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Bifidobacteria Isolated From Infants and Cultured on Human Milk Oligosaccharides Affect Intestinal Epithelial Function

Chichlowski, Maciej; De Lartigue, Guillaume; German, J. Bruce; Raybould, Helen E.; Mills, David A.

Journal of Pediatric Gastroenterology and Nutrition: September 2012 - Volume 55 - Issue 3 - p 321–327
doi: 10.1097/MPG.0b013e31824fb899
Hepatology and Nutrition

Objectives: Human milk oligosaccharides (HMOs) are the third most abundant component of breast milk. Our laboratory has previously revealed gene clusters specifically linked to HMO metabolism in selected bifidobacteria isolated from fecal samples of infants. Our objective was to test the hypothesis that growth of selected bifidobacteria on HMO stimulates the intestinal epithelium.

Methods: Caco-2 and HT-29 cells were incubated with lactose (LAC)- or HMO-grown Bifidobacterium longum subsp infantis (B infantis) or B bifidum. Bacterial adhesion and translocation were measured by real-time quantitative polymerase chain reaction. Expression of pro- and anti-inflammatory cytokines and tight junction proteins was analyzed by real-time reverse transcriptase. Distribution of tight junction proteins was measured using immunofluorescent microscopy.

Results: We showed that HMO-grown B infantis had a significantly higher rate of adhesion to HT-29 cells compared with B bifidum. B infantis also induced expression of a cell membrane glycoprotein, P-selectin glycoprotein ligand-1. Both B infantis and B bifidum grown on HMO caused less occludin relocalization and higher expression of anti-inflammatory cytokine, interleukin-10 compared with LAC-grown bacteria in Caco-2 cells. B bifidum grown on HMO showed higher expression of junctional adhesion molecule and occludin in Caco-2 cells and HT-29 cells. There were no significant differences between LAC or HMO treatments in bacterial translocation.

Conclusions: The study provides evidence for the specific relation between HMO-grown bifidobacteria and intestinal epithelial cells. To our knowledge, this is the first study describing HMO-induced changes in the bifidobacteria–intestinal cells interaction.

Supplemental Digital Content is available in the text

Foods for Health Institute, University of California, Davis, CA.

Address correspondence and reprint requests to David A. Mills, PhD, University of California, One Shields Ave, Davis, CA 95616 (e-mail:

Received 15 July, 2011

Accepted 30 December, 2011

The study was supported by award no. F32AT006642 from the National Center for Complementary & Alternative Medicine, the University of California Discovery Grant Program, the California Dairy Research Foundation, USDA NRI-CSREES award 2008-35200-18776, NIEHS Superfund P42 ES02710, and by NIH awards R01HD059127, R01HD061923, and R21AT006180.

Supplemental digital content is available for this article. Direct URL citations appear in the printed text and are provided in the HTML and PDF versions of this article on the journal's Website (

The authors report no conflicts of interest.

Breast-feeding is associated with multiple benefits in infants (1). There are well-documented differences in the intestinal microbiota between human milk–fed and formula-fed infants (2). The predominance of beneficial bacteria, mainly bifidobacteria, in the gut microbiota of breast-fed infants is thought to result in part from the fermentation of oligosaccharides, nondigestible carbohydrates consisting of several types of linked monosaccharides (3). Human milk oligosaccharides (HMOs) are the third most abundant component of human milk (4) and comprise >200 HMO structural species (5,6). Our research group has previously demonstrated that HMOs can selectively nourish a protective bifidobacterial microbiota isolated from fecal samples of breast-fed infants (7–10). Using genomic analysis of Bifidobacterium longum subsp infantis (B infantis), our laboratory has revealed gene clusters specifically linked to oligosaccharide metabolism that are expressed only during growth on HMO but not during growth on lactose (LAC) or commercial prebiotics (fructooligosaccharides or galactooligosaccharides) (11–14). It was shown that other bifidobacteria (eg, B bifidum, B longum subsp longum, and B breve) also grow on HMO as the sole carbon source (10); however, other mechanisms of HMO metabolism were proposed (12,15).

The ability to adhere to the intestinal epithelium may play an important role in gut colonization because it prevents the peristaltic elimination of bacteria (16,17). Adhesion promotes the modulation of the immune system (18,19) and prevents pathogens from attaching to the gut mucosa (20). Garrido et al (13) characterized a family of solute binding proteins in B infantis, which have an affinity for mammalian glycans. Strain- and species-specific adhesion of bifidobacteria to epithelial cells is not a new phenomenon and has been described elsewhere (21–23); however, previous reports described minimal invasion of bifidobacteria, suggesting that bifidobacteria do not translocate (24).

Intestinal barrier function requires tight junctional (TJ) complexes, which are made up of complex lipoprotein structures (25); disruption of TJs leads to compromised intestinal integrity (26–30). Ewaschuk et al (31) demonstrated that soluble factors produced by B infantis increase and expression of occludin, a primary TJ protein in the human intestinal epithelial cell line, suggesting improved barrier function (31,32). There are numerous examples in the literature showing the role of probiotics in TJ repair and maintenance (33–35); these effects seem to be mediated by upregulation of TJ proteins.

