Influence of early gut microbiota on the
maturation of childhood mucosal and systemic
Y M Sjögren, Sara Tomicic, Anna Lundberg, Malin Fagerås-Böttcher, B Björkstén, E
Sverremark-Ekström and Maria Jenmalm
Linköping University Post Print
N.B.: When citing this work, cite the original article.
This is the pre-reviewed version of the following article:
Y M Sjögren, Sara Tomicic, Anna Lundberg, Malin Fagerås-Böttcher, B Björkstén, E
Sverremark-Ekström and Maria Jenmalm, Influence of early gut microbiota on the maturation
of childhood mucosal and systemic immune responses, 2009, Clinical and Experimental
Allergy, (39), 12, 1842-1851.
which has been published in final form at:
Postprint available at: Linköping University Electronic Press
Influence of early gut microbiota on the maturation of childhood
mucosal and systemic immune responses.
Gut flora and immune responses.
Ylva M. Sjögren1, Sara Tomicic2, Anna Lundberg2, Malin F. Böttcher2, Bengt
Björkstén3, Eva Sverremark-Ekström1, Maria C. Jenmalm2
1The Department of Immunology, the Wenner-Gren Institute, Stockholm University,
2 The Division of Paediatrics, the Department of Clinical and Experimental Medicine, Faculty
of Health Sciences, Linköping University, Linköping, Sweden.
3 The Institute of Environmental Medicine, Karolinska Institutet, Stockholm, Sweden.
Ylva M. Sjögren
The Department of Immunology
The Wenner-Gren Institute
Arrhenius laboratory of Natural Sciences F5
Svante Arrhenius väg 16-18
106 91 Stockholm
Phone: +46 8 16 44 36
Fax: +46 8 6129542
Introduction: Among sensitized infants those with high, as compared with low levels, of
salivary secretory IgA (SIgA) are less likely to develop allergic symptoms. Also, early
colonization with certain gut microbiota, e.g. Lactobacilli and Bifidobacterium species, might
be associated with less allergy development. Although animal and in vitro studies emphasize
the role of the commensal gut microbiota in the development of the immune system, the
influence of the gut microbiota on immune development in infants is unclear.
Objective: To assess whether early colonization with certain gut microbiota species
associates with mucosal and systemic immune responses i.e. salivary SIgA and the
spontaneous toll like receptor (TLR) 2 and TLR4 mRNA expression and LPS-induced
cytokine/chemokine responses in peripheral blood mononuclear cells (PBMCs).
Methods: Faecal samples were collected at one week, one month and two months after birth
from 64 Swedish infants, followed prospectively to five years of age. Bacterial DNA was
analyzed with real-time PCR using primers binding to Clostridium difficile, four species of
bifidobacteria, two lactobacilli groups and Bacteroides fragilis. Saliva was collected at age
six and twelve months and at two and five years and SIgA was measured with ELISA. The
PBMCs, collected twelve months after birth, were analyzed for TLR2 and TLR4 mRNA
expression with real-time PCR. Further, the PBMCs were stimulated with LPS and
cytokine/chemokine responses were measured with Luminex.
Results: The number of Bifidobacterium species in the early faecal samples correlated
significantly with the total salivary SIgA levels at six months. Early colonization with
Bifidobacterium species, lactobacilli groups or C. difficile did not influence TLR2 and TLR4
expression in PBMCS. However, PBMCs from infants colonized early with high amounts of
Bacteroides fragilis expressed lower levels of TLR4 mRNA spontaneously. Furthermore,
LPS-induced production of inflammatory cytokines and chemokines, e.g. IL-6 and CCL4
(MIP-1β), were inversely correlated to the relative amounts of Bacteroides fragilis in the
early faecal samples.
Conclusion: Bifidobacterial diversity may enhance the maturation of the mucosal SIgA
system and early high colonization with Bacteroides fragilis might down-regulate LPS
responsiveness in infancy.
Gut microbiota, lactobacilli, bifidobacteria, Clostridium difficile, Bacteroides fragilis,
SIgA, TLR2, TLR4, infant
APRIL A proliferation-inducing factor
cDNA complementary deoxyribonucleic acid
ELISA Enzyme linked immunosorbent assay
G- Gram negative
G+ Gram positive
GALT Gut associated lymphoid tissue
MAMPs Microbial associated molecular pattern
mRNA messenger ribonucleic acid
PBMCs Peripheral blood mononuclear cells
PRR Pattern recognition receptor
PSA Polysaccharide A
RT PCR Reverse transcriptase polymerase chain reaction
SIgA Secretory IgA
Th T helper cell
TLR Toll-like receptor
TNF Tumour necrosis factor
TSLP Thymic stromal-derived lymphoprotein
A reduced microbial pressure in Westernized countries is postulated to underlie
the increase in allergy development during the past decades. Alterations in the early gut
microbiota may precede allergy development (1,2). Children developing allergy are,
compared to those who remain non-allergic, not as often colonized with bifidobacteria and
enterococci but more frequently colonized with clostridia including Clostridium (C.) difficile
early in life (1,2). Bifidobacterium colonization at species level might also be associated with
allergy (3-5). Furthermore, children who develop allergy during their five years of life were
already during the first week of life less often colonized with lactobacilli (L.) group I,
comprising L. rhamnosus, L. casei and L. paracasei, as compared to children not developing
allergy (3). However, not all studies demonstrate a relationship between the early gut
microbiota and allergy development (6). Whether pre- and postnatal administration of
probiotic bacteria associates with decreased incidence of allergic disease is unclear. Less IgE-
associated eczema (7) and less allergy development up to two years (8) but not five years of
age (9) are reported in probiotic-treated infants. However, others studies find no any
association between probiotic administration and allergy development (10). In order to
understand whether the early gut microbiota is associated with allergy development the
possible mechanisms, explaining how the early gut microbiota influence infant immune
responses and thus subsequent allergy development, need to be investigated.
