Bile acids mimic oxidative stress induced upregulation of thioredoxin reductase in colon cancer cell lines

Sandra Lechner1, Ulf Müller-Ladner1, Klaus Schlottmann1, Barbara Jung2, Michael McClelland2, Josef Rüschoff3, John Welsh2, Jürgen Schölmerich1 and Frank Kullmann1,1

1 Department of Internal Medicine I, University of Regensburg, D-93042 Regensburg, Germany,
2 Sidney Kimmel Cancer Center, San Diego, CA 92092, USA,
3 Institute of Pathology, Klinikum Kassel, D-34125 Kassel, Germany


    Abstract
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 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Bile acids have been suggested to play an important role in the etiology of colon and gastric cancer after gastrectomy, but the molecular biology of these effects is poorly understood. We evaluated the effect of different bile acids on human gastric and colon carcinoma cells and identified genes by RNA arbitrarily primed PCR for differential display that are modulated following treatment with hydrophobic bile acids. Thioredoxin reductase (TR) mRNA was upregulated after treatment with taurochenodeoxycholic acid (TCDCA) in St 23132 cells. This raised the question whether deoxycholic acid (DCA) would have regulative effects on TR in HT-29 cells. After an incubation time of 6 h with DCA, TR mRNA expression was increased up to threefold. Ursodeoxycholic acid had no influence on TR mRNA expression. The upregulation of TR after DCA incubation was almost identical to incubation with 12-O-tetradecanoylphorbol-13-acetate. This implies that hydrophobic bile acids mediate oxidative stress in gastrointestinal cancer cells, which was confirmed by measurement of oxidative burst after treatment with DCA. The results suggest that hydrophobic bile acids induce oxidative stress in gastrointestinal cancer resulting in a compensatory upregulation of TR mRNA, one of the key components in the complex anti-oxidant defense system within eukaryotic cells. The activation of at least parts of the redox signaling system is potentially related to the cytotoxicity and the stimulation of the cell death machinery induced by toxic bile acids.

Abbreviations: CA, cholic acid; DCA, deoxycholic acid; LCA, lithocholic acid; ROS, reactive oxygen species; TCDCA, taurochenodeoxycholic acid; TPA, 12-O-tetradecanoylphorbol-13-acetate; TR, thioredoxin reductase; UDCA, ursodeoxycholic acid


    Introduction
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Bile acids have been suggested to play an important role in the etiology of colon cancer (1) and gastric cancer after gastrectomy (2,3), which is supported by epidemiological data, mutagenetic studies, signaling studies and investigations on the effect of bile salts in carcinogen treated rats (4). However, the molecular biology of all these effects is poorly understood. Therefore, the identification of key crucial transcriptional interaction sites of different bile acids might provide substantial information on the as yet unexplained impact of bile acids on tumor promotion in the gastrointestinal tract.

Different bile acids have different, complex effects on tumor growth in experimental animal models. There are bile acids with tumor-promoting activity (5) whereas others showed cytoprotective effects (6). For example, studies on colon tumor have demonstrated that cholic acid (CA) increases tumor incidence in azoxymethane-treated rats (7). In contrast, the tertiary and less hydrophobic bile acid ursodeoxycholic acid (UDCA) had no effect by itself, but it decreases tumor incidence in combination with CA in azoxymethane-treated rats (7). Similar findings were obtained by Narisawa et al. (6) showing that in N-methylnitrosourea-treated F344 rats the continuous feeding of a small dose of UDCA may prevent colon carcinogenesis. Furthermore, it has been found that UDCA is able to reduce the number of aberrant crypt foci in azoxymethane-treated rats (8), while CA treatment increased proliferation of aberrant crypt foci but not of normal crypts (9). The co-carcinogenic effect of bile acids in colon carcinogenesis was also shown in germ-free rats (10).

Shekels et al. (11) have elucidated that the cytotoxicity of bile acids correlates with their relative hydrophobicity. The hydrophobic, secondary bile acids deoxycholic acid (DCA) and lithocholic acid (LCA) are supposed to be cytotoxic for normal colonic crypt cells, resulting in an increased compensatory proliferation of colonic epithelial cells, which is associated with an increased risk of colon cancer (12). These findings support the hypothetical role of DCA as a promoter of colon cancer (13). The higher proliferation rate of colonic mucosal cells could be due to the fact that DCA is able to increase the frequency of mitotic events (14).

