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J. Microbiol. Biotechnol. (2013), 23(4), 518–526http://dx.doi.org/10.4014/jmb.1205.05018First published online January 19, 2013pISSN 1017-7825 eISSN 1738-8872

Effect of Probiotics Lactobacillus and Bifidobacterium on Gut-DerivedLipopolysaccharides and Inflammatory Cytokines: An In Vitro Study Usinga Human Colonic Microbiota Model

Rodes, Laetitia1, Afshan Khan

1, Arghya Paul

1, Michael Coussa-Charley

1, Daniel Marinescu

1,

Catherine Tomaro-Duchesneau1, Wei Shao

1, Imen Kahouli

1,2, and Satya Prakash

1*

1Biomedical Technology and Cell Therapy Research Laboratory, Department of Biomedical Engineering and Artificial Cells andOrgans Research Centre, Faculty of Medicine, McGill University, Quebec, H3A 2B4, Canada2Department of Experimental Medicine, McGill University, Quebec, H3A 2B4, Canada

Received: May 9, 2012 / Revised: November 28, 2012 / Accepted: December 2, 2012

Gut-derived lipopolysaccharides (LPS) are critical to

the development and progression of chronic low-grade

inflammation and metabolic diseases. In this study, the

effects of probiotics Lactobacillus and Bifidobacterium on

gut-derived lipopolysaccharide and inflammatory cytokine

concentrations were evaluated using a human colonic

microbiota model. Lactobacillus reuteri, L. rhamnosus, L.

plantarum, Bifidobacterium animalis, B. bifidum, B. longum,

and B. longum subsp. infantis were identified from the

literature for their anti-inflammatory potential. Each

bacterial culture was administered daily to a human colonic

microbiota model during 14 days. Colonic lipopolysaccharides,

and Gram-positive and negative bacteria were quantified.

RAW 264.7 macrophage cells were stimulated with

supernatant from the human colonic microbiota model.

Concentrations of TNF-α, IL-1β, and IL-4 cytokines

were measured. Lipopolysaccharide concentrations were

significantly reduced with the administration of B. bifidum

(-46.45 ± 5.65%), L. rhamnosus (-30.40 ± 5.08%), B. longum

(-42.50 ± 1.28%), and B. longum subsp. infantis (-68.85 ±

5.32%) (p < 0.05). Cell counts of Gram-negative and

positive bacteria were distinctly affected by the probiotic

administered. There was a probiotic strain-specific effect on

immunomodulatory responses of RAW 264.7 macrophage

cells. B. longum subsp. infantis demonstrated higher

capacities to reduce TNF-α concentrations (-69.41 ± 2.78%;

p < 0.05) and to increase IL-4 concentrations (+16.50 ± 0.59%;

p < 0.05). Colonic lipopolysaccharides were significantly

correlated with TNF-α and IL-1β concentrations (p < 0.05).

These findings suggest that specific probiotic bacteria,

such as B. longum subsp. infantis, might decrease colonic

lipopolysaccharide concentrations, which might reduce

the proinflammatory tone. This study has noteworthy

applications in the field of biotherapeutics for the prevention

and/or treatment of inflammatory and metabolic diseases.

Key words: Lactobacilli, bifidobacteria, probiotics, cytokine,

lipopolysaccharide, inflammation

Gut-derived lipopolysaccharides (LPS) are critical to the

development and/or progression of a variety of human

conditions, such as chronic low-grade inflammation [5],

metabolic diseases [5, 6, 8], and liver diseases [1, 34]. LPS

are potent immunomodulatory components derived from

the cell wall of Gram-negative bacteria [28], which can

translocate to the blood circulation [28]. The immune

recognition of LPS is mediated through the cluster of

differentiation-14 / toll-like receptor-4 receptor complex in

epithelial cells of the intestine and immune cell types,

predominantly macrophages, B cells, and dendritic cells

[12, 28, 43, 45]. An intracellular signaling cascade is

subsequently promoted, leading to the secretion of

proinflammatory cytokines, such as interleukin (IL)-4, IL-

1β and tumor necrosis factor (TNF)-α [5, 33]. IL-4, IL-1β,

and TNF-α have all been linked with the initiation and/or

maintenance of metabolic diseases [9].