Animal studies have emphasized the importance of the gut microbiota in
educating the immune system. The gut associated lymphoid tissue (GALT) in germ-free (GF)
animals is underdeveloped (11,12) with few IgA+ B cells (13). In addition, also the spleen of
GF mice contains fewer numbers of CD4+ T cells (14). Colonization with lactobacilli strains
increases the numbers of IgA+ B cells (13) and a polysaccharide from Bacteroides fragilis
(PSA) could restore the proportion of splenic CD4+ T cells (14). Also, PSA administration
increases IL-10 production (15). The early colonization appears to be of particular
importance as Bifidobacterium infantis could restore Th1 responses in neonatal but not in
adult ex-germ-free mice (16). Additionally, the serum levels of IgA appears to increase in
piglets colonized shortly after birth but remain low in GF piglets (17).
Studies in humans indicate that the early colonization with certain bacteria
influence systemic immune responses. For example, Bacteroides fragilis colonization in
infancy appears to increase the number of circulating IgA and IgM antibody producing cells
(18). Furthermore, infants who received a mixture of probiotic strains from birth had higher
plasma levels of C-reactive protein, total IgA, total IgE and IL-10 at six months than infants
in the placebo group (19). Although the composition of the gut microbiota at six months does
not appear to influence the salivary IgA levels at that age (20), it is conceivable that the
microbiota that colonizes the gut shortly after birth might influence immune development.
The innate immune compartment responds to different microbial associated
molecular patterns (MAMPs) by pattern recognition receptors (PRRs) expressed on immune
cells, e.g. dendritic cells, and mucosal epithelia (21). The capacity to respond to PRR signals
is important for adaptive immune responses such as IL-12 dependent direction of naïve T
cells into Th1 cells (22). The PRR Toll like receptor (TLR) 2 responds to MAMPs such as
lipoteichoic acid from Gram positive (G+) bacteria, while the TLR4 recognizes the endotoxin
lipopolysaccharide (LPS) from Gram negative (G-) bacteria together with a complex with
CD14 and MD-2 (23). Soluble CD14 appears to be higher in the plasma of infants early
colonized with Staphylococcus (S.) aureus than in the plasma of non-colonized infants (24).
However, it is not known how the early infant gut microbiota, consisting of both G+ and G-
bacteria, influences TLR responsiveness.
High salivary secretory IgA (SIgA) may protect sensitized children from
developing allergic symptoms and non-allergic children tend to have higher salivary SIgA
levels than allergic children (25). Recently, we also demonstrated that Swedish children have
less SIgA early in life compared to Estonian age-matched children (Tomicic et al
unpublished). As Estonian children are frequently colonized with lactobacilli (26), we
hypothesize that the early gut microbiota, notably lactobacilli and bifidobacteria at species
level, could influence the maturation of salivary SIgA production. The possible allergy-
protective effects from the increased pre- and postnatal microbial exposure in farming
environments might increase the expression of several PRRs e.g. TLR2 and TLR4 (27,28).
Thus, we also hypothesized that early exposure to certain intestinal microbes, the G-
Bacteroides fragilis and the G+ C. difficile, bifidobacteria and lactobacilli, modulates TLR
expression and LPS responsiveness. In addition, as Bacteroides fragilis colonization/PSA
administration induces IFNγ production in GF mice (14), we hypothesized that early
Bacteroides fragilis colonization influences spontaneous and PHA-induced IFNγ production
The birth cohort, comprising 123 Swedish children born between March 1996 and
October 1999, has been described in detail elsewhere (29). Briefly, the children were born at
term and had an uncomplicated perinatal period. Inclusion in the present study was based on
availability of faecal samples at one week, one month and/or at two months of age. In all, 64
infants were included. From the majority of these children blood samples collected at twelve
months, and/or salivary samples, collected at six months, twelve months, two years and/or
five years were also available. The study was approved by the Regional Ethics Committee for
Human Research at Linköping University. The parents of all children gave their informed
consent in writing. Clinical examinations of the children were made at three or six and twelve
months and at two and five years. At these occasions, skin prick tests were performed, and
questionnaires were completed regarding, in example, symptoms of allergy and use of
antibiotics. As development of allergic disease in relation to their early gut microbiota has
been investigated in a previous study in these children (3), this will not be discussed here.
A majority of the children had a history of atopic disease in the immediate family
(78%, Table 1). In total, three children were delivered with caesarean section. Most infants
were exclusively breastfed during their first three months (83%) and only two infants
received antibiotics during this time.
Analysis of bacterial DNA in the faecal samples
The analysis of the bacterial DNA has been described previously (3). In short,
faecal samples were collected into sterile plastic containers by the parents when the infants
were one week (collected at day five or six), one month and two months old. The samples
were stored at -70ºC until analysis.
Qiamp DNA Stool Mini Kit™ (Qiagen, Hilden, Germany) was used for the
isolation of DNA from 180-220 mg faeces and the included protocol for increasing the
bacterial DNA over human DNA was used. The concentration of nucleic acids was measured
with BIO-RAD Smartspec (Bio-Rad laboratories, Hercules, CA, USA) at 260 nm using BIO
RAD trUView Disposable Cuvettes (Bio-Rad laboratories, Hercules, CA, USA).