To elucidate the complex role of bile acids in tumorigenesis, the identification of tumor-promoting or tumor-inhibiting genes which are up- or downregulated when different bile acids are applied is essential. In support of such an idea, recent work has already suggested that bile acids can regulate protein kinase C expression (15), HLA class I gene expression (16) or transcription factors such as AP-1 (17) or estrogen responsive ring finger protein (efp) (18). Various strategies have been developed to examine differential gene expression (19–22). RNA arbitrarily primed PCR (RAP-PCR) for differential display has been proven to be efficient and reliable for numerous experimental settings (23–26). The purpose of the study was to analyze differential gene expression by RAP-PCR in gastrointestinal cancer cell lines restricted to bile acid treatment and characterize the functional property of the dysregulated gene in this context.


    Methods
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 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Experimental design
Primarily we started with the effects of the primary bile acid taurochenodeoxycholic acid (TCDCA) on the gastric cancer cell line St 23132 to stimulate events that might occur during gastric reflux. The experimental design is shown in detail in Figure 1Go. In total, eight different cell populations were used which reflects in general a chronic incubation with TCDCA (100 µM) over a time period of 14 days or an incubation with a higher concentration of TCDCA (500 µM) over a short time period (48 h).



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Fig. 1. Experimental design of TCDCA incubation in St 23132 (gastric cancer) cells.

 
Materials
12-O-tetradecanoylphorbol-13-acetate (TPA), DCA, DMSO, TCDCA were purchased from Sigma (St Louis, MO). UDCA was obtained from Calbiochem (Bad Soden, Germany). The sodium salt of bile acids was dissolved in phosphate-buffered saline (PBS, PAA Laboratories GmbH, Linz, Austria), while a stock solution of TPA (1 mg/ml) prepared in DMSO was diluted with acetone.

Cell culture
Cell lines were obtained from DSMZ (Deutsche Sammlung für Mikroorganismen und Zellkulturen GmbH, Braunschweig, Germany). The gastric cancer cell line St 23132 was grown in RPMI 1640 (Gibco BRL, Grand Island, NY), and the colon cancer cell line HT-29 in Dulbecco’s MEM (Biochrom, Berlin, Germany). Media were supplemented with 10% fetal calf serum (FCS, Gibco BRL) and 1% penicillin/streptomycin (PAA Laboratories GmbH). Cells were grown in monolayers at 37°C in 10% CO2. For further investigations cells were treated at 80% confluency with bile acids (100 or 500 µM TCDCA, 50 or 500 µM DCA or UDCA) or DMSO (0.1%), TPA (10, 100 or 1000 ng/ml medium), acetone (0.1%) or without fetal calf serum for 3 or 6 h prior to analysis.

RNA extraction
Total cellular RNA was extracted using the RNeasy spin column purification kit (Qiagen, Hilden, Germany). To remove contaminating genomic DNA, the total RNA was treated with DNase I (0.2 U/µl; Boehringer Mannheim, Germany) for 45 min at 37°C in a Tris–MgCl2 buffer (each 0.01 M, pH 8.0) and in the presence of an RNase-inhibitor (1.2 units/µl; Boehringer Mannheim). After DNase treatment, another set of RNeasy spin columns was used for cleaning the treated RNA. RNA concentrations were measured spectrophotometrically at 260 nm and adjusted, and equal aliquots were then electrophoresed on 1% agarose gels stained with ethidium bromide to compare large and small rRNAs qualitatively and to exclude degradation.

