Specific response of a novel and abundant Lactobacillus amylovorus-like phylotype to dietary prebiotics in the guts of weaning piglets.
ABSTRACT Using 16S rRNA gene-based approaches, we analyzed the responses of ileal and colonic bacterial communities of weaning piglets to dietary addition of four fermentable carbohydrates (inulin, lactulose, wheat starch, and sugar beet pulp). An enriched diet and a control diet lacking these fermentable carbohydrates were fed to piglets for 4 days (n = 48), and 10 days (n = 48), and the lumen-associated microbiota were compared using denaturing gradient gel electrophoresis (DGGE) analysis of amplified 16S rRNA genes. Bacterial diversities in the ileal and colonic samples were measured by assessing the number of DGGE bands and the Shannon index of diversity. A higher number of DGGE bands in the colon (24.2 +/- 5.5) than in the ileum (9.7 +/- 4.2) was observed in all samples. In addition, significantly higher diversity, as measured by DGGE fingerprint analysis, was detected in the colonic microbial community of weaning piglets fed the fermentable-carbohydrate-enriched diet for 10 days than in the control. Selected samples from the ileal and colonic lumens were also investigated using fluorescent in situ hybridization (FISH) and cloning and sequencing of the 16S rRNA gene. This revealed a prevalence of Lactobacillus reuteri in the ileum and Lactobacillus amylovorus-like populations in the ileum and the colon in the piglets fed with fermentable carbohydrates. Newly developed oligonucleotide probes targeting these phylotypes allowed their rapid detection and quantification in the ileum and colon by FISH. The results indicate that addition of fermentable carbohydrates supports the growth of specific lactobacilli in the ilea and colons of weaning piglets.
- SourceAvailable from: Ignacio Badiola[Show abstract] [Hide abstract]
ABSTRACT: This study aimed to provide novel insights into the gastrointestinal microbial diversity from different gastrointestinal locations in weaning piglets using PCR-restriction fragment length polymorphism (PCR-RFLP). Additionally, the effect of different feed additives was analyzed. Thirty-two piglets were fed with four different diets: a control group and three enriched diets, with avilamycin, sodium butyrate, and a plant extract mixture. Digesta samples were collected from eight different gastrointestinal segments of each animal and the bacterial population was analysed by a PCR-RFLP technique that uses 16S rDNA gene sequences. Bacterial diversity was assessed by calculating the number of bands and the Shannon-Weaver index. Dendrograms were constructed to estimate the similarity of bacterial populations. A higher bacterial diversity was detected in large intestine compared to small intestine. Among diets, the most relevant microbial diversity differences were found between sodium butyrate and plant extract mixture. Proximal jejunum, ileum, and proximal colon were identified as those segments that could be representative of microbial diversity in pig gut. Results indicate that PCR-RFLP technique allowed detecting modifications on the gastrointestinal microbial ecology in pigs fed with different additives, such as increased biodiversity by sodium butyrate in feed.BioMed research international. 01/2014; 2014:269402.
Dataset: De Angelis 2006
- [Show abstract] [Hide abstract]
ABSTRACT: The commensal bacteria Lactobacillus are widely used as probiotic organisms conferring a heath benefit on the host. They have been implicated in promoting gut health via the stimulation of host immunity and anti-inflammatory responses, as well as protecting the intestinalmucosa against pathogen invasion. Lactobacilli grow by fermenting sugars and starches and produce lactic acid as their primary metabolic product. For efficient utilisation of varied carbohydrates, lactobacilli have evolved diverse sugar transport and metabolic systems, which are specifically induced by their own substrates. Many bacteria are also capable of sensing and responding to changes in their environment. These sensory responses are often independent of transport or metabolism and are mediated through membrane-spanning receptor proteins. We employed DNA-based pyrosequencing technology to investigate the changes in the intestinal microbiota of piglets weaned to a diet supplemented with either a natural sugar, lactose or an artificial sweetener (SUCRAM®, consisting of saccharin and neohesperidin dihydrochalcone (NHDC); Pancosma SA). The addition of either lactose or saccharin/NHDC to the piglets' feed dramatically increased the caecal population abundance of Lactobacillus, with concomitant increases in intraluminal lactic acid concentrations. This is the first report of the prebiotic-like effects of saccharin/NHDC, an artificial sweetener, being able to influence the commensal gut microbiota. The identification of the underlying mechanism(s) will assist in designing nutritional strategies for enhancing gut immunity and maintaining gut health.The British journal of nutrition 01/2014; · 3.45 Impact Factor
APPLIED AND ENVIRONMENTAL MICROBIOLOGY, July 2004, p. 3821–3830
Copyright © 2004, American Society for Microbiology. All Rights Reserved.
Vol. 70, No. 7
Specific Response of a Novel and Abundant Lactobacillus
amylovorus-Like Phylotype to Dietary Prebiotics in the
Guts of Weaning Piglets
Sergey R. Konstantinov,1* Ajay Awati,2Hauke Smidt,1Barbara A. Williams,2
Antoon D. L. Akkermans,1and Willem M. de Vos1
Laboratory of Microbiology, Agrotechnology and Food Sciences Group, Wageningen University,1and
Animal Nutrition Group, Wageningen Institute of Animal Sciences,2Wageningen, The Netherlands
Received 29 December 2003/Accepted 4 March 2004
Using 16S rRNA gene-based approaches, we analyzed the responses of ileal and colonic bacterial commu-
nities of weaning piglets to dietary addition of four fermentable carbohydrates (inulin, lactulose, wheat starch,
and sugar beet pulp). An enriched diet and a control diet lacking these fermentable carbohydrates were fed to
piglets for 4 days (n ? 48), and 10 days (n ? 48), and the lumen-associated microbiota were compared using
denaturing gradient gel electrophoresis (DGGE) analysis of amplified 16S rRNA genes. Bacterial diversities in
the ileal and colonic samples were measured by assessing the number of DGGE bands and the Shannon index
of diversity. A higher number of DGGE bands in the colon (24.2 ? 5.5) than in the ileum (9.7 ? 4.2) was
observed in all samples. In addition, significantly higher diversity, as measured by DGGE fingerprint analysis,
was detected in the colonic microbial community of weaning piglets fed the fermentable-carbohydrate-enriched
diet for 10 days than in the control. Selected samples from the ileal and colonic lumens were also investigated
using fluorescent in situ hybridization (FISH) and cloning and sequencing of the 16S rRNA gene. This revealed
a prevalence of Lactobacillus reuteri in the ileum and Lactobacillus amylovorus-like populations in the ileum and
the colon in the piglets fed with fermentable carbohydrates. Newly developed oligonucleotide probes targeting
these phylotypes allowed their rapid detection and quantification in the ileum and colon by FISH. The results
indicate that addition of fermentable carbohydrates supports the growth of specific lactobacilli in the ilea and
colons of weaning piglets.
The diets, microbiota, and gastrointestinal (GI) tract inter-
actions of mammals are extremely complex and are the result
of millions of years of coevolution between the higher verte-
brates and their microbiota. As a consequence, any major
changes in lifestyle and diet are likely to place stress on the
stability of these interactions and affect the entire GI tract
ecophysiology. In contrast to the gradual weaning of human
babies, piglets within a production environment are weaned at
an early stage with solid feed and transported to production
farms. This combination of stress factors can lead to diarrhea,
a reduced growth rate, and in some cases even death (54). In
order to enhance growth and suppress the activity of the gut
microbiota, antimicrobial compounds have been fed to wean-
ing pigs for more than 4 decades (6). Nowadays, the emergence
of antibiotic resistance in the human commensal bacteria has
raised concerns about the impact of anitimicrobial compounds
for agricultural use (53) and has accelerated the search for
alternative nutritional strategies, such as the addition of pro-
biotics and prebiotics. These approaches have become an in-
creasingly important consideration in swine nutrition because
of accumulating evidence of their potential benefits in animals
and humans and the possibility that they could replace antibi-
otics in feed (5, 21, 61, 64). The development of such dietary
strategies requires a combination of in vitro, in vivo, and chal-
lenge studies involving both expertise in animal nutrition and
an evaluation of the composition and activity of the indigenous
microbiota throughout the GI tract.