Bacterial DNA was analyzed with real-time PCR using primers binding to C.
difficile, B. adolescentis, B. longum, B. bifidum, B. breve, lactobacilli group I (comprising L.
rhamnosus, L. casei and L. paracasei), lactobacilli group II (comprising L. gasseri and L.
johnsonii) and Bacteroides fragilis. Sequences and concentrations of primers are described in
(3). The primers were used due to their specificity in binding to the specific bacterial DNA,
as well as for their suitability in SYBR Green PCR chemistry. Reference bacterial DNA, used
as standard and positive control, was purchased from LGC Standards (Borås, Sweden) and
BIOTECHON Diagnostics (Potsdam, Germany). The SYBR Green real-time PCR was
performed using 96-well detection plates in ABI prism 7000 (Applied Biosystem, Stockholm,
Sweden). All samples were performed in triplicates. Each well contained 2xPower SYBR
Green mastermix (Applied Biosystems, Stockholm, Sweden), forward and reverse primer
(MWG-Biotech, Edersburg, Germany), DNA and water. The Absolute Quantification
protocol in 7000 System software version 1.2.3f2 (Applied Biosystems) was employed and
the amplification was performed using the default program of 40 cycles, which also included
melting curve analysis. The software calculated the amount of specific bacterial DNA from
the standard curve, constructed from known amounts of reference bacterial DNA (5ng diluted
in 10-fold dilution series down to 50 fg). To avoid detecting false positives, triplicates with
CT values above 35 were considered as negative. The amount of the specific bacterial DNA
was then related to the total amount of nucleic acids in each sample. The specific bacterial
DNA is thus expressed as percent specific bacterial DNA of total nucleic acids and referred
to as relative amounts of specific bacterial DNA. The detection limit was 5*10-6 % specific
bacterial DNA of total nucleic acids. Negative samples were assigned a value ten times below
the detection limit i.e. 5*10-7 % specific bacterial DNA of total nucleic acids, and used in the
Analysis of total secretory IgA in saliva
Saliva was collected at six and twelve months and at two and five years of age
from the buccal cavity using a hand-pump connected to a thin plastic tube and immediately
frozen at -20oC. Before analysis of SIgA, the samples were heated at 56 oC for 30 minutes
and then centrifuged at 5000g for 15 minutes. Total SIgA was analyzed with ELISA using an
anti-human secretory component antibody (Dakopatts AB) as coating antibody, as previously
described (25). Human IgA (Sigma Immunochemicals) was diluted in seven steps for the
standard curve. The SIgA, bound to the coating antibodies, was detected with alkaline
phosphatase conjugated goat anti-human IgA antibodies (Sigma Immunochemicals) and
FAST pNPP substrate. The detection range was 16 to 1000 ng/mL for SIgA.
Venous blood samples were drawn into heparinized tubes (Becton Dickinson,
Stockholm, Sweden) at 12 months of age. Peripheral blood mononuclear cells (PBMCs) were
isolated on a Ficoll Paque density gradient (Pharmacia Biotech, Uppsala, Sweden). The cells
were thereafter cryopreserved according to standard methodology in 10% dimethyl sulfoxide
(Sigma-Aldrich, Stockholm, Sweden), 50% foetal calf serum and 40 % RPMI-1640 (Life
Technologies AB, Täby, Sweden).
Reverse transcription (RT) PCR of mRNA and quantification of gene expression
250 µL of cell suspension (1x106 viable cells/mL (as checked by Trypan blue
exclusion) in AIM-V serum free medium (Life Technologies AB) with 20 µM
mercaptoethanol (Sigma-Aldrich)) was cultivated for 24h in 37°C with 5% CO2 (Forma
CO2-incubator model 3862, Forma Scientific Inc., Marietta, Ohio, USA) with no added
stimulus. Thereafter the cells were lysed with RLT lysis buffer (Qiagen, Hilden, Germany).
The cell lysates were stored in -70° C until RNA isolation. Total RNA was isolated using
RNeasy™ 96 Protocol (Qiagen, Hilden, Germany) according to manufacturer´s instructions.
Briefly, the cells were lysed by RLT lysis buffer, mixed with ethanol and applied to
RNeasy™ 96 well plates. Contaminants were washed away by buffers and the RNA was
eluted in 2 x 30µl of RNase free water.
RNA was converted to complementary DNA (cDNA) using High Capacity
Archive Kit (Applied Biosystems, Foster City, CA, USA) according to manufacturer´s
instructions. Briefly, RNA was mixed with MultiScribe reverse transcriptase, random
primers, dNTPs, reverse transcription buffers and RNAse free water in 40µl reactions and run
for 10 min at 25° followed by 37°C for 120 min. The gene expression analysis was performed
with quantitative real-time PCR. Taqman® Gene Expression and Taqman® Fast Universal
PCR Master Mix were purchased from Applied Biosystems. The assay id:s were TLR2
Hs00610101_m1 and TLR4 Hs00152939_m1. Primers, probes, Master Mix, water and cDNA
was mixed and the samples were run on an Applied Biosystems 7500 Fast Real-Time PCR
system. The thermal cycling conditions were 95°C for 20s, followed by 40 cycles of 95°C
for 3s and 60°C for 30s. rRNA was used as internal controls, i.e. the amount of the expressed
gene was calculated relative to the amount of rRNA in each sample. Standards were used to
create a standard curve from which the amounts were calculated in each run using the
standard curve method as described in User Bulletin no 2 (Applied Biosystems). The inter-
assay variation was <6% for both genes and the slopes of the standard curves varied between
-3.3 to -3.9. Each sample was run in duplicates and a CV of maximum 15% was allowed.