RNA arbitrarily primed PCR
RAP-PCR of total cellular RNA was performed as previously described (27). Three amounts of RNA (500 ng, 250 ng and 100 ng) were used as template. First strand synthesis was carried out in 10 µl reactions (2 U/reaction RNAse inhibitor, 50 mM Tris–HCl, pH 8.3, 50 mM KCl, 4 mM MgCl2, 10 mM dithiothreitol, 0.2 mM dNTPs, 2 µM first strand arbitrary primer, 18.75 U/reaction murine leukemia virus reverse transcriptase, Promega, Madison, WI) and incubated for 60 min at 37°C. Second strand synthesis was performed with 4 U/reaction AmpliTaq Stoffel Fragment (Perkin Elmer, Norwalk, CT), 4 mM MgCl2, 0.2 mm each dNTP, 2 µCi/reaction {alpha}[32P] dCTP, and 4 µM arbitrary second primer and subsequently cycled through 35 low stringency cycles. Three µl of the PCR reaction were mixed with 12 µl of formamide-dye buffer and denatured at 68°C for 15 min. One to two µl of these solutions were loaded onto 8 M urea/6% polyacrylamide sequencing gels. Electrophoresis was performed for 4–6 h at 50 W in 1X Tris–borate EDTA buffer. All three reactions at each starting concentration of RNA were loaded side by side. Gels were then transferred to 3MM Whatman paper, dried under vacuum at 80°C, and directly placed against Kodak BioMaxTM autoradiography film (Kodak, Stuttgart, Germany) at room temperature. Several luminescence labels (autoradiogram markers, Stratagene, San Diego, CA) were attached to the gel to facilitate alignment of the autoradiograms with the gels in case the isolation of one of the fragments was desired. Multiple exposures for a variable time followed, depending on the intensities of interesting bands of the original fingerprint.

Arbitrary primers used were US6 (5'-GTGGTGACAG-3') for first strand and Nuclear 1+ (5'-ACGAAGAAGAG-3'), OPN28 (5'-GCACCAGGGG-3'), Kinase A2+ (5'-GGTGCCTTTGG-3'), OPN24 (5'-AGGGGCACCA-3') for second strand synthesis. One RAP-PCR reaction was performed leaving out the reverse transcriptase as a control of DNA contamination.

Isolation and purification of differentially amplified PCR products
Gel slices carrying the fragment of interest were then excised with a razor blade and placed in 50 µl TE (10 mM Tris–HCl, 1 mM EDTA, pH 8.0). The DNA was eluted by incubating at 65°C for 3 h. After 100-fold dilution with water, 10 µl of the eluates were taken for reamplification of the desired product using the primers of the original fingerprint and the conditions outlined above for 20 cycles. The PCR products were routinely checked by denaturing PAGE running the reamplified product next to the original fingerprint to verify its size and purity.

After a differentially amplified RAP-PCR product is detected, eluted, and reamplified, it is most often contaminated with a mixture of several different products of similar size. To identify the regulated transcripts, we used native polyacrylamide gels to separate the sequences of the reamplified mixture based on SSCP as described previously (28). Once the correct band is identified by this procedure, it is cut from the SSCP gel and reamplified a second time.

Cloning and sequencing
After verifying its correct size and purity on 4% agarose gels, the reamplified products from the SSCP gels were cloned into pCR®-II Topo using the TOPO-TA-Cloning® Kit (Invitrogen, De Schelp, Netherlands). After blue-white screening of the clones, 10 white colonies per desired product and one blue colony were picked and suspended in 50 µl water. Aliquots of these bacterial suspensions were checked by high-stringency PCR for the presence and the correct length of inserts using the T7 and the M13 reverse sequencing primers. Clones with the correct length were subsequently grown overnight in 5 ml LB medium containing 50 µg/ml of ampicillin for plasmid isolation.

Five clones per transcript were sequenced with the Applied Biosystems 373 automatic sequencer using the Perkin Elmer (Norwalk, CT) DNA sequencing kit. The resulting sequences were aligned to the GenBank database using BLASTSearch (29).

Confirmation of differential gene expression by semiquantitative RT-PCR
Semiquantitative RT-PCR was performed using the QuantumRNA module (Ambion, Austin, TX) for confirmation of differential gene expression. 18 S rRNA primers, which amplify a 488 bp fragment, were used in conjunction with TR-specific primers (TR_F 5'-CGATCTGCCCG TTGTGTTTG-3' and TR_B 5'-CAAGTAACGTGGTCTTTCACCAGTG-3'). These were designed encoding a 601 bp fragment of the TR mRNA (bp 32 to bp 633). Relative transcript abundance can be compared following standardization of the co-amplified 18 S rRNA RT-PCR product as an internal standard. PCR conditions were chosen to allow amplification in the exponential phase. For that purpose competing primers for 18 S rRNA amplification of which the 3' end is blocked, so called competimers, were titrated to allow amplification of the abundant 18 S cDNA template in the exponential phase. AMV reverse transcriptase (Gibco BRL, Grand Island, NY) was used for cDNA synthesis and AmpliTaq (Gibco BRL) for amplification. The following cycling conditions were chosen: 94°C 5 min – 25 cycles of 94°C 30 s, 59°C 30 s, 72°C 1 min – 72°C 7 min.

