Content uploaded by Maria Leonor Fernandez Murga
Author content
All content in this area was uploaded by Maria Leonor Fernandez Murga on Aug 26, 2015
Content may be subject to copyright.
Published Ahead of Print 8 July 2013.
10.1128/AEM.00869-13.
2013, 79(18):5472. DOI:Appl. Environ. Microbiol.
Leonor Fernández-Murga and Yolanda Sanz
Ester Sánchez, Ester Donat, Carmen Ribes-Koninckx, Maria
with Celiac Disease in Children
Duodenal-Mucosal Bacteria Associated
http://aem.asm.org/content/79/18/5472
Updated information and services can be found at:
These include:
REFERENCES
http://aem.asm.org/content/79/18/5472#ref-list-1at:
This article cites 53 articles, 16 of which can be accessed free
CONTENT ALERTS
more»articles cite this article),
Receive: RSS Feeds, eTOCs, free email alerts (when new
http://journals.asm.org/site/misc/reprints.xhtmlInformation about commercial reprint orders:
http://journals.asm.org/site/subscriptions/To subscribe to to another ASM Journal go to:
on June 10, 2014 by guesthttp://aem.asm.org/Downloaded from on June 10, 2014 by guesthttp://aem.asm.org/Downloaded from
Duodenal-Mucosal Bacteria Associated with Celiac Disease in Children
Ester Sánchez,
a
Ester Donat,
b
Carmen Ribes-Koninckx,
b
Maria Leonor Fernández-Murga,
a
Yolanda Sanz
a
Microbial Ecology and Nutrition Research Group, Institute of Agrochemistry and Food Technology, National Research Council (IATA-CSIC), Valencia, Spain
a
; Hospital
Universitario La Fe, Valencia, Spain
b
Celiac disease (CD) is an immune-mediated enteropathy triggered by the ingestion of cereal gluten proteins. This disorder is
associated with imbalances in the gut microbiota composition that could be involved in the pathogenesis of CD. The aim of this
study was to characterize the composition and diversity of the cultivable duodenal mucosa-associated bacteria of CD patients
and control children. Duodenal biopsy specimens from patients with active disease on a gluten-containing diet (n ⴝ 32), patients
with nonactive disease after adherence to a gluten-free diet (n ⴝ 17), and controls (n ⴝ 8) were homogenized and plated on plate
count agar, Wilkins-Chalgren agar, brain heart agar, or yeast, Casitone, and fatty acid agar. The isolates were identified by partial
16S rRNA gene sequencing. Renyi diversity profiles showed the highest diversity values for active CD patients, followed by non-
active CD patients and control individuals. Members of the phylum Proteobacteria were more abundant in patients with active
CD than in the other child groups, while those of the phylum Firmicutes were less abundant. Members of the families Enterobac-
teriaceae and Staphylococcaceae, particularly the species Klebsiella oxytoca, Staphylococcus epidermidis, and Staphylococcus
pasteuri, were more abundant in patients with active disease than in controls. In contrast, members of the family Streptococ-
caceae were less abundant in patients with active CD than in controls. Furthermore, isolates of the Streptococcus anginosus and
Streptococcus mutans groups were more abundant in controls than in both CD patient groups, regardless of inflammatory sta-
tus. The findings indicated that the disease is associated with the overgrowth of possible pathobionts that exclude symbionts or
commensals that are characteristic of the healthy small intestinal microbiota.
C
eliac disease (CD) is a chronic intestinal disorder caused by a
deregulated immune response to gluten proteins from wheat,
rye, and barley and their cross-related varieties in genetically sus-
ceptible individuals. CD presents a set of diverse clinical features,
which typically includes fatigue, weight loss, diarrhea, and ane-
mia. Damage to the intestinal mucosa in patients with CD is char-
acterized by intraepithelial lymphocytosis, crypt hyperplasia, and
villous atrophy (1). In CD patients, the pathological response to
gluten proteins involves both adaptive and innate immunity. It is
known that gliadin-specific CD4
⫹
T cells develop an inflamma-
tory reaction by production of Th1 cytokines (e.g., gamma inter-
feron [IFN-␥]) at the mucosal level, which also induces CD8
⫹
cells to kill epithelial cells, contributing to tissue damage (2). In
addition, a new subset of T cells, termed Th17 cells, was shown to
contribute to CD pathogenesis by producing proinflammatory
cytokines (such as interleukin-17 [IL-17], IFN-␥, and IL-21), al-
though these cells can also produce mucosa-protective and regu-
latory factors (IL-22 and transforming growth factor )(3, 4).
Some gluten peptides that are not recognized by T cells can induce
tissue damage by activating components of innate immunity;
thus, peptide p31-43/49 activates the production of IL-15 and
natural killer cell receptor-mediated cytotoxicity by intraepithelial
lymphocytes, contributing to tissue injury (5–7). Improvement of
the pathological lesions occurring in the intestinal mucosa of
sensitive individuals is usually observed after gluten with-
drawal from the diet; however, compliance with this dietary
recommendation is complex, and other alternative strategies
are being investigated (8).
HLA class II molecules DQ2 and DQ8 are the major risk factors
predisposing individuals to CD and account for 35% of the genetic
risk (9). Although the role of these molecules has been well estab-
lished in the pathogenesis of CD, their frequency in the general
population is approximately 30%, whereas only 1 to 3% of indi-
viduals actually develop the disease (10). These data would suggest
that the presence of HLA molecules is a necessary factor but is not
sufficient alone for disease development. Although gluten is the
main environmental trigger of CD, its intake does not fully explain
disease development, and thus, other environmental factors are
thought to be involved. In recent years, early microbial infections
(11, 12) and imbalances in the composition of the gastrointestinal
microbiota (13–20) have also been associated with CD. Molecular
techniques have shown that, compared to the fecal and duodenal
microbiota of healthy individuals, the fecal and duodenal micro-
biota of CD patients is characterized by the presence of higher
numbers of Gram-negative bacteria (bacteroides and enterobac-
teria) and lower numbers of Gram-positive bacteria, like bifido-
bacteria (19, 20). In vitro assays have shown that this altered mi-
crobiota and some enterobacteria isolated from CD patients could
activate proinflammatory pathways, while some bifidobacteria
could inhibit the inflammatory or toxic effects induced by the
same isolated enterobacteria and gluten peptides (21–24). Altera-
tions in the intestinal microbiota are also involved in the patho-
genesis of chronic inflammatory bowel disease (IBD) (25, 26) and
other immune-related disorders (27–29). For instance, IBD pa-
tients have altered duodenal bacterial populations in comparison
to healthy controls (30–32). Nevertheless, neither the specific bac-
teria involved in pathologies affecting the small intestine nor their
possible pathogenic modes of action are fully understood.
Received 16 March 2013 Accepted 23 June 2013
Published ahead of print 8 July 2013
Address correspondence to Yolanda Sanz, yolsanz@iata.csic.es.
Copyright © 2013, American Society for Microbiology. All Rights Reserved.
doi:10.1128/AEM.00869-13
5472 aem.asm.org Applied and Environmental Microbiology p. 5472–5479 September 2013 Volume 79 Number 18
on June 10, 2014 by guesthttp://aem.asm.org/Downloaded from
This study was designed to establish whether live culture-de-
pendent bacteria associated with the duodenal mucosa of patients
with active and nonactive CD and controls differ in composition
and biodiversity, as reported in previous molecular studies, with a
view to exploring their potentially pathogenic features in the fu-
ture.
