Cholic acid accumulation and its diminution by short-chain fatty acids in bifidobacteria

Peter Kurdi1, Hiroshi Tanaka2,{dagger}, Hendrik W. van Veen3,{ddagger}, Kozo Asano4, Fusao Tomita4 and Atsushi Yokota5

1 Northern Advancement Center for Science & Technology, Kita 7 Nishi 2, Kita-ku, Sapporo 060-0807, Japan
2 Snow Brand Milk Products Co., Ltd, 1-1-2, Minamidai, Kawagoe 350-1165, Japan
3 Department of Microbiology, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, 9751 NN Haren, The Netherlands
4 Laboratory of Applied Microbiology, Division of Applied Bioscience, Graduate School of Agriculture, Hokkaido University, Kita 9 Nishi 9, Kita-ku, Sapporo 060-8589, Japan
5 Laboratory of Microbial Resources and Ecology, Division of Applied Bioscience, Graduate School of Agriculture, Hokkaido University, Kita 9 Nishi 9, Kita-ku, Sapporo 060-8589, Japan

Correspondence
Atsushi Yokota
yokota{at}chem.agr.hokudai.ac.jp


   ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Cholic acid (CA) transport was investigated in nine intestinal Bifidobacterium strains. Upon energization with glucose, all of the bifidobacteria accumulated CA. The driving force behind CA accumulation was found to be the transmembrane proton gradient ({Delta}pH, alkaline interior). The levels of accumulated CA generally coincided with the theoretical values, which were calculated by the Henderson–Hasselbalch equation using the measured internal pH values of the bifidobacteria, and a pKa value of 6·4 for CA. These results suggest that the mechanism of CA accumulation is based on the diffusion of a hydrophobic weak acid across the bacterial cell membrane, and its dissociation according to the {Delta}pH value. A mixture of short-chain fatty acids (acetate, propionate and butyrate) at the appropriate colonic concentration (117 mM in total) reduced CA accumulation in Bifidobacterium breve JCM 1192T. These short-chain fatty acids, which are weak acids, reduced the {Delta}pH, thereby decreasing CA accumulation in a dose-dependent manner. The bifidobacteria did not alter or modify the CA molecule. The probiotic potential of CA accumulation in vivo is discussed in relation to human bile acid metabolism.


Abbreviations: BSH, bile salt hydrolase; CA, cholic acid; cFSE, carboxyfluorescein succinimidyl ester; DiSC3(5), 3,3'-dipropylthiadicarbocyanine iodide; SCFA, short-chain fatty acid

{dagger}Present address: Ciphergen Biosystems K.K., Yokohama 240-0005, Japan.

{ddagger}Present address: Department of Pharmacology, University of Cambridge, Tennis Court Road, Cambridge CB2 1PD, UK.


   INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Bile, which is produced by liver cells, is composed mainly of bile salts, and is secreted into the duodenum via the bile duct. Bile salts are glycine and taurine conjugates of bile acids, and act as natural ionic detergents. In the intestine, the bile salts play an essential role in emulsifying lipids, which enables intra-luminal lipolysis and the absorption of lipolytic products by enterocytes. Cholic acid (CA) is one of the most common free bile acids in the intestine, and is produced mostly by the deconjugation of bile salts, such as taurocholic acid and glycocholic acid. Deconjugation is carried out by the bile salt hydrolases (BSHs) of the indigenous members of the genera Bacteroides (Masuda, 1981), Bifidobacterium (Tanaka et al., 1999), Clostridium (Masuda, 1981) and Lactobacillus (Tanaka et al., 1999). The free bile acids are further modified by various intestinal micro-organisms to produce secondary bile acids, such as deoxycholic acid and lithocholic acid (Baron & Hylemon, 1997; Kitahara et al., 2000).