Radioactive PCR products of simultaneously amplified TR and 18 S cDNA templates were analysed on a 4–20% TBE gel (Novex, Frankfurt, Germany). Evaluation was done by phosphor imaging densitometry (Phosphor Imager, Molecular Dynamics, Sunnyvale, CA) and subsequent data analysis was performed using the Ambis software (ImageQuant, Molecular Dynamics).

Western blotting TR
Rabbit polyclonal antisera were raised against one unique internal peptide sequence (MNGPEDLPKSYDYD) of TR (accession number Q16881), encompassing amino acids 1–14. Two rabbits were immunized and antisera were collected two months later. Peptide synthesis and immunization was performed by Pineda (Berlin, Germany). Each animal produced antibodies that recognized the peptide.

Cells were lysed with ice cold modified RIPA buffer containing a completeTM mini protease inhibitor cocktail tablet (Boehringer, Mannheim, Germany). Total cell extracts were centrifuged (13 000 g) for 30 min at 4°C. Protein concentrations were measured with the bicinchoninic acid assay from Sigma (Deisenhofen, Germany), and 10 µg whole cell protein was mixed with 5x Laemmli sample buffer. Proteins were denatured by heating to 95°C for 10 min and separated on a 10% SDS-Polyacrylamide-Minigel (SDS-Page). After electrophoresis, transfer of proteins to nitrocellulose membranes (0.2 µm, Novex, San Diego, CA) was performed in a NovexXCell IITM blotting apparatus (San Diego, CA) for 1.5 h at 25 V. The membranes were blocked with 5% milk powder in 1x TTBS (1x TBS + 0.05% Tween) overnight. Afterwards the membranes were incubated for 2 h with TR antiserum (1:10 000) in 1x TBS containing 3% dry milk powder. After washing three times with 1x TBS, the secondary antibody (goat-anti-rabbit-HRP-conjugate, Bio-Rad, Munich, Germany) in 1x TBS containing 3% milk powder was applied. Following a 1 h incubation, the membrane was washed several times in 1x TBS for 1 h, then detection was performed using the NOWA chemiluminescence detection system (EnerGene, Regensburg, Germany).

Flow cytometric analysis
After treatment with bile acids (50 or 500 µM DCA/UDCA for 6 h) HT-29 cells (5 x 106) were incubated with 1 µM dihydrorhodamine 123 (DHR 123, Molecular Probes, Leiden, The Netherlands) at 37°C for 15 min. For positive control cells were stimulated with 0.1 µM TPA at 37°C for another 15 min. After harvesting cells were washed twice with HBSS (Hank’s balanced salt solution supplemented with 10 mM Hepes, Sigma). In the presence of reactive oxygen species (ROS) such as O2–, H2O2 or OH-, DHR 123 turned into the green fluorescent rhodamine 123, which was measured at 515–535 nm with a Coulter Epics XL-MCL flow cytometer (Coulter Electronics, Krefeld, Germany). Data analysis was done using the SystemTM II software (Coulter Electronics, Krefeld). Per experiment 10 000 cells were analyzed (30,31).


    Results
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Differential gene expression upon treatment of gastric cancer cells with TCDCA
The regulative effects of the bile acid TCDCA on gastric carcinoma cells (St 23132) were analyzed demonstrated by applying RAP-PCR for differential display. A couple of fingerprints, obtained with different primer combinations, were performed and analyzed. The autoradiography revealed 47 transcripts which were found only after treatment with 500 µM TCDCA for 48 h. Fourteen differentially expressed transcripts were observed after long-term incubation with 100 µM TCDCA, of which 11 transcripts were upregulated after incubation with 100 µM TCDCA for 14 days as well as after incubation with 500 µM TCDCA for 48 h. First we analyzed three transcripts which were upregulated after 48 h treatment with 500 µM TCDCA (indicated in Figure 2Go).



