Establishment of normal gut microbiota is compromised under excessive hygiene conditions.

Abstract

Early gut colonization events are purported to have a major impact on the incidence of infectious, inflammatory and autoimmune diseases in later life. Hence, factors which influence this process may have important implications for both human and animal health. Previously, we demonstrated strong influences of early-life environment on gut microbiota composition in adult pigs. Here, we sought to further investigate the impact of limiting microbial exposure during early life on the development of the pig gut microbiota.
Outdoor- and indoor-reared animals, exposed to the microbiota in their natural rearing environment for the first two days of life, were transferred to an isolator facility and adult gut microbial diversity was analyzed by 16S rRNA gene sequencing. From a total of 2,196 high-quality 16S rRNA gene sequences, 440 phylotypes were identified in the outdoor group and 431 phylotypes in the indoor group. The majority of clones were assigned to the four phyla Firmicutes (67.5% of all sequences), Proteobacteria (17.7%), Bacteroidetes (13.5%) and to a lesser extent, Actinobacteria (0.1%). Although the initial maternal and environmental microbial inoculum of isolator-reared animals was identical to that of their naturally-reared littermates, the microbial succession and stabilization events reported previously in naturally-reared outdoor animals did not occur. In contrast, the gut microbiota of isolator-reared animals remained highly diverse containing a large number of distinct phylotypes.
The results documented here indicate that establishment and development of the normal gut microbiota requires continuous microbial exposure during the early stages of life and this process is compromised under conditions of excessive hygiene.

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Establishment of Normal Gut Microbiota Is Compromised
under Excessive Hygiene Conditions
Bettina Schmidt1, Imke E. Mulder1, Corran C. Musk1, Rustam I. Aminov1, Marie Lewis2, Christopher R.
Stokes2, Mick Bailey2, James I. Prosser3, Bhupinder P. Gill4, John R. Pluske5, Denise Kelly1*
1Gut Immunology Group, Rowett Institute of Nutrition and Health, University of Aberdeen, Aberdeen, United Kingdom, 2Veterinary Pathology, Infection and Immunity,
School of Clinical Veterinary Science, University of Bristol, Bristol, United Kingdom, 3Institute of Biological and Environmental Sciences, University of Aberdeen, Aberdeen,
United Kingdom, 4Agricultural and Horticultural Development Board, Milton Keynes, United Kingdom, 5School of Veterinary and Biomedical Sciences, Murdoch
University, Murdoch, Western Australia, Australia
Abstract
Background: Early gut colonization events are purported to have a major impact on the incidence of infectious,
inflammatory and autoimmune diseases in later life. Hence, factors which influence this process may have important
implications for both human and animal health. Previously, we demonstrated strong influences of early-life environment on
gut microbiota composition in adult pigs. Here, we sought to further investigate the impact of limiting microbial exposure
during early life on the development of the pig gut microbiota.
Methodology/Principal Findings: Outdoor- and indoor-reared animals, exposed to the microbiota in their natural rearing
environment for the first two days of life, were transferred to an isolator facility and adult gut microbial diversity was
analyzed by 16S rRNA gene sequencing. From a total of 2,196 high-quality 16S rRNA gene sequences, 440 phylotypes were
identified in the outdoor group and 431 phylotypes in the indoor group. The majority of clones were assigned to the four
phyla Firmicutes (67.5% of all sequences), Proteobacteria (17.7%), Bacteroidetes (13.5%) and to a lesser extent,
Actinobacteria (0.1%). Although the initial maternal and environmental microbial inoculum of isolator-reared animals was
identical to that of their naturally-reared littermates, the microbial succession and stabilization events reported previously in
naturally-reared outdoor animals did not occur. In contrast, the gut microbiota of isolator-reared animals remained highly
diverse containing a large number of distinct phylotypes.
Conclusions/Significance: The results documented here indicate that establishment and development of the normal gut
microbiota requires continuous microbial exposure during the early stages of life and this process is compromised under
conditions of excessive hygiene.
