OBJECTIVE:

The gut microbiota may contribute to the onset and maintenance of irritable bowel syndrome (IBS). In this study, the microbiotas of patients suffering from IBS were compared with a control group devoid of gastrointestinal (GI) symptoms.

METHODS:

Fecal microbiota of patients (n = 27) fulfilling the Rome II criteria for IBS was compared with age- and gender-matched control subjects (n = 22). Fecal samples were obtained at 3 months intervals. Total bacterial DNA was analyzed by 20 quantitative real-time PCR assays covering approximately 300 bacterial species.

RESULTS:

Extensive individual variation was observed in the GI microbiota among both the IBS- and control groups. Sorting of the IBS patients according to the symptom subtypes (diarrhea, constipation, and alternating predominant type) revealed that lower amounts of Lactobacillus spp. were present in the samples of diarrhea predominant IBS patients wheras constipation predominant IBS patients carried increased amounts of Veillonella spp. Average results from three fecal samples suggested differences in the Clostridium coccoides subgroup and Bifidobacterium catenulatum group between IBS patients (n = 21) and controls (n = 15). Of the intestinal pathogens earlier associated with IBS, no indications of Helicobacter spp. or Clostridium difficile were found whereas one case of Campylobacter jejuni was identified by sequencing.

CONCLUSIONS:

With these real-time PCR assays, quantitative alterations in the GI microbiota of IBS patients were found. Increasing microbial DNA sequence information will further allow designing of new real-time PCR assays for a more extensive analysis of intestinal microbes in IBS.

INTRODUCTION

Irritable bowel syndrome (IBS) is a common gastrointestinal (GI) disorder characterized by abdominal pain and disturbances in the bowel function. The causes of IBS are poorly known. As the symptoms of IBS vary from constipation to diarrhea, causes underlying this syndrome may well be heterogeneous. Psychosomatic factors, altered GI motility, and visceral hypersensitivity have been proposed as possible reasons behind IBS (¹). Also, certain enteric pathogens have been associated with the onset of IBS (^2,3,4). Recently, presence of low-level inflammation in the GI mucosa of the IBS patients was observed in colonoscopic biopsy specimens of IBS patients (⁵). The GI normal bacteria may likewise contribute to the onset and maintenance of IBS. Indications of the GI tract microbiota abnormalities have been obtained by comparing the fecal bacterial populations of IBS patients and healthy controls (⁶) and by analysis of bowel fermentation patterns (^7,8,9). Significance of the GI tract bacteria in IBS has, however, remained inadequately studied.

The GI tract normal microbiota works as a barrier against pathogens and stimulates the host immune system; furthermore, the microbial metabolism influences the host both in beneficial and disadvantageous ways. The GI microbiota of an adult human comprises more than 500 species, with predominance of obligate anaerobes. Culture-independent studies have established that only a fraction of the organisms present in the feces are cultivable (^10,11); therefore, the results obtained by cultivation are likely to be biased. In this study, we wanted to test whether the previously suggested presence of an altered microbiota in IBS could be verified with modern molecular approaches. Recent advances in the bacterial taxonomy based on molecular methods, especially comparison of the 16S ribosomal RNA genes, facilitate the design of molecular tools targeting taxonomically valid bacterial groups. Here, we applied real-time PCR for comparing the fecal microbiota of patients fulfilling the Rome II criteria for IBS (¹²) with age- and sex-matched controls devoid of GI symptoms. The PCR assays used were designed to enable detection and quantification of 20 different bacterial species or groups, with the overall coverage of approximately 300 species.

MATERIALS AND METHODS

Subjects

Experienced physicians recruited 27 well-diagnosed IBS patients to the study (Table 1). Patients fulfilled the Rome II criteria for IBS (¹²), except for 3 subjects who reported slightly less than 12 wk of abdominal pain during the preceding year. The study was conducted in connection with a clinical 6-month trial, and the participants comprised the placebo group in this trial. During the trial, participants in the placebo group ingested one 200 mg placebo capsule daily, consisting of microcrystalline cellulose, magnesium stearate, and gelatine as encapsulating material. Altogether, 22 patients completed the clinical study.