Within the human intestinal microbiota, the lactobacilli and bifidobacteria have attracted much attention with regard to their potential probiotic effects. Although many Lactobacillus and Bifidobacterium species have been associated with various health-promoting and beneficial properties (Ouwehand et al., 2002), their interactions with free bile acids are not well characterized. The growth inhibition of intestinal bacteria by free bile acids has been demonstrated (Binder et al., 1975), but the effects of free bile acids on the physiology of intestinal bacteria have not been elucidated. In our previous reports (Kurdi et al., 2000; Yokota et al., 2000), we showed that Lactococcus lactis actively extrudes CA from the cell in an ATP-dependent manner, whereas various Lactobacillus species from the intestine, dairy products and other environments are capable of accumulating CA when they are energized by glucose. The mechanism underlying CA accumulation seems to be not transporter-mediated, but depends on the diffusion of hydrophobic CA across the bacterial cell membrane according to the transmembrane proton gradient ({Delta}pH, alkaline interior), which is formed upon energization with glucose. These findings led us to investigate the interactions of CA with bifidobacteria in the intestines of infants and healthy adults. In addition, we studied the effects on CA accumulation of short-chain fatty acids (SCFAs), which are normally present in the human large intestine as a mixture of acetate, propionate and butyrate.


   METHODS
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INTRODUCTION
METHODS
RESULTS
DISCUSSION
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Bacterial strains.
The Bifidobacterium strains used in this study (Table 1) were obtained from the Japan Collection of Microorganisms (JCM, Wako, Japan) and Snow Brand Milk Products Co. Ltd (SBT, Kawagoe, Japan). The Eubacterium lentum-like strain c-25 was kindly provided by Professor Dr Hiroshi Oda (Department of Bacteriology, Faculty of Medicine, Kagoshima University, Kagoshima, Japan).


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Table 1. CA transport in Bifidobacterium strains

 
CA transport in Bifidobacterium spp.
CA transport experiment.
The bacteria were grown until mid-exponential phase in half-strength MRS (1/2; MRS) broth (Difco) that was supplemented with 0·25 g L-cysteine/HCl l-1 under anaerobic conditions at 37 °C, using a mixed gas (N2/CO2/H2; 8 : 1 : 1). Preparation of the de-energized, washed cell suspension and the CA transport experiments were carried out essentially as described previously (Kurdi et al., 2000). The cells were harvested, washed with 50 mM potassium phosphate (pH 7·0), 1 mM MgSO4 and 0·1 U horseradish peroxidase ml-1 (Buffer 1), and de-energized with 10 mM of 2-deoxyglucose in Buffer 1 that was supplemented with 1·0 U horseradish peroxidase ml-1 (Buffer 2). The cells were washed three times with Buffer 1 to completely remove the 2-deoxyglucose and resuspended at 3 mg protein ml-1 in 150 mM potassium phosphate (pH 7·0), 1 mM MgSO4 and 1·0 U horseradish peroxidase ml-1 (Buffer 3), to an OD660 value of approximately 10. Buffer 3 was used to prevent substantial extracellular acidification, which would greatly affect the results of the transport experiment. It was confirmed that this buffer allowed a drop of only 0·1 of a pH unit in these experiments. The resulting cell suspension was equilibrated by stirring for 10 min from time 0 in 0·116 mM (16 mCi mmol-1, 592 MBq mmol-1) [carboxyl-14C]CA (Perkin Elmer) at 37 °C under anaerobic conditions (mixed gas). Under these conditions, once the cells were energized by 10 mM (final concentration) glucose, 100 µl samples were mixed with 3 ml of Stop Buffer (Kurdi et al., 2000) and the samples were filtered quickly through 0·45 µm cellulose acetate filters (Schleicher & Schuell). The cells on the filters were immediately washed with 3 ml of Stop Buffer using filtration. The membranes were placed into Eppendorf tubes, and the levels of radioactivity were counted in a scintillation counter after the addition of 1·4 ml of the scintillation cocktail (Emulsifier Scintillator Plus; Perkin Elmer). The control cells (no added glucose) were treated with 2 µM valinomycin and 1 µM nigericin at time 0, to ensure complete de-energization. A sample of the reaction mixture without cells was also measured as the background level, which was subtracted from each of the test readings.

When the effects of the SCFAs were examined, the sodium salts were added to a final concentration of 117 mM, i.e. 66 mM acetate, 26 mM propionate and 25 mM butyrate. These concentrations approximate the respective levels of the acids in the ascending colon (Cummings, 1997). The effect of 39 mM SCFAs with the same component ratio was also investigated, to check the dose response. The SCFA mixtures were added 8 min after energization with glucose; the addition of the SCFA mixtures did not change the pH of the medium.