Cytokine and chemokine analysis
After thawing, 0.25 mL of 1*106 viable cells/mL cell suspension in AIM-V serum
free medium were cultured in 37°C with 5% CO2. The cells were either cultured with 10
ng/mL LPS Salmonella enterica serotype thyphimurium (Sigma-Aldrich) or with medium
alone. The viability was checked by trypan blue exclusion. After 24h the supernatants were
collected and stored at -70°C until analysis. The cell supernatants were analysed with a
multiplex assay kit, according to the manufacturer’s instructions (Human cytokine 9-plex
panel, Bio-Rad Laboratories, Hercules, CA, USA). The assay detects the following analytes;
IL-6, CXCL8 (IL-8), IL-10, IL12p70, IL-17, IL-1β, CCL2 (MCP-1), CCL4 (MIP-1β), and
TNF. The samples were analysed on a Luminex100 instrument (Biosource, Nivelles, Belgium)
and the data was analysed with the software StarStation 2.0 (Applied Cytometry Systems,
Sheffield, UK). The lower detection limit was 10 pg/mL for IL-6 and CCL4, 6 pg/mL for IL-
10, CXCL8 and TNF, 2 pg/mL for IL12p70, 3 pg/mL for IL-17, 4 pg/mL for IL-1β, and 18
pg/mL for CCL2. Comparisons of LPS-induced cytokine and chemokine responses were
made after the control value, i.e. responses from cells cultured in medium alone, was
For IFNγ analyses, 1 mL of cell suspension (1*106 viable cells/mL in AIM-V
serum free medium supplemented with 20µM mercaptoetanol (Sigma-Aldrich)) was grown in
duplicates alone or with 2µg/mL phytohaemagglutamin (PHA) (Sigma-Aldrich). The culture
conditions and analyses have been described in more detail a previous paper (30). After one
and six days of culture, the supernatants were collected and stored at -20ºC. Interferon γ was
measured with an ELISA kit (CLB Pelikine CompactTM, Research Diagnostics Inc., Flandern,
NJ, USA). The detection limit was 25 pg/mL. Spontaneous IFNγ production was detected
after six days, but not one day of culture.
Spearman’s rank coefficient was calculated to investigate whether relative
amount of bacterial DNA and/or number of Bifidobacterium species in faecal samples
correlated with concentrations of total salivary SIgA and the relative mRNA expression of
TLR2 and TLR4. It was also calculated to understand whether the bacterial amounts
influenced the production of cytokines and chemokines from PBMCs after microbial
stimulation. Mann-Whitney U test was performed to evaluate whether infants colonized with
the specific bacteria had different levels of total SIgA in saliva compared with un-colonized
infants. In addition, Mann-Whitney U test was calculated to understand whether the
colonized and un-colonized groups expressed different levels of TLR2 and TLR4 mRNA and
produced different levels of IFNγ at 12 months. Fisher´s exact test was calculated to
understand whether colonization with Bacteroides fragilis was associated with spontaneous
IFNγ production. Many statistical tests were performed and thus p-values close to 0.05 might
be false significances. As this study was not sufficiently powered to detect very low p-values,
we only report significant values observed at several time points, or significant values
observed at one occasion with at least a tendency at another time point. This approach would
decrease the risk of including false significances. The study should be viewed as exploratory
and consequently p<0.05 were chosen as statistically significant.
Gut microbiota during the first two months
Bifidobacteria and Bacteroides fragilis were commonly present already in one-
week old infants, whereas the other bacteria tended to become more frequently detected as
the infants grew older (table 2). Lactobacilli occurred in lower amounts than bifidobacteria
and Bacteroides fragilis. Few infants were colonised with C. difficile.
Total salivary secretory IgA associates with the early gut microbiota.
The number of Bifidobacterium species in faeces collected one week, one month
and two months after birth correlated with total salivary SIgA at six months (r=0.51 to 0.58,
p=0.02 to 0.045, Fig. 1 exemplifies the one month colonization) but not in older children.
When analyzing the different Bifidobacterium species separately, it was shown that infants
colonized with B. adolescentis at one and two months had significantly higher levels of SIgA
at six months (Fig. 2a and median; 12.0 (4.3-19-7) µg/mL, respectively) compared to non-
colonized infants (Fig. 2a and median; 5.3 (2.0-10.0) µg/mL, respectively). Furthermore, the
intensity of one and two month B. adolescentis colonization (expressed as percentage B.
adolescentis DNA of all faecal nucleic acids), was associated with higher SIgA at six months
(r=0.66 and 0.55, p=0.007 and 0.03, respectively). In addition, SIgA levels at six months
tended to be higher in infants colonized with B. breve at one week and one month (median;
9.0 (6.2-17.0) µg/mL and Fig 2a, respectively) than in non-colonized infants (median; 4.3
(2.0-15.5) µg/mL, p=0.02 and Fig 2a, respectively). The intensity of B. bifidum colonization
at one week, one month and two months after birth was correlated with the SIgA production
at twelve months (r=0.41 to 0.47, p=0.01 to 0.04, Fig. 3 exemplifies one month colonization).
The levels of SIgA at six months and five years were significantly associated with
the colonisation with lactobacilli group I at one month after birth (Fig. 2a and d). Also,
increased levels of SIgA at five years were associated with colonization with C. difficile at
one and two months after birth (p=0.008 to 0.03, Fig. 2d). The few individuals colonized with
this bacterium could greatly bias the results, however. The salivary SIgA levels at two years
of age were not associated with the early gut microbiota (Fig. 2c).
Early colonization with Bacteroides fragilis associates with spontaneous TLR4 mRNA
expression, LPS responsiveness and spontaneous IFNγ production at 12 months of age.
The expression of TLR4 mRNA was negatively correlated to the intensity of
Bacteroides fragilis colonization during the first week and month after birth (r=-0.47, p=0.02
and Fig 4). None of the other analyzed bacteria associated with the spontaneous expression of
TLR2 or TLR4 (data not shown).