In addition, previous reports suggest that bifidobacteria showing higher adhesion to intestinal epithelial cells also had a higher anti-inflammatory capacity (36). For example, B infantis induced intestinal production of anti-inflammatory cytokine interleukin (IL)-10 while reducing production of proinflammatory cytokine tumor necrosis factor (TNF) (37,38). Lipopolysaccharide (LPS) induces exaggerated activation of nearly all proinflammatory cytokines, chemokines, immune receptors, and cell surface adhesion molecules (39). Preising et al (36) showed that bifidobacteria were able to inhibit LPS-induced secretion of IL-8, an early mediator of inflammatory responses, in Caco-2 cells. Those authors also reported variations between 2 epithelial cell lines, Caco-2 and HT-29, in this response to bifidobacteria.

Stimuli such as cytokines have been reported to influence not only the expression of TJ proteins but also their association with cytoskeleton (rearrangement) or paracellular permeability (40). Whether other genes (besides TJs) involved in bacteria–host interactions are upregulated in epithelial cells upon contact with bifidobacteria remains to be elucidated. An example of a possible gene candidate is the α1-proteinase inhibitor (SERPING), which is synthesized by the intestinal epithelial cells, and is likely to act as an immunomodulatory factor (41). Also, P-selectin glycoprotein ligand-1 (SELPLG) is a glycoprotein that binds to the cell adhesion molecule and plays a key role in the inflammatory response (42). Microarray studies in our laboratory have previously shown that incubation of HMO with B infantis activated the transcriptional upregulation of SELPLG (43).

The present study focused on characterizing the interaction between HMO-grown bifidobacteria and cultured human intestinal epithelial cells. We hypothesized that growth on HMO would facilitate colonization and that would lead to the protected modulation of the host, including altered gene expression. The effects of HMO on the bifidobacterial–intestinal epithelium interaction are presently not known. Here, the analyzed interaction included adhesion, translocation, production of inflammatory cytokines, and expression and distribution of TJ proteins. To our knowledge, this is the first study describing HMO-induced changes in the bifidobacteria–intestinal epithelial cells interaction.

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Bacterial Strains and Culture Conditions

The bifidobacterial strains used in the present study were B bifidum American Type Culture Collection (ATCC) 29521 (B bifidum) and B infantis ATCC 15697. Cultures from stocks frozen at −80°C in glycerol were grown overnight anaerobically at 37°C in the semisynthetic de Man-Rogosa-Sharpe broth (Becton Dickinson, Franklin Lakes, NJ) supplemented with 1% (wt/vol) L-cysteine hydrochloride (44). This medium was supplemented with 2% (wt/vol) LAC (Sigma Aldrich, St Louis, MO) or HMO as the sole carbon source. Media was autoclaved separately from the carbohydrates, which was added to the cooled broth after sterile filtration (Millipore Corp, Bedford, MA). All of the bacterial cultures used in the adhesion experiment were grown for 48 hours (stationary growth).

HMO used in the present study was provided by the laboratories of Dr Bruce German. The purification process of HMO was performed as described previously (45).

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Epithelial Cell Lines

Enterocyte-like human colon adenocarcinoma (Caco-2) cells show marked characteristics of human intestinal epithelial cells, including the ability to differentiate as well as to polarize and form TJ complexes (46). In the present study, Caco-2 cells were obtained from the ATCC HTB-37, passages 31–37, and were maintained in our laboratory by culture in Dulbecco modified Eagle medium (DMEM) containing 10% heat-inactivated (30 minutes at 56°C) fetal calf serum, 1% nonessential amino acids, 50-IU/mL penicillin, and 50-μg/mL streptomycin. Cells were cultured routinely in 75-cm2 flasks at 37°C in a 5% CO2 constant-humidity environment with medium replaced every 2 to 3 days. Another human colon adenocarcinoma, HT-29 (47) cell culture (ATCC HTB-38, passages 39–44) was maintained in the same conditions except the McCoy 5A medium (Invitrogen, Carlsbad, CA) was used. Monolayers were subcultured at 80% confluence exposing the monolayers to 0.25% trypsin and 0.9 mmol/L ethylenediamintetraacetic acid (EDTA) (Invitrogen) using split ratio of 1:10. Epithelial cell monolayers and Transwells (described below) were treated by LPS (Sigma Aldrich) at 100 ng/mL.

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Adhesion Assay

For adhesion assays, Caco-2 and HT-29 cells were seeded in 24-well plate (2 cm2/well; BD Falcon, Franklin Lakes, NJ). Adhesion experiments were performed 15 days after confluence, a time when morphological and functional differentiation is complete (48,49). The viable cell number, counted in a Neubauer chamber, was about 6 × 105 cells/well. Cell monolayers were carefully washed twice with sterile phosphate buffered saline (PBS; pH 7.3) before bacterial cells were added. B infantis and B bifidum from the exponentially grown 48-hour-old cultures were collected by centrifugation (4000g for 10 minutes), washed, and resuspended in DMEM for assays with Caco-2 cells and in McCoy medium for assays with HT-29 cells. For reference purposes (100% values), 1-mL aliquots of the original bacterial cell suspensions used in the adhesion assay were centrifuged, the cells resuspended in 200 μL trypsin/EDTA plus 200 μL PBS and then frozen and stored at −20°C until quantification of the bacteria. Approximately, 1 × 108 cells of each strain were incubated with a monolayer of fully differentiated cells. All of the incubations were performed in biologically independent triplicates. Plates were then incubated at 37°C, 5% CO2 for 2 hours, after which all of the monolayers were washed gently 3 times with PBS to release unbound bacteria. The epithelial cells were detached from the plastic surface by incubation with 200 μL trypsin/EDTA per well (10 minutes, 37°C). To perform quantification of adherent bacteria, samples of bacteria plus epithelial cells were incubated at 37°C for 30 minutes in Gram-positive lysis buffer consisting of 20 mmol/L Tris-Cl, 2 mmol/L sodium EDTA, 1.2% Triton X-100, and lysosyme (final conc 20 mg/mL). The adhesion capacity was determined using real-time quantitative polymerase chain reaction (qPCR; online-only appendix, and was expressed as the number of adherent bacteria divided by total number of bacteria added, multiplied by 100.