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Fig. 2. RNA fingerprint following bile acid treatment. RAP-PCR, generated with the arbitrary primers US6– Nuclear 1+; loaded are two RNA concentrations (500 ng, 250 ng). (A) RPMI 14 days; (B) 100 µM TCDCA 14 days; (C) RPMI 28 days, (D) 100 µM TCDCA 14 days – RPMI 14 days; (E) RPMI 26 days – 500 µM TCDCA 2 days; (F) 100 µM TCDCA 14 days – RPMI 12 days 500 µM TCDCA 2 days. {downarrow} F9XVI (thioredoxin-reductase); {downarrow}{downarrow} unknown transcript; {downarrow}{downarrow}{downarrow} glycinamide-ribonucleotide-synthetase.

 
The sequence of the first product aligned almost completely (99%) over 174 bp with the mRNA of the thioredoxin reductase (TR) (gene bank accession number: S79851), the second RAP-PCR product had no sequence homology to the known mRNA sequences, and the third fragment matched almost completely over 187 bp (one mismatch) with the mRNA of the glycinamide ribonucleotide synthetase (gene bank accession number: D32051).

Confirmation of the upregulation of TR by TCDCA in gastric cancer cells
To confirm the effect of TCDCA on TR mRNA expression in St 23132, the cell line was grown in medium containing 500 µM TCDCA for 24 h. This incubation resulted in a 3.9-fold induction of TR mRNA (Figure 3Go). In addition, the cells were treated with 500 µM DCA, 500 µM CA and 0.1% DMSO for 6 h. Figure 3Go shows the strong influence of the hydrophobic bile acid DCA on TR expression while CA treatment resulted only in a slight upregulation of TR mRNA.



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Fig. 3. Semiquantitative RT-PCR (St 23132 cells). Expression of the TR transcript in St 23132 cells after incubation with 500 µM TCDCA for 24 h, 500 µM DCA, 500 µM CA or 0.1% DMSO for 6 h. The data were generated by comparison of the intensity of the TR bands vs. the intensity of the 18 S control band. The quotient TR intensity/18 S intensity as mean ± standard error is shown. Four independent experiments were performed.

 
TR expression in colonic carcinoma cells following treatment with bile acids
Since TR is a key component of the redox signaling system, this leads to the question whether bile acids have regulative effects on TR expression in colonic cancer cells. To study the regulation of the TR expression in colon cancer cells, HT-29 cells were treated with the secondary bile acid DCA, one of the physiologically occurring bile acids of the large intestine, and with UDCA. Cells were incubated with 50 or 500 µM of each bile acid for 3 or 6 h before total cellular RNA was prepared. We observed no significant upregulation of TR mRNA after 3 h incubation of HT-29 cells with 50 or 500 µM DCA. Extension of the incubation time to 6 h revealed a clear increase of TR expression already after treatment with 50 µM and especially with 500 µM DCA (Figure 4A and BGo). The treatment with DCA led not only to increased TR mRNA but also to increased protein levels (Figure 5Go).




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Fig. 4. Semiquantitative RT-PCR after bile acid treatment (HT-29 cells). (A) Expression of the TR transcripts in HT-29 cells after incubation with DMSO 0.1% for 3 and 6 h, DCA 50 µM and 500 µM for 3 and 6 h, UDCA 50 µM and 500 µM for 3 and 6 h, acetone 0.1% for 6 h, TPA 10, 100 and 1000 ng/ml for 6 h and culture in serum-free medium for 24 h. The data were generated by comparison of the intensity of the TR bands vs. the intensity of the 18 S control band. The quotient TR intensity/18 S intensity is shown as mean ± standard error. (B) Quantitative RT-PCR of the TR mRNA expression in HT-29 cells after incubation with different bile acids (DCA, UDCA for 3 and 6 h). The 601 bp fragment corresponds to a part of the TR mRNA, the 488 bp fragment to a part of the 18 S rRNA (internal control). The quotient TR/18 S resulted in the relative expression level of TR mRNA.

 


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Fig. 5. Western blot analysis for TR protein levels in HT-29 cells following treatment with DCA for 6 h. Lane 1: RPMI only; lane 2: 50 µM DCA; lane 3: 100 µM DCA; lane 4: size marker.

 
Treatment with 50 or 500 µM UDCA for 3 or 6 h did not alter significantly the expression level of TR mRNA in comparison to DMSO or acetone (Figure 4A and BGo).