Calculation of the accumulated CA.
The absolute amount of CA that was associated with the cells was expressed in nmol (mg protein)-1. The CA accumulation factor, which is defined as the ratio of the internal CA concentration to the external CA concentration, was also calculated. Calculation of the internal CA concentration was based on the assumption of an internal cell volume of 3 µl (mg protein)-1 (Poolman et al., 1987). Non-specific binding of CA to the cell surface and/or to the cell membrane was estimated from the positive deviation of the calculated internal CA concentration of the control series (no glucose added; de-energized with ionophores) from the extracellular CA concentration of 0·116 mM. The non-specific binding value estimated in this way was subtracted from the calculated internal CA concentration values in the energized series. Thus, the accumulation factor was obtained from these corrected intracellular CA concentrations, where the accumulation factor for the control series was set at 1·0.

The protein content of the cell suspensions was determined using the DC Protein Assay Kit (Bio-Rad) according to the manufacturer's instructions and BSA as the standard. The cell suspensions were boiled for 5 min in 1 M NaOH and then centrifuged; the resulting supernatants were used in the assays.

Measurement of intracellular pH.
Internal pH measurements were performed as described previously (Kurdi et al., 2000), using the internally conjugated fluorescent pH probe carboxyfluorescein succinimidyl ester (cFSE; Molecular Probes) (Breeuwer et al., 1996). Briefly, the cells were cultured until mid-exponential phase, harvested and washed twice in Buffer 1. The cells were resuspended to an OD660 value of approximately 0·5 in Buffer 3 and incubated at 37 °C for 30 min in the presence of the precursor probe carboxyfluorescein diacetate succinimidyl ester. To eliminate unbound probe, the cells were incubated with glucose for 1 h and then washed once in Buffer 3. The cells were subsequently resuspended in Buffer 3, and the intracellular pH measurements were carried out. The effects of SCFAs on the internal pH were examined by the addition of SCFA mixtures at final total concentrations of 117 or 39 mM.

Transmembrane electrical potential ({Delta}{Psi}) measurements.
Changes in {Delta}{Psi} during energization were monitored using the fluorescent dye 3,3'-dipropylthiadicarbocyanine iodide [DiSC3(5); Molecular Probes], which is a cationic probe that crosses the cell membrane, and the fluorescence of which is quenched as the membrane potential develops (negative interior). The harvested cells were washed twice with ice-cold Buffer 4 (Buffer 1 that contained 65 U catalase ml-1 instead of peroxidase), then resuspended in Buffer 5 (Buffer 3 with 65 U catalase ml-1 in place of peroxidase), to an OD660 value of approximately 10, and stored on ice. The replacement of peroxidase with catalase was important for reproducible measurements of {Delta}{Psi} in the bifidobacteria because (i) peroxidase quenched the fluorescence of the DiSC3(5) probe, even before energization of the cells (see below), while catalase did not have this effect, and (ii) the addition of peroxidase or catalase was critical for bifidobacterial metabolism of glucose under experimental anaerobic conditions. The cells were added to a stirred cuvette that contained Buffer 5 (final OD660 value of 0·05) and DiSC3(5) (final concentration of 0·5 µM). Glucose (10 mM final concentration) was then added under anaerobic conditions (mixed gas was introduced into the cuvette headspace), to energize the cells. Fluorescence measurements were performed with an LS50B fluorimeter (Perkin Elmer) with excitation and emission wavelengths of 651 and 675 nm, respectively (slit widths of 4·0 nm).

CA metabolism by bifidobacteria.
The Bifidobacterium strains were cultured in 3 ml of 1/2; MRS broth, as described in the transport experiment section, while the positive control, E. lentum-like strain c-25, was grown in GAM Broth ‘Nissui’ (Nissui Pharmaceutical). Both of these media contained 0·15 mM sodium cholate; the cultures were incubated for 48 h under anaerobic conditions (mixed gas). The culture broths were acidified with concentrated HCl to pH 2, and the bile acids were extracted with ethyl acetate. The bile acids were separated by TLC using Silica gel 60 (Merck) and cyclohexane/ethyl acetate/acetic acid (7 : 23 : 3, v/v; Eneroth, 1963) as the solvent. The bile acid spots on the TLC plate were visualized by spraying with the colouring reagent 5 % (w/v) phosphomolybdic acid {H3[P(Mo3O10)4].nH2O}, which was dissolved in ethanol, and then heated in an oven at 110 °C for 10 min.