MATERIALS AND METHODS
Subjects. Biopsy samples from three groups of children were included in
this study: 32 from patients with active CD (mean age, 5.1 years; range, 2
to 14 years) on a normal gluten-containing diet, 17 from patients with
nonactive CD (mean age, 5.9 years; range, 3 to 8 years) after following a
gluten-free diet for at least 2 years, and 8 from control children (mean age,
6.9 years; range, 3 to 13 years) with no known gluten intolerance. The
control group consisted of children who were investigated for weight loss,
growth retardation, or functional intestinal disorders of non-CD origin;
and their non-CD status was confirmed by showing a normal villous
structure by examination of the biopsy specimen. CD was diagnosed ac-
cording to the criteria given by the European Society for Pediatric Gastro-
enterology, Hepatology and Nutrition (33). The children included in the
study had not been treated with antibiotics for at least 1 month before
sampling.
The study was conducted in accordance with the ethical rules of the
Helsinki Declaration (Hong Kong revision, September 1989), according
to EEC Good Clinical Practice guidelines (document 111/3976/88, July
1990), and under the guidelines of current Spanish law which regulates
clinical research in humans (Royal Decree 561/1993). The study protocol
was approved by the Committee on Ethical Practice from CSIC and the
Hospital Universitario La Fe (Valencia, Spain). Written informed consent
was obtained from the parents of the children included in the study. The
clinical characteristics of the children are shown in Table 1.
Sample preparation and bacterial isolation. Duodenal biopsy speci-
mens (approximately 10 mg) were obtained by capsule endoscopy, kept
under anaerobic conditions (AnaeroGen; Oxoid, Hampshire, United
Kingdom), and analyzed in less than2htoavoid alterations in bacterial
viability. Biopsy specimens were homogenized in 200 l of a phosphate-
buffered saline (PBS) solution (130 mM sodium chloride, 10 mM sodium
phosphate, 0.05% cysteine, pH 7.2) by pipetting and thorough agitation
in a vortex mixer (10 s). Each homogenized sample was randomly plated
on two different culture media (100 l).
The following media were used: plate count agar (PCA; Scharlau, Bar-
celona, Spain) (34), Wilkins-Chalgren agar (Scharlau, Barcelona, Spain)
(35), brain heart agar (BH; Scharlau, Barcelona, Spain) (36), and yeast,
Casitone, and fatty acid agar (YCFA) (37). PCA plates were incubated
under aerobic conditions at 37°C for 48 h, whereas Wilkins-Chalgren, BH,
and YFCA plates were incubated under anaerobic conditions at 37°C for
48 h using anaerobic jars and an AnaeroGen system (Oxoid, Hampshire,
United Kingdom), which generates an atmosphere of ⬍1% oxygen sup-
plemented with carbon dioxide within 30 min, facilitating the culture of
fastidious and obligate anaerobes. All the viable and cultivable bacteria
recovered from duodenal biopsy samples (mucus and mucosa-associated
bacteria) were isolated and restreaked onto the same agar media. For
preliminary identification of the isolates, conventional microbiological
methods were used, including analysis of colony and cellular morphology
and Gram staining. All isolates were stored at ⫺80°C in the presence of
glycerol (20%, vol/vol) until use for further characterization.
DNA extraction. For DNA extraction, bacterial isolates were grown in
the same isolation broth media and harvested at the late log growth phase.
The bacterial suspensions were centrifuged for 5 min at 6,000 ⫻ g, the
pellets were resuspended in 100 l of suspension buffer (10 mM Tris-HCl,
1 mM sodium EDTA, pH 8.0) with lysozyme (50 mg/ml) (Sigma, St.
Louis, MO), and the homogenates were incubated at 37°C for 1 h. The
bacterial DNA extraction procedure was adapted from a standard cetylt-
rimethylammonium bromide (CTAB) DNA purification method (38).
DNA samples were stored at ⫺20°C until used as the templates for PCR.
Identification of bacterial isolates. The bacterial DNA of each isolate
was partially amplified with 16S rRNA gene target primers 968f (5=-AAC
GCGAAGAACCTTA-3=) and 1401r (5=-CGGTGTGTACAAGACCC-3=)
(39). When necessary, complete 16S rRNA amplification and sequencing
were performed with the primers 27␦-f (5=-AGAGTTTGATCCTGGCTC
AG-3=)(40) and 1401r. Amplification reactions were carried out in a
50-l volume containing 10 mM Tris-HCl (pH 8.3), 2.5 mM MgCl
2
,1
M each primer, 200 M deoxynucleoside triphosphates, and 2.5 U of
Taq polymerase (Ecotaq; Ecogen, Spain). The amplification program was
1 cycle at 94°C for 5 min; 30 cycles at 94°C for 1 min, 50°C for 1 min, and
72°C for 2 min; and finally, 1 cycle at 72°C for 7 min. The amplification
products were subjected to gel electrophoresis in 1% agarose gels, purified
using GFX PCR DNA and a Gel Band DNA purification kit (GE Health-
care, Buckinghamshire, United Kingdom), and sequenced in an ABI
Prism-3130XL genetic analyzer (Applied Biosystems, CA). Search analy-
TABLE 1 Clinical characteristics of study subjects
a
Characteristic Active CD (n ⫽ 32) Nonactive CD (n ⫽ 17) Control (n ⫽ 8)
Mean (SD) age (yr) 5.1 (3.2) 5.9 (1.2) 6.9 (4.2)
No. (%) of study subjects
Sex (M/F) 14 (43.7)/18 (56.3) 8 (47.1)/9 (52.9) 4 (50)/4 (50)
Symptoms
Abdominal pain 5 (15.6) 0 (0) 2 (25)
Diarrhea 3 (9.4) 0 (0) 5 (62.5)
Weight loss 5 (15.6) 3 (17.6) 1 (12.5)
Anemia 9 (28.1) 2 (11.8) 0 (0)
Iron deficiency 17 (53.1) 0 (0) 0 (0)
Presence of antigliadin antibodies (AGA
⫹
)
32 (100) 0 (0) 0 (0)
Presence of antitransglutaminase antibodies (tTG
⫹
)
32 (100) 0 (0) 0 (0)
Duodenal biopsy
b
M0-1 0 (0) 17 (100) 8 (100)
M3 32 (100) 0 (0) 0 (0)
HLA type DQ2 and DQ8 32 (100) 17 (100) NA
c
a
Data are expressed as absolute numbers (percentages related to the total numbers) for all characteristics except age, which is expressed as the mean (standard deviation). M, male;
F, female.
b
Modified Marsh classification of CD (1): M0, normal mucosa; M0-1, infiltrative lesions, seen in patients on a gluten-free diet (suggesting that minimal amounts of gliadin are
being ingested), patients with dermatitis herpetiformis, and family members of patients with CD; M2, hyperplastic type, occasionally seen in patients with dermatitis herpetiformis;
M3, ⬎40 intraepithelial lymphocytes per 100 enterocytes, crypts increased, and villi with atrophy (partial or complete villous atrophy), seen in cases of typical CD.
c
NA, not applicable.