In the past, the microbial community in the GI tracts of pigs
has been studied intensively, but most attention was paid to
easily cultivable commensal bacteria and a number of oppor-
tunistic pathogens (10, 57). Many of the strictly anaerobic GI
tract bacteria are still difficult to cultivate and therefore remain
undetectable by conventional techniques (59, 63). Recent phy-
logenetic analysis based on the in vitro amplification of the 16S
rRNA gene and other phylogenetic markers by PCR tech-
niques have revealed dramatically higher diversity than de-
scribed previously by cultivation (20, 28, 41). While molecular
approaches based on PCR can introduce different types of bias
(22, 65), a combination of PCR and fingerprinting techniques,
such as denaturing gradient gel electrophoresis (DGGE) and
terminal restriction fragment length polymorphism, has led to
new insights into GI microbial ecology and the effects of dif-
ferent dietary strategies and host factors on bacterial-commu-
nity composition (69).
It has been recognized that a stable indigenous microbiota in
the intestine can prevent colonization by pathogens (46, 62).
This so-called colonization resistance may be of utmost impor-
tance for animals, especially at stressful times, such as weaning.
The promotion of colonization resistance through the addition
of prebiotics has been suggested as a comparatively easy way to
improve enteric health (4, 7, 66). Prebiotics have been used to
induce the colonization of bacteria, such as lactobacilli and
* Corresponding author. Mailing address: Laboratory of Microbiol-
ogy, Agrotechnology and Food Sciences Group, Wageningen Univer-
sity, Hesselink van Suchtelenweg 4, 6703 CT Wageningen, The Neth-
erlands. Phone: 31 317 483118. Fax: 31 317 483829. E-mail:
bifidobacteria, considered to be beneficial for the host (13, 14,
61). Stimulation of the Lactobacillus population within the GI
tracts of piglets is of specific importance, not only due to the
potential effect of the bacteria on gut function and health (57,
58), but also because of their possible antagonistic activities
toward other bacteria (19, 51). Lactobacilli establish early in
the piglet intestine, and although succession occurs throughout
the pig’s lifetime, they remain a predominant part of the in-
testinal bacterial community (3, 38, 57, 60). Numerous studies
have suggested that some prebiotics may specifically stimulate
intestinal lactobacilli. The application of lactobacilli as probi-
otics or therapeutic supplements has also been studied (44, 45).
However, little is known of the response of the bacterial com-
munity to such dietary interventions.
This work describes changes in the predominant ileal and
colonic bacterial populations in weaning piglets that were fed
a diet containing four added fermentable carbohydrates,
namely, inulin, lactulose, wheat starch, and sugar beet pulp.
The data indicate that the incorporation of these four ingre-
dients in the diet results in outgrowth of lactobacilli in the
small intestine and higher diversity in the colon. Two particular
phylotypes related to Lactobacillus amylovorus and Lactobacil-
lus reuteri were the most prevalent throughout the guts of
piglets fed with the prebiotics, as demonstrated by DGGE of
16S rRNA gene amplicons in combination with sequence anal-
ysis. Newly developed DNA oligonucleotide probes targeting
these key species allowed their rapid detection and quantifica-
tion in the ilea and colons of piglets by fluorescent in situ
MATERIALS AND METHODS
Animals, diets, and sampling. All of the procedures involving animals were
conducted in accordance with the Dutch law on experimental animals and were
approved by the Animal Experimental Committee of Wageningen University.
Three identical but independent feeding experiments including a total of 108
piglets (crossbred Hypor ? Pietrain) were started immediately at the time of
weaning (25 to 28 days old). Each experiment used 36 piglets. At the start of the
experiment (day 1), four piglets were sacrificed. The remaining 32 piglets were
offered one of two diets (16 piglets per diet): the HF diet containing four added
fermentable carbohydrates, namely, lactulose, inulin, sugar beet pulp, and wheat
starch, and the LF diet with a low concentration of fermentable carbohydrates
(Table 1). The diets were composed in such a way that the total energy and
protein contents were comparable. On days 4 and 10 of each experiment, eight
piglets were sacrificed per treatment.
The samples were divided into aliquots, one of which was used for genomic-
DNA extraction, followed by 16S rRNA gene-targeted PCR-DGGE analysis,
cloning, and sequencing. In parallel, aliquots from the same samples were fixed
for FISH and for determination of the lactic acid concentration.
DNA isolation. DNA isolation from lumen samples (0.2 g) was done by using
the Fast DNA Spin kit (Qbiogene, Inc., Carlsbad, Calif.). Agarose gel (1.2%
[wt/vol]) electrophoresis in the presence of ethidium bromide was used to check
visually for DNA quality and yield.
PCR amplification. All primers used in this study are listed in Table 2. Primers
S-D-Bact-0968-a-S-GC and S-D-Bact-1401-a-A-17 were used to amplify the V6
to V8 regions of the 16S rRNA gene. PCR was performed using the Taq DNA
polymerase kit from Life Technologies (Gaithersburg, Md.). PCR mixtures (50
?l) contained 0.5 ?l of Taq polymerase (1.25 U), 20 mM Tris-HCl (pH 8.5), 50
mM KCl, 3.0 mM MgCl2, 200 ?M each deoxynucleoside triphosphate, 5 pmol of
the primers, 1 ?l of DNA diluted to ?1 ng/?l, and UV-sterilized water. The
samples were amplified in a T1 thermocycler (Whatman Biometra, Go ¨ttingen,
Germany), and the cycling consisted of 94°C for 5 min and 35 cycles of 94°C for
30 s, 56°C for 20 s, 68°C for 40 s, and 68°C for 7 min (final extension). Aliquots
(5 ?l) were analyzed by electrophoresis on 1.2% (wt/vol) agarose gels containing
ethidium bromide to check for product size and quantity.
To investigate the Lactobacillus-specific GI tract bacterial community by
DGGE, a specific nested-PCR approach was chosen. For the initial amplifica-
tion, S-D-Bact-0011-a-A-17 and S-G-Lab-0677-a-A-17 primers were employed
(18), using the following cycling conditions: predenaturation at 94°C for 5 min;
35 cycles of 94°C for 30 s, 66°C for 20 s, and 68°C for 40 s; and a final extension
at 68°C for 7 min. The PCR products were then used as templates in nested
PCRs with S-G-Lab-0159-a-S-20 and S-*-Univ-0515-a-A-24-GC. The cycling
program was identical to the one used for the amplification of the V6 to V8
regions of the 16S rRNA gene.
DGGE analysis. The amplicons obtained from the lumen-extracted DNA were
separated by DGGE according to the specifications of Muyzer et al. (36) using
a Dcode TM system (Bio-Rad Laboratories, Hercules, Calif.). Electrophoresis
was performed in an 8% polyacrylamide gel (37.5:1 acrylamide-bisacrylamide;
dimensions, 200 by 200 by 1 mm) using a 38 to 48% denaturing gradient (35) for
separation of PCR products obtained with primers S-D-Bact-0968-a-S-GC and
S-D-Bact-1401-a-A-17, while gradients of 30 to 60% were employed for the
separation of the S-G-Lab-0159-a-S-20- and S-*-Univ-0515-a-A-24-GC-gener-
ated amplicons. The gels were electrophoresed for 16 h at 85 V in 0.5? TAE
buffer (48) at a constant temperature of 60°C and subsequently stained with
Analysis of DGGE gels. Analysis of all DGGE samples was done as described
previously (27). Briefly, all gels were scanned at 400 dots/in and analyzed using
Molecular Analyst/PC software (version 1.12; Bio-Rad). First, a number of bands
were assessed per lane using the band-searching algorithm in the program. A
manual check was done, and the DGGE fragments constituting ?1% of the total
area of all bands were omitted from further analysis. Second, as a parameter for
the structural diversity of the microbial community, the Shannon index of general
diversity, H? (9, 27, 32, 50), was calculated using the following function: H? ? ??