Citation: Schmidt B, Mulder IE, Musk CC, Aminov RI, Lewis M, et al. (2011) Establishment of Normal Gut Microbiota Is Compromised under Excessive Hygiene
Conditions. PLoS ONE 6(12): e28284. doi:10.1371/journal.pone.0028284
Editor: Markus M. Heimesaat, Charite ´, Campus Benjamin Franklin, Germany
Received July 25, 2011; Accepted November 4, 2011; Published December 2, 2011
Copyright: ? 2011 Schmidt et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported by a joint grant from the Department for Environment, Food and Rural Affairs (DEFRA) and the Meat and Livestock
Commission (MLC) to BS and IEM (LS3658/CSA 6738), and the Rural and Environmental Science and Analytical Services (RESAS) to DK and RIA. The funders had no
role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: D.Kelly@abdn.ac.uk
Introduction
The mammalian gut is colonized by a highly complex, diverse
and dynamic microbiota. Although considered sterile during
gestation, at delivery the gut is exposed to microbes during passage
through the birth canal. ‘Environmental’ bacteria are then
ingested from the vagina, feces, skin and the early-life environment
[1]. Following birth, bacterial transfer to the neonatal intestine is
continuous throughout the suckling and nursing periods. The
resulting microbiota is very diverse and reflects the microbial
communities associated with the birth and rearing environments,
as well as maternal contact [2,3].
Convergence towards a stable commensal gut microbiota is
thought to be established in adult life [4], and although significant
temporal variability in the microbiota has recently been
documented [5], microbiota composition undoubtedly has life-
long consequences for the host [6]. Experimental evidence has
highlighted its crucial role in regulating complex mechanisms of
host development, lipid metabolism, pathogen response, tissue
repair and immune homeostasis [7,8,9,10,11,12,13]. Both host-
dependent and host-independent factors affect microbial compo-
sition and include host genetics, nutrition, mode of delivery,
gestational age, rearing environment and antibiotic exposure
[14,15,16]. For example, microbial colonization in infants
delivered by caesarean section occurs later than in naturally-
delivered infants and compositional differences in intestinal
microbiota appear to persist throughout life [14,17]. Rearing
environment and exposure to antibiotics also have profound effects
on the adult gut mucosa-adherent microbiota and immune
development in the pig [18]. Analysis of 16S rRNA gene
sequences has revealed major differences in mucosa-adherent
microbial diversity in the ileum of adult pigs reared in different
environments [18]. The gut microbiota of pigs housed in natural
outdoor environments was dominated by Firmicutes, in particular
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by Lactobacillus spp., whereas animals housed under hygienic
conditions in indoor environments displayed reduced numbers of
lactobacilli and higher numbers of Proteobacteria including
potentially pathogenic phylotypes.
The aim of the current study was to elucidate the impact of
limiting microbial exposure during development, by maintaining
animals in environments of excessive hygiene, on the composition
and dynamics of the adult pig microbiota. Piglets were originally
colonized in outdoor (extensive) and indoor (intensive) rearing
systems with distinct microbial communities and then reared in
high-hygiene isolators. Mucosa-associated ileal microbiota was
analyzed by comparison of 16S rRNA gene sequences from the
two groups.
Materials and Methods
Ethics Statement
All animal studies were performed according to the regulations
and guidance provided under the UK Home Office Animals
(Scientific Procedures) Act 1986. Experimental protocols were
approved by the University of Bristol Ethical Review Group and
the Home Office under project license number PPL 30/2482.
Experimental animals and tissue sampling
Five Large White6Landrace sows (Sus scrofa) were housed either
in an outdoor (extensive, OIs) or an indoor (intensive, InIs) rearing
facility. The sows were artificially inseminated by the same boar to
minimize genetic variation among the offspring. Two days after
birth, two piglets per sow (N=5 per group, ten piglets in total)
were transferred to isolators (specific pathogen-free, positive-
pressure units supplied with a high-efficiency particulate air
(HEPA) filter) through a dunk tank containing 1% w/v
bactericidal and virucidal disinfectant solution (Virkon; Antec
International Ltd, Sudbury, UK). Up until day 28, the piglets were
fed a commercial, bovine milk-formula (Piggimilk; Parnutt Feeds,
Sleaford, UK) dispensed by an automated liquid feeding system.
From day 29 onwards, all piglets were fed creep feed (Multiwean,
SCA NUTRITION Ltd) ad libitum. The experiment was
performed using two consecutive replicates.
At day 56, all piglets were sacrificed by injection of sodium
pentobarbitone (Euthesate, Willows Francis Veterinary Ltd). The
ileum, defined as the region corresponding to 75% in length from
the pyloric sphincter, was excised. 16S rRNA gene libraries were
constructed using bacterial DNA derived from this site as it
represents a key region involved in microbial antigen sampling,
immune induction and effector activity.
Mucosal microbiota analysis
Ileal tissue was cut open and contents were removed. Tissue was
then washed with ice-cold phosphate buffered saline (PBS) and
incubated overnight in ice-cold PBS/0.1% Tween 20 (Sigma-
Aldrich Inc., Gillingham, UK) solution with continuous shaking.