Table 1 - Characteristics of the Patients (n = 27) and Healthy Controls (n = 22).

Full table

All patients had undergone clinical investigation and endoscopy or barium enema of the GI tract 0–1 yr prior to the study. Patients pregnant, lactating, with organic intestinal diseases or other severe systematic diseases, previous major or complicated abdominal surgery, severe endometriosis, or dementia, or otherwise inadequate cooperation were excluded. Subjects were also excluded if they had received antimicrobial medication during the previous 2 months. Patients with lactose intolerance were included if they reported to follow a low-lactose or lactose-free diet.

All the patients gave their written informed consent and were told that they could withdraw from the study at any time. The Human Ethics Committee at The Joint Authority for the Hospital District of Helsinki and Uusimaa (HUS) approved the study protocol.

Healthy control subjects devoid of GI symptoms were recruited to form an age- and gender-matched control group for the IBS patients (Table 1). Intestinal disturbances (including lactose intolerance and celiac disease) and ongoing antibiotic treatments were considered as exclusion criteria for control group.

Study Protocol

During the 6-month study period, the participants gave three fecal samples: at the beginning of the study, at 3 months, and at 6 months. The participants were instructed to follow their ordinary diet and way of living. Patients who had a regular IBS medication (n = 10; mainly commercial fiber analogues, antidiarrheals, laxatives) were allowed to continue the medication throughout the study.

Fecal samples were stored anaerobically immediately after defecation, aliquoted, and stored at -70°C within 4 h of delivery. For real-time PCR analysis, total DNA was isolated from the feces as described by Apajalahti et al. (¹³). Even though 22 IBS patients completed the clinical study, all three samples could be analyzed with real-time PCR from only 21 patients due to technical issues in sample handling.

Bacterial Strains and Growth Conditions/Control DNAs

Genomic DNAs from Atopobium parvulum ATCC 33793, Bacteroides fragilis DSM 2151, Bifidobacterium adolescentis DSM 20083, Bifidobacterium bifidum DSM 20456, Bifidobacterium longum DSM 20219, B. pseudocatenulatum DSM 20438, Campylobacter jejuni Neqas 6037, Clostridium difficile ATCC 9689, C. perfringens ATCC 13124, Desulfovibrio desulfuricans ATCC 7757, Enterococcus faecalis DSM 20478, Escherichia coli DSM 6897, Fusobacterium nucleatum ATCC 25586, Helicobacter pylori DSM 4867, Lactobacillus acidophilus ATCC 4356, L. casei ATCC 393, Ruminococcus productus DSM 2950, and Veillonella parvula ATCC 10790 were used as positive and negative controls in real-time PCR. With Fusobacterium prausnitzii specific PCR assay, however, the PCR amplified 16S rRNA gene of F. prausnitzii ATCC 27766 was used instead of the genomic DNA.

Measurement of DNA Concentration

DNA concentrations were measured from the standard DNA and fecal DNA preparations with a Versafluor fluorometer (Bio-Rad, USA).

Real-Time PCR

The oligonucleotides and optimized PCR conditions used in this study are summarized in Table 2. Papers describing the targets for the assays in detail are listed in Table 2. Briefly, the following assays were designed genus-specific: Bifidobacterium spp., Campylobacter spp., Desulfovibrio spp., Enterococcus spp., Fusobacterium spp. (with F. prausnitzii omitted), and Veillonella spp. The PCR primers for Bacteroides-Prevotella-Porphyromonas as well as Helicobacter-Flexispira-Wolinella were set to amplify bacterial targets from the above-mentioned genera. Assays for Atopobium group, Clostridium coccoides group, C. perfringens group, and E. coli group targeted phylogenetically related species classified in several genera. The remaining tests were species specific or detected a small number of closely related species.