In addition, substantial evidence has demonstrated that

gut dysbiosis [25, 34, 46] might be responsible for increased

endotoxemia [37], which further induces chronic low-grade

inflammation [5, 33] and participates in the pathophysiology

*Corresponding authorPhone: +1-514-398-3676; Fax: +1-514-398-7461;E-mail: [email protected]

519 Rodes et al.

of metabolic [5, 6, 8] and liver [1, 34] diseases. The

manipulation of the gut microbiota composition to reduce

endotoxemia and prevent and/or treat human diseases has

already been performed [6-8, 17, 18, 40, 50]. Studies using

antibiotics to selectively eliminate Gram-negative bacteria

have reported significant decreases in cecal and plasma

LPS concentrations [6, 17, 18]. Strategies based on the use

of prebiotics have demonstrated normalized low-grade

inflammation and improved glucose tolerance. Those effects

were associated with increased gut bifidobacterial content [7,

8]. Interestingly, recent studies have suggested that probiotic

supplements might reduce endotoxemia and chronic low-

grade inflammation [40, 50]. However, to our knowledge,

there have been no previous studies examining the effect

of probiotics Lactobacillus and Bifidobacterium on gut-

derived LPS and inflammatory cytokines.

The most widely used probiotic bacteria belong to

Bifidobacterium and Lactobacillus spp. Those species have

been largely investigated for their health promoting properties

in human diseases where gut dysbiosis is inferred to play a

pathogenic role. They are assumed to re-equilibrate the gut

microbiota composition towards health promoting bacterial

populations [40, 50]. Among many hypothesized mechanisms

of action, Bifidobacteria and Lactobacilli have been

extensively studied for their immunomodulatory activities

[13-15, 19, 22, 24, 27, 29, 30, 32, 35, 36, 39, 41, 47-49].

Bacterial cells and components interact with a wide variety

of cells present in the intestines, such as epithelial, dentritic,

and macrophage cells, which further induce the secretion

of pro- and anti-inflammatory cytokines [4, 10, 11, 24, 27,

29, 39, 49]. Frequently, in vitro studies are performed using

bacterial cultures exposed to intestinal cell lines, such as

intestinal epithelial cells-6 [4], Caco-2 cells [4], and RAW

264.7 macrophage cells [10, 11]. Those studies are useful

to investigate the immunomodulatory properties of bacterial

preparations from a pure bacterial culture. However, they

neglect the effects induced in the existing gut microbiota.

In this study, we investigated the effect of probiotics

Lactobacillus and Bifidobacterium on colonic LPS and

inflammatory cytokine concentrations, using an established

model of human colonic microbiota [38] and RAW 264.7

macrophage cell lines.

MATERIALS AND METHODS

Bacterial Cultures and Growth Conditions

Probiotic bacteria tested were identified from the literature for their

anti-inflammatory potential (Table 1). Bacterial strains tested included

L. reuteri NCIMB 701359, L. rhamnosus ATCC 53103, L. plantarum

ATCC 14917, B. animalis ATCC 25527, B. bifidum ATCC 29521,

B. longum ATCC 15707, and B. longum subsp. infantis ATCC

Table 1. List of probiotic strains of Lactobacillus and Bifidobacterium tested in this study.

Bacterial species Strains screened in this study Literature on anti-inflammatory potential

Lactobacillus reuteri NCIMB 701359 [22, 29, 30, 49]

Bifidobacterium bifidum NCIMB 702715 [13, 24]

Lactobacillus rhamnosus ATCC 53103 [15, 19, 36]

Bifidobacterium animalis NCIMB 702242 [27, 39, 47]

Lactobacillus plantarum NCIMB 11974 [32, 41, 48, 49]

Bifidobacterium longum NCIMB 702259 [13, 49]

Bifidobacterium longum subsp. infantis NCIMB 702205 [14, 35]

Fig. 1. Schematic representation of the experimental design:Administration of probiotics Lactobacillus and Bifidobacteriumin a human colonic microbiota model and quantification ofcolonic lipopolysaccharides, and Gram-negative/positive bacterialcell counts and inflammatory cytokines. A human colonic microbiota model was set up. After inoculation with

fresh human feces, a model stabilization period of 14 days was performed.