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INTRODUCTION
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CA accumulation in Bifidobacterium cells
The CA levels of Bifidobacterium breve JCM 1192T cells equilibrated between the external medium and the de-energized cells (Fig. 1a), since the hydrophobic protonated CA was able to diffuse freely across the cell membrane (Kamp & Hamilton, 1993). The addition of glucose as a fermentable substrate energized the cells, after which the amount of CA in the cells started to increase, as compared to control cells that did not receive glucose (Fig. 1a). All of the tested bifidobacteria accumulated CA [1·2–3·6 nmol (mg protein)-1], and none of the strains showed detectable CA extrusion when energized with glucose (Table 1). The apparent (i.e. measured) CA accumulation factor, which was defined as the ratio of the internal CA concentration to the external concentration, was between 2·7 and 8·5 (Table 1). These values were similar to those observed for lactobacilli (Kurdi et al., 2000). In addition, we measured the CA transport in B. breve JCM 1192T at 1·0 and 2·0 mM external CA concentrations, and observed CA accumulation factors of about 5 and 4, respectively (data not shown).



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Fig. 1. CA transport (a) follows the kinetics of the internal pH changes (b), but not those of the membrane potential (c) when ionophores are added to energized B. breve JCM 1192T cells. (a) De-energized, washed cells were incubated with 0·116 mM [carboxyl-14C]CA in the absence ({blacksquare}) or in the presence of 10 mM glucose (Glc, {bullet}). Ionophores, valinomycin (Val, {triangleup}) and nigericin (Nig, {triangledown}) were added to the energized cells at final concentrations of 2 and 1 µM, respectively. (b) The cells (OD660~0·5) were pre-loaded with cFSE and energized with 10 mM Glc for approximately 10 min, before the addition of Val or Nig at final concentrations of 5 or 200 nM, respectively. (c) Washed cells were added (OD660~0·05) to the cuvette that contained the DiSC3(5) probe (0·5 µM). The cells were energized with Glc (10 mM) for about 10 min, followed by the addition of Val (5 nM) or Nig (200 nM). The fluorescence intensity is expressed in arbitrary units (a.u.). All the experiments were carried out at an external pH of 7·0. The data shown are representative of at least three experiments that gave similar results.

 
Bioenergetics of CA accumulation
The observation that CA accumulation is an energy-dependent process led us to investigate the contribution of the components of the proton motive force, {Delta}{Psi} and {Delta}pH, to CA accumulation. Therefore, we studied the effects of valinomycin (dissipates {Delta}{Psi}) and nigericin (abolishes {Delta}pH) on CA accumulation in bifidobacteria. The results of these experiments using B. breve JCM 1192T and Bifidobacterium bifidum JCM 1255T (data not shown) revealed that the addition of 2 µM valinomycin increased the amount of accumulated CA (Fig. 1a), as compared to the control energized cells that were not treated with valinomycin. However, the addition of 1 µM nigericin reduced the amount of accumulated CA in the energized cells to the equilibration level (Fig. 1a). These observations indicate that the pH gradient ({Delta}pH) is the driving force behind CA accumulation.

To further confirm the involvement of {Delta}pH in the accumulation process, the internal pH of the JCM 1192T cells was measured with the fluorescence probe cFSE (Fig. 1b). The internal pH started to increase (from pH 7·15 to around pH 7·5) 5 min after energization by glucose. This pH level was maintained until the addition of ionophores. The formation of the {Delta}pH coincided with CA accumulation (Fig. 1a). The addition of valinomycin resulted in an increase in the internal pH, while the addition of nigericin abolished {Delta}pH (Fig. 1b). These changes in internal pH and {Delta}pH may be responsible for the alteration of accumulated CA levels in the JCM 1192T cells upon the addition of ionophores (Fig. 1a).

Monitoring of the changes of {Delta}{Psi} using DiSC3(5) (Fig. 1c) revealed that the development of {Delta}{Psi} upon energization with glucose and CA accumulation occurred simultaneously. However, nigericin addition increased the {Delta}{Psi}, while valinomycin addition totally abolished it. These results clearly demonstrate that the {Delta}pH component and not the {Delta}{Psi} component of the proton motive force is the driving force behind the CA accumulation process.