In concordance with the TLR4 mRNA findings, the intensity of Bacteroides
fragilis colonization at one week, one month and two months were all inversely related to the
LPS induced production of IL-6 and CCL4 (MIP-1β) (r=-0.62 to -0.79, p=0.002 to 0.04 and
r= -0.75 to -0.86, p=0.001 to 0.005, Fig. 5 exemplifies one month colonization). Also, LPS
induced IL-8 and IL-17 were inversely related to the intensity of Bacteroides fragilis
colonization at one week after birth (r=-0.75, p=0.008 and r=-0.68, p=0.02). In contrast, the
intensity of B. adolescentis colonization at two months correlated or tended to correlate to
TNF, IL-10 and IL-12p70 (r=0.64, p=0.03, r=0.67, p=0.03 and r=0.55, p=0.08 respectively)
with similar tendencies at one month (r=0.54, p=0.07, r=0.46, p=0.13 and r=0.58, p=0.05
respectively). None of the other investigated bacteria yielded any conclusive significant
correlations, i.e. significant at one occasion with at least a tendency at another time point.
Early Bacteroides fragilis colonization was also investigated for its possible role
in modulating IFNγ production at twelve months. Infants colonized with Bacteroides fragilis
at one week and one month did not produce IFNγ spontaneously more often at twelve months
(9/22 and 7/20) than infants not colonized with Bacteroides fragilis (2/10 and 2/13). All the
infants who produced IFNγ spontaneously at twelve months and were un-colonized at one
week and one month had become colonized with Bacteroides fragilis at age two months,
however. Thus, among the 21 infants colonized with Bacteroides fragilis at two months 10
produced IFNγ spontaneously at twelve months compared to no spontaneous IFNγ-producers
among the 14 un-colonized infants (p=0.019). There was no difference in IFNγ production
after PHA stimuli between the colonized and un-colonized infants.
The Bifidobacterium flora during the first two months after birth was associated
with enhanced production of salivary SIgA at six and twelve months. The SIgA levels at six
months were mainly associated with the diversity of the Bifidobacterium flora and
colonization with B. adolescentis, whereas the colonization with B. bifidum was associated
with the SIgA levels at twelve months. The present study, in which bifidobacteria were
identified at species level suggests that the diversity of Bifidobacterium species might have a
larger impact on the maturation of IgA responses rather than the number of bacteria within
the Bifidobacterium genera (20,31). The colonization shortly after birth might be of particular
importance for maturation of immune responses, as others have reported that colonization
with bifidobacteria at six months did not associate with salivary SIgA levels at that age (20).
Early colonization with a strain of E. coli can induce faecal antigen-specific SIgA levels
already the first weeks after birth (32) and early colonization with toxigenic S. aureus
influences plasma IgA levels at 4 months but not at 18 months (31), thus further supporting
the role of early colonization for maturation of IgA responses. As early colonization with
bifidobacterial species was not associated with salivary SIgA levels at two and five years,
other factors are probably more important for later induction of salivary SIgA responses.
Recurrent respiratory infections stimulate IgA formation as suggested by the observation that
infants exposed to more than three infections during infancy have higher SIgA levels than
less infected infants (Sandin et al unpublished). Whether the gut microbiota and recurrent
infections account for early and late SIgA formation respectively, needs to be further studied.
Early Bacteroides fragilis colonization associates with number of IgA producing cells (18)
however we found no association between Bacteroides fragilis colonization and salivary
Both T cell dependent and T cell independent IgA class switching occurs in the
human gut (reviewed in (33)). Interestingly, the gut microbiota has been shown to activate
intestinal epithelial cells (IECs) through TLRs leading to the production of a proliferation
inducing ligand (APRIL) and thymic stromal-derived lymphoprotein (TSLP) (34). The TSLP
in turn influences dendritic cells to produce additional APRIL, which is important for T cell
independent IgA2 class switching (34). Furthermore, IgA production is profoundly reduced in
germ-free mice (13). Also, some studies indicate that GALT IgA-producing B cells can
migrate to other part of the mucosal tissue, such as salivary glands (reviewed in (35)). Thus,
it is conceivable that lactic acid bacteria in the gut may influence IgA production in saliva.
Alternatively, the studied gut microbiota species are epiphenomena correlating with unknown
factors that the infants are exposed to, e.g. in the oral mucosa, and this is what effects the
SIgA production in saliva. We previously showed that bifidobacterial diversity was
associated with house dust exposure to microbes (endotoxin) (3).
As higher levels of SIgA might protect sensitized infants from allergy
development (25), it is tempting to speculate that a more diverse early Bifidobacterium flora
could prevent allergy development partially by increasing salivary SIgA. Studies from our
group also indicate that salivary SIgA levels, already at age three, six and twelve months, are
lower among two year-old sensitized infants with allergic symptoms compared to
asymptomatic sensitized infants ((25), Tomicic et al unpublished). Thus, early induction of
SIgA levels could be of relevance for allergy development. We previously showed that
infants developing allergy were less often colonized with B. adolescentis during their first
two months of life (3). However, we could not conclusively show that development of allergy
up to five years of age was associated with less Bifidobacterium diversity in early infancy (3).
In addition, others showed that B. bifidum colonization was more common among infants
who developed allergy at six months (36) and B. adolescentis colonization could be more
common in allergic compared to non-allergic infants and children (5,37). Thus, the role of the
Bifidobacterium flora in association with allergy development is controversial and needs
further investigation. Speculatively, the observed association between the early
Bifidobacterium flora and SIgA levels during the first year could indicate that an overall more
diverse gut microbiota increases salivary SIgA. Actually, a more diverse early gut microbiota
is more common among non-allergic infants compared to allergic infants (38).
The colonization with lactobacilli group I (L. rhamnosus, L. casei and L.
paracesei) at one month was also associated with higher levels of SIgA in saliva at six
months and five years. This is further supported by the observations that Swedish infants
appear to be less often colonized with lactobacilli and also have lower levels of SIgA in
saliva than Estonian infants ((26), Tomicic unpublished).