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Bacterial Translocation

For experiments testing translocation of HMO-grown bifidobacteria, Caco-2 or HT-29 cells were seeded on Transwell polycarbonate cell culture inserts (24-mm diameter; 3.0-μm pore size; Corning Inc, Ithaca, NY) at 2 to 3 × 105 cells/cm2 and grown for 14 days postconfluence to achieve fully differentiated monolayers. Translocation studies were conducted in Hanks balanced salt solution, which reduces bacterial growth by approximately 1000-fold compared with DMEM while maintaining bacterial viability (50). This allows a more accurate estimation of the numbers of bacteria crossing the epithelial monolayer. Complete differentiation was confirmed by measurement of transepithelial electrical resistance using a Millicell-ERS voltohmmeter fitted with chopstick electrodes (Millipore Corp) and cells were used when value was >1000 ohm/cm2. Culture medium was removed by washing monolayers twice with Hanks balanced salt solution at 37°C. After equilibration for 30 minutes, any monolayers for which transepithelial electrical resistance has not returned to within 10% of the value before removal of experimental media was discarded. Bifidobacteria were inoculated into the apical chamber of the Transwell to a final concentration of 108 bacteria per milliliter. After a 2-hour incubation period (37°C; 5% CO2, 95% room air), during which translocation of bacteria was allowed to occur, the concentration of bacteria in the basolateral chamber was determined by qPCR (online-only appendix,

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Statistical Analysis

Data are expressed as mean ± standard deviation of the results of 4 independent trials conducted in triplicate. All of the statistical tests used a 2-sided significance level of 0.05. A paired t test was used to compare LAC and HMO experimental groups for each outcome measure. All of the reported significance levels represent 2-tailed P value. Expression of the cytokines and TJs was determined using ΔCt (online-only appendix, Translocation was expressed as 103 bacteria per square centimeter of the monolayer.

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Adhesion of HMO-grown Bifidobacteria in Caco-2 and HT-29 Cells

Numerous studies in our and other laboratories demonstrated the ability of 2 bifidobacterial strains, B infantis and B bifidum, to grow on HMO as a sole carbon source. (10). Based on those results, both B infantis and B bifidum were selected for experiments focused on the interaction between intestinal epithelial cells and bifidobacteria grown on HMO. Adhesion activity of B infantis and B bifidum grown on LAC or HMO was evaluated by a real-time PCR-based method (Fig. 1). In comparison with traditional techniques (eg, viable counts), the analytical approach using real-time PCR is rapid, accurate, and particularly useful for studying bacterial adhesion, especially when dealing with different phenotypes of the genus Bifidobacterium (23). Different adhesion could be observed between 2 bifidobacterial species tested. The levels of adhesion ranged from 8.1% to 20.4% for Caco-2 cells (Fig. 1A) and from 12.5% to 26.5% for HT-29 cells (Fig. 1B). B bifidum grown on LAC was the most efficient strain in terms of adhesion to Caco-2 epithelial cells (20.4%), whereas HMO-grown B infantis incubated with HT-29 cells adhered at the rate of 26.5%. There were numerical trends between HMO and LAC treatments; however, the only significant difference was observed for B infantis incubated with HT-29 cells (P < 0.05). The largest alteration in the binding among bacteria incubated with HT-29 cells was shown with B infantis (8.5% for LAC and 26.5% for HMO) (Fig. 1B).



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HMO-grown Bifidobacteria Alter TJ Expression

Caco-2 and HT-29 cells were incubated with LAC- or HMO-grown bifidobacteria and expression of occludin, zona occludens (ZO)-1, and junctional adhesion molecule (JAM-A) was measured. These particular proteins were selected because they have been implicated in TJ barrier function and are known to be expressed in intestinal cells (51). Caco-2 incubated with B bifidum cultured on HMO showed higher expression of occludin (3.6-fold induction), whereas incubation with both HMO-grown bifidobacterial species resulted in enhanced expression of JAM-A compared with LAC-grown bacteria —these changes were from 5.7-fold induction for B bifidum to 1.7 for B infantis (Fig. 2A). There were no differences in ZO-1 expression between the LAC and HMO treatments in Caco-2 cells. In HT-29 cells, B infantis grown on HMO had 2-fold increase of ZO-1 expression compared with LAC. Occludin expression was not altered in HT-29 cells incubated with HMO-grown bifidobacteria; however, JAM-A was 4-fold higher for HMO-grown B bifidum.