From day 0, Lactobacillus or Bifidobacterium probiotic strains were

administered daily to the human colonic microbiota model. A control

fermentation system was performed without administration of probiotic

bacteria. Statistical analysis was analyzed to determine the effect of time

vs. day 0. On days 0, 7, and 14, samples were extracted from the colonic

model for quantification of colonic LPS and Gram-negative/positive

bacterial concentrations. RAW 264.7 macrophage cells were stimulated

with bacterial supernatant extracted from the colonic model. Positive and

negative controls consisted of RAW 264.7 cells exposed to, respectively,

2 µg/ml LPS, and DMEM supplemented with 10% FBS. Concentrations of

TNF-α, IL-1β, and IL-4 cytokines were measured.

PROBIOTICS, LIPOPOLYSACCHARIDES, AND INFLAMMATION 520

15697. L. reuteri NCIMB 701359 was purchased from NCIMB

(Aberdeen, Scotland). The other bacterial cultures were purchased

from Cedarlane (Burlington, Canada). A 1% (v/v) inoculum of each

bacterial strain was passaged daily in de Man, Rogosa, and Sharpe

(MRS) broth (Fisher Scientific, Ottawa, Canada) at 37oC in 5%

CO2. Bacterial cell viability was determined on MRS agar triplicate

plates at 3 different dilutions. For Lactobacilli enumeration, agar

plates were incubated for 24 h at 37oC in 5% CO2. For Bifidobacteria

enumeration, agar plates were supplemented with 0.05% (w/v) cysteine

(Fisher Scientific) and incubated in anaerobic jars with anaerobe

atmosphere-generating bags (Oxoid, Nepean, Canada) for 48 h at 37oC.

The Human Colonic Microbiota Model

A semi-continuous human colonic microbiota fermentation system

was operated, as described previously [38]. The system was operated

with a retention time of 144 h. Following the fermentation system

start-up, a stabilization period of 14 days was performed to allow

the bacterial communities to stabilize in order to be representative of

the gut microbiota. On day 0, administration of pure cultures of

Bifidobacterium or Lactobacillus strains was initiated, where 5 × 108 CFU

was administered daily to the human colonic microbiota model during

14 days. A control fermentation system was performed without

daily administration of probiotics. A schematic representation of the

experimental procedure is given in Fig. 1.

Quantification of Gram-Positive and Gram-Negative Bacteria

Gram-negative and positive bacteria were quantified by plate

counting. On days 0, 7, and 14, samples were extracted from the

human proximal colonic microbiota model and serially diluted in

physiological solution (8.5 g/l NaCl). Bacterial cell viability was

determined on selective agar triplicate plates at 3 different dilutions.

McConkey agar (Becton Dickinson, Mississauga, Canada) was used

for the quantification of Gram-negative bacteria. Phenyl ethyl

alcohol agar (Becton Dickinson) was used for the quantification of

Gram-positive bacteria. The agar media were supplemented with

0.05% (w/v) cysteine for the quantification of anaerobes. The agar

plates were incubated for 24 h at 37oC in 5% CO2 for aerobes.

Anaerobic incubation was performed in anaerobic jars with anaerobe

atmosphere-generating bags for 48 h at 37oC. Total Gram-positive

and Gram-negative bacterial cell counts were calculated as the sum

of, respectively, the anaerobic and aerobic Gram positive and negative

bacterial cell counts.

Lipopolysaccharides Quantification

Samples from the human colonic microbiota model were extracted

on days 0, 7, and 14 and instantly frozen at -20oC in glass vials.

LPS were quantified using the QCL 1000, a commercially available

quantitative chromogenic Limulus amebocyte lysate test from Lonza

(Cedarlane). Samples were assayed at different dilutions against a

standard curve of LPS concentrations. Kit instructions were followed

to pursue the Limulus amebocyte lysate assay procedure.

RAW 264.7 Macrophage Cells Maintenance

RAW 264.7 cells were obtained from Cedarlane. The cells were grown

and maintained in Dulbecco’s modified Eagle’s medium (DMEM,

Invitrogen, Carlsbad, Canada) supplemented with 10% fetal bovine

serum (FBS) from Invitrogen. Cell cultures were performed at 5%

CO2 in a humidified atmosphere at 37oC. Cells were grown to 95%

to 100% confluency and passaged every 3-4 days.