Duodenal-Mucosal Bacteria in Celiac Disease
September 2013 Volume 79 Number 18 aem.asm.org 5473
on June 10, 2014 by guesthttp://aem.asm.org/Downloaded from
ses to determine the closest relatives of the partial 16S rRNA gene se-
quences retrieved were conducted in GenBank using the Basic Local
Alignment Search Tool (BLAST) algorithm, and sequences with more
than 97% similarity were considered to be of the same species.
Data and statistical analyses. The Renyi diversity index was used to
explore differences in the mucosa-associated bacteria among active and
nonactive CD patients and control children. This index provides three
further diversity index values: species richness (S), the Shannon diversity
index (H=), and the Simpson dominance index (1-D), which were deter-
mined using Paleontological Statistics (PAST) software (41).
Differences in the relative abundance of the duodenal mucosa-associ-
ated bacteria (estimated as isolates belonging to a specific taxon related to
all isolates recovered from samples from each child group) were estab-
lished by applying chi-square tests and, when appropriate, the two-tailed
Fisher’s exact test. Analyses were carried out with Statgraphics software
(Manugistics, Rockville, MD), and statistical differences were established
at a P value of less than 0.05.
RESULTS
Subjects. The clinical characteristics of the groups of children in-
cluded in the study are shown in Table 1. No statistically signifi-
cant differences in the gender ratio representation in the study
were detected. Patients with active CD on a normal gluten-con-
taining diet showed clinical symptoms of the disease, positive CD
serology markers (antigliadin antibodies and antitransglutami-
nase antibodies), and signs of severe enteropathy by duodenal
biopsy examination classified as type 3 according to the Marsh
classification of CD (M3) (1). Patients with nonactive CD who
had been on a gluten-free diet for at least 2 years showed negative
CD serology markers and normal mucosa or infiltrative lesions
classified as type 0-1 according to the Marsh classification of CD.
The study included 32 biopsy specimens from children with active
CD (mean age, 5.1 years), 17 biopsy specimens from children with
nonactive CD (mean age, 5.9 years), and finally, 8 biopsy speci-
mens from children without known gluten intolerance (mean age,
6.9 years) who were included in the control group for comparative
purposes.
Influence of culture media on bacterial taxa recovered. Four
different culture media, including PCA, Wilkins-Chalgren agar,
BH, and YFCA, were used for isolating bacteria from biopsy spec-
imens from the CD patients and controls. The same proportion of
biopsy specimens (50%) from patients with active CD, patients
with nonactive CD, and control children were cultured in each
medium, and therefore, the suitability of each medium to recover
duodenal bacteria could be analyzed independently of subject
health status. A total of 29 CFU was recovered in PCA (1.0 ⫾ 1.4
CFU/10 mg of biopsy specimen, on average), 52 CFU was recov-
ered in Wilkins-Chalgren agar (1.9 ⫾ 1.8 CFU/10 mg of biopsy
specimen), 141 CFU was recovered in BH (4.4 ⫾ 6.9 CFU/10 mg
of biopsy specimen), and 81 CFU was recovered in YFCA (2.6 ⫾
3.8 CFU/10 mg of biopsy specimen).
The abundance of cultivable bacterial species associated with
the mucosa of the subjects included in this study is shown in Table
2. Some differences in the bacterial phyla, genera, and species iso-
lated from the different culture media were detected.
When the isolates were classified into different phyla, differ-
ences were found for Proteobacteria, whose members were more
frequently recovered in PCA, followed by YFCA, Wilkins-Chal-
gren agar, and BH; significant differences were detected between
PCA and BH (P ⬍ 0.01) and between YCFA and BH (P ⫽ 0.02).
Differences among the culture media were not detected for iso-
lates belonging to the phyla Actinobacteria and Firmicutes.
In relation to families and species, members of the family
Staphylococcaceae were more frequently isolated in PCA and
Wilkins-Chalgren agar than in BH (P ⬍ 0.01) and YCFA (P ⫽ 0.01
and P ⫽ 0.03, respectively). Of the staphylococcal species, Staph-
ylococcus epidermidis was more frequently isolated in PCA and
Wilkins-Chalgren agar than in YCFA (P ⫽ 0.01), and Staphylococ-
cus pasteuri was isolated significantly more frequently in PCA than
in BH (P ⬍ 0.01).
Members of the family Streptococcaceae were more frequently
isolated in BH and YFCA than in PCA (P ⬍ 01 and P ⫽ 0.02,
respectively) and in BH than in Wilkins-Chalgren agar (P ⬍ 0.01).
Within this family, the Streptococcus anginosus group was signifi-
cantly more abundant in biopsy samples cultured in BH than in
those cultured in Wilkins-Chalgren agar (P ⫽ 0.02).
Finally, members of the Clostridiaceae family were more fre-
quently isolated in Wilkins-Chalgren agar than in BH (P ⫽ 0.02),
and those of the Enterobacteriaceae family were more frequently
isolated in PCA and YFCA than in BH (P ⫽ 0.03 and P ⫽ 0.02,
respectively).
The species richness (S), Shannon species diversity (H=), and
Simpson species dominance (1-D) indexes were calculated for
PCA (S ⫽ 11, H= ⫽ 2.18, and 1-D ⫽ 0.86), Wilkins-Chalgren agar
(S ⫽ 15, H= ⫽ 2.35, and 1-D ⫽ 0.87), BH (S ⫽ 27, H= ⫽ 2.69, and
1-D ⫽ 0.91), and YFCA (S ⫽ 22, H= ⫽ 2.71, and 1-D ⫽ 0.91), in
order to apply the Renyi index. Renyi diversity profiles showed
that the use of PCA and Wilkins-Chalgren agar led to the recovery
of bacteria with lower species diversity than the use of either BH or
YFCA. Renyi diversity profiles also showed that the curves for
PCA and Wilkins-Chalgren agar intersected each other, and the
same was observed for the curves for BH and YFCA; therefore, the
diversity of these pairs could not be compared (data not shown).
Duodenal mucosa-associated bacteria in CD patients and
controls. The proportion of biopsy specimens inoculated in each
culture medium was similar (⬃25%) for each group of individuals
(patients with active CD, patients with nonactive CD, and con-
trols), and therefore, the total number of bacteria recovered in the
different media was considered to represent the differences among
the study groups, regardless of the different culture media used. A
total of 146 isolates were recovered from biopsy specimens from
active CD patients (4.6 ⫾ 4.8 CFU/10 mg of sample, on average),
84 were recovered from biopsy specimens from nonactive CD
patients (5.1 ⫾ 4.1 CFU/10 mg of sample), and 71 were recovered
from biopsy specimens from the control group (8.9 ⫾ 11.7
CFU/10 mg of sample).
The relative abundance of cultivable bacteria associated with
the duodenal mucosa of the different child groups and the differ-
ences in abundance between groups are shown in Table 3.Inre-
lation to the phyla, members of the phylum Proteobacteria were
more abundant in biopsy samples from patients with active CD
than in those from controls (P ⬍ 0.01) and nonactive CD patients
(P ⬍ 0.01), while the relative abundance of members of the Fir-
micutes in biopsy samples from patients with active CD was less
than that in samples from controls (P ⬍ 0.01) and nonactive CD
patients. Members of the phylum Actinobacteria were also more
abundant in biopsy samples from patients with active CD than in
samples from nonactive CD patients (P ⫽ 0.02).