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B bifidum and B infantis Grown on HMO Prevent Occludin Relocation Into the Cytoplasm

We used immunofluorescence assay to evaluate the localization of occludin in epithelial cells incubated with bifidobacteria grown on LAC or HMO. Under normal conditions (no bacterial treatment), occludin was localized in a pattern consistent with their distribution in TJs in Caco-2 cells (supplemental Fig. 1,, whereas cells incubated with HMO-grown bifidobacteria caused mild occludin relocalization to the cytoplasm. LAC treatment caused considerable relocalization of occludin and it was characterized by discontinuities in membrane staining and submembranous internalization of these proteins. Thus, both B infantis and B bifidum grown on HMO prevented the significant intracellular redistribution of occludin. HT-29 cells showed the similar pattern of occludin distribution (data not shown).

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Immunomodulatory and Inflammatory Markers in the Epithelial Cells Are Affected by HMO-grown Bifidobacteria

To determine whether growth of bifidobacteria on HMO induces the immunomodulation in epithelial cells, we analyzed the expression of 2 factors, SELPLG and SERPING. B infantis grown on HMO significantly upregulated the expression of SELPLG (7-fold induction vs LAC) compared with B bifidum (Fig. 3). No significant differences were observed in the expression of SERPING in B infantis or B bifidum. Expression of these 2 immunomodulatory markers was not affected by HMO treatment in HT-29 cells (data not shown). We further assessed the expression of pro- and anti-inflammatory cytokines in Caco-2 and HT-29 cells incubated with bifidobacteria grown on HMO and LAC. The results for expression of the genes for IL-8, IL-10, and TNF are shown in Figure 4. Caco-2 cells incubated with HMO-grown B bifidum and B infantis had significantly increased expression of IL-10 (fold induction vs LAC = 4.6 and 2.5, respectively). HMO-grown B infantis showed increased expression of IL-8, whereas B bifidum did not show induction of IL-8 expression in Caco-2 cells compared with cells incubated with LAC-grown bacteria (Fig. 4A). B infantis grown on HMO lowered the expression of proinflammatory TNF (0.6-fold vs LAC). HMO-grown bifidobacteria had no effect on secretion of measured cytokines in HT-29 cells with values comparable with that of LAC-grown bifidobacteria (Fig. 4B). LPS of Gram-negative bacteria triggers an inflammatory response and the subsequent overexpression of proinflammatory cytokines, for example, IL-8 (52). LPS treatment increased expression of all of the inflammatory cytokines, including 5-fold induction for IL-8 for both Caco-2 and HT-29 cells; however, no significant differences were noted between LAC and HMO treatments (data not shown).





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Translocation of HMO-grown Bifidobacteria in Caco-2 and HT-29 Cells

To examine whether the observed changes in the expression and localization of TJs, as well as immune activation, are associated with alterations in the epithelial barrier, we tested translocation of bifidobacteria. We observed minimal translocation of LAC and HMO-grown bifidobacteria in Caco-2 and HT-29 cells (Fig. 5). No significant difference in translocation was observed between bacteria grown on LAC or HMO in monolayers not treated with LPS. Translocation in HT-29 cells was at the lower rate (0.3–0.45 × 103 cells per square centimeter of the monolayer) compared with Caco-2 cells. No correlation between adhesion and translocation was observed for 2 bifidobacterial species incubated with Caco-2 or HT-29 cells. LPS treatment dramatically increased translocation in both Caco-2 and HT-29 cells; however, there were no significant differences between the bacteria or the sugar source treatments (Fig. 5).



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The present study characterizes for the first time the interactions between HMO-grown bifidobacteria and intestinal epithelial cells. Bacterial adhesion is critical to these interactions because the ability to adhere also represents a significant prerequisite for the transient intestinal colonization (17). HMOs are naturally evolved substrates that provide a structure-specific growth advantage to coevolved bifidobacteria and result in beneficial interactions with the host (10). Unlike LAC, HMOs pass mainly unabsorbed and undigested through the small intestine into the colon, where they are fermented to short-chain fatty acids creating an acidic environment (53). We used LAC as a standard comparative in the present study to exemplify different phenotypes of bifidobacteria consuming those sugars because both LAC and HMO are milk's constituents delivered during lactation. In addition, the core structures of HMOs consist of LAC (5).

The ability of selected bifidobacteria to consume complex oligosaccharides from human milk likely enables this genus to be one of the most abundant colonizers of the breast-fed infant gut (7,8). Previous studies have shown that among 3 gut commensals (B infantis, Lactobacillus gasseri, and Escherichia coli), only B infantis was able to ferment HMO as a sole carbon source and achieve high cell densities (9,54). Although growth of B bifidum on HMO was weaker than B infantis, direct consumption of HMO was observed previously (10). Those 2 species were shown to metabolize HMO via different mechanisms (7,8,10,55). In the present study, both B infantis and B bifidum were selected for analysis of the interaction with the intestinal epithelial cells.

We used 2 epithelial cell lines to test the abilities of HMO-grown B bifidum and B infantis to adhere to the intestinal cells. The first adhesion model is based on the human colon adenocarcinoma cell line Caco-2, whose characteristics simulate structural and functional distinctiveness of mature enterocytes in vitro (46). The second cell line used in the present study, HT-29, maintains a constant proliferation rate with practically no further differentiation (24,56). We used incubation period of 2 hours because it was previously shown sufficient to enable epithelial cells to adapt to the presence of bacteria without acidification of the medium (57).