Detached bacteria were harvested by centrifugation at 10,0006g
for 10 min at 4uC. Total DNA from the pellet was isolated using a
DNA Spin Kit for SoilH (QBiogene Inc., Cambridge, UK)
according to the manufacturer’s protocol. PCR amplification of
16S rRNA genes was carried out with the universal primer set S-
D-Bact-0008-a-S-20(59-AGAGTTTGATCMTGGCTCAG-39)
and S-*-Univ-1492-a-A-19 (59-ACGGCTACCTTGTTACGA-
CTT-39) [19]. PCR cycling conditions were: one cycle at 94uC
for 5 min, followed by 25 cycles at 94uC for 30 s, 57uC for 30 s,
72uC for 2 min, with a final extension at 72uC for 10 min. PCR
products were purified with the WizardH SV Gel & PCR Clean-up
System (Promega, Southampton, UK), cloned into the pCR-4
cloning vector and transformed into E. coli TOP 10 chemically-
competent cells according to the manufacturer’s instructions
(TOPO TA Cloning Kit; Invitrogen, Paisley, UK). Recombinant
colonies were picked and archived in 96-well plates. Inserts were
sequenced at the RINH genomics facility (University of Aberdeen,
UK)using theprimer
(59CCGTCAATTCATTTGAGTTT-39) and S-*-Univ-0519-a-
A-18 (59-GWATTACCGCGGCKGCTG-39) [19]. All clone
libraries were constructed under identical conditions to minimize
sample-to-sample variation. The methods used here are prone to
under-sampling, thus the relative differences in gut bacterial
composition between the samples is presented. However, on the
basis of the clone number analysed, the data presented strongly
suggests that microbial diversity was high in both treatment
groups. This data is discussed in the context of diversity analysis of
naturally-reared littermates from the same experiment for which
clone libraries were adequately sampled [18].
setS-*-Univ-0907-a-A-20
Sequence alignment and phylogenetic analysis
The 16S rRNA gene reads were assembled using the Lasergene
6 package (DNASTAR Inc.; Infogen Bioinformatics, Broxburn,
UK). Assembled sequences were tested for possible chimeras using
Chimera Check v2.7 (online analysis at RDP-II website, http://
rdp.cme.msu.edu/) and Bellerophon (http://foo.maths.uq.edu.
au/,huber/bellerophon.pl [20]). Sequences with no close match
in RDP-II were additionally subjected to Basic Local Alignment
Search Tool (BLAST) analysis (http://www.ncbi.nlm.nih.gov/
BLAST). Chimeric and poor quality sequences were excluded
from further phylogenetic analyses.
The resulting 16S rRNA gene sequences were aligned using
Multiple Sequence Comparison by Log-Expectation (MUSCLE,
http://www.ebi.ac.uk /Tools/muscle) and the alignments were
inspected manually. The distance matrix (generated from the
multiple sequence alignment) was calculated using the Dnadist
application of the Phylogeny Inference Package and Jukes-Cantor
distance of 0.01. This stringent phylotype definition at 99% cut-off
was used because evidence suggests that bacteria with nearly-
identical 16S rRNA gene sequences may represent different
phylotypes and species [21].
Rarefaction and collector’s curves of observed phylotypes,
richness estimates and diversity indices were determined with the
DOTURprogram (http://schloss.micro.umass.edu/software/
dotur.html [22]) using Jukes-Cantor corrected distance matrix.
The bias-corrected Chao 1 richness estimator was calculated after
1000 randomizations of sampling without replacement. Collector’s
curves of observed and estimated (Chao 1 and the abundance-
based coverage estimator, ACE) richness were constructed.
Diversity values were estimated using the Shannon (H) and
Simpson indices (D). The Simpson reciprocal index was calculated
as 1/D, and another version of the Simpson diversity index as 1-D.
The Good’s coverage percentage was calculated with the formula
[12(n/N)]6100, where n is the number of phylotypes in a sample
represented by one clone (singletons) and N is the total number of
sequences in that sample [23].
A similarity search of the 16S rRNA gene sequences against
database entries was performed using the Basic Local Alignment
Search Tool (BLAST) program at the National Center for
Biotechnology Information website (http://www.ncbi.nlm.nih.
gov/BLAST). Sequences were assigned to respective bacterial
phylotypes using a .99% sequence similarity criterion.
Phylotype comparisons were made among groups of animals
using the Mann-Whitney U test. Multiple comparisons were
carried out using the Kruskal-Wallis test, with P,0.05 considered
statistically significant.
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Nucleotide sequence accession numbers
The nucleotide sequences obtained in this study were submitted
to the GenBank database under accession numbers JN882713 to
JN884815.