Quantification of Inflammatory Cytokines Secreted

On days 0, 7, and 14, samples were removed from the human

colonic microbiota model and centrifuged for 20 min at 2,000 ×g

and 4oC. The supernatant, containing the free LPS, was collected.

RAW 264.7 cells adjusted to a density of 5 × 105 cells/ml were

exposed for 24 h to the bacterial supernatant. The positive control

consisted of RAW 264.7 macrophage cells stimulated with 2 µg/ml

LPS from Escherichia coli (Sigma-Aldrich, Oakville, Canada). The

negative control consisted of RAW 264.7 macrophage cells exposed

to DMEM supplemented with 10% FBS. After 24 h of stimulation,

the culture supernatant was collected and stored at -20oC until

cytokines analysis. The induction of TNF-α, IL-1β, and IL-4 cytokines

was assayed using commercial mouse ELISA kits (eBioscience, San

Diego, USA). Kit instructions were followed to pursue the solid-

phase enzyme-linked immunoassay procedure.

Statistical Analysis

Values are reported as the mean ± SD of triplicates. All data were

analyzed using Prism software (Prism, Version 5.0). The D’Agostino

and Pearson normality test was performed to assess the Gaussian

distribution of the data. Bartlett’s test was performed to assess the

homogeneity of variances. Statistical analysis was analyzed using

one-way ANOVA followed by Bonferroni post-hoc tests or Kruskal-

Wallis test followed by Dunn’s post hoc tests. Correlations were

performed using Spearman’s rank correlation. Value of p < 0.05 were

considered significant.

RESULTS

Effects of Probiotics Lactobacillus and Bifidobacterium

on Colonic Lipopolysaccharides Concentrations

Colonic LPS concentrations in the human colonic microbiota

model were determined before and after 7 and 14 days

of daily administration of probiotics Lactobacillus and

Bifidobacterium (Table 2). L. reuteri, B. bifidum, L.

rhamnosus, B. longum, and B. longum subsp. infantis

reduced LPS concentrations in a time-dependent manner.

Reductions in LPS concentrations on day 14 averaged

23.81 ± 3.18% with L. reuteri (1.80 × 104 ± 2.92 × 103 EU/ml

on day 0 vs. 1.37 × 104 ± 2.55 × 103 EU/ml on day 14),

46.45 ± 5.65% with B. bifidum (1.80 × 104 ± 1.42 × 103 EU/ml

on day 0 vs. 9.63 × 103 ± 1.06 × 103 EU/ml on day 14),

30.40 ± 5.08% with L. rhamnosus (1.92 × 104 ± 1.59 ×

103 EU/ml on day 0 vs. 1.33 × 104 ± 3.50 × 102 EU/ml on

day 14), 42.50 ± 1.28% with B. longum (1.86 × 104 ± 1.87

× 103 EU/ml on day 0 vs. 1.07 × 104 ± 1.29 × 103 EU/ml on

day 14), and 68.85 ± 5.32% with B. longum subsp. infantis

(2.09 × 104 ± 2.03 × 103 EU/ml on day 0 vs. 6.50 × 103 ±

5.30 × 102 EU/ml on day 14). The effect was significant

with B. bifidum, L. rhamnosus, B. longum, and B. longum

subsp. infantis (p < 0.05). Conversely, L. plantarum

administration significantly increased LPS concentrations

by 28.84 ± 4.44% on day 7 (1.79 × 104 ± 8.46 × 102 EU/ml

on day 0 vs. 2.31 × 104 ± 3.18 × 103 EU/ml on day 7)

(p < 0.05). The effect was not maintained after 14 days of

daily L. plantarum administration.