Pilog Pi, where Piis the importance probability of the bands in a lane. H? was
calculated on the basis of the bands on the gel tracks that were applied for the
generation of the dendrograms by using the intensities of the bands as judged by
peak height in the densitometric curves. Pi, was calculated as follows: Pi? ni/n?,
where niis the height of a peak and n? is the sum of all peak heights in the
densitometric curve. The similarity between the DGGE profiles was determined
by calculating a band similarity (Dice) coefficient, SD[SD? 2nAB/(nA? nB),
where nAis the number of DGGE bands in lane 1, nBrepresents the number of
DGGE bands in lane 2, and nABis the number of common DGGE bands] (27,
Statistical analysis. For statistical analysis, the number of DGGE bands, the
Shannon index of general diversity, and the band similarity coefficient (SD) were
calculated. Differences between these parameters for the two diets were tested
for significance using Tukey’s Studentized range test of multiple comparisons
(56) according to the following equation: Y ? ? ? Di? εij, where Y is the result,
? is the mean, D is the effect of the diet, and εijis the error term. All statistical
analyses were performed using the SAS GLM procedure (55).
TABLE 1. Compositions of the diets
Diet or ingredient
Lactulose (?50% DM)c
aLF, diet with a low concentration of fermentable carbohydrates.
bHF, diet containing lactulose, inulin, sugar beet pulp, and wheat starch.
cDM, dry matter.
3822 KONSTANTINOV ET AL.APPL. ENVIRON. MICROBIOL.
Generation and screening of 16S rRNA gene clone libraries. PCR was per-
formed with a Taq DNA polymerase kit from Life Technologies using primers
S-D-Bact-0011-a-S-17 and S-D-Bact-1492-a-A-19. Amplification was carried out
as described previously (27). The PCR product was purified with the QIAquick
PCR purification kit (Westburg, Leusden, The Netherlands) according to the
manufacturer’s instructions. The purified PCR product was cloned into pGEM-T
(Promega, Madison, Wis.) using competent Escherichia coli JM109 as a host. The
colonies of ampicillin-resistant transformants were transferred with a sterile
toothpick to 15 ?l of Tris-EDTA buffer and boiled for 15 min at 95°C. PCR was
immediately performed with the vector-specific primers T7 and SP6 to check the
sizes of the inserts using the cell lysate as a template. Plasmids containing an
insert of ?1.6 kb were used to amplify the V6 to V8 regions of the 16S rRNA
gene. The amplicons were compared with the bands of DGGE profiles that
comprised ?1% of the total area of all bands. Clones representing an insert
corresponding to a dominant band were grown in Luria Broth liquid medium (5
ml) with ampicillin (100 ?g ml?1). Plasmid DNA was isolated using the Wizard
Plus purification system (Promega) and used for sequence analysis of the cloned
16S rRNA gene by using a Sequenase (T7) sequencing kit (Amersham Life
Sciences, Slough, United Kingdom) according to the manufacturer’s specifica-
tions and using either the T7 and SP6 primers or S-Univ-1100-a-A-15 labeled
with IRD-800. Sequences were automatically analyzed on a LiCor (Lincoln,
Neb.) DNA Sequencer 4000L and corrected manually. The sequences were also
compared to those available in public databases by using BLAST analysis (2).
The partial and complete 16S rRNA gene sequences were checked for chimeric
constructs by the Ribosomal Database Project CHECK_CHIMERA program
(31). None of the sequences were found to be PCR-generated chimeras.
Cloning and sequencing of DGGE bands after Lactobacillus-specific PCR
amplification. Representative bands were excised from DGGE gels using a
QIAEXII Gel extraction kit (Westburg) according to the instructions in the
manual. After reamplification using the original S-G-Lab-0159-a-S-20 and S-*-
Univ-0515-a-A-24-GC primer set, cloning and sequencing analysis were carried
out as previously described.
Design and validation of oligonucleotide probes for FISH analysis. Nearly
complete 16S rRNA sequences of L. amylovorus-like and L. reuteri-like isolates
(this study) and closely related L. amylovorus-like (OTU171) and L. reuteri-like
(OTU173) isolates from pig intestine (28) were aligned, and probes targeting
these sequences were designed using the ARB software package (http://www
.arb-home.de). Probes were designed taking into consideration the types and
positions of nucleotide mismatches with sequences of related species with a G?C
content of ?50% and a length of ?18 nucleotides. Sequence comparisons using
the ARB, Check Probe, and BLAST programs confirmed that the targeted
regions were conserved among the 16S rRNA sequences of OTU171 (L. amy-
lovorus-like) isolated from pig intestine and Lactobacillus kitasatoi isolated from
chickens (34) (Fig. 1, band B) and to L. reuteri-like OTU173 (Fig. 1, band A).
Probe OTU171-0088-a-A-18 was found to also match the partial sequence of
Lactobacillus galinarum (National Center for Biotechnology Information acces-
sion no. X97898). However, a comparison among the L. galinarum, OTU171, and
L. kitasatoi sequences showed 99 to 100% homology among them based on 600
bp (E. coli positions 20 to 620).
The reference strains L. amylovorus DSMZ 20531 and L. reuteri DSMZ 20015
were used as positive controls (Table 3). Ten reference Lactobacillus strains and
Enterococcus faecalis DSMZ 20478, frequently found in the GI tract, were used
as negative controls to evaluate the specificities of the newly designed probes.
The temperature of hybridization was 50°C, and if needed, formaldehyde was
added to increase the specificity (17).
The reference strains used in this study were obtained from the sources
indicated in Table 3. The strains were cultivated as recommended by the culture
collections in the respective catalogues. Exponentially grown cells were harvested
at 5,000 ? g for 10 min, washed with 0.2-?m-pore-size-filtered phosphate-buff-
ered saline (PBS; per liter, 8 g of NaCl, 0.2 g of KCl, 1.44 g of Na2HPO4, and
0.24 g of KH2PO4, pH 7.2), and diluted 1:3 with 4% paraformaldehyde in PBS.
After fixation at 4°C for 16 h, the cells were stored in 50% ethanol-PBS at ?80°C
for subsequent FISH analysis.
Collection and preparation of ileal- and colonic-lumen samples for FISH.
Ileal- and colonic-lumen samples from 108 experimental piglets were processed
as described previously (12). In short, 0.5 g of lumen samples was resuspended in
4.5 ml of PBS and vortexed with five or six glass beads (diameter, 3 mm) for at
least 3 min to homogenize the sample. After centrifugation at 700 ? g for 1 min,
1 ml of the supernatant was added to 3 ml of 4% paraformaldehyde in PBS and
stored for 16 h at 4°C. After being washed twice with PBS, the fixed cells were
stored in 50% ethanol-PBS at ?80°C until further use.
Enumeration of bacteria by FISH. For microscopic analysis, fixed cells were
spotted on gelatin-coated glass slides and dried for 20 min at 50°C. The optimal
cell concentration for counting using the different probes was determined using
dilution series of the lumen samples. After drying of the slides, the cells were
dehydrated for 3 min in 50, 70, and finally 96% ethanol-H2O. Ten microliters of
hybridization buffer (0.9 M NaCl, 20 mM Tris-HCl [pH 7.5], 0.1% [wt/vol]
sodium dodecyl sulfate) containing 10 ng of Cy3-labeled lactobacillus probes/?l
or 5 ng of fluorescein-isothiocyanate-labeled S-D-Bact-0338-a-A-17/?l (Table 2)
was added to each well, followed by incubation at 50°C for 16 h. After hybrid-
ization, the slides were washed in 50 ml of hybridization buffer for 10 min. For
total cell counts, DAPI (4?,6-diamino-2-phenylindole) at a final concentration of
100 ng/ml was added to the washing buffer. After the slides were rinsed in
double-distilled water, they were immediately air dried and mounted in Vecta-
shield (Vector Laboratories, Burlingame, Calif.). Digital images of the slides
were analyzed, and fluorescence-positive cells were counted using Qwin image
analysis software (Leica Microsystems, Rijswijk, The Netherlands). For each
analysis, 25 microscopic fields were counted.