In relation to families, members of the Enterobacteriaceae were
more abundant in patients with active CD than in nonactive CD
Sánchez et al.
5474 aem.asm.org Applied and Environmental Microbiology
on June 10, 2014 by guesthttp://aem.asm.org/Downloaded from
TABLE 2 Cultivable bacterial taxa from active and nonactive CD patients and control subjects isolated in PCA, Wilkins-Chalgren, BH, and YCFA
Closest relative
No. (%) of clones
a
PCA (n ⫽ 29) Wilkins-Chalgren agar (n ⫽ 50) BH (n ⫽ 142) YCFA (n ⫽ 81)
Phylum Actinobacteria 2 (6.9) 1 (2.0) 12 (8.5) 6 (7.4)
Actinomycetaceae 0 0 7 (4.9) 4 (4.9)
Actinomyces odontolyticus 0 0 7 (4.9) 4 (4.9)
Corynebacteriaceae 0 0 1 (0.7) 0
Corynebacterium accolens 0 0 1 (0.7) 0
Micrococcaceae 2 (6.9) 1 (2.0) 3 (2.1) 1 (1.2)
Kocuria kristinae 2 (6.9) 1 (2.0) 1 (0.7) 1 (1.2)
Rothia mucilaginosa 0 0 2 (1.4) 0
Propionibacteriaceae 0 0 1 (0.7) 1 (1.2)
Propionibacterium acnes 0 0 1 (0.7) 1 (1.2)
Phylum Firmicutes 20 (69.0) 43 (86.0) 124 (87.3) 65 (79.0)
Carnobacteriaceae 0 1 (2.0) 5 (3.5) 1 (1.2)
Granulicatella adiacens 0 1 (2.0) 5 (3.5) 1 (1.2)
Clostridiaceae 0
AB
5 (10.2)
A
2 (1.4)
B
2 (2.5)
AB
Clostridium bifermentans 0 3 (6.0) 1 (0.7) 0
Clostridium butyricum 0 2 (4.0) 0 0
Clostridium perfringens 0 0 1 (0.7) 2 (2.5)
Enterococcaceae 1 (3.5) 0 1 (0.7) 0
Enterococcus faecalis 1 (3.5) 0 1 (0.7) 0
Lactobacillaceae 0 0 0 2 (2.5)
Lactobacillus fermentum 0 0 0 2 (2.5)
Staphylococcaceae 10 (34.5)
A
14 (28.0)
A
14 (9.9)
B
9 (11.1)
B
Staphylococcus aureus 0 0 0 3 (3.7)
Staphylococcus epidermidis 6 (20.7)
A
9 (18.0)
A
13 (9.2)
AB
3 (3.7)
B
Staphylococcus hominis 0 2 (4.0) 0 1 (1.2)
Staphylococcus pasteuri 4 (13.8)
A
3 (6.0)
AB
1 (0.7)
B
2 (2.5)
AB
Streptococcaceae 9 (31.0)
A
22 (44.0)
A
90 (63.4)
B
40 (49.4)
AB
Streptococcus anginosus group 0
AB
0
A
17 (12.0)
B
8 (9.9)
AB
Streptococcus australis 0 0 4 (2.8) 0
Streptococcus bovis group 0 0 2 (1.4) 1 (1.2)
Streptococcus gallolyticus 0 0 1 (0.7) 0
Streptococcus mitis group 6 (20.7) 8 (16.0) 21 (14.8) 8 (9.9)
Streptococcus mutans group 0 0 1 (0.7) 4 (4.9)
Streptococcus pneumoniae 0 0 11 (7.8) 3 (3.7)
Streptococcus salivarius group 1 (3.5) 11 (22.0) 23 (16.2) 18 (22.22)
Streptococcus sanguinis group 2 (6.9) 3 (6.0) 14 (9.9) 6 (7.4)
Streptococcus suis 0 0 2 (1.4) 0
Veillonellaceae 0 0 3 (2.1) 2 (2.5)
Veillonella atypical 0 0 0 1 (1.2)
Veillonella dispar 0 0 1 (0.7) 0
Veillonella parvula 0 0 2 (1.4) 1 (1.2)
Unclassified Bacillales 0 1 (2.0) 3 (2.1) 0
Gemella haemolysans 0 1 (2.0) 2 (1.4) 0
Gemella sanguinis 0 0 1 (0.7) 0
Phylum Proteobacteria 7 (24.14)
A
6 (12.0)
AB
5 (3.5)
B
10 (12.6)
A
Burkholderiaceae 1 (3.5) 0 0 0
Burkholderia cepacia 1 (3.5) 0 0 0
Neisseriaceae 1 (3.5) 0 0 0
Neisseria flavescens 1 (3.5) 0 0 0
Enterobacteriaceae 5 (17.2)
A
5 (10.0)
AB
5 (3.5)
B
10 (12.3)
A
Enterobacter cloacae 2 (6.9) 2 (4.0) 0 2 (2.5)
Escherichia coli 0 0 3 (2.1) 3 (3.7)
Klebsiella oxytoca 3 (10.3) 3 (6.0) 3 (2.1) 6 (7.4)
Pseudomonadaceae 0 1 (2.0) 0 0
Pseudomonas stutzeri 0 1 (2.0) 0 0
a
Data are expressed as absolute numbers of isolated clones belonging to one specific taxonomic group (phylum, family, or species) and, in parentheses, the percentage related to the
total number of isolates from each culture medium (PCA, Wilkins-Chalgren agar, BH, and YCFA). Different letters within a row denote statistically significant differences at P ⬍
0.05, estimated by using a two-by-two chi-square test and, when appropriate, Fisher’s exact test.