The TLR4 mRNA expression was lower in infants colonized with high relative
amounts of the G- bacterium Bacteroides fragilis at one week and one month after birth.
Furthermore, LPS-induced CCL4 (MIP1β), CXL8 (IL-8), IL-6 and IL-17 levels were
inversely correlated to the intensity of Bacteroides fragilis colonization. Partially supporting
our findings, it has been observed that increased home dust exposure to components from G-
bacteria, i.e. 3-hydroxy fatty acids, is inversely associated with the production of IL-6 and
TNF after mitogen stimulation (39). Lappalainen also showed that exposure to muramic acid,
from G+ bacteria, correlated with IL-6 and TNF production (39). Supporting this, we
observed that the relative amounts of the G+ B. adolescentis tended to correlate with the
production of TNF, IL-10 and IL-12p70 after LPS stimulation. Yet, the spontaneous
expression of TLR4 mRNA was not associated with early B. adolescentis colonization
possibly implying that these associations were found by chance. The spontaneous TLR2
mRNA expression was not associated with colonization with any of the investigated bacteria.
This could indicate that these early gut microbiota species do not associate with TLR2
expression and the childhood TLR2 expression may be more influenced by prenatal microbial
exposure (27) than the early gut microbiota.
High LPS exposure induces endotoxin (LPS) tolerance (40) and endotoxin
tolerant mice have decreased splenic TLR2 and TLR4 (41). Speculatively, the observed
decreased LPS responsiveness in infants colonized with high levels of Bacteroides fragilis
could thus be due to endotoxin tolerance caused by the high exposure to the G- Bacteroides
fragilis. Also, PSA, which is another component from Bacteroides fragilis has shown
immunomodulatory properties (14,15). This component is able to balance the production of
inflammatory cytokines, e.g. TNF and IL-17, in mice with induced colitis (15). As PSA
increases the production of IL-10, this has been proposed to be the explanation for regulating
inflammatory responses (15). We could not show that infants colonized with high amounts of
Bacteroides fragilis produce higher levels of IL-10 after LPS stimuli, however. The
decreased TLR4 mRNA expression associated with intense Bacteroides fragilis colonization
could be an additive effect to the regulation of the inflammatory cytokines. Spleen cells from
ex-germfree mice colonized with Bacteroides fragilis/PSA produce more IFNγ after
αCD3/αCD28 stimulation than spleen cells from GF mice (14). In a very different system, we
showed that infants colonized with Bacteroides fragilis during their first two months more
often produced IFNγ spontaneously than un-colonized infants. However, early colonization
with Bacteroides fragilis was not associated with higher PHA-induced IFNγ production. As
spleen cells from conventional and ex-GF mice colonized with Bacteroides fragilis produce
similar levels of IFNγ after αCD3/αCD28 stimuli (14), the other gut microbiota species of
infants may influence their PHA-induced IFNγ production. In our previous paper, we could
not find any relationship between early Bacteroides fragilis colonization and allergy
development (3) postulating that the immune responses associated with Bacteroides fragilis
colonization in infants play a minor role in the development of allergy.
In conclusion, the early Bifidobacterium flora was associated with salivary SIgA
levels during the first year of life in this exploratory study. Thus bifidobacterial diversity may
enhance the maturation of the mucosal SIgA system. In addition, early colonization with
elevated amounts of Bacteroides fragilis was associated with decreased LPS responsiveness
indicating that Bacteroides fragilis could influence systemic immune responses.
We are grateful to all children and parents participating in this study and to the
following people at Linköping University Hospital; research nurse Lena Lindell, laboratory
technologist Ann-Marie Fornander and Dr Karel Duchén for clinical evaluation of the
This work was supported by the Ekhaga foundation, the Cancer and Allergy
foundation, Mjölkdroppen, the Magnus Bergvall foundation, the Konsul Th C Berghs
foundation, the Swedish Research Council (57X-15160-05-2 and 74X-20146-01-2), the
National Swedish Association against Allergic Diseases, the National Heart and Lung
Association and the Swedish Foundation for Health Care Sciences and Allergy Research.
Table 1. Demographic data of the subjects.
Born with Caesarean section
Any atopic heredity
Exclusively breastfed ≥ 3 months 53 (83%)a
Oral antibiotics ≤ 3 months
Number of family members
Household area/individual (m2)
a = missing data for one subject regarding exclusive breastfeeding.
b = missing data for three subjects regarding number of family members and thus household
3 (3-8) b
29 (18-75) b
Table 2. Proportion of infants (%) colonized with Bifidobacterium, Lactobacillus, C.
difficile and Bacteroides fragilis during the first two months of life. The numbers in the
headers show the number of infants with available faecal samples at the different time
points. The figures, after the different bacteria, show the percent of infants who were
colonized with the bacteria.
Bacteria n=52 n=56 n=56
B. longum 88 91 91
B. adolescentis 71 66 75
B. bifidum 56 63 63
B. breve 40 43 50
Lactobacilli group I 31 57 57
Lactobacilli group II 33 43 54
C. difficile 6 11 21
Bacteroides fragilis 60 55 55
1 week 1 month 2 months
Fig. 1. Total salivary secretory IgA (SIgA) at six months is associated with a
more diverse Bifidobacterium (B.) flora. The y-axis shows total salivary secretory IgA
(SIgA) (µg/ml) at six months and the x-axis shows the number of Bifidobacterium species at
Fig. 2. Colonization with bifidobacteria, lactobacilli and Clostridium difficile
at one month of age in relation to salivary secretory IgA (SIgA) at age six months (a),
twelve months (b), two years (c) and five years (d). The positive sign denote colonized
infants and the negative sign those lacking the bacteria. The numbers in brackets show the
number of infants from whom both faecal and salivary samples were available. The short
lines demonstrate the median values. Colonization with Bacteroides fragilis and B. longum
did not influence the SIgA production and these bacteria are therefore not shown.