Lactic acid analysis. The lactic acid concentration in ileal lumen was analyzed
by high-performance liquid chromatography (Jasco Instruments) using a column
(Supelcogel; C-610H; 30 cm by 7.8-mm internal diameter) and a precolumn
(Supelcoguard; C-610H; 5 cm by 4.6-mm internal diameter) with 1% H2SO4as
TABLE 2. Oligonucleotide primers and probes used in this study
Primer or probeSequence (5?–3?) Reference
AGA GTT TGA T(C/T)(A/C) TGG CTC AG
GGA AAC AG(A/G) TGC TAA TAC CG
CGC CGG GGG CGC GCC CCG GGC GGG GCG GGG GCA CGG GGG
G ATC GTA TTA CCG CGG CTG CTG GCA
CAC CGC TAC ACA TGG AG
CGC CCG GGG CGC GCC CCG GGC GGG GCG GGG GCA CGG GGG
GAA CGC GAA GAA CCT TAC
CGG TGT GTA CAA GAC CC
GGG TTG CGC TCG TTG
GGT TAC CTT GTT ACG ACT T
TAA TAC GAC TCA CTA TAG G
GAT TTA GGT GAC ACT ATA G
GCT GCC TCC CGT AGG AGT
GGT ATT AGC A(C/T)C TGT TTC CA
CGC TTT CCC AAC GTC ATT
CCA TCG TCA ATC AGG TGC
aNomenclature according to Alm et al. (1).
VOL. 70, 2004EFFECTS OF PREBIOTICS ON PIGLET INTESTINAL MICROBIOTA3823
the mobile phase. The concentrations were determined by UV detection at 210
Nucleotide sequence accession numbers. The sequences reported in this study
were deposited in GenBank under the following accession numbers: AY493201
throughout the experimental period. The average weaning
weight of the piglets was 7.5 kg. At the end of the experiment,
no significant difference in body weight gain between the di-
etary treatments was observed.
Effects of fermentable carbohydrates on bacterial diversity
in the ileum and colon. A comparative 16S rRNA gene-tar-
geted DGGE fingerprinting analysis of bacterial communities
was performed for ileal- and colonic-lumen samples of piglets
that were fed two different diets. Samples from piglets that
were sacrificed on the day of weaning (n ? 12) and 4 (n ? 48)
and 10 (n ? 48) days after weaning were analyzed. The number
of DGGE bands and the Shannon index of general diversity
were assessed for each sample and subjected to statistical anal-
ysis according to GI tract location across all time periods. For
all samples, a comparison between the two locations along the
GI tract revealed a statistically higher number of DGGE bands
(24. 2 ? 5. 5; P ? 0.05) in the colonic lumen compared to the
ileal lumen (9.7 ? 4. 2). The Shannon index of diversity in the
colon (1.43 ? 0.3; P ? 0.05) was also higher than in the ileum
(0. 86 ? 0. 26). No significant differences in the number of
DGGE bands or diversity were detected on day 4 in the ilea or
in the colons of piglets fed the different diets. However, by day
10 after weaning, the diversity was significantly higher in the
colonic samples from HF piglets, as evidenced by the number
of bands (27. 5 ? 5.6; P ? 0.05), than in those of the LF group
(20.5 ? 6.3). There was no statistical difference between the
number of DGGE bands and the diversity index in the ileal
samples by day 10.
The influence of the diet on the bacterial-community struc-
ture in the ileal and colonic lumens of HF and LF group piglets
on days 4 and 10 was further elucidated. By day 4, there were
no DGGE bands detected in one dietary group that were
completely absent in the other (data not shown). In contrast, a
simple visual comparison of the DGGE banding patterns by
FIG. 1. Effects of the fermentable-carbohydrate-containing diet on the ileum bacterial community by day 10 of the experiment. (A) DGGE of
PCR products of V6 to V8 regions of 16S rRNA gene sequences retrieved from lumen samples on day 10 after weaning. Lanes: 1 to 6, piglets on
HF diet; 7 to 12, piglets on LF diet; M, marker. Fragments A, B, and C were identified from 16S rRNA gene clone libraries. (B) Monitoring of
the Lactobacillus-like community of piglets. DGGE analysis of amplicons generated by nested PCR with primers S-G-Lab-0159-a-S-20 and
S-*-Univ-0515-a-A-24-GC, originating from piglets on the HF diet (lanes 1 to 6) and piglets on the LF diet (lanes 7 to 12). The dominant fragments
in Lactobacillus-like patterns were identified by the clones corresponding to L. acidophilus, L. reuteri, and L. amylovorus.
3824 KONSTANTINOV ET AL.APPL. ENVIRON. MICROBIOL.
day 10 of the experiment revealed a marked difference be-
tween the ileal samples of the two dietary groups. A represen-
tative DGGE analysis of the PCR fragments generated with
primers S-D-Bact-0968-a-S-GC and S-D-Bact-1401-a-A-17 is
shown in Fig. 1A. Two particularly strong bands (A and B)
were present in the ileal samples from piglets fed with the HF
diet on day 10. Band B was also detected in 18 out of the 24
colonic-lumen samples from pigs fed the HF diet on day 10 and
was absent in pigs fed with the LF diet (data not shown). The
ileal samples of the LF group, on the other hand, were dom-
inated by another band located in the uppermost part of the
DGGE gels (Fig. 1A, band C). Band C was not present in the
HF diet samples. To obtain an objective interpretation of the
electrophoretic patterns of the HF and LF ilea, the samples
were subjected to a numerical analysis based on the Dice
similarity coefficient, followed by cluster analysis. The similar-
ity was visualized using the unweighted pair group method with
averaging algorithm (Fig. 2). Cluster analysis revealed that all
24 samples from HF diet-fed piglets formed a coherent cluster
with similarity indices above 60%. Within this cluster, 20 out of
24 samples grouped together, with similarity indices higher
than 75%. The low similarities between HF and LF samples
confirmed the visual differences in their DGGE fingerprints
(Fig. 1). The average similarity index of the HF colon samples
was 45% (data not shown). Taken together, the results ob-
tained after DGGE analysis demonstrate that the bacterial
compositions in the ilea and colons of piglets were modulated
by the HF diet by day 10.
Identification of cloned 16S rRNA gene sequences in DGGE
patterns. In order to identify changes in the bacterial diversity
detected by DGGE analysis, the 16S rRNA genes from the
ileal-lumen samples of four HF-fed piglets (10 days) and four
LF-fed piglets were amplified and cloned into E. coli, and 15
clones per sample were partially sequenced (Table 4). To iden-
tify the dominant bands, A and B, that appeared in 90% of
samples 10 days after starting the HF diet, together with a third
distinct band (C) in the LH diet (Fig. 1A), the V6 to V8 regions
of the 16S rRNA gene were amplified from the cell lysates of
a total of 120 transformants. The mobilities of these amplicons
during DGGE were compared to those obtained from rRNA
gene sequences retrieved from samples from the piglets fed for
10 days with the HF and LF diets. Sixty-one percent of the
clones were assigned to one of the dominant bands in the
DGGE profiles, while 39% did not match any of the detectable
bands. The clones from the eight different clone libraries cor-
responding to bands A, B, and C were completely sequenced.
The 16S rRNA gene sequences of the clones representing band
A were identified as L. reuteri-like, or OTU173, while band B
showed similarity to OTU171, or L. amylovorus-like (28).
Clones matching the position of band C were 98% similar to
Sarcinia ventriculi (Fig. 1A).
FIG. 2. Similarity index of DGGE profiles obtained from ileal-
lumen microbiota of 48 piglets fed either HF or LF diet for 10 days.
The normalization of the DGGE gels was done with respect to the
reference standards included in three gels containing the ileal-lumen
samples of HF and LF from the three replicate experiments. The Dice
coefficient of similarity between banding patterns of different gels was
calculated. This allowed the generation of a dendrogram, and the
samples were grouped according to the similarity of their community
profiles. UPGMA, unweighted pair group method with averaging.