Table 2 - Oligonucleotides Used in this Study for the Real-Time PCR Assays.

Full table

Quantitative PCR was performed with an iCycler iQ apparatus (Bio-Rad, USA) associated with the iCycler optical system interface software (version 2.3; Bio-Rad). All PCRs were performed in triplicate in a volume of 25 l, using 96-well optical grade PCR plates and an optical sealing tape (Bio-Rad). An aliquot of 50 ng of each fecal DNA preparation was subjected to real-time PCR reactions.

Reaction mixtures for the optimized SYBR Green I-based assays consisted of 1:75,000 dilution of SYBR Green I (Molecular Probes), 10 mM Tris-HCl (pH 8.8), 150 mM KCl, 0.1% Triton X-100, 2–5 mM MgCl₂, 100 M each dNTP, 0.5 M each primer and 0.02 U Dynazyme II l^-1 (Finnzymes, Finland), and either 5 l template or water (no-template control). Concentrations of the reaction components for each PCR assay are summarized in Table 2. The thermal cycling conditions used were an initial DNA denaturation step at 95°C for 5 min followed by 35 cycles of denaturation at 95°C for 15 s, primer annealing at optimal temperature (See Table 1) for 20 s, extension at 72°C for 30 s, and an additional incubation step at 80–85°C for 30 s to measure the SYBR Green I fluorescence. Finally, melt curve analysis was performed by slowly cooling the PCRs from 95 to 60°C (0.3°C per cycle) with simultaneous measurement of the SYBR Green I signal intensity.

The reaction mixtures of 5'-nuclease (TaqMan) assays consisted of 10 mM Tris-HCl (pH 8.8), 150 mM KCl, 0.1% Triton X-100, 2–4 mM MgCl₂, 200 M each dNTP, 1 M each primer, 80 nM of fluorescent probe and 0.03 U Dynazyme II l^-1 (Finnzymes, Finland), and either 5 l template or water (see Table 2 for detailed information on the concentrations of the reaction components). An initial denaturation at 95°C was followed by 30–40 cycles of denaturation at 95°C for 15 s, and a combined incubation step for primer annealing and extension, during which the fluorescent signal was also measured.

Sequencing of PCR Products Obtained with Primers for Campylobacter spp.

In addition to real-time PCR targeting the Campylobacter genus, sequencing of the PCR products (approximately 246 bp) obtained from the assay in question was used for putative identification of the campylobacteria present. Two sequencing reactions with forward and reverse primers of the Campylobacter spp. specific PCR (see Table 2) were performed from each PCR sample using the Big Dye Terminator chemistry (Applied Biosystems, USA). The thermal cycling protocols were carried out in a Peltier Thermal Cycler PTC-200 (MJ Research, USA). The sequencing reactions were purified according to the protocol provided by the sequencing kit manufacturer, and sequenced in ABI 310 Genetic Analyzer (Applied Biosystems, USA). The sequences were processed with SequencherTM 3.0 sequence analysis software (Gene Codes Corporation, USA) and a database search was then performed against the GeneBank sequences.

Data Analysis

All real-time PCR reactions were performed in triplicate. For each assay, results obtained by PCR were converted to the average estimate of target bacterial genomes present in 1 g of feces (wet weight). Majority of the PCR tests were set to detect several bacterial species, most likely with varying ribosomal DNA copy numbers and genome sizes, therefore, estimated average genome sizes for each target bacteria group were used while differences in the rrn copy numbers had to be omitted. The following genome sizes were used: 2.3 Mb for Lactobacillus spp., 2 Mb for Bifidobacterium spp., 3 Mb for Enterococcus spp., 4 Mb for C. coccoides group, 2.2 Mb for Fusobacterium spp., 3 Mb for Veillonella spp., 3 Mb for C. perfringens group, 3 Mb for Desulfovibrio spp., 3 Mb for Atopobium group, 4 Mb for Bacteroides-Prevotella-Porphyromonas group, 4.6 Mb for E. coli subgroup, and 5.2 Mb for B. fragilis. The amounts of F. prausnitzii genomes in fecal samples were estimated assuming that the rrn copy number of the species in question was 10.