TR expression upon treatment with TPA
In order to answer the question whether an oxidative stress inducing agent like TPA is able to induce TR mRNA expression, we treated HT-29 cells with TPA. After incubating cells with TPA (1, 10 or 100 ng/ml medium) a 2.11-fold increase (TPA 100 ng/ml medium) in TR expression was obvious, similar to the increase after treatment with 500 µM DCA for 6 h (Figure 4AGo).

Flow cytometric analysis of the oxidative stress induced by bile acids
It is discussed that bile acid treatment, especially treatment with the cytotoxic bile acid DCA, results in oxidative stress for cells. Using the DHR assay for determining the cellular redox state we could show an elevated amount of ROS after treatment with DCA. The level of the increase is assessed by rhodamine 123 fluorescence (Figure 6Go).



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Fig. 6. Flow cytometry analysis of oxidative burst with the fluorescent dye rhodamine 123 in HT-29 cells (10 000 cells). Cells above the horizontal line are regarded as having oxidative burst. (A) Cells cultured in normal medium. (B) Cells in 500 µM DCA-supplemented medium, cultured for 6 h. (C) Cells in 500 µM DCA-supplemented medium, cultured for 6 h, followed by a 15 min culturing in 0.1 µM TPA-supplemented medium.

 
DCA in combination with TPA in comparison to TPA treatment alone leads to a significantly higher fluorescence rate indicating more reactive oxygen species (Figure 7AGo). In contrast, UDCA was not able to cause oxidative stress (Figure 7BGo), which correlates with the TR mRNA expression analysis.




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Fig. 7. (A) Quantification of oxidative burst with the fluorescent dye rhodamine 123 in HT-29 cells after treatment with TPA 0.1 µM for 15 min, 50 or 500 µM DCA for 6 h with or without additional TPA treatment (0.1 µM for 15 min). The difference of treated/non-treated cells as mean ± standard error is shown. Four independent experiments were performed. *P < 0.01 using Mann–Whitney U test. (B) Quantification of oxidative burst with the fluorescent dye rhodamine 123 in HT-29 cells after treatment with TPA 0.1 µM for 15 min, 50 or 500 µM UDCA for 6 h with or without additional TPA treatment (0.1 µM for 15 min). The difference of treated/non-treated cells as mean ± standard error is shown. Four independent experiments were performed.

 

    Discussion
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 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
There is convincing evidence to suggest that bile acids, which are physiologically present in the gastrointestinal tract, have cytotoxic and neoplastic effects. The mechanisms responsible for these properties after contact of intestinal epithelial cells with bile acids are not clear. In the present study we examined differences in gene expression in carcinoma cells after treatment with bile acids. Using RAP-PCR for differential display following treating gastric carcinoma cells (St 23132) with TCDCA, we found a frequent higher proportion of differentially expressed transcripts in cells incubated with 500 µM TCDCA for 48 h. This suggests that differential gene expression in gastric cancer cells is present predominantly at high concentrations of TCDCA. Nevertheless, following gastric surgery and loss of the pyloric sphincter, soluble bile salt concentrations (2.5 to 32 mM) exceed that of normally occurring concentration of bile acids in stomach contents (0.05 mM to 0.17 mM) (32). Therefore, the concentrations of TCDCA, which we had chosen in our experimental design, were physiological at least under these circumstances.

Of the differentially expressed fragments, we were able to isolate an upregulated transcript which was identified as thioredoxin reductase (TR). In the following experiments we demonstrated that the hydrophobic bile acid DCA, a secondary bile acid, induced upregulation of TR mRNA in colon cancer cells (HT-29), while the more hydrophilic bile acid UDCA had no effect on TR mRNA expression.