Results
Bacterial diversity is elevated in pigs naturally colonized
but reared in isolators
Rarefaction curves (Fig. 1), which estimates species richness as a
function of the number of clones sampled, were generated by
plotting the number of phylotypes (operational taxonomic units,
OTUs) against the number of newly identified sequences. Neither
of the rarefaction curves reached a plateau at the genus (95%) and
species levels (99%), indicating that even after sampling over 1000
sequences for each treatment group, the number of OTUs was
likely to increase with additional sampling. Rarefaction curves of
the individual animals within the treatment groups are illustrated
in Fig. S1 and S2. Interestingly, within both treatment groups the
individual diversity varied greatly between animals, suggesting
strong genotypic influences. The high number of OTUs
encountered in some individuals contributed to the overall high
diversity observed in both treatment groups. Additional sampling
is required to determine the true gut bacterial diversity in these
adult animals, but the data presented strongly suggests that the
microbial diversity is high in both treatment groups.
Collector’s curves were constructed as plots of the cumulative
number of OTUs recorded as a function of sampling effort (number
of clones sampled from each clone library). Sequences with a
similarity .99% were considered to belong to the same OTU.
Collector’s curves of the observed and estimated phylotype richness
are shown in Fig. 2. Each curve reflects the series of observed or
estimated richness values obtained as clones were added to the data
set in a random order. The curves rose less steeply as a decreasing
proportion of new phylotypes were encountered in the treatment
groups. The number of unseen phylotypes was represented by the
gap between the observed phylotypes and the number of phylotypes
estimated by Chao1 and ACE. The difference between the
estimated and observed phylotype richness was high in both isolator
treatment groups. Novel phylotypes continued to be identified
throughout sampling. The relatively constant estimates of the
number of unobserved phylotypes in each treatment group as
observed richness increased indicated that the estimated richness
was likely to increase further with additional sampling. The overall
speciesrichnessintheOIsgroupwasestimatedat440phylotypesby
Chao1 and 438 by ACE (Table 1 and Fig. 2). Estimated phylotype
richness was slightly lower in the InIs group with 431 phylotypes as
estimated by Chao1 and 416 phylotypes by ACE. Good’s coverage
was 89.5% for the OIs sequence set and 89% for the InIs sequence
set, indicating that eleven additional phylotypes would be expected
for every 100 additional sequenced clones. This contrasts with the
lower diversity values previously reported for naturally-reared
littermates from the same study [18] (Table 1). Compared to their
isolator-reared littermates, naturally-reared OUT and IN animals
had Chao1 values of 254.9 and 259 and ACE values of 208.1 and
280.2, respectively. The lower diversity in these animals was also
reflected in the library coverage, as both libraries had a Good’s
coverage of greater than 92%. Sequences were subjected to BLAST
searches against GenBank entries to assign them to the lower
taxonomic ranks.
Taxonomic placement of sequences into 4 major phyla
The 16S rRNA genes from the mucosa-associated ileal samples
were subjected to the RDP Classifier analysis (with a 95%
confidence threshold). Based on the classification results, the clone
sequences were assigned to four phyla: Firmicutes (67.5% of all
sequences), Proteobacteria (17.7%), Bacteroidetes (13.5%), and
Actinobacteria (0.1%) (Table 2). 1.2% of the sequences remained
unclassified by the RDP Classifier. These results largely corre-
spond to the distribution across the bacterial phyla in the
naturally-reared animals (OUT and IN), as previously reported
[18], where clones were assigned to Firmicutes (69.7% of all
sequences), Proteobacteria (17.7%), Bacteroidetes (11.4%), and
Actinobacteria (0.5%). However, when comparing outdoor sow-
reared and outdoor-isolator reared animals directly, an increase in
Bacteroidetes from 1.08% to 16.5% and Proteobacteria from
4.63% to 19.5%, coinciding with a reduction in Firmicutes from
94% to 62.5%, was noted (Table 2). Within the Firmicutes, the
Lactobacillales were the most affected taxon with a reduction from
81.6% to 15.2%. When comparing indoor sow-reared and indoor-
isolator reared animals, an increase in Bacteroidetes (3.72% to
10.2%) and a decrease in Proteobacteria (28.26% to 15.7%) was
observed.
Firmicutes.
67.5% of all sequences (1161 clones) were
affiliated with the Firmicutes phylum. Bacilli (41.8%) and
Clostridia (56.5%) were the most abundant bacterial classes
within this phylum, with Erysipelotrichi (1.5%) in low abundance
(Table 2).