Although the adhesive phenotype is frequently present in the Bifidobacterium sp (52,58,59), of importance for the present study are the specific changes in adhesion induced by HMO growth. Previous reports demonstrated a strongly adhesive phenotype of B bifidum MIMBb75 (60), which was able to adhere to both Caco-2 and HT-29 cells (23). Our observations support these data; however, unique for the present study, HMO had a significant impact on the binding abilities of tested bifidobacteria. As seen in Figure 1, a high percentage of B infantis grown on HMO adhered to HT-29 epithelial cell lines. The proliferation rate of HT-29 cells is constant and is higher than that in Caco-2 cells (56); thus, the interaction between bifidobacteria and HT-29 cells is expected to be greater than the interaction with Caco-2 cells. Interestingly, HMO-grown B infantis induced expression of SELPLG (annotated as a cell membrane glycoprotein) in Caco-2 cells but not in HT-29 cells. These results suggest a synergistic effect on the gut ability to sense and modulate genes playing a role in the binding and signaling to bacterial gut commensals. Here, we show that HMO-grown B infantis (but not B bifidum) facilitates colonization and leads to a protective modulation of the host's intestinal epithelium.

Ewaschuk et al (31) showed that B infantis–conditioned medium lowered the mucosal permeability, changed expression of TJ proteins, and prevented occludin redistribution into the cytoplasm. Our goal was to test whether growth on HMO affects the permeability of epithelium and expression and localization of TJ proteins, which are positioned around the apical end of the lateral cell membrane (31). The redistribution of TJ from the intercellular junctions into the intracellular compartment would be an implication of a defective barrier function. We analyzed the distribution of occludin using an immunofluorescence method (supplemental Fig. 1, Under baseline conditions (no bacteria), occludin was not found in the cytoplasm of the epithelial cells. When B infantis or B bifidum were incubated with epithelial monolayers, relocalization of the occludin to the cytoplasm occurred; however, this relocalization of occludin was less apparent when bacteria were grown on HMO compared with LAC. Incubation with LAC-grown bifidobacteria was characterized by discontinuities in membrane staining and submembranous internalization of these proteins.

Previous studies strongly suggest that alterations in TJ composition and protein localization may have a role in the pathogenesis of chronic inflammatory conditions (25). Defining the immunomodulatory capacity of HMO-grown bifidobacteria seems relevant in to understand their contribution to the establishment of the mucosal tolerance and immune responses in the early stages of life. Several studies have evaluated the effects of different bifidobacteria in the production of cytokines by intestinal epithelial cells (36,61,62); however, reports in the literature are often inconclusive (36). For example, researchers noticed minimal impact of probiotic strains B animalis subsp lactis Bb12 and, to a lesser extent, B longum NCC2705 (60) on the cytokine expression induction. In another study, 8 of 19 bifidobacteria, including 2 B bifidum strains, were demonstrated to stimulate the production of IL-8 (63). Our findings show that B infantis attenuates baseline IL-8 secretion in HT-29 cells (64). As expected, the LPS treatment increased expression of IL-8 in Caco-2 cells (65,66); however, the expression of IL-8 was not affected by HMO treatment compared with LAC in the present study.

Previous studies have demonstrated that TNF increases permeability by inducing redistribution of various TJ proteins by internalization (40,67). Here, the expression of TNF by either Caco-2 or HT-29 cells was not significantly altered in the HMO group; however, it is possible that numerical trends toward higher expression of TNF by HMO-grown B bifidum could turn the mucosal immune system on standby and prevent the release of severe inflammation. Other cytokines such as IL-10 have been shown to decrease permeability. We observed higher expression of IL-10 in Caco-2 incubated with HMO-grown B bifidum and B infantis, suggesting potential anti-inflammatory properties of HMO growth. Overall, the response of Caco-2 and HT-29 cells to LPS treatment was consistent with the previously published research (65,66) and HMO did not have any significant effects on the levels of cytokine expression in LPS-treated cells.

Both bifidobacterial species tested in the present study, regardless of whether they were grown on HMO or LAC, translocated at the minimal rate through the intestinal epithelial cells, confirming previous reports that the genus Bifidobacterium is generally noninvasive (24). Despite the observed changes in TJ expression and localization, growth on HMO did not seem to affect the bacterial translocation regardless of the LPS presence.

The concept of pre- and probiotics has attracted increasing attention in recent years. A number of publications show anti-inflammatory effects of probiotics in vivo and in vitro (68,69), and live probiotics and commensals have been shown to affect monolayer barrier function in cultured human epithelial cells (20). Our results support the concept of synbiotics as a synergistic combination of probiotics and prebiotics (70). According to our data, the growth on HMO as a sole carbon source enhances epithelial binding and can induce anti-inflammatory response in the intestinal epithelial cells; however, it did not have any effects in the presence of the massive inflammatory stimulation by LPS. Is it possible that B infantis, as well as other bifidobacteria, exerts the beneficial effect on human physiology in a prophylactic fashion, serving to enhance barrier function? Further studies are needed to establish the role of HMO-grown bifidobacteria in the infant's developing gut.

