and RelA translocation was analyzed by immunofluorescence. As a positive control, cells were stimulated with TNF (10 ng/ml) for 30 min (D). Unstimulated cells served as negative control (A). Double staining with Hoechst dye was performed to depict nuclear DNA (EH).

 

    DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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The aim of this study was to investigate the molecular mechanisms of BA-induced gene expression in CECs. We could demonstrate that the unconjugated BA DCA and the taurine conjugate TDCA induced expression of the chemokine IL-8 in HT-29 cells. The BA-induced IL-8 mRNA expression and secretion appeared to be time and dose dependent.

Cytokine-mediated IL-8 expression in CECs has been shown to be regulated by the transcription factor NF-{kappa}B (19). Therefore, to dissect the molecular mechanism of BA-induced IL-8 gene expression, we investigated the effects of DCA and TDCA on NF-{kappa}B activation. It is interesting that we found differences between DCA and TDCA in their potential to induce the classical NF-{kappa}B pathway. DCA induced I{kappa}B{alpha} serine 36 phosphorylation, RelA nuclear translocation, and increased nuclear NF-{kappa}B-binding activity, whereas TDCA revealed no significant effects.

RelA translocation has been reported (26) previously in response to DCA in another colon tumor cell line (HCT-116). However, other authors (15) were unable to demonstrate increased NF-{kappa}B binding activity in LoVo adenocarcinoma cells on stimulation with chenodeoxycholic acid. Moreover, a recent study (40) using the rat small intestinal cell line (IEC-6) found NF-{kappa}B activation also in response to TDCA stimulation. It is unclear whether these discrepancies are due to the use of different bile salts, different concentrations of BA, or the different cell lines used. To exclude that the BA effects on IL-8 gene expression and the NF-{kappa}B signal transduction were specific for certain cell lines, we performed key experiments with primary human CECs. As in HT-29 cells, DCA and TDCA were capable of inducing IL-8 secretion in primary CECs, whereas only DCA induced RelA translocation. This confirms that there seems to be a classical NF-{kappa}B-dependent and an independent pathway in BA-induced IL-8 expression in CECs.

To elucidate the mechanism of action of TDCA on gene expression in CECs, we analyzed RelA phosphorylation, because we recently demonstrated that it plays a critical role in bacteria-induced gene expression in CECs (10, 11). It is interesting that TDCA induced RelA serine 536 phosphorylation, and this effect was blocked by molecular inhibition of IKK{beta}. It has been shown previously (10, 17, 44) that NF-{kappa}B can be activated by cytokines or bacterial products through phosphorylation of the RelA/p65 subunit at serine 536 without altering the level of the phosphorylation of I{kappa}B and nuclear localization of RelA. However, this mechanism has not been described for BA so far.

The functional relevance of these different mechanisms of action on the NF-{kappa}B signaling cascade between DCA and TDCA was confirmed by experiments investigating BA-induced IL-8 expression after inhibition of the proteasome or the I{kappa}B kinase IKK{beta}. Although the pharmacological inhibitor MG132 completely blocked DCA-induced IL-8 expression, only moderate effects were seen in cells stimulated with TDCA. However, in contrast, adenoviral delivery of a dominant-negative form of IKK{beta} completely blocked TDCA-induced IL-8 expression. These data suggest that TDCA induces IL-8 gene expression primarily through RelA phosphorylation, whereas DCA utilizes mainly signal-induced I{kappa}B degradation and RelA nuclear translocation.

Currently, we can only speculate on the potential molecular mechanisms causing the differences between DCA and TDCA on NF-{kappa}B signal transduction. It seems likely that they are attributable to the decreased permeability of the cytoplasmatic membrane of the taurine-conjugated BA TDCA compared with the lipophilic, unconjugated BA DCA. In line with this hypothesis, TDCA is capable of inducing classic NF-{kappa}B activation in hepatoma cells (36). In contrast to CECs, these cells express a transporter for taurine-conjugated BA. As an alternative mechanism, recent studies (32, 45) indicate that BA-mediated signal transduction can be mediated through receptors located on the cell membrane such as the EGF receptor. Further studies are required to elucidate what mechanisms cause the different effects between DCA and TDCA on the NF-{kappa}B signaling cascade and whether these mechanisms are specific for CECs.

Constitutive oxidant defense levels are relatively low in human CECs compared with the liver (9). Thus in vivo activation of NF-{kappa}B by DCA, which represents the major fraction of BA in the colon, may be needed to protect CECs from oxidative stress caused by bile salts (7, 22) that would otherwise lead to toxic effects and apoptosis. It has been shown before in a hepatoma cell line that some BA attenuate their inherited cytotoxic effects by activating a PI3-kinase-dependent survival signal that is mediated by PKC{zeta} and NF-{kappa}B (36). Payne et al. (26) previously found that apoptosis induced by DCA is potentiated by inhibition of NF-{kappa}B, and there is further evidence that NF-{kappa}B is involved in the regulation or prevention of apoptosis. However, this seems to be a delicate balance, because an increase in the number of cells with activated NF-{kappa}B in the normal-appearing colonic mucosa of a patient with cancer has been found, leading Payne et al. (26) to suggest the hypothesis of induction of apoptosis-resistant cells in colon carcinogenesis. Furthermore, the increased activation of NF-{kappa}B has been shown to lead to cellular transformation (21), tumorgenicity (14, 38), or a metastatic phenotype (5).

Our results could indicate that, in contrast to TDCA, DCA may prevent its inherited cytotoxicity by simultaneously activating intrinsic cell survival signals in CECs, eventually, however, reducing their resistance to carcinogenesis.

The different effects of DCA and TDCA on the NF-{kappa}B signaling cascade may have further implications for dietary or pharmaceutical strategies to change BA concentrations and composition. Such therapeutic strategies have been designed for treatment or prevention of relapse of irritable bowel disease or for chemo prevention of cancer. A Western-type diet, with its high meat and fat content, will increase the intestinal BA concentration and, by virtue of its high taurine content, will also increase the percentage of taurine-conjugated BA (8, 16). It is estimated that diets rich in taurine-containing food, such as meat and seafood, can raise the amount of taurine-conjugated BA that passes daily through the colon >10-fold (12). However, it is possible to alter the BA concentrations and composition in fecal water through dietary intervention, with food rich in fibers (1, 31). Our results may provide a possible mechanism by which such diets could interfere with signaling pathways differentially affected by individual BA and influence carcinogenesis and inflammation.

In summary, in this study we demonstrated differences between DCA and TDCA in their molecular mechanisms of proinflammatory gene expression in a colonic tumor cell line as well as in primary CECs. The relevance of these in vitro findings under physiological and pathophysiological conditions is currently unknown and has to be further evaluated. Insights into the mechanisms responsible for the differences between BAs in their effects on the NF-{kappa}B signal transduction pathway may improve our understanding of the pathophysiology of intestinal inflammation and carcinogenesis.


    GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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This work was supported by grants from the Deutsche Forschungsgemeinschaft to C. Hellerbrand, H. Herfarth, and G. Rogler (SFB585) and by the Else Kröner Fresenius-Stiftung to C. Hellerbrand and H. Herfarth.


    FOOTNOTES
 

Address for reprint requests and other correspondence: C. Hellerbrand, Dept. of Internal Medicine I, Univ. of Regensburg, D-93042 Regensburg, Germany (E-mail: claus.hellerbrand{at}klinik.uni-regensburg.de).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 

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