The most abundant order in the Bacilli class was Lactobacillales
(480 clones), including Lactobacillacaeae, Leuconostocaceae, Streptococca-
ceae and, in lower abundance, Enterococcaceae and Aerococcaceae
(Fig. 3).
The Lactobacillaceae family in the OIs group (7.8% of OIs
sequences) consisted of only a small number of OTUs, including
Lactobacillus reuteri, L. amylovorous, L. johnsonii, L. brevis, L. pentosus and
L. plantarum. The InIs library contained 27.4% Lactobacillaceae-
affiliated clones, with similar phylotypes to the OIs group
including L. reuteri, L. amylovorous, L. johnsonii, L. brevis, L. pentosus
and L. plantarum. An additional phylotype, not detected in the OIs
group, was Pediococcus pentosaceus (CP000422) (Fig. 4).
Leuconostocaceae were represented by a total of 150 clones, 101 of
which were obtained from the InIs group and the remaining from
the OIs group. Within the InIs group, three OTUs were present.
All sequences had 99% similarity to Weissella paramesenteroides
(AB362621), W. hellenica (AB015642) and, in lower abundance, W.
cibaria (AB362617). Interestingly, in the OIs group only animals in
replicate 1 possessed members of the Leuconostocaceae family.
Similar phylotypes were obtained from the OIs group including
W. hellenica (AB015642) and two phylotypes related to W.
paramesenteroides (AB362621). W. paramesenteroides strain CTSPL5
(EU855224) was the predominant phylotype with 27 clones. None
of these phylotypes showed a significant difference between the
two treatment groups.
Streptococcaceae were represented in lower abundance with a total
of 87 clones, 70 of which were detected in the InIs group and the
remaining in the OIs group. Strains belonging to Streptococcus suis
(AF009481), Str. thermophilus strain Y-2 (DQ911624), Str. parauberis
(FJ009631) and Lactococcus lactis subsp. lactic (AE005176) were
identified. Uncultured bacterium clone SQ_aah81g09, which
possessed 99% similarity to Str. gallolyticus, was found in high
abundance in the InIs group (22 clones). Both Str. suis and Str.
gallolyticus have been implicated in a wide variety of infections in
pigs including pneumonia and septicemia [24,25] and have also
been described as human pathogens [26].
From 20 Enterococcaceae-related clones, 18 were detected in the
InIs group. Enterococcus gallinarum strain 22B (EF025908) was the
most abundant phylotype. E. gallinarum, a motile bacterium, is
primarily found in the gastrointestinal tract and in food products
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[27] and plays a role in invasive infections in humans, especially in
immune-compromised or chronically ill patients [28,29,30]. Four
additional OTUs were identified as Enterococcus sp. DJF_O30
(EU728749), E. avium (AF133535), E. faecalis (AB362601) and E.
italicus strain 1102 (EF535230). Two sequences related to E. faecalis
and E. italicus were identified in the OIs group. Due to high
variation between animals in the treatment groups, no statistically
significant difference in the proportion of Enterococci was
observed.
Clostridia.
Members of the Clostridia class were present in
all treatment groups (Table 2). The OIs group showed a higher
abundance of this class. A total of 353 clones were grouped into
Lachnospiraceae, with 140 clones obtained from the InIs group and
the remaining from the OIs group (Fig. 3). The Lachnospiraceae
family was represented by a large number of OTUs, mainly
uncultured clones. Sixteen distinct OTUs had less than 97%
sequence similarity to database entries. The most abundant
members obtained from the InIs group included uncultured
bacterium clone p-2176-s59-3 (AF371605), Eubacteriaceae bac-
teriumDJF_CR57k1(EU728737)
(AF371541). Similar phylotypes were detected in the OIs group
in equal numbers. The Peptostreptococcaceae family was another
abundant member of the Clostridia class, accounting for 7% of the
sequenced clones (120 sequences). Thirty-two clones originated
from the InIs group (3.9% of InIs sequences) and 88 clones from
the OIs group (9.7% of OIs sequences). In the InIs group, 32
clones were obtained exclusively from replicate 1 animals. One
predominant phylotype had 99% identity to uncultured bacterium
clone VWP_aaa01b10 (EU475070), isolated from the feces of
Visayan warty pigs [31]. This clone was also found in high
and clonep-2482-18B5
Figure 1. 16S rRNA gene library rarefaction curves from isolator-reared animals at multiple OTU cutoff levels. Rarefaction curves were
generated by plotting the number of phylotypes (OTUs) against the number of clones sequenced. At 99% cut-off, rarefaction analysis of clone
libraries suggested that both the InIs (A) and OIs (B) group possessed a highly diverse mucosa-associated bacterial community. Clearly, even after
sampling .1000 clones for each treatment group, the number of OTUs continued to increase even as OTU definitions relaxed towards 95% (genus
level).