Duodenal-Mucosal Bacteria in Celiac Disease
September 2013 Volume 79 Number 18 aem.asm.org 5475
on June 10, 2014 by guesthttp://aem.asm.org/Downloaded from
TABLE 3 Relative abundance of cultivable bacterial taxa from biopsy specimens from patients with active and nonactive CD and control subjects
isolated in PCA, Wilkins-Chalgren agar, BH, or YCFA
Closest relative
No. (%) of clones
a
Active CD (n ⫽ 146) Nonactive CD (n ⫽ 85) Control (n ⫽ 71)
Phylum Actinobacteria 15 (10.6)
A
2 (2.4)
B
4 (5.6)
AB
Actinomycetaceae 9 (5.8)
A
0
B
2 (2.8)
AB
Actinomyces odontolyticus 9 (5.8)
A
0
B
2 (2.8)
AB
Corynebacteriaceae 1 (0.7) 0 0
Corynebacterium accolens 1 (0.7) 0 0
Micrococcaceae 5 (3.5) 0 2 (2.8)
Kocuria kristinae 3 (2.0) 0 2 (2.8)
Rothia mucilaginosa 2 (1.3) 0 0
Propionibacteriaceae 0 2 (2.4) 0
Propionibacterium acnes 0 2 (2.4) 0
Phylum Firmicutes 104 (73.2)
A
76 (91.6)
B
66 (93.0)
B
Carnobacteriaceae 2 (1.3) 4 (4.8) 1 (1.4)
Granulicatella adiacens 2 (1.3) 4 (4.8) 1 (1.4)
Clostridiaceae 4 (2.8) 2 (2.4) 3 (4.2)
Clostridium bifermentans 1 (0.7) 0 3 (4.2)
Clostridium butyricum 0 2 (2.4) 0
Clostridium perfringens 3 (2.0) 0 0
Enterococcaceae 2 (1.4) 0 0
Enterococcus faecalis 2 (1.4) 0 0
Lactobacillaceae 0 0 2 (2.8)
Lactobacillus fermentum 0 0 2 (2.8)
Staphylococcaceae 32 (22.5)
A
8 (9.6)
B
2 (2.8)
B
Staphylococcus aureus 3 (2.0) 0 0
Staphylococcus epidermidis 28 (18.2)
A
6 (7.1)
B
2 (2.8)
B
Staphylococcus hominis 1 (7.8) 2 (2.4) 0
Staphylococcus pasteuri 12 (6.9)
A
1 (1.2)
B
0
B
Streptococcaceae 59 (41.6)
A
58 (69.9)
B
58 (81.7)
B
Streptococcus anginosus group 0
A
0
A
25 (35.2)
B
Streptococcus australis 4 (2.6) 0 0
Streptococcus bovis group 0 3 (3.6) 0
Streptococcus gallolyticus 0 1 (1.2) 0
Streptococcus mitis group 14 (9.1)
A
21 (25.0)
B
8 (11.3)
A
Streptococcus mutans group 0
A
0
A
5 (7.0)
B
Streptococcus pneumoniae 7 (4.6) 6 (7.1) 1 (1.4)
Streptococcus salivarius group 25 (16.2) 19 (22.2) 9 (12.7)
Streptococcus sanguinis group 9 (5.8) 6 (7.1) 10 (14.1)
Streptococcus suis 0 2 (2.4) 0
Veillonellaceae 3 (2.1) 2 (2.4) 0
Veillonella atypica 1 (0.7) 0 0
Veillonella dispar 1 (0.7) 0 0
Veillonella parvula 1 (0.7) 2 (2.4) 0
Unclassified Bacillales 2 (1.4) 2 (2.4) 0
Gemella haemolysans 1 (0.7) 2 (2.4) 0
Gemella sanguinis 1 (0.7) 0 0
Phylum Proteobacteria 23 (16.2)
A
5 (6.0)
B
1 (1.4)
B
Burkholderiaceae 0 0 1 (1.4)
Burkholderia cepacia 0 0 1 (1.4)
Enterobacteriaceae 22 (15.5)
A
4 (4.8)
B
0
B
Enterobacter cloacae 6 (3.9) 0 0
Escherichia coli 5 (3.3) 0 0
Klebsiella oxytoca 11 (7.1)
A
4 (4.8)
AB
0
B
Neisseriaceae 0 1 (1.2) 0
Neisseria flavescens 0 1 (1.2) 0
Pseudomonadaceae 1 (0.7) 0 0
Pseudomonas stutzeri 1 (0.7) 0 0
a
Data are expressed as the absolute numbers of isolated clones belonging to one specific taxonomic group (phylum, family, or species) and, in parentheses, the percentage related to
the total number of isolates from each group of children (patients with active CD, patients with nonactive CD, and controls). Different letters within a row denote statistically
significant differences at P ⬍ 0.05, estimated by using a two-by-two chi-square test and, when appropriate, Fisher’s exact test.
Sánchez et al.
5476 aem.asm.org Applied and Environmental Microbiology
on June 10, 2014 by guesthttp://aem.asm.org/Downloaded from
patients and control children (P ⫽ 0.03 and P ⬍ 0.01, respec-
tively). In particular, Klebsiella oxytoca isolates were more abun-
dant in patients with active CD than in control children (P ⫽
0.02). In addition, members of the family Staphylococcaceae were
more abundant in patients with active CD than in patients with
nonactive CD and control individuals (P ⫽ 0.02 and P ⬍ 0.01,
respectively). In particular, S. epidermidis and S. pasteuri isolates
were more abundant in patients with active CD than in patients
with nonactive CD (P ⫽ 0.03 and P ⫽ 0.04, respectively) and in
control children (P ⬍ 0.01 and P ⫽ 0.01, respectively). Further-
more, members of the family Streptococcaceae were less abundant
in patients with active CD than in patients with nonactive CD
and in control children (P ⬍ 0.01). Statistically significant differ-
ences in the abundance of some particular Streptococcus groups
were also detected; thus, the S. anginosus and Streptococcus mutans
groups were more abundant in control individuals than in pa-
tients with active CD (P ⬍ 0.01 and P ⫽ 0.02, respectively) and
nonactive CD (P ⬍ 0.01 and P ⫽ 0.02, respectively), whereas
members of the Streptococcus mitis group were more abundant in
patients with nonactive CD patients than those with active CD
(P ⫽ 0.01). In relation to the family Actinomycetaceae, the isolates
of the only species of that family identified (Actinomyces odonto-
lyticus) were more abundant in patients with active CD than in
those with nonactive CD patients (P ⫽ 0.04).
The species richness (S), Shannon species diversity (H=), and
Simpson species dominance (1-D) indexes were different between
patients with active CD (S ⫽ 27, H= ⫽ 2.73, and 1-D ⫽ 0.91),
patients with nonactive CD (S ⫽ 17, H= ⫽ 2.35, and 1-D ⫽ 0.82),
and controls (S ⫽ 13, H= ⫽ 2.06, and 1-D ⫽ 0.82), indicating
different species diversity between the child groups studied. Renyi
diversity profiles showed that active CD patients had the highest
biodiversity of duodenal cultivable bacteria, followed by nonac-
tive CD patients and controls (data not shown).
DISCUSSION
The study reported herein demonstrates that the microbiota asso-
ciated with the duodenal mucosa of CD patients has a character-
istic deviation from the normal microbiota structure, which may
characterize the disease. The alterations reported in the present
study are partly consistent with those previously detected by mo-
lecular techniques using specific primers or probes (19, 20). Thus,
our results support the hypothesis that normal components of the
microbiota are excluded and replaced by others that could act as
pathobionts in this specific disease environment. Although such
associations do not demonstrate causality between the altered mi-
crobial groups and the disease, they provide a rationale for further
studies on the possible pathogenic modes of action of such alter-
ations and specific bacteria in CD.
To obtain bacterial isolates that are representative of those in-
habiting the duodenal mucosa in both numbers and diversity,
four different culture media previously described in the literature
(34–37) were used. In general, the greatest species diversity and
quantitative recovery of mucosa-associated bacteria were ob-
tained using the BH and YFCA culture media. These differences
could be linked to the high nutritional requirements of duodenal
bacteria, which are better met by the compositions of these media;
incubation conditions may also have been more appropriate, as
they were more anaerobic than those used for PCA and Wilkins-
Chalgren agar. The diverse morphology of the small intestine fa-
vors a precise spatial relationship for strains within particular in-
testinal nutritional and microaerobic environments (42), and
therefore, it is rather complicated to completely reproduce the in
vivo environmental conditions. Also, even though the duodenum
environment is not strictly anaerobic, the possibility that some
anaerobic bacteria were lost due to oxygen exposure during sam-
ple manipulation cannot be disregarded.