Measurements of the internal pH values of eight Bifidobacterium strains with cFSE revealed a positive correlation between the CA accumulation factors and the internal pH values of the respective strains (Table 1). The higher the pH gradient (i.e. {Delta}pH), the higher the accumulation factor in most cases, which confirms that {Delta}pH is the driving force behind CA accumulation. The theoretical accumulation factors, which were calculated from the measured internal pH values using the Henderson–Hasselbalch equation (pH=pKa+log[A-]/[HA], where the pKa of CA was 6·4), were lower than the actual accumulation factors (Table 1). It is possible that active CA transporters contributed to the CA accumulation in certain strains (e.g. JCM 1192T) that had large differences between their measured and predicted accumulation factors. However, in strains with smaller differences between their predicted and measured accumulation factors, CA may have accumulated solely as the result of diffusion through the membrane, followed by {Delta}pH-dependent dissociation.

Effect of SCFAs on CA accumulation
Various mixtures of sodium acetate, sodium propionate and sodium butyrate, at final total concentrations of 117 mM (which corresponds to the concentration of these SCFAs in the ascending colon) or 39 mM, were used to test the effect of SCFAs on CA accumulation. CA accumulation in JCM 1192T cells was reduced by at least 50 % in the presence of the 117 mM SCFA mixture (Fig. 2a), as compared to cells that were incubated in the absence of SCFAs, while the 39 mM SCFA mixture produced a less pronounced reduction (~20 %) in CA accumulation. The addition of nigericin further decreased CA accumulation, which suggests that a certain {Delta}pH level was maintained in the JCM 1192T cells in the presence of 117 mM SCFAs. These SCFAs are weak acids with pKa values of 4·75, 4·87 and 4·81 for acetic, propionic and butyric acid, respectively. These weak acids, as is the case with CA, can be accumulated in bacterial cells (Russell, 1991), and can theoretically reduce the internal pH of bacterial cells (Diez-Gonzalez & Russell, 1997). As expected, measurements of the internal pH changes of energized JCM 1192T cells upon the addition of the SCFA mixtures revealed decreases in the internal pH (Fig. 2b, c). The {Delta}pH levels were reduced by about 60 and 22 % by the addition of SCFA mixtures at 117 mM (Fig. 2b) and 39 mM (Fig. 2c), respectively. These reductions correspond to the reductions in the amounts of accumulated CA in JCM 1192T cells following treatment with the SCFA mixtures. These results indicate that in the presence of SCFAs, acidification of the intracellular environment and the subsequent decrease in {Delta}pH reduce CA accumulation.



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Fig. 2. SCFA mixtures impair CA transport and reduce the internal pH of B. breve JCM 1192T. (a) De-energized, washed cells were incubated with 0·116 mM [carboxyl-14C]CA ({blacksquare}), and then energized with 10 mM glucose (Glc) in the absence ({bullet}) or in the presence of the SCFA mixtures. The SCFAs were applied at total concentrations of 117 mM ({blacktriangleup}; acetate 66 mM, propionate 26 mM, butyrate 25 mM) and 39 mM ({triangleup}; with the same component ratio). Nigericin (Nig, {triangledown}) was added at a final concentration of 1 µM. (b, c) The internal pH measurements were performed as described in the legend to Fig. 1. SCFAs at concentrations of 117 and 39 mM were used in (b) and (c), respectively. Nig was added at a final concentration of 200 nM. The magnitude of {Delta}pH is indicated. All of the experiments were conducted at an external pH of 7·0. The data shown are representative of at least three experiments that gave similar results.

 
CA metabolism by bifidobacteria
Although our experiments demonstrate that CA is accumulated spontaneously in energized bifidobacterial cells, the possibility exists that bifidobacteria metabolize CA. To test this hypothesis, bile acids were extracted from whole culture broths, in which bifidobacteria had been incubated with CA for 48 h, and analysed using the TLC method. As shown in Fig. 3, none of the Bifidobacterium strains used in the transport experiments produced deoxycholic acid or any other metabolite of CA, while deoxycholic acid formation was confirmed in the E. lentum-like strain c-25.