The Graph Prism version 3.02 (GraphPad Software, Inc) was used for statistical analyses of the data. Kruskal-Wallis signed rank test with Dunn's posttest was used for comparison of microbiota among the control-, diarrhea-predominant IBS, mixed-type IBS, and constipation-predominant IBS groups. Mann-Whitney U test was used for pairwise comparison of the IBS patients with healthy controls.

RESULTS

Fecal Microbiota of Control versus IBS Subjects at the Beginning of the Study

During a 6-month period, fecal microbiota of the IBS patients and healthy controls were analyzed at 3-month intervals. Using real-time PCR, a total of 20 bacterial groups (covering about 300 different species) was quantified from the fecal DNA samples. The target bacteria groups were selected either on basis of previous reports suggesting a connection with IBS for the bacteria in question, or due to the predominant nature of these bacteria in the gut ecosystem. In the beginning of the study, the patients were divided into diarrhea-, constipation-, or alternating-type IBS groups according to their symptoms. However, the typical symptoms experienced by an individual IBS patient may fluctuate between different time periods. Therefore, the fecal samples collected in the beginning of the study period were considered to represent best the typical microbiota in each IBS subtype.

Results for 17 real-time PCR tests are presented in Table 3 as mean values and upper confidence limits of target bacterial genomes in 1 g of feces (wet weight) for each subject group. In general, an extensive, though expected, variation was observed between the microbiota of individuals. Obviously, the natural, subject-specific deviation in bacterial counts can easily hide any real group-wise differences in mean bacterial counts. To overcome this, Kruskal-Wallis signed rank analysis was used for group-wise comparison of the subject groups (Table 4). Significantly lower amounts of lactobacilli (p < 0.019) were observed in connection with diarrhea predominant IBS than in constipation predominant IBS; these two patient groups seemed to form the extremities of the counts for Lactobacillus spp (Table 4, Fig. 1A). Also, counts for Veillonella spp. were significantly higher (p < 0.045) with the constipation predominant IBS group than with healthy controls (Table 4, Fig. 1B). The IBS patients also had higher levels (p < 0.04) of R. productus and C. coccoides species (Table 4). Other target bacteria analyzed within this study did not show statistically significant differences in the Kruskal-Wallis analysis between the four subject groups (Table 4); however, the diarrhea predominant IBS patients seemed to have lower amounts of Desulfovibrio spp. (Table 4), and Bifidobacterium spp. (Table 4, Fig. 1C) than healthy control subjects or patients diagnosed with constipation predominant or alternating type IBS.

Figure 1.

Box and whiskers plots presenting the amounts of genomic DNAs for (A) Lactobacillus spp., (B) Veillonella spp., and (C) Bifidobacterium spp. in 1 g fecal samples (wet weight) of control subjects and patients diagnosed with diarrhea, alternating type or constipation predominant IBS. The box extends from 25th percentile to 75th percentile, with a line at the median (50th percentile); the whiskers extending above and below the box show the highest and lowest values for the target genomic DNA.

Full figure and legend (25K)

Table 3 - Genomic Targets in the Feces of IBS Patients and Healthy Control Subjects.

Full table

Table 4 - Kruskal-Wallis Analysis for 0-Months Samples of the Control Subjects and Patients Suffering from Diarrhea, Alternating Type, or Constipation Predominant IBS.