TABLE 3. Bacterial strains, sources, media for cultivation, and FISH results
Lactobacillus rhamnosus LGG
aDSMZ, German Collection for Microorganisms and Cell Culture, Braunschweig, Germany; VTT, VTT culture collection, FIN-02044, Finland; ATCC, American
Type Culture Collection, Manassas, Va.
bMRS, lactobacillus MRS broth (Difco, Sparks, Md.); WW, Wilkins West broth.
c(I), L-S-OTU171-a-A-0088; (II), L-S-OTU173-a-A-0085; ?, hybridization signal; ?, no hybridization signal.
VOL. 70, 2004EFFECTS OF PREBIOTICS ON PIGLET INTESTINAL MICROBIOTA 3825
Although the amplification of the V6 to V8 regions using
general bacterial primers allowed the visualization of the major
differences between the HF and LF samples and the screening
of the 16S rRNA gene clone libraries, it yielded poor resolu-
tion of Lactobacillus populations. Therefore, specific amplifi-
cation of the Lactobacillus GI tract bacterial community was
used to screen the clones matching one of the dominant bands
after V6 to V8 16S region DGGE analysis. Lactobacillus-spe-
cific amplification, in combination with DGGE analysis, con-
firmed the predominance of one particular phylotype related
to L. amylovorus, while a band related to L. reuteri was not
consistently found in the samples from the piglets fed with the
HF diet (Fig. 1B). L. acidophilus was also present in HF and
LF samples irrespective of diet.
To confirm the visual match between the 16S rRNA gene
clones and the DGGE bands, the HF diet-specific bands were
excised from DGGE gels after Lactobacillus-specific PCR, re-
amplified, and sequenced. Sequencing analysis of the bands (A
and B) confirmed their identities as L. reuteri-like and L. amy-
lovorus-like (28). Interestingly, of the four HF diet samples, an
average of four other phylotypes related to Lactobacillus mu-
cosae (97%), L. galinarum (97%), and two different Lactoba-
cillus species clones, one oral (98%; accession no. AY005048)
and one isolated from swine production facilities (99%; acces-
sion no. AY017059), were found (Table 4). However, their
sequences did not match any visible DGGE bands. In compar-
ison, the Lactobacillus diversity in LF was predominated by
Lactobacillus acidophilus-like related sequences. The results of
the clone libraries showed increased lactobacillus diversity in
the HF samples, while DGGE analysis suggested a specific
outgrowth of L. amylovorus-like phylotypes in the terminal ilea
of weaning piglets.
Development and evaluation of FISH probes specific for L.
amylovorus and L. reuteri-like isolates. Potential probes were
identified based on the alignment of the complete 16S rRNA
sequences of the clones matching DGGE bands A and B (Fig.
1A) and related Lactobacillus spp. (Table 3).
The probes were experimentally validated by performing
FISH analysis on a range of Lactobacillus species and other
bacteria that are commonly found in large numbers in the pig
GI tract (28, 57). A constant temperature of 50°C for 16 h and
0% (vol/vol) formamide in the hybridization buffer were used,
resulting in specific hybridization only with the respective tar-
get strains (Table 3). Subsequently, the validated probes were
used to enumerate target bacteria in individual ileal and co-
lonic samples from piglets fed the different diets for 10 days.
To evaluate whether the microbiota was affected by the diet,
the total cell counts and the total bacterial and lactobacillus-
enterococcus counts for the HF and LF diets were compared
(Table 5). Within the HF, the lactobacillus-enterococcus
counts were significantly higher (P ? 0.05) than in the LF.
Hybridization with the L-*-OTU171-0088-a-A-18 probe de-
tected the OTU171 phylotype in 83% of the ileal samples and
75% of the piglet colonic samples from the HF diet, while no
hybridization-positive cells were obtained for the LF diet (Ta-
ble 5). In comparison, an L. reuteri-like related population was
detected in 75% of the ileal-lumen samples and 41% of the
colonic-lumen samples from the HF diet using the L-*-
Lactic acid concentration in ileal lumen. Lactic acid was
measured in the lumen samples from the terminal ilea of all
piglets from days 1, 4, and 10 after the introduction of the diet
(Fig. 3). By days 4 and 10, a significantly higher lactic acid
concentration was recovered in the samples from the HF diet
TABLE 4. Clones with similarity to known sequences in GenBanka
No.Closest relative in GenBank
No. of clones
L. reuteri (DSM 20016 T)
L. acidophilus subsp. johnsonii
Lactobacillus sp. oral clone CX036
Lactobacillus sp. strain CLE-4
S. ventriculi (DSM 286)
Leptotrichia sp. (oral clone)
aClones were retrieved from ileal-lumen samples of four HF-fed piglets and LF-fed piglets at 5 weeks of age. The clones are listed according to their abundances
in the eight different 16S rRNA gene clone libraries.
3826 KONSTANTINOV ET AL.APPL. ENVIRON. MICROBIOL.
than in those from the LF diet. As lactic acid is a common end
product of fermentation of lactobacilli, these results were in
agreement with the outgrowth of lactobacilli in the terminal
ileum, as demonstrated by 16S rRNA gene-based DGGE and
FISH analyses, and they suggest that the lactobacilli were not
only present but also metabolically active.
The bacteriological results reported here indicate that the
addition of specific fermentable carbohydrates to the diet can
lead to a shift in both the composition and activity of the
microbial communities of the small and large intestines of
weaning piglets. Two particular phylotypes related to L. amy-
lovorus and L. reuteri were the most prevalent populations in
the ilea of piglets fed the HF diet for 10 days, as demonstrated
by DGGE analysis and a phylotype-specific 16S rRNA-tar-
geted FISH analysis. In addition, bacterial diversity was in-
creased by day 10 in the colons of the HF group, as evidenced
by the higher number of DGGE bands and the Shannon index
of diversity in the corresponding samples. Given the present
global concern about antibiotic replacement in feed, these
results show that careful design of the diet can indeed stimu-
late supposedly beneficial bacteria. They also indicate that
other species can be suppressed. These findings, therefore, are
not only interesting for piglet microbiology and nutrition at the
time of weaning but also provide new insights into the specific
effects of prebiotics on the indigenous Lactobacillus commu-
nities of piglets.
The availability of fast sequencing techniques offers an un-
precedented opportunity to conduct comprehensive surveys of
pig microbial communities (20, 28). Results based on compar-
ative sequence analyses of the 16S rRNA and chaperonin-60
genes documented the complexity of the intestinal microbial
community and suggested that the majority of the bacterial
species colonizing the GI tract in pigs have not yet been char-
acterized. However, cloning and sequencing is time-consuming
and may limit the number of samples that can be processed.
Thus, the high sample throughput required to determine com-
munity responses to experimental treatments, such as intro-
duction of prebiotics or probiotics, needs to be achieved by the
analysis of multiple clone libraries. Alternatively, DGGE and a
similar technique called temperature gradient gel electro-
phoresis have been introduced into microbial ecology (35–37)
as one attempt to obtain an overview of the structural diversity
of microbial communities. As reported previously, DGGE and
temperature gradient gel electrophoresis are sensitive enough
to detect bacteria that constitute 1% of the total bacterial
community (67). The PCR-DGGE detection limit has also
been estimated previously by dilution series of pure cultures
(68). In addition, the primer pair used in the present study
FIG. 3. Luminal lactic acid concentrations in the terminal ilea of
piglets. The data are expressed as mean values plus standard errors of
the mean for all samples. Asterisks, significantly different concentra-
tions of lactic acid (P ? 0.05).
TABLE 5. FISH results
Ileum (n ? 24)Colon (n ? 24)Ileum (n ? 24)Colon (n ? 24)
Total cell counta
Total bacterial counta
2.1 ? 1.1 ? 108
1.89 ? 1.4 ? 108
1.5 ? 0.6 ? 108d
3.1 ? 2.4 ? 1010
2.7 ? 1.5 ? 1010
3.8 ? 2.1 ? 108
1.2 ? 1.2 ? 108
1.17 ? 1.1 ? 108
0.4 ? 0.3 ? 107
2.3 ? 1.4 ? 1010
1.94 ? 0.9 ? 1010
6.3 ? 3.1 ? 108
L. amylovorus-like (Probe, L-
No. of piglets colonized (%)
1.3 ? 108
0.64–2.4 ? 108
3.7 ? 109
1.3–5.6 ? 109
L. reuteri- like (Probe, L-S-
No. of piglets colonized (%)
7.7 ? 107
4.4–12 ? 107
5.8 ? 107
5.3–6.2 ? 107
5.5 ? 107
5.1–6.3 ? 107
aResults for total cell counts (DAPI staining), total bacterial counts (S-D-Bact-0338-a-A-17), and lactobacillus-enterococcus counts (S-G-Lab-0158-a-A-20).
bMean ? standard deviation (cells per gram). ND, no bacteria detected.
cSpecific counts for the probes L-S-OTU171-a-A-0088 (L. amylovorus-like) and L-S-OTU173-a-A-0085 (L. reuteri-like) for ileal- and colonic-lumen samples from
piglets fed for 10 days on HF or LF diet.
dSignificant differences (P ? 0.05) from the values compared (bold face).