Fig. 3. Total salivary secretory IgA (SIgA) at twelve months is associated
with the intensity of B. bifidum colonization. The y-axis shows total salivary secretory IgA
(SIgA) (µg/ml) at twelve months and the x-axis shows intensity of B. bifidum colonization
(expressed as percent B. bifidum DNA of total amount nucleic acids at one month. The dotted
line shows the detection limit at 5*10-6 % B. bifidum DNA of total amount nucleic acids. n.d.
= not detectable.
Fig. 4. The spontaneous expression of TLR4 mRNA at twelve months is
inversely correlated to the intensity of Bacteroides fragilis colonization at one month
after birth. The y-axis shows the relative TLR4 mRNA expression/rRNA in PBMCs
collected twelve months after birth and the x-axis shows the relative amount of Bacteroides
fragilis at one month. The dotted line shows the detection limit at 5*10-6 % Bacteroides
fragilis DNA of total amount nucleic acids. n.d. = not detectable.
Fig. 5. The LPS-induced production of CCL4 (MIP-1β) (a) and IL-6 (b) from
PBMCs, collected twelve months after birth, is inversely correlated to the intensity of
Bacteroides fragilis colonization one month after birth. The y-axis shows the concentration
of MIP-1β (a) and IL-6 (b) in supernatants from PBMCs collected twelve months after birth
and stimulated with LPS for 24 hours. The x-axis shows the relative percent of Bacteroides
fragilis at one month. The dotted line shows the detection limit, 5*10-6 % Bacteroides fragilis
DNA of total amount nucleic acids. n.d. = not detectable.
(1) Björkstén B, Sepp E, Julge K, Voor T, Mikelsaar M. Allergy development and the intestinal microflora
during the first year of life. J.Allergy Clin.Immunol. 2001;108:516-520.
(2) Penders J, Thijs C, van den Brandt PA, Kummeling I, Snijders B, Stelma F, et al. Gut microbiota
composition and development of atopic manifestations in infancy: the KOALA Birth Cohort Study. Gut
(3) Sjögren YM, Jenmalm MC, Böttcher MF, Björkstén B, Sverremark-Ekström E. Altered early infant gut
microbiota in children developing allergy up to 5 years of age. Clin.Exp.Allergy 2009;39:518-526.
(4) Gore C, Munro K, Lay C, Bibiloni R, Morris J, Woodcock A, et al. Bifidobacterium pseudocatenulatum is
associated with atopic eczema: a nested case-control study investigating the fecal microbiota of infants.
J.Allergy Clin.Immunol. 2008;121:135-140.
(5) Stsepetova J, Sepp E, Julge K, Vaughan E, Mikelsaar M, de Vos WM. Molecularly assessed shifts of
Bifidobacterium ssp. and less diverse microbial communities are characteristic of 5-year-old allergic children.
FEMS Immunol.Med.Microbiol. 2007;51:260-269.
(6) Adlerberth I, Strachan DP, Matricardi PM, Ahrné S, Orfei L, Åberg N, et al. Gut microbiota and
development of atopic eczema in 3 European birth cohorts. J.Allergy Clin.Immunol. 2007;120:343-350.
(7) Abrahamsson TR, Jakobsson T, Böttcher MF, Fredrikson M, Jenmalm MC, Björkstén B, et al. Probiotics in
prevention of IgE-associated eczema: a double-blind, randomized, placebo-controlled trial. J.Allergy
(8) Kukkonen K, Savilahti E, Haahtela T, Juntunen-Backman K, Korpela R, Poussa T, et al. Probiotics and
prebiotic galacto-oligosaccharides in the prevention of allergic diseases: a randomized, double-blind, placebo-
controlled trial. J.Allergy Clin.Immunol. 2007;119:192-198.
(9) Kuitunen M, Kukkonen K, Juntunen-Backman K, Korpela R, Poussa T, Tuure T, et al. Probiotics prevent
IgE-associated allergy until age 5 years in cesarean-delivered children but not in the total cohort. J.Allergy
(10) Kopp MV, Hennemuth I, Heinzmann A, Urbanek R. Randomized, double-blind, placebo-controlled trial of
probiotics for primary prevention: no clinical effects of Lactobacillus GG supplementation. Pediatrics
(11) Suzuki K, Ha SA, Tsuji M, Fagarasan S. Intestinal IgA synthesis: a primitive form of adaptive immunity
that regulates microbial communities in the gut. Semin.Immunol. 2007;19:127-135.
(12) Smith K, McCoy KD, Macpherson AJ. Use of axenic animals in studying the adaptation of mammals to
their commensal intestinal microbiota. Semin.Immunol. 2007;19:59-69.
(13) Ibnou-Zekri N, Blum S, Schiffrin EJ, von der Weid T. Divergent patterns of colonization and immune
response elicited from two intestinal Lactobacillus strains that display similar properties in vitro. Infect.Immun.
(14) Mazmanian SK, Liu CH, Tzianabos AO, Kasper DL. An immunomodulatory molecule of symbiotic
bacteria directs maturation of the host immune system. Cell 2005;122:107-118.
(15) Mazmanian SK, Round JL, Kasper DL. A microbial symbiosis factor prevents intestinal inflammatory
disease. Nature 2008;453:620-625.
(16) Sudo N, Sawamura S, Tanaka K, Aiba Y, Kubo C, Koga Y. The requirement of intestinal bacterial flora for
the development of an IgE production system fully susceptible to oral tolerance induction. J.Immunol.
(17) Butler JE, Weber P, Sinkora M, Baker D, Schoenherr A, Mayer B, et al. Antibody repertoire development
in fetal and neonatal piglets. VIII. Colonization is required for newborn piglets to make serum antibodies to T-
dependent and type 2 T-independent antigens. J.Immunol. 2002;169:6822-6830.