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The authors thank Man Ki Tsui for technical assistance.

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1. James DC, Lessen R. Position of the American Dietetic Association: promoting and supporting breastfeeding. J Am Diet Assoc 2009; 109:1926–1942.
2. Nakamura N, Gaskins HR, Collier CT, et al. Molecular ecological analysis of fecal bacterial populations from term infants fed formula supplemented with selected blends of prebiotics. Appl Environ Microbiol 2009; 75:1121–1128.
3. German JB, Freeman SL, Lebrilla CB, et al. Human milk oligosaccharides: evolution, structures and bioselectivity as substrates for intestinal bacteria. Nestle Nutr Workshop Ser Pediatr Program 2008; 62:205–218.
4. Kunz C, Rudloff S, Baier W, et al. Oligosaccharides in human milk: structural, functional, and metabolic aspects. Ann Rev Nutr 2000; 20:699–722.
5. Chichlowski M, German JB, Lebrilla CB, et al. The influence of milk oligosaccharides on microbiota of infants: opportunities for formulas. Annu Rev Food Sci Technol 2011; 2:331–351.
6. Niñonuevo MR, Lebrilla CB. Mass spectrometric methods for analysis of oligosaccharides in human milk. Nutr Rev 2009; 67:S216–S226.
7. LoCascio RG, Ninonuevo M, Kronewitter S, et al. A versatile and scalable strategy for glycoprofiling bifidobacterial consumption of human milk oligosaccharides. Microb Biotechnol 2009; 2:333–342.
8. LoCascio RG, Ninonuevo MR, Freeman SL, et al. Glycoprofiling of bifidobacterial consumption of human milk oligosaccharides demonstrates strain specific, preferential consumption of small chain glycans secreted in early human lactation. J Agric Food Chem 2007; 55:8914–8919.
9. Ward RE, Ninonuevo M, Mills DA, et al. In vitro fermentation of breast milk oligosaccharides by Bifidobacterium infantis and Lactobacillus gasseri. Appl Environ Microbiol 2006; 72:4497–4499.
10. Ward RE, Niñonuevo M, Mills DA, et al. In vitro fermentability of human milk oligosaccharides by several strains of bifidobacteria. Mol Nutr Food Res 2007; 51:1398–1405.
11. Sela DA, Chapman J, Adeuya A, et al. The genome sequence of Bifidobacterium longum subsp. infantis reveals adaptations for milk utilization within the infant microbiome. Proc Natl Acad Sci U S A 2008; 105:18964–18969.
12. Sela DA, Li Y, Lerno L, et al. An Infant-associated bacterial commensal utilizes breast milk sialyloligosaccharides. J Biol Chem 2011; 286:11909–11918.
13. Garrido D, Kim JH, German JB, et al. Oligosaccharide binding proteins from Bifidobacterium longum subsp. infantis reveal a preference for host glycans. PLoS ONE 2011; 6:e17315.
14. LoCascio RG, Desai P, Sela DA, et al. Broad conservation of milk utilization genes in Bifidobacterium longum subsp. infantis as revealed by comparative genomic hybridization. Appl Environ Microbiol 2010; 76:7373–7381.
15. Zivkovic AM, German JB, Lebrilla CB, et al. Human milk glycobiome and its impact on the infant gastrointestinal microbiota. Proc Natl Acad Sci U S A 2011;108(suppl 1):4653–8.
16. Erickson AK, Willgohs JA, McFarland SY, et al. Identification of two porcine brush border glycoproteins that bind the K88ac adhesin of Escherichia coli and correlation of these glycoproteins with the adhesive phenotype. Infect Immun 1992; 60:983–988.
17. Tuomola E, Crittenden R, Playne M, et al. Quality assurance criteria for probiotic bacteria. Am J Clin Nutr 2001; 73:393S–398S.
18. Schiffrin E, Brassart D, Servin A, et al. Immune modulation of blood leukocytes in humans by lactic acid bacteria: criteria for strain selection. Am J Clin Nutr 1997; 66:515S–520.
19. He F, Ouwehand AC, Isolauri E, et al. Comparison of mucosal adhesion and species identification of bifidobacteria isolated from healthy and allergic infants. FEMS Immunol Med Microbiol 2001; 30:43–47.
20. Resta-Lenert S, Barrett KE. Live probiotics protect intestinal epithelial cells from the effects of infection with enteroinvasive Escherichia coli (EIEC). Gut 2003; 52:988–997.
21. Bernet MF, Brassart D, Neeser JR, et al. Adhesion of human bifidobacterial strains to cultured human intestinal epithelial cells and inhibition of enteropathogen-cell interactions. Appl Environ Microbiol 1993; 59:4121–4128.
22. Crociani J, Grill JP, Huppert M. Adhesion of different bifidobacteria strains to human enterocyte-like Caco-2 cells and comparison with in vivo study. Lett Appl Microbiol 1995; 21:146–148.
23. Candela M, Seibold G, Vitali B, et al. Real-time PCR quantification of bacterial adhesion to Caco-2 cells: competition between bifidobacteria and enteropathogens. Res Microbiol 2005; 156:887–895.