doi:10.1371/journal.pone.0028284.g001
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abundance in the OIs group. Interestingly, this clone was only
recovered in high abundance in animals from replicate 1 and was
not obtained from replicate 2 animals. Replicate 2 animals shared
similar phylotypes distinct from replicate 1 animals including
uncultured bacterium clone BARB_aaa01h10 (EU475654) and
uncultured bacterium clone aaa02d03 (EU475689). Six percent of
all sequences were grouped into Ruminococcaceae and were mainly
represented by a diverse range of uncultured clones. The most
abundant clones included uncultured bacterium clone SJTU_
C_03_14 (EF403979), which was also obtained from the OIs
group, and uncultured bacterium clone p-2609-9F5 (AF371720) in
the OIs group. Eighteen clones were affiliated with the
Veillonellaceae
family, including
(EU779292) in the InIs group and clone D19 (AM500725) in
the OIs group. Thirty-nine clones belonging to the Clostridiaceae
family were obtained from both groups and were mainly
cloneVWP_aaa01c05
Figure 2. 16S rRNA gene library collector’s curves from indoor isolator-reared and outdoor isolator-reared animals. Collector’s curves
of the observed (black) and estimated (Chao1 (blue) and ACE (pink)) phylotype richness calculated at 99% OTU cut-off level from indoor isolator-
reared (A) and outdoor isolator-reared animals (B). The relatively constant estimates of the number of unobserved phylotypes in each treatment
group as observed richness increases indicate that estimated richness is likely to increase further with additional sampling.
doi:10.1371/journal.pone.0028284.g002
Table 1. Indices of diversity, richness and library coverage for
16S rRNA gene libraries (N=5).
MeasurementInIsOIsIN[18]
OUT[18]
Chao1 estimator of species richness431.2440.9259.0254
ACE abundance estimator416.2438.5280.2 208
Shannon diversity index (H)4.54.9 4.44.2
Simpson diversity index (1-D)0.98 0.99 0.980.97
Simpson reciprocal index (1/D)44.689.652.040.4
Good’s estimator of coverage (%) 89%89.5%93.592.5
Calculations were made based on OTU definition at 99% sequence identity.
doi:10.1371/journal.pone.0028284.t001
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represented by two clones including VWP_aaa03f12 (EU779318)
and RL179_aao56e08 (DQ796928). For all three bacterial families
(Veillonellaceae, Clostridiaceae and Ruminococcaceae) the majority of
clones were obtained from replicate 1 animals in both treatment
groups.
Bacteroidetes.
All 16S rRNA gene libraries contained
sequences related to Prevotellaceae, yet they were most prevalent
in the OIs group (Fig. 5). Thirteen OTUs encountered had less
than 97% similarity to database entries. Abundant members were
identified in both libraries, independent from the farm origin,
including uncultured bacterium clone p-1980-s959-5 (AF371890),
uncultured bacterium clone SPIM_d08_1 (EU467242), uncultured
bacterium clone p-2190-s959-3 (AF371875) and uncultured
bacterium clones ML_aaj26e06 and ML_aae88g11 (EU776786/
EU467242). Thirty-six additional OTUs were in low abundance,
all with a close similarity to uncultured bacterial clones, thereby
contributing to the high diversity in the isolator-reared animals.
The 19 clones classified as Bacteroidaceae were obtained from both
treatment groups (Fig. 5). Fifteen clones were retrieved from the
InIs libraries and four sequences from the OIs libraries. In the InIs
libraries, eight OTUs were present related to Bacteroides vulgatus
(CP000139) and B. plebius (AB200217). Fourteen clones affiliated
with Porphyromonadaceae were obtained exclusively from the OIs
libraries from two animals in the same replicate (Fig. 5). All 14
clones had only 97% similarity or less to previously isolated clones.
Proteobacteria.
sequences)
proteobacteria (302 clones) being the most abundant group.
Most of the clones classified as Pasteurellaceae (Table 2 and Fig. 6).
177 clones were obtained from the OIs and 127 clones from the
InIs mucosa-adherent libraries. Phylotype distribution was very
similar between the two treatment groups. The majority of
sequences were affiliated with Actinobacillus minor, A. porcinus and the
low abundance sequences with A. rossii. This clone has been
isolated from the intestine and reproductive tract of pigs and is
consideredasan opportunistic
spontaneous abortion. Within the Enterobacteriaceae family the
most abundant clones were identified as E. coli spp.