We also analyzed whether some of the media used proved bet-
ter at isolation of specific bacteria. In this regard, PCA and
Wilkins-Chalgren agar seemed to favor the growth and isolation
of members of the family Staphylococcaceae but hindered the
growth of members of the family Streptococcaceae. BH medium
favored the growth of members of the family Streptococcaceae but
hampered that of members of the family Enterobacteriaceae.
Wilkins-Chalgren agar also favored the recovery of members of
the family Clostridiaceae compared to the other media. We con-
firm that none of the media or incubation conditions tested were
suitable for the recovery of all viable bacteria detected in the sam-
ples analyzed when used alone, and therefore, various media must
be used to improve the recovery of bacteria that are representative
of the live bacteria inhabiting the duodenum.
We observed an increased diversity of the cultivable mucosa-
associated bacteria recovered from CD patients compared to the
diversity of bacteria recovered from the controls, and these differ-
ences were restored after adherence to a gluten-free diet. In con-
cordance with this finding, denaturing or temperature gradient
gel electrophoresis (DGGE and TGGE, respectively) analysis of
duodenal samples showed a higher bacterial diversity associated
with the small intestinal microbiota of CD patients (13, 18). How-
ever, several recent molecular studies (43–46) have reported that
reduced mucosal bacterial diversity is associated with inflamma-
tory bowel disease (IBD), although the conditions and techniques
used were not comparable to those used in the present study and
the section of the intestinal tract studied was not the same.
Considering the isolates from all subject groups under study,
our results show that the most abundant were those belonging to
the phylum Firmicutes, followed by those of the phyla Proteobac-
teria and Actinobacteria. This is in concordance with the findings
of a previous culture-independent study, where the same three
phyla dominated the proximal small intestine of CD patients, fol-
lowed by other phyla, such as Bacteroidetes or Fusobacteria (47).
Although our previous culture-independent studies also detected
increased numbers of duodenal and fecal Bacteroides spp. in CD
patients compared with controls (19, 20, 48), this bacterial group
was not isolated with the culture conditions applied, probably due
to exposure to oxygen during the process of homogenization of
biopsy specimens and the use of nonselective media for Bacte-
roides, which could have helped to limit the growth of less anaer-
obic and less nutritionally demanding bacteria. Culture-indepen-
dent studies indicate that the members of the normal human gut
microbiota mainly belong to two phyla, Firmicutes and Bacte-
roidetes, with a smaller number of bacteria belonging to the Pro-
teobacteria and Actinobacteria, although these conclusions are
mainly based on analyses of the fecal microbiota composition (45,
49). Previous data also suggest that only 12% of the total species
richness was detected by applying both molecular and cultivation-
based approaches (50). Remarkably, with both approaches, Firmi-
cutes represented the most abundant group, Proteobacteria were
relatively poorly detected by molecular approaches, and Bacte-
roidetes were less abundant when they were assessed with cultiva-
tion-based approaches than with molecular techniques (49–51).
Duodenal-Mucosal Bacteria in Celiac Disease
September 2013 Volume 79 Number 18 aem.asm.org 5477
on June 10, 2014 by guesthttp://aem.asm.org/Downloaded from
In relation to CD, differences in phylum representation were iden-
tified, and in particular, isolates belonging to the Proteobacteria
were more abundant in active CD patients than in nonactive CD
patients and controls. In this context, other studies have also as-
sociated an increase in the Proteobacteria and, in particular, an
increase in adherent-invasive Escherichia coli, Campylobacter con-
cisus, and enterohepatic Helicobacter with IBD (52).
In addition, active and nonactive CD seemed to be associated
with a decreased abundance of members of the family Streptococ-
caceae, specifically, the S. anginosus and S. mutans groups. The
active phase of the disease was also associated with increased pro-
portions of Enterobacteriaceae and Staphylococcaceae and, in par-
ticular, the species Klebsiella oxytoca, S. epidermidis, and S. pas-
teuri. In concordance with these observations, recent culture-
independent studies indicate that the duodenal and fecal
microbiotas of CD patients are characterized by higher numbers
or proportions of Escherichia coli and Staphylococcus species (19,
20). Furthermore, previous studies using cultured-dependent
techniques have shown increased levels of S. epidermidis (16)in
feces from both active and nonactive CD patients in comparison
with healthy controls and a lower prevalence of salivary S. mutans
in association with CD (53). It seems that dominant genera in the
normal microbiota of healthy individuals, which may act as sym-
bionts, like Streptococcus spp., are replaced in the CD patient mi-
crobiota by potential pathobionts, like Staphylococcus spp. (S. epi-
dermidis) and enterobacteria, which could contribute to breaking
down the normal dynamics and balance of the ecosystem.
To our knowledge, this is the first time that cultivable mucosa-
associated bacteria of patients with active and nonactive CD have
been studied, because previous studies were focused on the char-
acterization of CD microbiota using molecular tools, such as
DGGE and TGGE (13, 15, 54), fluorescence in situ hybridization
(FISH) (20), or real-time PCR (19). Culture-dependent studies
are intrinsically biased by the culture media used, the impact of
potential oxygen exposure, and the inability to detect viable but
noncultivable bacteria present in biological samples; notwith-
standing these limitations, the results obtained in the present
study are coherent with those of previous studies based on molec-
ular techniques, which overcome these limitations. Therefore, the
use of culture-dependent techniques has allowed the characteriza-
tion of the active fraction of the mucosal microbiota of CD pa-
tients and will facilitate future investigation into the possible
pathogenic role that isolated bacteria play in the development of
CD.
Conclusions. This study demonstrates that the duodenal-mu-
cosal microbiota of CD patients presents alterations in the diver-
sity and abundance of different cultivable bacterial taxa, which
could be a consequence of the pathogenesis of CD, which involves
massive destruction of the small bowel mucosa and the conse-
quent release of intracellular contents and serum into the gut. In
the active phase of the disease, the mucosa-associated microbiota
was characterized by a higher abundance of members of the phy-
lum Proteobacteria and the families Enterobacteriaceae and
Staphylococcaceae, apparently excluding members of the phylum
Firmicutes and the family Streptococcaceae, which are normal in-
habitants of the healthy small intestine. These alterations are at-
tenuated after long-term adherence to a gluten-free diet, but the
microbiota is not completely restored; in particular, a reduced
abundance of specific species of Streptococcus (S. anginosus and S.
mutans) also characterizes the microbiota of CD patients with
active and nonactive disease. These findings also suggest their po-
tential use as hallmarks of CD, regardless of inflammatory status.
ACKNOWLEDGMENTS
This work was supported by grants AGL2011-25169 and Consolider Fun-
C-Food CSD2007-00063 from the Spanish Ministry of Economy and
Competitiveness. The scholarship to E. Sánchez from the Institute
Danone is fully acknowledged.
REFERENCES
1. Dickson BC, Streutker CJ, Chetty R. 2006. Coeliac disease: an update for
pathologists. J. Clin. Pathol. 59:1008 –1016.