24. Moroni O, Kheadr E, Boutin Y, et al. Inactivation of adhesion and invasion of food-borne Listeria monocytogenes by bacteriocin-producing Bifidobacterium strains of human origin. Appl Environ Microbiol 2006; 72:6894–6901.
25. Chichlowski M, Hale LP. Bacterial-mucosal interactions in inflammatory bowel disease--an alliance gone bad. Am J Physiol Gastrointest Liver Physiol 2008; 295:G1139–G1149.
26. Ciccocioppo R, Finamore A, Ara C, et al. Altered expression, localization, and phosphorylation of epithelial junctional proteins in celiac disease. Am J Clin Pathol 2006; 125:502–511.
27. Coeffier M, Gloro R, Boukhettala N, et al. Increased proteasome-mediated degradation of occludin in irritable bowel syndrome. Am J Gastroenterol 2010; 105:1181–1188.
28. Gassler N, Rohr C, Schneider A, et al. Inflammatory bowel disease is associated with changes of enterocytic junctions. Am J Physiol Gastrointest Liver Physiol 2001; 281:G216–G228.
29. Zeissig S, Bürgel N, Günzel D, et al. Changes in expression and distribution of claudin 2, 5 and 8 lead to discontinuous tight junctions and barrier dysfunction in active Crohn's disease. Gut 2007; 56:61–72.
30. Khailova L, Dvorak K, Arganbright KM, et al. Bifidobacterium bifidum improves intestinal integrity in a rat model of necrotizing enterocolitis. Am J Physiol Gastrointest Liver Physiol 2009; 297:G940–949.
31. Ewaschuk JB, Diaz H, Meddings L, et al. Secreted bioactive factors from Bifidobacterium infantis enhance epithelial cell barrier function. Am J Physiol Gastrointest Liver Physiol 2008; 295:G1025–G1034.
32. Turner JR. Molecular basis of epithelial barrier regulation: from basic mechanisms to clinical application. Am J Pathol 2006; 169:1901–1909.
33. Zyrek AA, Cichon C, Helms S, et al. Molecular mechanisms underlying the probiotic effects of Escherichia coli Nissle 1917 involve ZO-2 and PKCzeta; redistribution resulting in tight junction and epithelial barrier repair. Cell Microbiol 2007; 9:804–816.
34. Seth A, Yan F, Polk DB, et al. Probiotics ameliorate the hydrogen peroxide-induced epithelial barrier disruption by a PKC- and MAP kinase-dependent mechanism. Am J Physiol Gastrointest Liver Physiol 2008; 294:G1060–1069.
35. Resta-Lenert S, Barrett KE. Probiotics and commensals reverse TNF-[alpha]- and IFN-[gamma]-induced dysfunction in human intestinal epithelial cells. Gastroenterology 2006; 130:731–746.
36. Preising J, Philippe D, Gleinser M, et al. Selection of bifidobacteria based on adhesion and anti-inflammatory capacity in vitro for amelioration of murine colitis. Appl Environ Microbiol 2010; 76:3048–3051.
37. Tanabe S, Kinuta Y, Saito Y. Bifidobacterium infantis suppresses proinflammatory interleukin-17 production in murine splenocytes and dextran sodium sulfate-induced intestinal inflammation. Int J Mol Med 2008; 22:181–185.
38. Sheil B, MacSharry J, O’Callaghan L, et al. Role of interleukin (IL-10) in probiotic-mediated immune modulation: an assessment in wild-type and IL-10 knock-out mice. Clin Exp Immunol 2006; 144:273–280.
39. Aderem A, Ulevitch RJ. Toll-like receptors in the induction of the innate immune response. Nature 2000; 406:782–787.
40. Bruewer M, Luegering A, Kucharzik T, et al. Proinflammatory cytokines disrupt epithelial barrier function by apoptosis-independent mechanisms. J Immunol 2003; 171:6164–6172.
41. Faust D, Hormann S, Friedrich-Sander M, et al. Butyrate and the cytokine-induced (1-proteinase inhibitor release in intestinal epithelial cells. Eur J Clin Invest 2001; 31:1060–1063.
42. Bode L, Rudloff S, Kunz C, et al. Human milk oligosaccharides reduce platelet-neutrophil complex formation leading to a decrease in neutrophil ß 2 integrin expression. J Leukoc Biol 2004; 76:820–826.
43. LoCascio RG. Glycomic and Genetic Characterization of the Metabolism of Human Milk Oligosaccharides by Bifidobacterium Species: Microbiology. Davis: University of California, Davis; 2009:241.
44. Barrangou R, Altermann E, Hutkins R, et al. Functional and comparative genomic analyses of an operon involved in fructooligosaccharide utilization by Lactobacillus acidophilus. Proc Natl Acad Sci U S A 2003; 100:8957–8962.
45. Gnoth MJ, Rudloff S, Kunz C, et al. Investigations of the in vitro transport of human milk oligosaccharides by a caco-2 monolayer using a novel high performance liquid chromatography-mass spectrometry technique. J Biol Chem 2001; 276:34363–34370.
46. Pinto M, Robine-Leon S, Appay MD, et al. Enterocyte-like differentiation and polarization of the human colon carcinoma cell line Caco-2 in culture. Biol Cell 1983; 47:323–330.
47. Cencic A, Langerholc T. Functional cell models of the gut and their applications in food microbiology—a review. Int J Food Microbiol 2010; 141:S4–S14.