Overall,
placed
17.7%
into
oftheclones(304
wereProteobacteria,with
c-
pathogen implicatedin
Discussion
The mucosal immune system of the pig undergoes rapid
changes during the neonatal period, similar to humans [32,33].
Hence, the pig is increasingly utilized as a translational model
[34,35,36]. We sought to evaluate the impact of limiting microbial
exposure during development on the composition of the pig gut
microbiota. Animals were naturally colonized during the first two
days after birth and then reared in isolators maintained to a very
high-hygiene status. Initially, all piglets remained with the sows,
housed in either indoor or outdoor environments, to promote
Table 2. Taxonomic composition of the mucosa-associated microbiota of indoor and outdoor isolator-reared animals and their
sow-reared counterparts.
PhylumBacterial taxa InIs OIs IN[18]
OUT[18]
% Bacteroidetes10.216.53.721.08
Family Prevotellaceae (%) 9.513.02.910.54
Family Bacteroidaceae (%) 0.70.20.400
Family Porphyromonadaceae (%)0 2.900.40
% Proteobacteria15.719.528.264.63
Class a-proteobacteria (%)0.3000.13
Class b-proteobacteria (%)000.200
Class c-proteobacteria (%)15.419.515.193.81
Family Pasteurellaceae (%)9.114.914.782.17
Family Enterobacteriaceae (%) 5.94.20.41.36
Family Pseudomonadaceae (%)0 0.300
Family Moraxellaceae (%) 0.2000
Class e-proteobacteria (%)0012.97 0.4
Family Helicobacteraceae (%)0010.46 0.4
Family Campylobacteraceae (%) 0.102.610
% Firmicutes73.162.566.2994.0
Class Erysipelotrichi (%) 1.20.80.900
Class Bacilli (%)42.815.218.8181.8
Order Bacillales (%) 0.60 0.20.2
Order Lactobacillales (%)42.215.218.681.6
Class Clostridia (%)28.746.546.6812.12
Family Lachnospiraceae (%)17.423.43.520.95
Family Veillonellaceae (%) 1.0 1.100.4
Family Clostridiaceae (%)2.8 2.313.172.72
Family Peptostreptococcaceae (%)3.9 9.7 24.447.49
Family Ruminococcaceae (%)2.7 8.80.900.54
doi:10.1371/journal.pone.0028284.t002
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‘natural’ (environmental and maternal) microbial colonization and
to ensure adequate colostrum intake during the first few days of
life.
Intriguingly, the 16S rRNA gene sequences generated from adult
isolator-reared pigs initially colonized during the early days of life
revealed a highly diverse microbiota which included some well-
Figure 3. Phylogenetic distribution and abundance of the Firmicutes phylum in the mucosa-associated microbiota from isolator-
reared animals (N=5).
doi:10.1371/journal.pone.0028284.g003
Figure 4. Abundance of Lactobacillaceae in the mucosa-adherent microbiota of isolator-reared animals (N=5).
doi:10.1371/journal.pone.0028284.g004
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Figure 5. Phylogenetic distribution and abundance of Bacteroidetes in the mucosa-associated microbiota of isolator-reared animals
(N=5).
doi:10.1371/journal.pone.0028284.g005
Figure 6. Phylogenetic distribution and abundance of Proteobacteria in the mucosa-associated microbiota of isolator-reared
animals (N=5).
doi:10.1371/journal.pone.0028284.g006
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known members of the mammalian gastrointestinal tract as well as
previously uncultured phylotypes. Although not well described in
the literature, the high sequence numbers and variability within the
Ruminococcaceae, Lachnospiraceae and Peptostreptoccocaceae families cer-
tainly contributed to the overall microbial diversity of these animals.
As a general concept, high microbial diversity is thought to be
beneficial to the ecosystem by reducing the opportunity for
colonization by infectious agents [37]. In the gut ecosystem, for
example, such rich diversity would yield a broad range of immune-
triggering compounds required to promote the development of the
mucosal immune system [38]. At the early stages of microbial
colonization/succession, high microbial diversity, largely reflecting
the birth environment, is to be expected as there are few barriers
limiting entry of bacteria into the gut ecosystem [39]. However,
development of the normal pig microbiota coincides with a natural
stabilization of the gut bacterial populations [4,16,35,40,41]
imposed by the strict environmental selection pressures operating
within the gut ecosystem. In agreement with this, we previously
showed that continuous outdoor environmental exposure in a
highly diverse ecosystem resulted in a stable gut microbiota with a
lower diversity than in indoor, intensively-reared and isolator-
reared littermates [18]. The increased mucosal diversity in isolator-
reared animals would therefore suggest that environmental and
immune-related control of the mucosa-adherent microbiota was
reduced by isolation of these animals and succession to the normal
stable microbiota was not achieved, with the microbiota remaining
more chaotic. Furthermore, despite the fact that the animals
originated from distinct microbial rearing environments, no
significant differences in the overall microbial composition were
observed, although this may also reflect the need for additional
sequence information.