The thioredoxin reductases (TRs) are enzymes belonging to the flavoprotein family of pyridine nucleotide–disulphide oxidoreductases that includes lipoamide dehydrogenase, glutathione reductase and mercuric ion reductase (33). TR catalyzes the NADPH-dependent reduction of thioredoxins (Trxs), a group of small (10–12 kDa), and widely distributed redox active peptides which have a conserved -Trp-Cys-Gly-Pro-Cys-Lys- catalytic site that undergoes reversible oxidation and reduction of the two Cys residues (34,35). The thioredoxin system plays an important role in maintaining the reducing intracellular milieu. It functions as electron donor for ribonucleotide reductase and is involved in other cellular events such as secretion, growth promotion (35–37), regulation of the transcription factors NF-{Phi}B and glucocorticoid receptor (38,39). The TR system can protect cells against oxidative stress (40), scavenge free radicals (41), and reduce H2O2 (42). As published previously, Trx has been shown to play an important role in regulating cancer cell growth and inhibiting apoptosis (43,44). TR and Trx comprise an ubiquitous system present in both pro- and eukaryotic organisms (45). Until today, two different human TRs have been cloned: a cytosolic enzyme (TR1) (46) and a mitochondrial enzyme (TR2) (47).

Our findings indicate that the taurin-conjugated form of the primary bile acid chenodeoxycholic acid and the secondary unconjugated bile acid DCA induce the cytosolic form of TR in gastric cancer cells as well as in colon cancer cells. Thus, it can be concluded that the expression level of TR depends on the hydrophobicity, concentration and conjugation status of the bile acid, and of course on the incubation time. Of interest, there was no difference in TR upregulation after 6 h treatment with DCA in gastric or colon cancer cells.

More recently, it has been shown that free oxygen radicals may be involved in the pathogenesis of bile acid hepatotoxicity. Treatment of freshly isolated rat hepatocytes with individual bile acids (100 to 200 µM for 4 h) decreased hepatocyte viability to 40% (48) or in our study with HT-29 cells after treatment with 500 µM DCA to 36% viable cells (data not shown). The decrease of cell viability could be reduced by preincubation with different antioxidants (48). Our results suggest that the secondary bile acid DCA is able to induce oxidative stress in colon cancer cells resulting in an upregulation of TR mRNA. Since TPA treatment resulted in oxidative stress, reflected by numerous biochemical responses, including significant generation of H2O2 and enhanced levels of myeloperoxidase, oxidized glutathione reductase activities, decrease in glutathione levels and superoxide dismutase activity (49), we analyzed the effects of TPA on the TR mRNA level. The expression level was almost identical to this of cells treated with 500 µM DCA for 6 h. Furthermore, bile acids are thought to cause oxidative damage by stimulating the generation of free oxygen radicals from mitochondria (50). Taking these findings into consideration, our results of the bile acid-induced elevated TR expression suggest that bile acids might induce the production of reactive oxygen species. To test this hypothesis, we measured the production of ROS in bile acid-treated HT-29 cells using DHR 123 and flow cytometry. These analyses showed that DCA treatment was able to induce ROS production and to increase the level of TPA induced ROS by simultaneous incubation. ROS are thought to contribute to mutagenesis, carcinogenesis and tumor promotion (51). It is of interest that only the cytotoxic, tumor-promoting bile acid DCA is able to induce the accumulation of ROS while the cytoprotective bile acid UDCA is not (Figure 7BGo). This is consistent with our results concerning the TR mRNA expression level: UDCA was not able to increase the TR transcript level (1.3-fold increase). The level of TR upregulation (three-fold) after DCA treatment (500 µM, 6 h) does correspond to the amplitude of oxidative stress (2.6-fold). Taken together, it could be shown that DCA is able to induce TR expression as well as oxidative stress, while UDCA does not alter TR expression nor induce oxidative stress.

Our data contribute to the finding of Payne et al. (52) concerning the activation of the two stress response proteins NF-{kappa}B and poly(ADP)ribose polymerase (PARP) by DCA treatment.

Our results suggest that hydrophobic bile acids induce oxidative stress in gastrointestinal cancer resulting in a compensatory upregulation of thioredoxin reductase mRNA, one of the key components in the complex anti-oxidant defense system of eukaryotic cells. The activation of at least parts of the redox signaling system is potentially related to the cytotoxicity and the stimulation of the cell death machinery induced by toxic bile acids.


    Notes
 
1 To whom correspondence should be addressed Email: frank.kullmann{at}klinik.uni-regensburg.de Back


    Acknowledgments
 
The authors thank Professor Alan F.Hofmann, University of California, San Diego, for helpful discussion and reading the manuscript. The work was supported by German Research Society grant (DFG no. Ku 1024/6-1, Mu 1383/1-3).


    References
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 

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Received March 5, 2002; revised March 27, 2002; accepted April 23, 2002.