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Fig. 3. Bifidobacteria do not metabolize CA. Thin-layer chromatogram of whole-cell extracts from 48 h-old cultures of various Bifidobacterium strains. The strains were cultured in the presence of 0·15 mM sodium cholate. Lanes: 1 and 14, deoxycholic acid (DCA); 2 and 15, CA; 3, B. breve JCM 1192T; 4, B. catenulatum JCM 1194T; 5, B. pseudocatenulatum JCM 1200T; 6, B. longum JCM 1217T; 7, B. infantis JCM 1222T; 8, B. bifidum JCM 1254; 9, B. bifidum JCM 1255T; 10, B. adolescentis JCM 7046; 11, B. longum SBT-2928; 12, Eubacterium lentum-like strain c-25 (positive control); 13, blank (no strain was inoculated into the medium). The results shown are representative of at least three experiments that gave similar results.

 

   DISCUSSION
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To date, there has been little information on the interactions between free bile acids and bifidobacteria, which are considered to be the most important indigenous bacteria in the large intestine, in terms of their abundance and health-promotion properties (Mitsuoka, 2002). Although one might anticipate that intestinal bacteria have active mechanisms for the extrusion of growth inhibitory compounds, such as bile acids, this notion is not consistent with our observations. We found that bifidobacteria do not have apparent bile acid extrusion activities. On the contrary, energized cells accumulate CA (Fig. 1a, Table 1). This activity is very similar to that described in lactobacilli (Kurdi et al., 2000). Bile acid transport was studied in Lactococcus lactis, which can actively export CA via a multi-specific organic anion transporter, which is driven by ATP (Yokota et al., 2000), while Escherichia coli utilizes the energy of the proton motive force to extrude chenodeoxycholic acid and taurocholate (Thanassi et al., 1997). Another intestinal microbe, Clostridium scindens (formerly known as Eubacterium sp. VPI 12708; Kitahara et al., 2000), takes up CA using the proton motive force (Mallonee & Hylemon, 1996) for the dehydroxylation of the CA molecule at the seventh carbon atom (White et al., 1980). A gene from Lactobacillus johnsonii 100-100 was identified that encodes an importer that takes up taurocholate for the intracellular BSH reaction (Elkins & Savage, 1998). Moreover, this strain appeared to contain another gene for a putative conjugated bile acid transporter (Elkins et al., 2001).

As is the case in lactobacilli (Kurdi et al., 2000), the results presented here indicate that the driving force for CA accumulation in bifidobacteria is the {Delta}pH component of the proton motive force, and not the {Delta}{Psi} component (Fig. 1). We hypothesize that CA accumulation in bifidobacteria occurs in the following way. (i) The hydrophobic CA molecules diffuse into the cytoplasm of the energized bifidobacterial cell and dissociate according to the ratio given by the Henderson–Hasselbalch equation under the given cytoplasmic pH value. (ii) Since the cytoplasmic pH value is higher than that outside (i.e. {Delta}pH), more dissociation takes place in the cytoplasm than in the external medium. The resulting negatively charged cholate anions cannot pass the membrane due to their polarity, and thus they are trapped inside the cell. (iii) CA influx continues until the concentration of the protonated CA molecule equilibrates on both sides of the cell membrane. In this situation, the total amount of CA (the sum of the protonated and dissociated species) is higher in the cytoplasm than in the external medium. (iv) The intracellular concentration of CA remains higher than that of the external environment, until the cells are energized and the CA concentration gradient across the membrane disappears, along with the disappearance of {Delta}pH. It is worth mentioning that substantial discrepancies were observed in some strains between the values for the measured CA accumulation factors and the values that were predicted by the above mechanism, i.e. the latter values were lower than the former values. Therefore, we cannot discount the possibility that an active CA uptake mechanism (e.g. driven by {Delta}pH) exists.