Full table

Typical Fecal Microbiota of Control versus IBS Subjects During the Follow-Up Period

To obtain an image of the typical fecal microbiota of the IBS patients, subjects with three samples were analyzed by studying the mean results of PCR assays for the samples. The required three fecal samples were obtained from only 21 out of 27 IBS patients and 15 out of 22 control subjects, which further reduced the group sizes for different IBS subtypes. Considering this reduction in subjects as well as the possibility for temporal variations often occurring in the symptoms of IBS, combining the IBS subjects as one group was thought reasonable. No significant differences were seen between the mean values for any of the target bacteria (data not shown). However, comparison of the two groups with the nonparametric Mann-Whitney U test revealed significant differences between the controls and IBS patients for C. coccoides group (p < 0.003) and B. catenulatum group (p < 0.039) (Table 5, Figs. 2A and B).

Figure 2.

Box and whiskers plots presenting the mean amounts of genomic DNAs for (A) C. coccoides group, and (B) B. catenulatum group in three fecal samples taken at 3 months intervals for control subjects and IBS patients. The box extends from 25th percentile to 75th percentile, with a line at the median (50th percentile); the whiskers extending above and below the box show the highest and lowest values for the target genomic DNA.

Full figure and legend (13K)

Table 5 - Mann-Whitney U Test Results for Quantitative Real-Time PCR Analysis from Three Samples (0, 3, and 6 Months after Beginning of the Study) of 21 IBS Patients and 15 Control Subjects.

Full table

Presence of Intestinal Pathogens

Occurrence of Campylobacter spp., Helicobacter spp., and C. difficile in the feces of IBS and control subjects was analyzed. No indications of the presence of Helicobacter spp. or C. difficile were found in either of the subject groups. The occurrence of Campylobacter spp. was indicated in the samples of five IBS patients, whereas no positive cases were observed among control subjects. Sequencing of the PCR products obtained from Campylobacter genus PCR (Table 6) confirmed this finding. However, just one patient diagnosed with an alternating type IBS harbored C. jejuni in feces (Table 6), whereas the other four patients with indications of Campylobacter spp. harbored either commensal (C. hominis) or mouth campylobacteria (Table 6).

Table 6 - Detection and Identification of Fecal Campylobacteria by Sequencing.

Full table

DISCUSSION

Although the knowledge of the functions of various microbes present in the GI tract is still limited, their overall importance to human health has become acknowledged. The gut normal microbiota is generally crucial to the host well being; yet bacteria belonging to the normal microbiota seem also capable of causing disease in some individuals or conditions. Involvement of the GI tract microbiota in the pathogenesis of the inflammatory bowel disease has been suggested (^14,15). Recently, an association was observed with sulfate reducing bacteria and ankylosing spondylitis (¹⁶). Altered GI microbiota with decreased bifidobacterial counts seem to even lie behind development of allergies (^17,18).

The gut normal microbiota may also have an important role in the development and maintenance of IBS. Evidence suggesting altered microbiota in IBS has been gathered by conventional microbiological methods (⁶), analysis of bowel fermentation patterns (^7,8,9), and by determination of the presence of single pathogenic microbes (¹⁹). A different fermentation pattern, particularly with an increased production of hydrogen, has been demonstrated for patients with IBS (⁷). Furthermore, application of an exclusion diet or antibiotics has been shown to reduce the symptoms of IBS and to shift the fermentation patterns towards normality (^7,8). Ingestion of probiotic administrations has been reported to improve the symptoms of IBS (^20,21,22), which further supports the theory of microbial imbalance.

Nevertheless, studies on the microbiota of IBS patients and the number of study subjects used have been surprisingly limited. A comprehensive scanning of the gut microbial groups as well as utilization of modern molecular methods in this context have also been lacking. Therefore, we analyzed the GI microbiota of IBS patients and healthy controls using a set of real-time PCR assays covering approximately 300 bacterial species. A total of 27 IBS patients and 22 controls devoid of GI symptoms were recruited and the microbiota of the patients was followed in three time points. Although the subject numbers may seem small, we feel that the almost total lack of preceding research makes our study, at least to some extent, a preliminary work. In addition, comparison of the results from various studies concerning alterations of fecal microbiota in IBS is relatively difficult. First, the diagnosis for IBS varies depending on the criteria used. A recent survey across eight European countries reported 11.5% (6.2–12%) overall prevalence for IBS, but only 2.9% met the Rome II criteria (²³). Therefore, usage of the strictest criteria for IBS most likely results in a different population of patients when compared to more expanded criteria. It is not always clear which standards have been used for selection of the IBS patients in various studies. Another point worth considering is the evolution of the bacterial taxonomy together with development of new, nucleic acid based methods for the analysis of bacterial groups. The conventional bacterial genera (e.g., Clostridium and Eubacterium) are no longer valid, and it is therefore sometimes difficult to determine which bacterial groups have actually been analyzed in certain studies.