2. Nilsen EM, Jahnsen FL, Lundin KE, Johansen FE, Fausa O, Sollid LM,
Jahnsen J, Scott H, Brandtzaeg P. 1998. Gluten induces an intestinal
cytokine response strongly dominated by interferon gamma in patients
with celiac disease. Gastroenterology 115:551–563.
3. Fernandez S, Molina IJ, Romero P, Gonzalez R, Pena J, Sanchez F,
Reynoso FR, Perez-Navero JL, Estevez O, Ortega C, Santamaria M.
2011. Characterization of gliadin-specific Th17 cells from the mucosa of
celiac disease patients. Am. J. Gastroenterol. 106:528 –538.
4. Monteleone I, Sarra M, Del Vecchio BG, Paoluzi OA, Franze E, Fina D,
Fabrizi A, MacDonald TT, Pallone F, Monteleone G. 2010. Character-
ization of IL-17A-producing cells in celiac disease mucosa. J. Immunol.
184:2211–2218.
5. Gianfrani C, Auricchio S, Troncone R. 2005. Adaptive and innate im-
mune responses in celiac disease. Immunol. Lett. 99:141–145.
6. Koning F, Schuppan D, Cerf-Bensussan N, Sollid LM. 2005. Patho-
mechanisms in celiac disease. Best. Pract. Res. Clin. Gastroenterol. 19:
373–387.
7. Wieser H, Koehler P. 2008. The biochemical basis of celiac disease. Cereal
Chem. 85:1–13.
8. Sanz Y. 2009. Novel perspectives in celiac disease therapy. Mini Rev. Med.
Chem. 9:359 –367.
9. Dubois PC, Trynka G, Franke L, Hunt KA, Romanos J, Curtotti A,
Zhernakova A, Heap GA, Adany R, Aromaa A, Bardella MT, van den
Berg LH, Bockett NA, de la Concha EG, Dema B, Fehrmann RS,
Fernandez-Arquero M, Fiatal S, Grandone E, Green PM, Groen HJ,
Gwilliam R, Houwen RH, Hunt SE, Kaukinen K, Kelleher D, Kor-
ponay-Szabo I, Kurppa K, MacMathuna P, Maki M, Mazzilli MC,
McCann OT, Mearin ML, Mein CA, Mirza MM, Mistry V, Mora B,
Morley KI, Mulder CJ, Murray JA, Nunez C, Oosterom E, Ophoff RA,
Polanco I, Peltonen L, Platteel M, Rybak A, Salomaa V, Schweizer JJ,
Sperandeo MP, et al. 2010. Multiple common variants for celiac disease
influencing immune gene expression. Nat. Genet. 42:295–302.
10. Niewinski MM. 2008. Advances in celiac disease and gluten-free diet. J.
Am. Diet. Assoc. 108:661– 672.
11. Stene LC, Honeyman MC, Hoffenberg EJ, Haas JE, Sokol RJ, Emery L,
Taki I, Norris JM, Erlich HA, Eisenbarth GS, Rewers M. 2006. Rotavirus
infection frequency and risk of celiac disease autoimmunity in early child-
hood: a longitudinal study. Am. J. Gastroenterol. 101:2333–2340.
12. Walters JR, Bamford KB, Ghosh S. 2008. Coeliac disease and the risk of
infections. Gut 57:1034 –1035.
13. Di Cagno R, De Angelis M, De Pasquale I, Ndagijimana M, Vernocchi
P, Ricciuti P, Gagliardi F, Laghi L, Crecchio C, Guerzoni ME, Gobbetti
M, Francavilla R. 2011. Duodenal and faecal microbiota of celiac chil-
dren: molecular, phenotype and metabolome characterization. BMC Mi-
crobiol. 11:219. doi:10.1186/1471-2180-11-219.
14. Sánchez E, Nadal I, Donat E, Ribes-Koninckx C, Calabuig M, Sanz Y.
2008. Reduced diversity and increased virulence-gene carriage in intesti-
nal enterobacteria of coeliac children. BMC Gastroenterol. 8:50. doi:10
.1186/1471-230X-8-50.
15. Sánchez E, Donat E, Ribes-Koninckx C, Calabuig M, Sanz Y. 2010.
Intestinal Bacteroides species associated with coeliac disease. J. Clin.
Pathol. 63:1105–1111.
16. Sánchez E, Ribes-Koninckx C, Calabuig M, Sanz Y. 2012. Intestinal
Staphylococcus spp. and virulent features associated with coeliac disease. J.
Clin. Pathol. 65:830 – 834.
17. Sanz Y, Sanchez E, Marzotto M, Calabuig M, Torriani S, Dellaglio F.
2007. Differences in faecal bacterial communities in coeliac and healthy
children as detected by PCR and denaturing gradient gel electrophoresis.
FEMS Immunol. Med. Microbiol. 51:562–568.
Sánchez et al.
5478 aem.asm.org Applied and Environmental Microbiology
on June 10, 2014 by guesthttp://aem.asm.org/Downloaded from
18. Schippa S, Iebba V, Barbato M, Di NG, Totino V, Checchi MP, Longhi
C, Maiella G, Cucchiara S, Conte MP. 2010. A distinctive ‘microbial
signature’ in celiac pediatric patients. BMC Microbiol. 10:175. doi:10
.1186/1471-2180-10-175.
19. Collado MC, Donat E, Ribes-Koninckx C, Calabuig M, Sanz Y. 2009.
Specific duodenal and faecal bacterial groups associated with paediatric
coeliac disease. J. Clin. Pathol. 62:264 –269.
20. Nadal I, Donat E, Ribes-Koninckx C, Calabuig M, Sanz Y. 2007.
Imbalance in the composition of the duodenal microbiota of children with
coeliac disease. J. Med. Microbiol. 56:1669 –1674.
21. Cinova J, de Palma G, Stepankova R, Kofronova O, Kverka M, Sanz Y,
Tuckova L. 2011. Role of intestinal bacteria in gliadin-induced changes in
intestinal mucosa: study in germ-free rats. PLoS One 6:e16169. doi:10
.1371/journal.pone.0016169.
22. de Palma G, Cinova J, Stepankova R, Tuckova L, Sanz Y. 2010. Pivotal
advance: bifidobacteria and Gram-negative bacteria differentially influ-
ence immune responses in the proinflammatory milieu of celiac disease. J.
Leukoc. Biol. 87:765–778.
23. Laparra JM, Sanz Y. 2010. Bifidobacteria inhibit the inflammatory re-
sponse induced by gliadins in intestinal epithelial cells via modifications of
toxic peptide generation during digestion. J. Cell. Biochem. 109:801– 807.
24. Laparra JM, Olivares M, Gallina O, Sanz Y. 2012. Bifidobacterium
longum CECT 7347 modulates immune responses in a gliadin-induced
enteropathy animal model. PLoS One 7:e30744. doi:10.1371/journal.pone
.0030744.
25. DuPont AW, DuPont HL. 2011. The intestinal microbiota and chronic
disorders of the gut. Nat. Rev. Gastroenterol. Hepatol. 8:523–531.
26. Kaur N, Chen CC, Luther J, Kao JY. 2011. Intestinal dysbiosis in inflam-
matory bowel disease. Gut Microbes 2:211–216.
27. Hormannsperger G, Clavel T, Haller D. 2012. Gut matters: microbe-host
interactions in allergic diseases. J. Allergy Clin. Immunol. 129:1452–1459.