521 Rodes et al.


on Colonic Gram-Negative and -Positive Bacterial Cell

Counts

Cell counts of Gram-negative bacteria (LPS-producing

bacteria) (Fig. 2A) and Gram-positive bacteria (non-LPS

producing bacteria) (Fig. 2B) in the human colonic

microbiota model were determined before and after 7 and

14 days of daily administration of probiotics Lactobacillus

and Bifidobacterium. There was a significant increase in

the Gram-negative bacterial cell count on day 7 with L.

reuteri (+21.78 ± 1.56%; 6.12 ± 0.07 log CFU/ml on day 0

vs. 7.46 ± 0.12 log CFU/ml on day 7; p < 0.05) and L.

plantarum (+7.71 ± 1.06%; 6.18 ± 0.05 log CFU/ml on day

0 vs. 6.66 ± 0.09 log CFU/ml on day 7; p < 0.05). The effect

was not significant on day 14. Conversely, there were

significant reductions on day 14 in Gram-negative bacteria

with B. longum (-7.28 ± 1.88%; 6.11 ± 0.06 log CFU/ml

on day 0 vs. 5.67 ± 0.06 log CFU/ml on day 14; p < 0.05)

and B. longum subsp. infantis (-9.55 ± 1.51%; 6.14 ±

0.06 log CFU/ml on day 0 vs. 5.55 ± 0.05 log CFU/ml on

day 14; p < 0.05). L. reuteri administration was associated

with a significant increase in total Gram-positive bacteria

on day 7 (+16.85 ± 3.64%; 6.47 ± 0.21 log CFU/ml on day

0 vs. 7.56 ± 0.01 log CFU/ml on day 7; p < 0.05). B.

animalis administration induced a significant reduction in

Gram-positive bacteria on day 14 (-18.37 ± 0.01%; 6.56 ±

0.11 log CFU/ml on day 0 vs. 5.36 ± 0.07 log CFU/ml on

day 14; p < 0.05). Colonic LPS concentrations were not

correlated with Gram-negative and -positive bacterial cell

counts.


on TNF-α, IL-1β, and IL-4 Cytokine Concentrations

To investigate the immunmodulatory effects of probiotics

Lactobacillus and Bifidobacterium, RAW 264.7 macrophage

cells were stimulated with bacterial supernatant (containing

most of the free LPS) sampled from the colonic model before

and after 7 and 14 days of daily probiotics administration.

The TNF-α (Fig. 3A), IL-1β (Fig. 3B), and IL-4 (Fig. 3C)

concentrations secreted by positive control (2 µg/ml LPS)

were significantly increased compared with the negative

control (DMEM supplemented with 10% FBS) (p < 0.05).

Fig. 2. Effects of probiotics Lactobacillus and Bifidobacteriumon colonic (A) Gram-negative and (B) Gram-positive bacterialcell counts in an in vitro human colonic model. Lactobacillus and Bifidobacterium probiotic strains were administered

daily to a human colonic microbiota model. A control fermentation system

was performed without administration of Lactobacillus and Bifidobacterium

strains. On days 0, 7, and 14, colonic colonic Gram-negative and Gram-

positive bacterial cell counts were quantified (log CFU/ml). Data represent

the mean ± SD (n = 3). *

Indicates statistically significant difference (p < 0.05)

obtained by a Kruskal-Wallis test followed by Dunn’s post hoc tests.

Table 2. Effects of probiotics Lactobacillus and Bifidobacterium on colonic lipopolysaccharides concentrations in a human colonicmicrobiota model.

Bacteria administered to the colonic model

Lipopolysaccharides concentration (EU/ml)

Day 0 Day 7 Day 14

Control 2.02 × 104 ± 2.53 × 103 1.84 × 104 ± 2.38 × 103 1.87 × 104 ± 3.22 × 103

L. reuteri 1.80 × 104 ± 2.92 × 103 1.79 × 104 ± 1.34 × 103 1.37 × 104 ± 2.55 × 103

B. bifidum 1.80 × 104 ± 1.42 × 103 1.17 × 104 ± 4.15 × 102 9.63 × 103 ± 1.06 × 103 *

L. rhamnosus 1.92 × 104 ± 1.59 × 103 1.55 × 104 ± 2.45 × 103 1.33 × 104 ± 3.50 × 102 *

B. animalis 1.95 × 104 ± 7.82 × 102 1.76 × 104 ± 2.10 × 103 1.81 × 104 ± 2.63 × 103

L. plantarum 1.79 × 104 ± 9.46 × 102 2.31 × 104 ± 3.18 × 103 * 1.77 × 104 ± 1.21 × 103

B. longum 1.86 × 104 ± 1.87 × 103 1.49 × 104 ± 1.23 × 103 1.07 × 104 ± 1.29 × 103 *

B. longum subsp. infantis 2.09 × 104 ± 2.03 × 103 6.87 × 103 ± 9.89 × 102 6.50 × 103 ± 5.30 × 102 *

Lactobacillus and Bifidobacterium probiotic strains were administered daily to a human colonic microbiota model. A control fermentation system was