(18) Grönlund MM, Arvilommi H, Kero P, Lehtonen OP, Isolauri E. Importance of intestinal colonisation in the
maturation of humoral immunity in early infancy: a prospective follow up study of healthy infants aged 0-6
months. Arch.Dis.Child.Fetal Neonatal Ed. 2000;83:186-92.
(19) Marschan E, Kuitunen M, Kukkonen K, Poussa T, Sarnesto A, Haahtela T, et al. Probiotics in infancy
induce protective immune profiles that are characteristic for chronic low-grade inflammation. Clin.Exp.Allergy
(20) Martino DJ, Currie H, Taylor A, Conway P, Prescott SL. Relationship between early intestinal
colonization, mucosal immunoglobulin A production and systemic immune development. Clin.Exp.Allergy
(21) Medzhitov R. Recognition of microorganisms and activation of the immune response. Nature
(22) Macatonia SE, Hosken NA, Litton M, Vieira P, Hsieh CS, Culpepper JA, et al. Dendritic cells produce IL-
12 and direct the development of Th1 cells from naive CD4+ T cells. J.Immunol. 1995;154:5071-5079.
(23) Kaisho T, Akira S. Toll-like receptor function and signaling. J.Allergy Clin.Immunol. 2006;117:979-87.
(24) Lundell AC, Adlerberth I, Lindberg E, Karlsson H, Ekberg S, Åberg N, et al. Increased levels of circulating
soluble CD14 but not CD83 in infants are associated with early intestinal colonization with Staphylococcus
aureus. Clin.Exp.Allergy 2007;37:62-71.
(25) Böttcher MF, Häggström P, Björkstén B, Jenmalm MC. Total and allergen-specific immunoglobulin A
levels in saliva in relation to the development of allergy in infants up to 2 years of age. Clin.Exp.Allergy
(26) Sepp E, Julge K, Vasar M, Naaber P, Björkstén B, Mikelsaar M. Intestinal microflora of Estonian and
Swedish infants. Acta Paediatr. 1997;86:956-961.
(27) Ege MJ, Bieli C, Frei R, van Strien RT, Riedler J, Ublagger E, et al. Prenatal farm exposure is related to the
expression of receptors of the innate immunity and to atopic sensitization in school-age children. J.Allergy
(28) Lauener RP, Birchler T, Adamski J, Braun-Fahrländer C, Bufe A, Herz U, et al. Expression of CD14 and
Toll-like receptor 2 in farmers' and non-farmers' children. Lancet 2002;360:465-466.
(29) Voor T, Julge K, Böttcher MF, Jenmalm MC, Duchén K, Björkstén B. Atopic sensitization and atopic
dermatitis in Estonian and Swedish infants. Clin.Exp.Allergy 2005;35:153-159.
(30) Böttcher MF, Jenmalm MC, Voor T, Julge K, Holt PG, Björkstén B. Cytokine responses to allergens
during the first 2 years of life in Estonian and Swedish children. Clin.Exp.Allergy 2006;36:619-628.
(31) Lundell AC, Hesselmar B, Nordström I, Saalman R, Karlsson H, Lindberg E, et al. High circulating
immunoglobulin A levels in infants are associated with intestinal toxigenic Staphylococcus aureus and a lower
frequency of eczema. Clin.Exp.Allergy 2009;39:662-70.
(32) Lodinova-Zadnikova R, Slavikova M, Tlaskalova-Hogenova H, Adlerberth I, Hanson LA, Wold A, et al.
The antibody response in breast-fed and non-breast-fed infants after artificial colonization of the intestine with
Escherichia coli O83. Pediatr.Res. 1991;29:396-399.
(33) Cerutti A. The regulation of IgA class switching. Nat.Rev.Immunol. 2008;8:421-434.
(34) He B, Xu W, Santini PA, Polydorides AD, Chiu A, Estrella J, et al. Intestinal bacteria trigger T cell-
independent immunoglobulin A(2) class switching by inducing epithelial-cell secretion of the cytokine APRIL.
(35) Brandtzaeg P. Do salivary antibodies reliably reflect both mucosal and systemic immunity?
(36) Suzuki S, Shimojo N, Tajiri Y, Kumemura M, Kohno Y. Differences in the composition of intestinal
Bifidobacterium species and the development of allergic diseases in infants in rural Japan. Clin.Exp.Allergy
(37) He F, Ouwehand AC, Isolauri E, Hashimoto H, Benno Y, Salminen S. Comparison of mucosal adhesion
and species identification of bifidobacteria isolated from healthy and allergic infants. FEMS
(38) Wang M, Karlsson C, Olsson C, Adlerberth I, Wold AE, Strachan DP, et al. Reduced diversity in the early
fecal microbiota of infants with atopic eczema. J.Allergy Clin.Immunol. 2008;121:129-134.
(39) Lappalainen MH, Roponen M, Hyvarinen A, Nevalainen A, Laine O, Pekkanen J, et al. Exposure to
environmental bacteria may have differing effects on tumour necrosis factor-alpha and interleukin-6-producing
capacity in infancy. Clin.Exp.Allergy 2008;38:1483-1492.
(40) Nomura F, Akashi S, Sakao Y, Sato S, Kawai T, Matsumoto M, et al. Cutting edge: endotoxin tolerance in
mouse peritoneal macrophages correlates with down-regulation of surface toll-like receptor 4 expression.
(41) Zhong B, Ma HY, Yang Q, Gu FR, Yin GQ, Xia CM. Decrease in toll-like receptors 2 and 4 in the spleen
of mouse with endotoxic tolerance. Inflamm.Res. 2008;57:252-259.
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