VOL. 70, 2004EFFECTS OF PREBIOTICS ON PIGLET INTESTINAL MICROBIOTA3827
(S-D-Bact-0968-a-S-GC and S-D-Bact-1401-a-A-17) was found
to amplify with the same efficiency 16S rRNA genes from the
complex soil bacterial communities (11). Individual DGGE
bands can be assigned to cultured organisms or retrieved ribo-
somal sequences (26, 27). This is usually not possible in acti-
vated sludge, sediments, soil, and other highly diverse micro-
bial systems because the banding patterns are too complex (9).
However, the number, precise positions, and intensities of the
bands reflect the number and relative abundance of dominant
rRNA sequence types in the sample and thus allow comparison
of microbial communities with each other. By applying this
approach to piglet GI tract lumen samples, a distinct diversity
value for each sample was obtained and changes in community
diversity over time in different experiments were observed. In
agreement with previous analyses of 16S rRNA gene libraries
obtained from pig ileum and colon samples (28), the results
reported in this study showed a significantly lower diversity in
the ileum than in the colon. Further elucidation of the diet
effect by using 16S rRNA gene PCR-DGGE analysis unveiled
the impact of the ileum microbiota in the utilization of the
fermentable carbohydrates. Marked differences in the bacteri-
al-community compositions between ileal samples from piglets
fed with HF or LF diets were demonstrated by day 10 of the
experiment (Fig. 1). While this has not been previously dem-
onstrated by culture-independent approaches, there are nu-
merous studies showing that the ilea of pigs harbor diverse and
active bacterial populations (reference 24; reviewed in refer-
ence 64). Furthermore, the increased diversity in the colons of
piglets fed the HF diet, as demonstrated by DGGE, is in
agreement with earlier data (27). Such a strong effect of the
diet on the porcine colon and fecal bacterial populations has
also been demonstrated when the animals were fed different
diets (29) containing fermentable carbohydrates and after in-
troduction of an exogenous Lactobacillus strain (51).
The combination of 16S rRNA gene-directed DGGE, clon-
ing, and sequencing in this study identified the phylogenetic
changes in the piglets’ microbiota and highlighted the out-
growth of L. amylovorus-like populations in the ilea of piglets
in the HF-fed group. However, because these approaches are
all based on PCR amplification methods, the results cannot be
converted into actual bacterial numbers. FISH, in combination
with microscopic analysis, has provided a powerful tool for
detecting and quantifying various bacterial genera, including
Lactobacillus, in human feces (16, 17). Sequences related to L.
amylovorus or phylotype OTU171 were recovered from the
colonic wall and lumen of a pig (41) and were found to be the
most abundant Lactobacillus phylotype in the GI tracts of
Danish pigs of different ages and with different feeding regimes
(28). The same phylotype was found independently to predom-
inate in the small-intestinal microbiota of weaning piglets
based on 16S rRNA gene sequence analyses and DGGE (25).
Since the present results suggest a significant stimulation of
this bacterium in the presence of the HF diet, a DNA oligo-
nucleotide probe targeting the phylotype was developed and
validated for FISH analysis. After validation, the probe was
used to quantify the number of hybridized cells in the ileal and
colonic lumens of piglets fed for 10 days with the HF or LF
diet. The FISH results showed that the sizes of OTU171-
related populations varied from 0.64 ? 108to 2.4 ? 108/g of
ileal lumen in 20 out of 24 piglets (Table 5B). The results were
in agreement with the DGGE analysis, where the phylotype
was detected as a dominant DGGE band in 80% of the ana-
lyzed piglets fed for 10 days with the HF diet.
Various studies of the effects of prebiotic oligosaccharides
on the colonic microbiota in humans have reported a stimula-
tion of lactobacilli by inulin and lactulose (reference 47; re-
viewed by Rastall and Gibson ). However, in many of the
in vitro and in vivo experiments, further characterization be-
yond the genus level has not been achieved (42). Populations
of lactobacilli related to L. amylovorus and L. reuteri have been
identified as common inhabitants of the human and animal
intestine. The properties of the type strains are also well es-
tablished. They are known for the ability to degrade starch. In
addition, both strains produce bacteriocins, potentially sup-
pressing other populations within the intestinal microbiota
(15). However, the extrapolation of functional properties from
well-characterized cultured strains to the related L. amylo-
vorus-like and L. reuteri-like phylotypes may not be justified. In
particular, a high level of 16S rRNA gene relatedness
(?97.5%) was found between the type strains of L. amylovorus,
Lactobacillus crispatus, L. gallinarum, and Lactobacillus kitasa-
tonis (34) and the most abundant L. amylovorus-like phylotype
detected in the study. Therefore, a study of the physiological
and genomic properties of a large collection of L. amylovorus-
like populations isolated from the pig intestine is under way in
our laboratory. The results will be described in a separate
In the present study, sugar beet pulp with a significant fer-
mentable-carbohydrate (including cell walls) content was also
included. The cell wall component of the diet has previously
been found not only to affect microbial fermentation in the GI
tracts of pigs (24), but also to play a role in stimulation or
inhibition of certain pathogens in the intestine (23). The ad-
dition of sugar beet pulp to the diets of pigs was reported to
reduce the population of coliforms (43), while others suggested
an increased proliferation of pathogenic E. coli if the piglets
were fed a fiber-enriched diet (33). The effect of dietary fiber
on the development of swine dysentery is also under discus-
sion. As shown by some reports, diets with low fiber and resis-
tant starches protect pigs from infection with Brachyspira hyo-
dysenteriae (8, 39, 40), while others were not able to confirm
these findings (29, 30). Our results suggest that the combina-
tion of fermentable dietary fiber and oligosaccharides may
specifically stimulate the L. amylovorus-like population along
the guts of weaning piglets.
This research was financially supported by the European Commu-
nities project HEALTHYPIGUT (QLK5-LT2000-00522).
We thank Wilma Akkermans-van Vliet, Dick Bongers, Cornelia
Malin, and Yanka Georgieva for their technical assistance. We are
grateful to Maria Saarela for providing the Lactobacillus VTT strains.
1. Alm, E. W., D. B. Oerther, N. Larsen, D. A. Stahl, and L. Raskin. 1996. The
oligonucleotide probe database. Appl. Environ. Microbiol. 62:3557–3559.
2. Altschul, S. F., W. Gish, W. Miller, E. W. Myers, and D. J. Lipman. 1990.
Basic local alignment search tool. J. Mol. Biol. 215:403–410.
3. Bateup, J., S. Dobbinson, K. Munro, M. A. McConnell, and G. W. Tannock.
1998. Molecular analysis of the composition of Lactobacillus populations
inhabiting the stomach and caecum of pigs. Microb. Ecol. Health Dis. 10:
4. Blaut, M. 2002. Relationship of prebiotics and food to intestinal microflora.
Eur. J. Nutr. 41:11–16.
3828 KONSTANTINOV ET AL.APPL. ENVIRON. MICROBIOL.
5. Buddington, R. K., K. Kelly-Quagliana, K. K. Buddington, and Y. Kimura.
2002. Non-digestible oligosaccharides and defense functions: lessons learned
from animal models. Br. J. Nutr. 87(Suppl. 2):S231–S239.
6. Cromwell, G. L. 2002. Why and how antibiotics are used in swine production.
Anim. Biotechnol. 13:7–27.