performed without probiotic administration. On days 0, 7, and 14, colonic LPS were quantified (EU/ml). Data represent the mean ± SD (n = 3). * Indicates

statistically significant difference (p < 0.05) obtained by a Kruskal-Wallis test followed by Dunn’s post hoc tests.


There were major reductions in TNF-α concentration with

B. animalis (-67.11 ± 0.56%; 1174.33 ± 8.87 µg/ml on day

0 vs. 386.20 ± 6.98 µg/ml on day 14; p < 0.05), L. plantarum

(-68.01 ± 1.88%; 1133.09 ± 62.29 µg/ml on day 0 vs.

362.49 ± 2.51 µg/ml on day 14; p < 0.05), B. longum

(-65.14 ± 1.45%; 1083.17 ± 30.48 µg/ml on day 0 vs.

377.62 ± 5.69 µg/ml on day 14; p < 0.05), and B. longum

subsp. infantis (-69.41 ± 2.78%; 1142.80 ± 88.43 µg/ml

on day 0 vs. 349.55 ± 5.32 µg/ml on day 14; p < 0.05). As

for IL-1β concentrations, reductions were most extreme on

day 14 with L. reuteri (-25.40 ± 2.55%; 637.13 ± 11.56 µg/ml

on day 0 vs. 475.33 ± 17.58 µg/ml on day 14; p < 0.05), B.

bifidum (-26.96 ± 2.52%; 650.17 ± 21.75 µg/ml on day 0

vs. 474.87 ± 10.29 µg/ml on day 14; p < 0.05), and L.

rhamnosus (-21.48 ± 2.39%; 647.10 ± 5.52 µg/ml on day

0 vs. 508.11 ± 18.50 µg/ml on day 14; p < 0.05). There

was a time-dependent increase in IL-4 concentration with

the administration of L. reuteri (+30.73 ± 7.07%; 214.57 ±

1.65 µg/ml on day 0 vs. 284.55 ± 35.55 µg/ml on day 14;

p < 0.05), L. plantarum (+46.12 ± 3.88%; 207.58 ± 8.90 µg/ml

on day 0 vs. 303.32 ± 4.95 µg/ml on day 14; p < 0.05)

and B. longum subsp. infantis (+16.50 ± 0.59%; 233.05 ±

2.11 µg/ml on day 0 vs. 271.50 ± 4.38 µg/ml on day 14;

p < 0.05).

Correlations Between Colonic Lipopolysaccharides

Concentrations and Secretion of Inflammatory Cytokines

There was a strong positive correlation between colonic

LPS concentrations and TNF-α (r = 0.447, P = 0.002; Fig. 4A)

and IL-1β (r = 0.504, P = 0.002; Fig. 4B) concentrations.

Conversely, colonic LPS concentrations were negatively

Fig. 3. Effects of probiotics Lactobacillus and Bifidobacteriumon the concentration of (A) TNF-α, (B) IL-1β, and (C) IL-4inflammatory cytokines secreted in RAW 264.7 macrophage cells.Lactobacillus and Bifidobacterium probiotic strains were administered

daily to a human colonic microbiota model. A control fermentation system

was performed without administration of probiotic bacteria. RAW 264.7

macrophage cells were stimulated with bacterial supernatant extracted

from the colonic model. Positive and negative controls consisted of RAW

264.7 cells exposed to, respectively, 2 µg/ml LPS and DMEM supplemented

with 10% FBS. Concentrations of TNF-α, IL-1β, and IL-4 cytokines were

measured (pg/ml). All values are reported as the mean ± SD (n = 3).

Statistical analysis was analyzed using one-way ANOVA followed by

Bonferroni post hoc tests to determine the effect of treatments vs. control

fermentation # (p < 0.05), and time vs. day 0 * (p < 0.05).