28. McLoughlin RM, Mills KH. 2011. Influence of gastrointestinal commen-
sal bacteria on the immune responses that mediate allergy and asthma. J.
Allergy Clin. Immunol. 127:1097–1107.
29. van Nimwegen FA, Penders J, Stobberingh EE, Postma DS, Koppelman
GH, Kerkhof M, Reijmerink NE, Dompeling E, van den Brandt PA,
Ferreira I, Mommers M, Thijs C. 2011. Mode and place of delivery,
gastrointestinal microbiota, and their influence on asthma and atopy. J.
Allergy Clin. Immunol. 128:948 –955.
30. Gophna U, Sommerfeld K, Gophna S, Doolittle WF, Veldhuyzen van
Zanten SJ. 2006. Differences between tissue-associated intestinal micro-
floras of patients with Crohn’s disease and ulcerative colitis. J. Clin. Mi-
crobiol. 44:4136 – 4141.
31. Kleessen B, Kroesen AJ, Buhr HJ, Blaut M. 2002. Mucosal and invading
bacteria in patients with inflammatory bowel disease compared with con-
trols. Scand. J. Gastroenterol. 37:1034 –1041.
32. Swidsinski A, Weber J, Loening-Baucke V, Hale LP, Lochs H. 2005.
Spatial organization and composition of the mucosal flora in patients with
inflammatory bowel disease. J. Clin. Microbiol. 43:3380 –3389.
33. Walker-Smith JA, Guandalini S, Schmitz J, Shmerling DH, Visakorpi
JK. 1990. Revised criteria for diagnosis of coeliac disease. Arch. Dis. Child.
65:909–911.
34. Hayashi H, Sakamoto M, Benno Y. 2002. Phylogenetic analysis of the
human gut microbiota using 16S rDNA clone libraries and strictly anaer-
obic culture-based methods. Microbiol. Immunol. 46:535–548.
35. Dubreuil L, Houcke I, Leroy I. 1996. In vitro activity of a new fluoro-
quinolone, marbofloxacin (RO 09–1168) against strictly anaerobic bacte-
ria and some bacteria from human fecal flora. Pathol. Biol. 44:333–336.
(In French.)
36. Canganella F, Zirletta G, Gualterio L, Massa S, Trovatelli LD. 1992.
Anaerobic facultative bacteria isolated from the gut of rabbits fed different
diets. Zentralbl. Mikrobiol. 147:537–540.
37. Duncan SH, Louis P, Thomson JM, Flint HJ. 2009. The role of pH in
determining the species composition of the human colonic microbiota.
Environ. Microbiol. 11:2112–2122.
38. Wilson K. 1988. Preparation of genomic DNA from bacteria, p 2.4.1–
2.4.5. In Ausubel FM, Brent R, Kingston RE, Moore DD, Smith JA, Seid-
man JG, Struhl K (ed), Current protocols in molecular microbiology.
Greene Publishing and Wiley Interscience, New York, NY.
39. Favier CF, Vaughan EE, De Vos WM, Akkermans AD. 2002. Molecular
monitoring of succession of bacterial communities in human neonates.
Appl. Environ. Microbiol. 68:219 –226.
40. Satokari RM, Vaughan EE, Akkermans AD, Saarela M, de Vos WM.
2001. Bifidobacterial diversity in human feces detected by genus-specific
PCR and denaturing gradient gel electrophoresis. Appl. Environ. Micro-
biol. 67:504 –513.
41. Tothmeresz B. 1995. Comparison of different methods for diversity or-
dering. J. Vegetation Sci. 6:283–290.
42. Espey MG. 2013. Role of oxygen gradients in shaping redox relationships
between the human intestine and its microbiota. Free Radic. Biol. Med.
55:130–140.
43. Frank DN, St Amand AL, Feldman RA, Boedeker EC, Harpaz N, Pace
NR. 2007. Molecular-phylogenetic characterization of microbial commu-
nity imbalances in human inflammatory bowel diseases. Proc. Natl. Acad.
Sci. U. S. A. 104:13780 –13785.
44. Friswell M, Campbell B, Rhodes J. 2010. The role of bacteria in the
pathogenesis of inflammatory bowel disease. Gut Liver 4:295–306.
45. Lepage P, Hasler R, Spehlmann ME, Rehman A, Zvirbliene A, Begun A,
Ott S, Kupcinskas L, Dore J, Raedler A, Schreiber S. 2011. Twin study
indicates loss of interaction between microbiota and mucosa of patients
with ulcerative colitis. Gastroenterology 141:227–236.
46. Ott SJ, Musfeldt M, Wenderoth DF, Hampe J, Brant O, Folsch UR,
Timmis KN, Schreiber S. 2004. Reduction in diversity of the colonic
mucosa associated bacterial microflora in patients with active inflamma-
tory bowel disease. Gut 53:685– 693.
47. Ou G, Hedberg M, Horstedt P, Baranov V, Forsberg G, Drobni M,
Sandstrom O, Wai SN, Johansson I, Hammarstrom ML, Hernell O,
Hammarstrom S. 2009. Proximal small intestinal microbiota and identi-
fication of rod-shaped bacteria associated with childhood celiac disease.
Am. J. Gastroenterol. 104:3058 –3067.
48. de Palma G, Nadal I, Medina M, Donat E, Ribes-Koninckx C, Calabuig
M, Sanz Y. 2010. Intestinal dysbiosis and reduced immunoglobulin-
coated bacteria associated with coeliac disease in children. BMC Micro-
biol. 10:63. doi:10.1186/1471-2180-10-63.
49. Eckburg PB, Bik EM, Bernstein CN, Purdom E, Dethlefsen L, Sargent
M, Gill SR, Nelson KE, Relman DA. 2005. Diversity of the human
intestinal microbial flora. Science 308:1635–1638.
50. Kovatcheva-Datchary P, Zoetendal EG, Venema K, de Vos WM, Smidt
H. 2009. Tools for the tract: understanding the functionality of the gas-
trointestinal tract. Therap. Adv. Gastroenterol. 2:9 –22.
51. Rajilic-Stojanovic M, Smidt H, de Vos WM. 2007. Diversity of the
human gastrointestinal tract microbiota revisited. Environ. Microbiol.
9:2125–2136.
52. Mukhopadhya I, Hansen R, El-Omar EM, Hold GL. 2012. IBD—what
role do Proteobacteria play? Nat. Rev. Gastroenterol. Hepatol. 9:219 –230.
53. Acar S, Yetkiner AA, Ersin N, Oncag O, Aydogdu S, Arikan C. 2012.
Oral findings and salivary parameters in children with celiac disease: a
preliminary study. Med. Princ. Pract. 21:129 –133.
54. Nistal E, Caminero A, Vivas S, Ruiz de Morales JM, Saenz de Miera LE,
Rodriguez-Aparicio LB, Casqueiro J. 2012. Differences in faecal bacteria
populations and faecal bacteria metabolism in healthy adults and celiac
disease patients. Biochimie 94:1724 –1729.
Duodenal-Mucosal Bacteria in Celiac Disease
September 2013 Volume 79 Number 18 aem.asm.org 5479
on June 10, 2014 by guesthttp://aem.asm.org/Downloaded from