Modulation of T Regulatory and Dendritic Cell Phenotypes Following Ingestion of Bifidobacterium longum, AHCC® and Azithromycin in Healthy Individuals

The probiotic Bifidus BB536 (BB536), which contains Bifidobacterium longum, has been shown to have enhanced probiotic effects when given together with a standardized extract of cultured Lentinula edodes mycelia (AHCC®, Amino Up Co. Ltd., Sapporo, Japan). BB536 and AHCC® may modulate T cell and dendritic cell (DC) phenotypes, and cytokine profiles to favour anti-inflammatory responses following antibiotic ingestion. We tested the hypothesis that orally administered BB536 and/or AHCC®, results in modulation of immune effector cells with polarisation towards anti-inflammatory responses following antibiotic usage. Forty healthy male volunteers divided into 4 equal groups were randomised to receive either placebo, BB536, AHCC® or a combination for 12 days in a double-blind manner. After 7 days volunteers also received 250 mg azithromycin for 5 days. Cytokine profiles from purified CD3+ T cells stimulated with PDB-ionomycin were assessed. CD4+ CD25+ forkhead box P3 (Foxp3) expression and peripheral blood DC subsets were assessed prior to treatment and subsequently at 7 and 13 days. There was no difference in cytokine secretion from stimulated CD3+ T cells between treatment groups. Compared with baseline, Foxp3 expression (0.45 ± 0.1 vs. 1.3 ± 0.4; p = 0.002) and interferon-gamma/interleukin-4 (IFN-γ/IL-4) ratios were increased post-treatment in volunteers receiving BB536 (p = 0.031), although differences between groups were not significant. For volunteers receiving combination BB536 and AHCC®, there was an increase in myeloid dendritic cells (mDC) compared with plasmacytoid DC (pDC) counts (80% vs. 61%; p = 0.006) at post treatment time points. mDC2 phenotypes were more prevalent, compared with baseline, following combination treatment (0.16% vs. 0.05%; p = 0.002). Oral intake of AHCC® and BB536 may modulate T regulatory and DC phenotypes to favour anti-inflammatory responses following antibiotic usage.


Cell density Determination
Cell density and viability were determined by trypan blue (Sigma-Aldrich, UK) dye exclusion. This method is based on the principle that viable cells are not permeable to certain dyes, whereas non-viable cells are. Cell suspension was mixed thoroughly via pipetting before adding 5 μl of cell suspension to 20 μl of trypan blue solution [0.4% (w/v) trypan blue in PBS]. The trypan blue-cell suspension was transferred to both chambers of a Brightline® Haemocytometer (Sigma-Aldrich, UK). Cell concentration per ml and the total number of cells were determined using the following calculations: Cells per ml = Total cell count × 10 4 × 5 (dilution factor) Total cell no. = Cells per ml × original vol. of cell suspension

Assay of Cytokine Concentration Using Flow Cytometry
Cytokines at various time points were assayed using flow cytometry and the cytometric bead array technique. The BenderMedSystems multiplex flowcytomix kit (Bender Medystems, Ebioscience, San Diego, CA, USA) was chosen to perform all cytokine assays. This kit employs a series of particles with discrete fluorescence intensities to simultaneously detect multiple soluble analytes, and when combined with flow cytometry, a multiplexed assay is created. The BenderMedSystems Human Th1/Th2 Cytokine Kit (Bender Medystems, USA) was used to quantitatively measure IL-2, IL-4, IL-5, IL-6, IL-10, TNF-α and IFN-γ protein levels in a single sample.
Firstly, the human cytokine standards were prepared. One vial of lyophilized human cytokine standards was reconstituted with 200 μl of assay diluents (provided in kit) to prepare a 10× bulk standard. The reconstituted standard was allowed to equilibrate for 20 min before agitating the vial to mix thoroughly. The 10× bulk standard was aliquoted to 10 μl Eppendorfs and stored at −20°C. 12 × 75 mm fluorescein-activated cell sorter (FACS) tubes (BD Biosciences, San Jose, CA, USA) were labelled and arranged in the following order: Top Standard, 1:2, 1:4, 1:8, 1:16, 1:32, 1:64, 1:128, and 1:256. 90 μl of assay diluents was added to the top standard tube while 30 μl of assay diluents was added to each of the remaining tube. 10 μl of 10× bulk standard was then transferred to the top standard tube and mixed thoroughly by inverting tube.
Serial dilutions followed by transferring 30 μl from the top standard to the 1:2 dilution tube and mixed thoroughly before transferring 30 μl from the 1:2 tube to 1:4 tube and so on to the 1:256 tube mixing thoroughly each time. The assay diluents reagent served as the negative control.
Next, mixed human cytokine capture beads were prepared. The capture beads were bottled individually (i.e. each cytokine in a different bottle) and were pooled together immediately before use. The number of assay tubes including standards and controls was determined and labelled. The capture bead suspension was vigorously vortexed for a few seconds before use. 3 μl aliquot of each capture bead for each assay tube to be analysed was added to a single tube labelled 'mixed capture beads' and vortexed to mix thoroughly. 15 μl of the mixed capture beads was added to the appropriate assay tubes followed by 15 μl of the human PE detection reagent (included in the kit). 15 μl of the human cytokine standard dilutions was then added to the control assay tubes while 15 μl of test samples was added to the test assay tubes. The assay tubes were incubated for 3 h at room temperature and protected from direct exposure to light before washing with 500 μl of wash buffer (included in kit) and centrifuged at 200 × g for 5 min. The supernatant was carefully aspirated and discarded from each assay tube and the bead pellets re-suspended in 300 μl of wash buffer before reading on a flow cytometer. Results were analysed using the Bender MedSystems Flow Cytomix Software Package (Bender Medystems, eBioscience, USA).