Fig. 4. Correlations between colonic lipopolysaccharide concentrationsand (A) TNF-α, (B) IL-1β, and (C) IL-4 inflammatory cytokinesconcentrations. Lactobacillus and Bifidobacterium probiotic strains were administered

daily to a human colonic microbiota model. A control fermentation

system was performed without probiotic administration. On days 0, 7, and

14, colonic LPS were quantified (EU/ml) and RAW 264.7 macrophage

cells were stimulated with bacterial supernatant extracted from the colonic

model. The concentrations of TNF-α, IL-1β, and IL-4 cytokines were

measured (pg/ml). Correlations between colonic LPS concentrations and

inflammatory cytokine concentrations were performed using Spearman’s

rank correlation.

523 Rodes et al.

correlated with IL-4 concentrations, but the effect was not

significant (r = -0.160, P = 0.170; Fig. 4C).

DISCUSSION

Compelling evidence demonstrates that gut microbiota dysbiosis

may promote systemic chronic low-grade inflammation [5,

33], and initiate metabolic [5, 6, 8] and liver [1, 34] diseases

through a mechanism that involves increased exposure to

Gram-negative bacteria-derived LPS. Current therapies to

modulate the human gut microbiota to reduce colonic LPS

concentrations have been developed with the aim to

improve the inflammatory status [6-8, 17, 18, 50]. However,

to our knowledge, there have been no previous studies

examining the effect of probiotics Lactobacillus and

Bifidobacterium on gut-derived LPS and inflammatory

cytokines. This was the objective of this study.

The effects of L. reuteri NCIMB 701359, L. rhamnosus

ATCC 53103, L. plantarum ATCC 14917, B. animalis

ATCC 25527, B. bifidum ATCC 29521, B. longum ATCC

15707, and B. longum subsp. infantis ATCC 15697 on

colonic LPS, and TNF-α, IL-1β, and IL-4 inflammatory

cytokine concentrations were evaluated using a human

colonic microbiota model [38] and RAW 264.7 macrophage

cells. Although the model of human colonic microbiota

used in this study neglects differences in individual gut

microbiotas and diseases, it allows the experimentation of

a complex culture of the gut microbiota under well-

standardized conditions. Results demonstrated that the

administration of B. bifidum, L. rhamnosus, B. longum,

and B. longum subsp. infantis to the human proximal

colonic microbiota model significantly reduced LPS

concentrations in a time-dependent manner. No data on

intestinal or cecal LPS have been reported in probiotic

studies, which impedes further comparison. Nevertheless,

preclinical studies have already shown that Bifidobacterium

and Lactobacillus bacteria can decrease plasma LPS

concentrations [40, 50]. Other studies using antibiotics

have reported significant decreases in cecal and plasma

LPS concentrations [6, 17, 18]. Strategies based on the use

of prebiotics have shown reduced endotoxemia, improved

glucose tolerance, and normalized low-grade inflammation

[7, 8]. Since colonic bacterial LPS are derived from Gram-

negative bacteria, colonic Gram-negative and -positive

bacterial cell counts were monitored. Reductions in Gram-

negative bacterial cell counts were significant following

administration of B. animalis, B. longum, and B. longum

subsp. infantis. Surprisingly, neither the Gram-negative

nor -positive bacterial cell count was correlated with LPS

concentrations. We hypothesize that uncultivable bacteria,

which are not quantified using plate counting, might

account for the discrepancy. Molecular techniques, such as

real-time polymerase chain reaction should be used to

quantify Gram-negative/positive bacterial cell counts in

future studies [3].