SCFA production by intestinal bacteria is a very important process in the large intestine, and provides energy for enterocytes (Cummings & Macfarlane, 1997). One of the major benefits of the fermentation of prebiotics is that it yields SCFAs (Cummings et al., 2001), which in turn decrease the intestinal pH. CA accumulation was impaired when mixtures of SCFAs (117 or 39 mM) were added to the experimental reaction mixture (Fig. 2), although the residual amounts of accumulated CA in these cells were clearly higher than those found in the non-energized control cells. The presence of SCFAs at the physiological concentrations found in the colon appears to exert a severe environmental stress, from the bioenergetical standpoint, on bifidobacterial cells. Even in the presence of SCFAs, CA accumulation may occur in energized Bifidobacterium cells, and this process may operate in vivo. This hypothesis was strengthened by our results which indicate that B. breve JCM 1192T accumulated CA even when external CA concentrations were 1·0 and 2·0 mM (around its physiological concentrations; estimated from Ewe & Karbach, 1989). Moreover, CA accumulation at 1·0 mM external CA concentration was observed in JCM 1192T cells even in the presence of 117 mM SCFAs by a factor of 2·4 (data not shown).

Our experiments on CA metabolism by bifidobacteria revealed that Bifidobacterium strains were unable to chemically modify the CA molecule (Fig. 3), which is in agreement with a previous report (Takahashi & Morotomi, 1994). This absence of any chemical modification of CA concurs with our hypothetical mechanism for CA accumulation, which suggests that CA accumulation by bifidobacteria results from the co-existence of a membrane {Delta}pH and a weak acid in the same environment. The participation of a putative active CA uptake system appears to be unlikely, since the tested bifidobacteria did not utilize CA. Therefore, CA accumulation appears to be the result of energization.

The conjugated bile acid taurocholic acid is not accumulated in the BSH-negative Lactobacillus salivarius subsp. salicinius strain JCM 1044 (Kurdi et al., 2000) due to its hydrophilicity (pKa=1·4). Thus, only unconjugated free bile acids are accumulated in lactobacilli and bifidobacteria. According to the published distributions of BSH activities, most of the bifidobacterial strains are BSH-positive (Tanaka et al., 1999). Therefore, in bifidobacteria, conjugated bile acids seem to be the source of free bile acids, which are supposed to be formed inside the cells from conjugated bile acids by BSH activities (Tanaka et al., 2000). It is possible that the CA that is formed from taurocholic acid and glycocholic acid is kept inside bifidobacterial cells in the intestine, for as long as the bacteria are energized.

One possible consequence of CA entrapment in bifidobacterial cells would be the decreased formation of deoxycholic acid in the large intestine. The bifidobacteria do not metabolize CA (Fig. 3; Takahashi & Morotomi, 1994), and thus are unable to produce deoxycholic acid following CA accumulation. Deoxycholic acid and lithocholic acid, which are formed via 7{alpha}-dehydroxylation from CA and chenodeoxycholic acid, respectively, by certain intestinal Clostridium and Eubacterium species (Baron & Hylemon, 1997), are cytotoxic and possible tumour promoters (Reddy et al., 1976; Reddy & Watanabe, 1979). Thus, the accumulation of CA may contribute to the decreased occurrence of colon carcinogenesis. Another possible impact of CA accumulation is a decrease in recycled CA during enterohepatic circulation due to the enhanced excretion of CA from the human host via the faeces. Under these conditions, the synthesis of bile acids from blood cholesterol increases, to compensate for the lost amounts of bile acids, thereby decreasing the blood cholesterol level. Although these features appear quite attractive, experimental evidence for the probiotic relevance of CA accumulation in bifidobacterial cells is lacking. Therefore, in vivo experiments that evaluate these possibilities are urgently needed.


   ACKNOWLEDGEMENTS
 
The authors wish to thank Professor Dr W. N. Konings (Department of Microbiology, University of Groningen, The Netherlands) and Dr I. Mierau (NIZO Food Research, Ede, The Netherlands) for encouragement and valuable suggestions during the experiments. This work was partly funded by the Nestlé Science Promotion Committee. The authors thank the Radioisotope Laboratory of the Graduate School of Agriculture, Hokkaido University for CA transport experiments.


   REFERENCES
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Baron, S. F. & Hylemon, P. B. (1997). Biotransformation of bile acids, cholesterol, and steroid hormones. In Gastrointestinal Microbiology, vol. 1, Gastrointestinal Ecosystems and Fermentations, pp. 470–510. Edited by R. I. Mackie & B. A. White. New York: International Thomson Publishing.

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Received 29 March 2003; accepted 28 April 2003.



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