Purification of CD3 + Cells from PBMCs
Cryopreserved cells, which had been stored for 2 weeks to 18 months, were thawed by placing the cryovials in a water bath at 37°C for 5 min. An equal volume of FBS pre-warmed to 37°C was gently added to the vial. The suspension was allowed to equilibrate in the water bath for 5 min and was then gently layered over 10 ml of RPMI 1640 (Sigma-Aldrich, UK) at 37°C. The cell suspension was allowed to equilibrate for 5 min, during which time the denser DMSO cell suspension settled to the bottom of the centrifuge tube. The tube was gently inverted twice to mix the suspension. After centrifugation for 10 min at 500 × g, cells were re-suspended in PBS plus 10% FCS, counted, and scored for viability by trypan blue exclusion.

Principle of Immunomagnetic Separation
First, CD3 + cells were magnetically labelled with CD3 MicroBeads (Miltenyi Biotec, Auburn, CA, USA). The cell suspension was loaded onto a MACS® Column (Miltenyi Biotec, USA) and placed into the magnetic field of a MACS Separator (Miltenyi Biotec, USA). The magnetically labelled CD3+ cells were retained on the column. The unlabelled cells were run through and this cell fraction was depleted of CD3 + cells. After removal of the column from the magnetic field, the magnetically retained CD3 + cells were eluted as the positively selected cell fraction. To isolate CD3 + T cells, 10 7 PBMCs were mixed with 20 μL of human CD3+ cell isolation MACS microbeads (Miltenyi Biotec, USA) in a total volume of 80 μL and incubated on ice for 30 minutes. During incubation an LS column (Miltenyi Biotec, USA) was primed by washing twice with MACS buffer (Miltenyi Biotec, USA). On completion of the incubation period, the MACS microbeads labelled cell suspension was loaded onto the LS column followed by washing twice with MACS buffer. On removal from the magnetic field CD3 + T cells retained in the column were eluted as the positively selected fraction. This fraction was then washed in MACS buffer twice and then resuspended in RPMI with glutamine and BSA.

Stimulation of Cytokine Release from CD3 + Cells
For activation, CD3 + cells were resuspended at a concentration of 4 × 10 5 cells per well in complete culture medium, e.g. RPMI 1640 supplemented with 2 mM L-glutamine, penicillin, streptomycin, 20% human serum and of phorbol 12,13-dibutyrate (PDB) (10 nM)/ionomycin (0.5 μM). 100 μl were distributed per well in duplicate. Complete culture medium was then added to bring the final volume to 200 μl. The plates were then incubated for 24 h at 37°C prior to centrifugation at 5000 × g for 15 min to collect the supernatants. These were placed into marked cryovials and snap frozen in liquid nitrogen before storage at −80°C.