7. Cummings, J. H., and G. T. Macfarlane. 2002. Gastrointestinal effects of
prebiotics. Br. J. Nutr. 87(Suppl. 2):S145–S151.
8. Durmic, Z., D. W. Pethick, J. R. Pluske, and D. J. Hampson. 1998. Changes
in bacterial populations in the colon of pigs fed different sources of dietary
fibre, and the development of swine dysentery after experimental infection.
J. Appl. Microbiol. 85:574–582.
9. Eichner, C. A., R. W. Erb, K. N. Timmis, and I. Wagner-Do ¨bler. 1999.
Thermal gradient gel electrophoresis analysis of bioprotection from pollut-
ant shocks in the activated sludge microbial community. Appl. Environ.
10. Ewing, W. N., and D. J. A. Cole. 1994. The living gut: an introduction to
micro-organisms in nutrition. Context Publications, Dungannon, Ireland.
11. Felske, A., A. D. L. Akkermans, and W. M. De Vos. 1998. Quantification of
16S rRNAs in complex bacterial communities by multiple competitive re-
verse transcription-PCR in temperature gradient gel electrophoresis finger-
prints. Appl. Environ. Microbiol. 64:4581–4587.
12. Franks, A. H., H. J. M. Harmsen, G. C. Raangs, G. J. Jansen, F. Shut, and
G. W. Welling. 1998. Variation of the bacterial population in human feces
measured by fluorescent in situ hybridization with group-specific 16S rRNA-
targeted oligonucleotide probes. Appl. Environ. Microbiol. 64:3336–3345.
13. Gibson, G. R. 1999. Dietary modulation of the human gut microflora using
the prebiotics oligofructose and inulin. J. Nutr. 129:1438S–1441S.
14. Gibson, G. R., and M. B. Roberfroid. 1995. Dietary modulation of the human
colonic microbiota: introducing the concept of prebiotics. J. Nutr. 125:1401–
15. Hammes, W. P., N. Weiss, and W. Holzapfel. 1991. The genera Lactobacillus
and Carnobacterium, p. 1535–1594. In A. Bolows, H. G. Truper, M. Dworkin,
W. Harder, and K. H. Schleifer (ed.), The procariotes: a handbook on the
biology of bacteria: ecophysiology, isolation, identification, applications, vol.
2. Springer-Verlag, New York, N.Y.
16. Harmsen, H. J. M., P. Elfferich, F. Schut, and G. W. Welling. 1999. A 16S
rRNA-targeted probe for detection of lactobacilli and enterococci in faecal
samples by fluorescent in situ hybridization. Microb. Ecol. Health Dis. 11:
17. Harmsen, H. J. M., G. C. Raangs, T. He, J. E. Degener, and G. W. Welling.
2002. Extensive set of 16S rRNA-based probes for detection of bacteria in
human feces. Appl. Environ. Microbiol. 68:2982–2990.
18. Heilig, H., E. G. Zoetendal, E. E. Vaughan, P. Marteau, A. D. L. Akkermans,
and W. M. de Vos. 2002. Molecular diversity of Lactobacillus spp. and other
lactic acid bacteria in the human intestine as determined by specific ampli-
fication of 16S ribosomal DNA. Appl. Environ. Microbiol. 68:114–123.
19. Henriksson, A., L. Andre ´, and P. Conway. 1995. Distribution of lactobacilli
in the porcine gastrointestinal tract. FEMS Microbiol. Ecol. 16:55–60.
20. Hill, J. E., R. P. Seipp, M. Betts, L. Hawkins, A. G. Van Kessel, W. L. Crosby,
and S. M. Hemmingsen. 2002. Extensive profiling of a complex microbial
community by high-throughput sequencing. Appl. Environ. Microbiol. 68:
21. Hopkins, M. J., and G. T. Macfarlane. 2003. Nondigestible oligosaccharides
enhance bacterial colonization resistance against Clostridium difficile in vitro.
Appl. Environ. Microbiol. 69:1920–1927.
22. Ishii, K., and M. Fukui. 2001. Optimization of annealing temperature to
reduce bias caused by a primer mismatch in multitemplate PCR. Appl.
Environ. Microbiol. 67:3753–3755.
23. Jensen, B. B., O. Hojberg, L. L. Mikkelsen, M. S. Hedemann, and N. Canibe.
2003. Enhancing intestinal function to treat and prevent intestinal disease, p.
103–121. Proceedings of the 9th International Symposium on Digestible
Physiology in Pigs. University of Alberta, Banff, Canada.
24. Jensen, B. B., and H. Jorgensen. 1994. Effect of dietary fiber on microbial
activity and microbial gas production in various regions of the gastrointes-
tinal tract of pigs. Appl. Environ. Microbiol. 60:1897–1904.
25. Kluess, J., A. Akkermans, S. Konstantinov, S. Kuhla, M. Kwella, and W. B.
Souffrant. 2003. The microbial community and its metabolic activities in the
small intestine of weaning piglets, p. 613–618. In W. B. Souffrant and C. C.
Metges (ed.), Progress in research on energy and protein metabolism.
Wageningen Academic Publishers, Rostock, Germany.
26. Konstantinov, S. R., N. Fitzsimons, E. E. Vaughan, and A. D. L. Akkermans.
2002. From composition to functionality of the intestinal microbial commu-
nities, p. 59–84. In G. W. Tannock (ed.), Probiotics and prebiotics: where are
we going? Caister Academic Press, Wymondham, United Kingdom.
27. Konstantinov, S. R., W.-Y. Zhu, B. A. Williams, S. Tamminga, W. M. de Vos,
and A. D. L. Akkermans. 2003. Effect of fermentable carbohydrates on faecal
bacterial communities as revealed by DGGE analysis of 16S rDNA. FEMS
Microbiol. Ecol. 43:225–235.
28. Leser, T. D., J. Z. Amenuvor, T. K. Jensen, R. H. Lindecrona, M. Boye, and
K. Moller. 2002. Culture-independent analysis of gut bacteria: the pig gas-
trointestinal tract microbiota revisited. Appl. Environ. Microbiol. 68:673–
29. Leser, T. D., R. H. Vindecrona, T. K. Jensen, B. B. Jensen, and K. Moller.
2000. Changes in the colon of pigs fed different experimental diet and after
infection with Brachyspira hyodysenteriae. Appl. Environ. Microbiol. 66:3290–
30. Lindecrona, R. H., T. J. Jensen, B. B. Jensen, T. Lezer, M. Jiufeng, and K.
Moller. 2003. The influence of diet on the development of swine dysentery
upon experimental infection. Anim. Sci. 76:81–87.
31. Maidak, B. L., J. R. Cole, J. C. T. Parker, G. M. Garrity, B. N. Larsen,
T. G. L. Li, M. J. McCaughey, G. J. Olsen, R. Overbeek, S. Pramanik, T. M.
Schmidt, J. M. Tiedje, and C. R. Woese. 1999. A new version of the RDP
(Ribosomal Database Project). Nucleic Acids Res. 27:171–173.
32. McCracken, V. J., J. M. Simpson, R. I. Mackie, and H. R. Gaskins. 2001.
Molecular ecological analysis of dietary and antibiotic-induced alterations of
the mouse intestinal microbiota. J. Nutr. 131:1862–1870.
33. McDonald, D. E., D. W. Pethick, B. P. Mullan, and D. J. Hampson. 2001.
Increasing viscosity of the intestinal contents alters small intestinal structure
and intestinal growth, and stimulates proliferation of enterotoxigenic Esch-
erichia coli in newly-weaned pigs. Br. J. Nutr. 86:487–498.
34. Mukai, T., K. Arihara, A. Ikeda, K. Nomura, F. Suzuki, and H. Ohori. 2003.
Lactobacillus kitasatonis sp. nov., from chicken intestine. Int. J. Syst. Evol.
35. Muyzer, G., T. Brinkhoff, U. Nu ¨bel, C. Santegoeds, H. Scha ¨fer, and C.
Wawer. 1998. Denaturing gradient gel electrophoresis (DGGE) in microbial
ecology, p. 1–27. In A. D. L. Akkermans, J. D. van Elsas, and F. J. de Bruijn
(ed.), Molecular microbial ecology manual, vol. 3.4.4. Kluwer Academic
Publishers, Dordrecht, The Netherlands.