48. Jovani M, Barbera R, Farre R, et al. Calcium, iron, and zinc uptake from digests of infant formulas by Caco-2 cells. J Agric Food Chem 2001; 49:3480–3485.
49. Gretchen JM, Michael LS, Raymond PG. Characterization of Caco-2 and HT29-MTX cocultures in an in vitro digestion/cell culture model used to predict iron bioavailability. J Nutr Biochem 2009; 20:494–502.
50. Clark E, Hoare C, Tanianis-Hughes J, et al. Interferon gamma induces translocation of commensal Escherichia coli across gut epithelial cells via a lipid raft-mediated process. Gastroenterology 2005; 128:1258–1267.
51. Anderson JM, Van Itallie CM. Physiology and function of the tight junction. Cold Spring Harb Perspect Biol 2009; 1:a002584.
52. Riedel CU, Foata F, Goldstein DR, et al. Interaction of bifidobacteria with Caco-2 cells--adhesion and impact on expression profiles. Int J Food Microbiol 2006; 110:62–68.
53. Ogawa K, Ben RA, Pons S, et al. Volatile fatty acids, lactic acid, and pH in the stools of breast-fed and bottle-fed infants. J Pediatr Gastroenterol Nutr 1992; 15:248–252.
54. Ninonuevo MR, Ward RE, LoCascio RG, et al. Methods for the quantitation of human milk oligosaccharides in bacterial fermentation by mass spectrometry. Anal Biochem 2007; 361:15–23.
55. Katayama T, Sakuma A, Kimura T, et al. Molecular cloning and characterization of Bifidobacterium bifidum 1,2-{alpha}-L-fucosidase (AfcA), a novel inverting glycosidase (glycoside hydrolase family 95). J Bacteriol 2004; 186:4885–4893.
56. Zweibaum A, Pinto M, Chevalier G, et al. Enterocytic differentiation of a subpopulation of the human colon tumor cell line HT-29 selected for growth in sugar-free medium and its inhibition by glucose. J Cell Physiol 1985; 122:21–29.
57. Boesten R, Schuren F, Willemsen L, et al. Bifidobacterium breve–HT-29 cell line interaction: modulation of TNF-( induced gene expression. Beneficial Microbes 2011; 2:115–128.
58. Gagnon M, Kheadr EE, Le Blay G, et al. In vitro inhibition of Escherichia coli O157:H7 by bifidobacterial strains of human origin. Int J Food Microbiol 2004; 92:69–78.
59. Perez PF, Minnaard Y, Disalvo EA, et al. Surface properties of bifidobacterial strains of human origin. Appl Environ Microbiol 1998; 64:21–26.
60. Guglielmetti S, Tamagnini I, Mora D, et al. Implication of an outer surface lipoprotein in adhesion of Bifidobacterium bifidum to Caco-2 cells. Appl Environ Microbiol 2008; 74:4695–4702.
61. Haller D, Bode C, Hammes WP, et al. Non-pathogenic bacteria elicit a differential cytokine response by intestinal epithelial cell/leucocyte co-cultures. Gut 2000; 47:79–87.
62. Parlesak A, Haller D, Brinz S, et al. Modulation of cytokine release by differentiated Caco-2 cells in a compartmentalized coculture model with mononuclear leucocytes and nonpathogenic bacteria. Scand J Immunol 2004; 60:477–485.
63. Morita H, He F, Fuse T, et al. Adhesion of lactic acid bacteria to Caco-2 cells and their effect on cytokine secretion. Microbiol Immunol 2002;46:293–7.
64. O’Hara AM, O’Regan P, Fanning A, et al. Functional modulation of human intestinal epithelial cell responses by Bifidobacterium infantis and Lactobacillus salivarius. Immunology 2006; 118:202–215.
65. Pozo-Rubio T, Mujico JR, Marcos A, et al. Immunostimulatory effect of faecal Bifidobacterium species of breast-fed and formula-fed infants in a peripheral blood mononuclear cell/Caco-2 co-culture system. Br J Nutr 2011; 106:1216–1223.
66. Candela M, Perna F, Carnevali P, et al. Interaction of probiotic Lactobacillus and Bifidobacterium strains with human intestinal epithelial cells: adhesion properties, competition against enteropathogens and modulation of IL-8 production. Int J Food Microbiol 2008; 125:286–292.
67. Li Q, Zhang Q, Wang M, et al. Interferon-[gamma] and tumor necrosis factor-[alpha] disrupt epithelial barrier function by altering lipid composition in membrane microdomains of tight junction. Clin Immunol 2008; 126:67–80.
68. Quigley EMM. Therapies aimed at the gut microbiota and inflammation: antibiotics, prebiotics, probiotics, synbiotics, anti-inflammatory therapies. Gastroenterol Clin North Am 2011; 40:207–222.
69. Grimoud J, Durand H, de Souza S, et al. In vitro screening of probiotics and synbiotics according to anti-inflammatory and anti-proliferative effects. Int J Food Microbiol 2010; 144:42–50.
70. Roberfroid MB. Prebiotics and synbiotics: concepts and nutritional properties. Br J Nutr 1998; 80:S197–202.

bifidobacteria; inflammation; intestinal epithelial cells; milk oligosaccharides

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