Flow Cytometric Determination of Foxp3 Expression
PBMCs were stained for cell surface markers for 30 minutes with 2.5 μl phycoerythrin Texas red conjugate [energy couple dye (ECD)] antihuman CD4 (eBioscience, San Diego, CA, USA) and 5μl phycoerythrin (PE) antihuman CD25 (eBioscience, USA). Cell surface markers for CD4 and CD25 were determined by staining for 30 minutes with 2.5 μl of ECD antihuman CD4 and 5μl fluorescein isothiocyanate (FITC) antihuman CD25 (eBioscience, USA). The cells were then washed with RPMI and 2% FCS; 2% formaldehyde was used for fixation of PBMCs for 10 minutes at room temperature (RT). The PBMCs were then washed once in phosphate buffered saline (PBS) containing 2% FCS, twice in PBS/0.5% Tween with 0.05% azide and 3% FCS. 2.5μl FITC antihuman Foxp3 (intracellular) was added to the corresponding tubes and incubated for 2 hours at 4°C (shaking gently every 20 min).
The PBMC pellet was then washed twice in PBS/0.5% Tween, 0.05% azide and 3% FCS. The pellet was then resuspended in 400 μl of 0.5% paraformaldehyde fixative solution for FC analysis.
The BD™CompBeads (BD Biosciences, USA) anti-mouse Ig, κ polystyrene microparticles were used to optimise fluorescence compensation settings for multicolour flow cytometric analyses. The set provided two populations of microparticles, the BD™CompBeads anti-mouse Ig, κ particles, which bind any mouse κ light chain-bearing immunoglobulin, and the BD™CompBeads negative control (FCS), which has no binding capacity. When mixed together with a fluorochrome-conjugated mouse antibody, the BD™CompBeads provide distinct positive and negative (background fluorescence) stained populations which can be used to set compensation levels manually or using instrument set-up software.
For each flow cytometric analysis sample an appropriate corresponding tube was set up with a minus fluorescence control (to identify and eliminate all possible non-specific antibody binding and to ensure accurate data acquisition and interpretation was carried out).
For the analysis, lymphocytes were gated from total PBMCs. Following this, a gate was created from cells demonstrating CD4 high SS low . Subsequent gate was created to select cells demonstrating CD4 high Foxp3 high . A final gate selecting cells with CD25 high Foxp3 high allowed the assay of CD4 + CD25 + Foxp3 + cells ( Figure S1).
Peripheral blood DCs can be divided specific phenotypic subsets based on the expression of CD11c. Furthermore, CD11c + DC can be subdivided into at least two other distinct phenotypes: a major CD1c + subset and a minor CD1c − subset [1]. Further definition and characterisation of peripheral blood DC subsets have been facilitated by identification of subset-specific surface markers CD141 (BDCA-3), CD303 (BDCA-2), and CD304 (BDCA-4).

Detection of Dendritic Cell Subsets by Flow Cytometry.
In the absence of pathogenic stimuli, CD303 and CD304 are co-expressed on CD11c − pDCs, whereas both CD1c + and CD141 + markers are associated with CD11c + mDCs. These markers can, therefore, be used to further define mDC subsets, CD11c + CD1c + CD141 − (mDC1) and CD11c + CD1c − CD141 + (mDC2), respectively. Importantly, there is no CD1c or CD141 expression on pDC, which can be used to differentiate mDC and pDC phenotypes [1]. Under steady state conditions, both mDC and pDC exhibit immature phenotypes with low expression of MHC molecules and the costimulatory molecules CD40, CD80, and CD86 [1][2][3]. A scheme of surface markers identification for mDCs and pDCs is shown in Figure S2.

Isolation of DC Subsets
Dendritic cell subsets were isolated using flow cytometry analysis of cell surface markers. A twotube method was utilised to differentiate live from dead cells and to separate pDCs from mDCs. This technique was required in order to allow the detection of fluorochromes on particular channels using a flowcytometer (MoFlow XDP, Beckman Coulter Inc., Brea, CA, USA) Using specific antibody combinations. Firstly, PBMCs were incubated for 15 min at 4°C with 1 μl of a viability detection agent (LIVE⁄DEAD Fixable Dead Cell Stain Kit, Life Technologies, Carlsbad, CA, USA). Cells were then washed in 1 ml of Hank's balanced salt solution before addition of the antibody cocktail. The concentrations of the various antibodies used in the FACS analysis are shown in Table S1. Following addition of the antibodies, the cells were incubated for 30 min at 4°C prior to three washes in Hank's balanced salt solution. Cells were the fixed in 2% formaldehyde prior to analysis.