Immunomodulatory properties of probiotics Lactobacillus

and Bifidobacterium were assessed using RAW 264.7

macrophage cells. Macrophages are known to play an

important role in the regulation of innate inflammatory

responses to intestinal bacteria, particularly to the bacterial

toxin LPS [16]. Macrophage recognition of bacterial

conserved elements is mediated predominantly by toll-like

receptors and nucleotide-binding oligomerization domain

molecules [28, 43]. This further triggers the production of

inflammatory cytokines and chemokines [5, 33, 43]. In

addition, macrophages, along with dendritic cells, present

foreign antigens to cells of the acquired immune system

[12, 43, 45]. Thus, both the innate and acquired immune

systems are integrated for efficient inflammatory responses

to microorganisms. It is believed that the activation of

innate immune signaling pathways by toll-like receptors

may help induce and/or maintain chronic inflammation

due to continuous exposure to LPS [23]. Previous in vitro

studies have investigated the inflammatory responses of

RAW 264.7 macrophage cells to pure bacterial cultures

[10, 11]. Those studies were useful in determining the

direct effects of a bacterial strain on macrophage activation.

However, they disregarded the effects of a mixed culture

induced in the gut microbiota ecosystem. Soluble components,

predominantly LPS, released from bacterial cells into the

gut environment are responsible for the majority of

inflammatory cytokines secreted [4, 11, 14, 26]. Lamina

propria-resident macrophages have been shown to suppress

or have anergic responses to LPS stimuli [42, 44]. However,

studies involving the inflamed mucosa have not observed

this feature, probably due to the recruitment of blood

monocytes-derived macrophages [20, 21]. In addition, it

was recently suggested that the inflammatory profile

elicited by murine RAW 264.7 macrophage cells depends

primarily on the initial dosage of LPS challenge [31]. In

the present study, murine macrophage cells were stimulated

with bacterial supernatant, which contained the free LPS,

derived from our human colonic microbiota model. This

novel approach, which allows to investigate probiotic

immunomodulatory activities in the existing gut microbiota,

has already demonstrated accuracy and reproducibility [2].

Cells density and LPS concentration (positive control)

were based on published publications [10, 11]. TNF-α, IL-

1β, and IL-4 inflammatory cytokines were selected for

quantification because they are promoted following the

immune recognition of gut-derived LPS [5, 33]. In addition,

they have been linked with inflammatory and metabolic

diseases [9]. Results demonstrated that the exposure of

macrophage cells to bacterial supernatant and LPS solution

stimulated major secretions of TNF-α, IL-1β, and IL-4

cytokines compared with the negative control. In addition,

probiotics Lactobacillus and Bifidobacterium induced


significant biological reductions in the TNF-α concentration.

The reductions were extreme with B. animalis, L. plantarum,

B. longum, and B. longum subsp. infantis. Reductions in

the IL-1β concentration were extreme with L. reuteri, L.

rhamnosus, and B. bifidum. The IL-4 concentration was

significantly increased with L. reuteri, L. plantarum, and

B. longum subsp. infantis. Overall, B. longum subsp.

infantis demonstrated a higher capacity to reduce colonic

LPS concentrations, to decrease TNF-α proinflammatory

cytokines, and to increase IL-4 anti-inflammatory cytokines

secretions. Interestingly, the colonic LPS concentrations

were positively and significantly correlated with the TNF-

α and IL-1β concentrations. We suggest that specific

probiotic strains of Lactobacillus and Bifidobacterium can

decrease colonic LPS concentrations, which might further

reduce the proinflammatory tone.

This is the first in vitro study to investigate the effects of

probiotics Lactobacillus and Bifidobacterium on colonic

LPS and inflammatory cytokine concentrations using a

human colonic microbiota model. The findings revealed that

specific probiotic strains, such as B. longum subsp. infantis,

can decrease colonic LPS concentrations, which might

further reduce the secretion of inflammatory cytokines in

macrophage cells. This study has noteworthy applications

in the field of biotherapeutics for the prevention and/or

treatment of inflammatory and metabolic diseases.

Acknowledgments

This work was supported by a research grant from the

Canadian Institutes of Health Research (CIHR) MOP No.

64308 to S. Prakash. L. Rodes and W. Shao acknowledge

the Doctoral Training Award from the Fonds de Recherche

Santé Québec (FRSQ). A. Paul acknowledges the Alexander

Graham Bell Canada Graduate Scholarship from the Natural

Sciences and Engineering Research Council (NSERC) and

the FRSQ Postdoctoral Training Award. M. Coussa-Charley

acknowledges the FRSQ Master’s award. C. Tomaro-

Duchesneau acknowledges the Alexander Graham Bell

Canada Graduate Scholarship from NSERC.

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