In terms of species composition, previous studies have shown
that the intestine of neonatal piglets is initially colonized by large
numbers of E. coli and Streptococcus spp. [42]. Generally,
Enterobacteriaceae are considered to be the early colonizers of the
gut [43,44] and are associated with the mucus layer [45].
Streptococcus spp. were also identified in the microbiota of
isolator-reared treatment groups in the current study, but
Enterobacteria were only rarely recovered.
L. reuteri, L. amylovorous, L. johnsonii, L. brevis, L. pentosus and L.
plantarum were found in isolator-reared animals although they were
present in lower numbers relative to littermates reared-outdoors
[18]. Leser et al. [46] reported a similar range of phylotypes
associated with the ileum in pigs from different rearing
environments. Furthermore, developmental shifts in the dominant
lactobacilli species in the pig gut have also been documented [47].
Hence, certain species of lactobacilli may be better adapted to the
gut at the various developmental stages and following dietary
change. In the current study, the lactobacilli acquired at the very
early life stages may not have been sufficiently adapted to the post-
weaning gut environment and isolator-reared animals were
restricted in their opportunity to acquire other, more adapted
species, unlike their outdoor-reared littermates.
Taken together, the experimental evidence presented in this
study illustrates that development in environments of excessive
hygiene hinders the progression towards an adult-type gut
microbiota, despite the acquisition of a highly diverse microbiota
in early life. In particular, we noted that Firmicutes were reduced
in isolator-reared animals when compared to outdoor-reared
littermates. Conversely, Bacteroidetes and Proteobacteria were
increased in isolator conditions. This identifies early life as a
crucial developmental period during which continual exposure to
environmental microbes is required to drive the ‘stabilization’ of
the gut microbiota towards an adult phenotype. Consistent with
this viewpoint, the gut microbiota in childhood is generally
considered unstable and highly susceptible to environmental
influences. Given the dramatic increases in the incidence of
immune-mediated diseases in Western society and the strong
association with altered microbial diversity [10,48] it is important
to consider that the microbiota of children in Western countries is
adversely affected and limited by low microbial diversity in the
environment. Recent evidence has emerged that children
migrating from the developing world to the Western world take
on the same susceptibility risk to IBD as the population of the
adoptive country, unlike adult migrants [49]. Clearly, lifestyle and
hygiene alter gut microbial diversity, but equally loss of important
ancestral microbes from our environments may have important
health consequences [50]. The current work strengthens the
notion that optimal acquisition of the adult microbiota requires
continuous microbial exposure, biodiverse environmental ecosys-
tems and processes of selection, succession and stabilization in the
context of the developing and maturing gut. Future work focusing
on childhood microbiota development in diverse environments is
important in defining the optimum microbiota and the natural
successional patterns of the adult microbiota. This knowledge may
provide greater insight into the importance of microbial
biodiversity and reversal of current human disease trends.
Supporting Information
Figure S1
curves from outdoor isolator-reared animals (OIs;
N=5). Rarefaction curves were generated by plotting the number
of phylotypes (OTUs) against the number of clones sequenced. At
99% cut-off, rarefaction analysis suggested that the individual
animals within the OIs group possessed a highly diverse mucosa-
associated bacterial community.
(TIF)
Individual 16S rRNA gene library rarefaction
Figure S2
curves from indoor isolator-reared animals (InIs; N=5).
Rarefaction curves were generated by plotting the number of
phylotypes (OTUs) against the number of clones sequenced. At
99% cut-off, rarefaction analysis suggested that the individual
animals within the InIs possessed a highly diverse mucosa-
associated bacterial community.
(TIF)
Individual 16S rRNA gene library rarefaction
Acknowledgments
We thank Pauline Young at the RINH Genomics Facility (University of
Aberdeen) and Beth Logan from the Gut Immunology Group (RINH,
University of Aberdeen) for sequencing of bacterial clones.
Author Contributions
Conceived and designed the experiments: DK CRS MB JRP BPG.
Performed the experiments: BS CRS ML MB. Analyzed the data: BS
CCM. Wrote the paper: BS IEM DK. Technical and scientific discussion
of the project: RIA JIP.
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