36. Muyzer, G., E. C. de Waal, and G. A. Uitterlinden. 1993. Profiling of complex
populations by denaturing gradient gel electrophoresis analysis of polymer-
ase chain reaction-amplified genes coding for 16S rRNA. Appl. Environ.
37. Muyzer, G., and K. Smalla. 1998. Application of denaturing gradient gel
electrophoresis (DGGE) and temperature gradient gel electrophoresis
(TGGE) in microbial ecology. Antonie Leeuwenhoek 73:127–141.
38. Naito, S., H. Hayashidani, K. Kaneko, M. Ogawa, and Y. Benno. 1995.
Development of intestinal lactobacilli in normal piglets. J. Appl. Bacteriol.
39. Pluske, J. R., D. W. Pethick, D. E. Hopwood, and D. J. Hampson. 2002.
Nutritional influence on some major enteric disease in pigs. Nutr. Res. Rev.
40. Pluske, J. R., P. M. Siba, D. W. Pethick, Z. Durmic, B. P. Mullan, and D. J.
Hampson. 1996. The incidence of swine dysentery in pigs can be reduced by
feeding diets that limit the amount of fermentable substrate entering the
large intestine. J. Nutr. 126:2920–2933.
41. Pryde, S. E., A. J. Richardson, C. S. Stewart, and H. J. Flint. 1999. Molecular
analysis of the microbial diversity present in the colonic wall, colonic lumen,
and cecal lumen of a pig. Appl. Environ. Microbiol. 65:5372–5377.
42. Rastall, R. A., and G. R. Gibson. 2002. Prebiotic oligosaccharides: evaluation
of biological activities and potential future developments, p. 107–148. In
G. W. Tannock (ed.), Probiotics and prebiotics: where are we going?, vol. 1.
Caister Academic Press, Wymondham, United Kingdom.
43. Reid, C. A., and K. Hilman. 1999. The effect of retogradation and amylose/
amylopectin ratio on starches and carbohydrates fermentation and microbial
populations in the porcine colon. Anim. Sci. 68:503–510.
44. Reid, G. 1999. The scientific basis for probiotic strains of Lactobacillus. Appl.
Environ. Microbiol. 65:3763–3766.
45. Reid, G., J. Howard, and B. S. Gan. 2001. Can bacterial interference prevent
infection? Trends Microbiol. 9:424–428.
46. Rolfe, R. 1997. Colonization resistance. In R. I. Mackie, B. A. White, and
R. E. Isaacson (ed.), Gastrointestinal microbiology, vol. 2. Chapman and
Hall, New York, N.Y.
47. Rycroft, C. E., M. R. Jones, G. R. Gibson, and R. A. Rastall. 2001. A
comparative in vitro evaluation of the fermentation properties of prebiotic
oligosaccharides. J. Appl. Microbiol. 91:878–887.
48. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a
laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold
Spring Harbor, N.Y.
49. Sanguinetti, C. J., E. Dias Neto, and A. J. G. Simpson. 1994. Rapid silver
staining and recovery of PCR products separated on polyacrylamide gels.
50. Shannon, C. E., and W. Weaver. 1963. The mathematical theory of commu-
nication. University of Illinois Press, Urbana, Ill.
51. Simpson, J. M., V. J. McCracken, H. R. Gaskins, and R. I. Mackie. 2000.
Denaturing gradient gel electrophoresis analysis of 16S ribosomal DNA
amplicons to monitor changes in fecal bacterial populations of weaning pigs
after introduction of Lactobacillus reuteri strain MM53. Appl. Environ. Mi-
52. Simpson, J. M., V. J. McCracken, B. A. White, H. R. Gaskins, and R. I.
Mackie. 1999. Application of denaturant gradient gel electrophoresis for the
analysis of the porcine gastrointestinal microbiota. J. Microbiol. Methods
53. Smith, D. L., A. D. Harris, J. A. Johnson, E. K. Silbergeld, and J. G. Morris,
Jr. 2002. Animal antibiotic use has an early but important impact on the
VOL. 70, 2004EFFECTS OF PREBIOTICS ON PIGLET INTESTINAL MICROBIOTA 3829
emergence of antibiotic resistance in human commensal bacteria. Proc. Natl.
Acad. Sci. USA 99:6434–6439.
54. Spreeuwenberg, M. A. M., J. M. A. J. Verdonk, H. R. Gaskins, and M. W. A.
Verstegen. 2001. Small intestine epithelial barrier function is compromised in
pigs with low feed intake at weaning. J. Nutr. 131:1520–1527.
55. Statistical Analysis Systems Institute. 1989. SAS/STAT user’s guide, version
6, 4th ed., vol. 2. Statistical Analysis Systems Institute, Cary, N.C.
56. Steel, R. G. D., and J. H. Torrie. 1980. Principles and procedures of statistics,
a biometrical approach, 2nd ed. McGraw-Hill, Tokyo, Japan.
57. Stewart, C. S. 1997. Microorganisms in hindgut fermentors, p. 142–186. In
R. I. Mackie, B. A. White, and R. E. Isaacson (ed.), Gastrointestinal micro-
biology, vol. 2. Chapman and Hall, New York, N.Y.
58. Tannock, G. 1999. Probiotics. A critical review. Horizon Scientific Press,
Wymondham, Norfolk, United Kingdom.
59. Tannock, G. W. 2001. Molecular assessment of intestinal microflora. Am. J.
Clin. Nutr. 73:410S–414S.
60. Tannock, G. W., R. Fuller, and K. Pedersen. 1990. Lactobacillus succession
in the piglet digestive tract demonstrated by plasmid profiling. Appl. Envi-
ron. Microbiol. 56:1310–1316.
61. Teitelbaum, J. E., and W. A. Walker. 2002. Nutritional impact of pre- and
probiotics as protective gastrointestinal organisms. Annu. Rev. Nutr. 22:107–
62. Van der Waaij, D. 1989. The ecology of the human intestine and its conse-
quences for the overgrowth of pathogens such as Clostridium difficile. Annu.
Rev. Microbiol. 43:69–87.
63. Vaughan, E. E., F. Schut, H. G. H. J. Heilig, E. G. Zoetendal, W. M. de Vos,
and A. D. L. Akkermans. 2000. A molecular view of the intestinal ecosystem.
Curr. Issues Intest. Microbiol. 1:1–12.
64. Verstegen, M. W., and B. A. Williams. 2002. Alternatives to the use of
antibiotics as growth promotors for monogastric animals. Anim. Biotechnol.
65. von Wintzingerode, F., U. B. Go ¨bel, and E. Stackebrandt. 1997. Determina-
tion of microbial diversity in environmental samples: pitfalls of PCR-based
rRNA analysis. FEMS Microbiol. Rev. 21:213–229.
66. Williams, B. A., M. W. A. Verstegen, and S. Tamminga. 2001. Fermentation
in the large intestine of single-stomached animals and its relationship to
animal health. Nutr. Res. Rev. 14:207–227.
67. Zoetendal, E. G., A. D. L. Akkermans, and W. M. de Vos. 1998. Temperature
gradient gel electrophoresis analysis of 16S rRNA from human fecal samples
reveals stable and host-specific communities of active bacteria. Appl. Envi-
ron. Microbiol. 64:3854–3859.
68. Zoetendal, E. G., K. Ben-Amor, A. D. Akkermans, T. Abee, and W. M. de
Vos. 2001. DNA isolation protocols affect the detection limit of PCR ap-
proaches of bacteria in samples from the human gastrointestinal tract. Syst.
Appl. Microbiol. 24:405–410.
69. Zoetendal, E. G., C. T. Collier, S. Koike, R. I. Mackie, and H. R. Gaskins.
2004. Molecular ecological analysis of the gastrointestinal microbiota: a
review. J. Nutr. 134:465–472.
3830 KONSTANTINOV ET AL.APPL. ENVIRON. MICROBIOL.