Generally, variation observed in this study between subjects was extensive. This was anticipated, as the host-specificity of the GI microbiota has been described previously (^24,25). However, despite this variation, some differences in the GI microbiota were seen between healthy controls and IBS patients. Balsari et al. (⁶) reported qualitatively similar fecal floras for IBS patients and healthy controls; quantitatively, however, a decrease was observed in the coliforms, lactobacilli, and bifidobacteria in the IBS patients' samples. Likewise, we could also observe the overall qualitative similarities of the fecal microbiota for IBS patients and controls (Table 3). Unlike Balsari et al. (⁶), we could not observe differences in the total Bifidobacterium spp. between the IBS patients and healthy controls; however, differences were observed in the B. catenulatum group when the pooled results from 3 fecal samples of 21 patients and 15 healthy controls were analyzed (Table 5, Fig. 2B). Similarly, bacterial counts for C. coccoides subgroup were significantly lower for the IBS patients than for the control group subjects (Table 5, Fig. 2B). We also saw a decrease in lactobacilli among patients diagnosed as diarrhea predominant (Table 4, Fig. 1A), which is well in accordance with Balsari et al. (⁶). Instead, constipation predominant or alternating type IBS were not characterized by changes in the amounts of lactobacilli. In a recent study, constipation predominant subjects showed an increased tendency for methane excretion whereas diarrhea predominant subjects did not excrete methane; furthermore, the severity of the constipation had a positive correlation with methane excretion (⁸). Indeed, patients with IBS suffer from a diverse range of symptoms and therefore, it is reasonable to assume that different microbiota could be associated with the IBS subgroups.

A role has been suggested for enteric pathogens, especially C. jejuni, in the onset of IBS (^2,4,26). Postinfection IBS is typically associated with diarrheal symptoms and a better recovery rate than noninfective IBS cases (²⁷). In the beginning of the study, 44% of the IBS patients were diagnosed as diarrhea predominant (Table 1); still, no support for postinfective IBS cases among the IBS patients was found when the patients were interviewed for diagnosis. In this study, we could not establish a connection between C. difficile or Helicobacter spp. and IBS. Five out of 25 patients were positive for Campylobacter spp. in real-time PCR analysis, and one case with C. jejuni carriage was identified. The observed prevalence of 4% for C. jejuni is in accordance with the previously reported relationship between this pathogen and IBS (^4,26); however, this is a solitary finding. Asymptomatic carriage rates of 0.1% have been reported for pathogenic Campylobacter spp. by Hellard et al. (²⁸), indicating that the long-term colonization of the GI tract by a pathogenic Campylobacter species is a rare event but on the other hand, healthy individuals have been shown to harbor frequently a commensal C. hominis in their GI tract (²⁹). Su et al. (³⁰) could show a high prevalence of functional dyspepsia associated with H. pylori infection, female gender, and stress in IBS. Locke et al. (³¹) could not observe a connection between H. pylori and IBS; nevertheless, there was a weak association with CagA antibody positivity and IBS. We could not detect any Helicobacter spp. from the feces of either IBS patients or healthy subjects, although an average seroprevalence rate of 31.4% (CI 95% 31.1–39.5) has been reported for H. pylori antibodies for randomly selected Finnish subjects aged 15–74 yr (³²). Total lack of positive signals for Helicobacter spp. in both the IBS and control groups could therefore indicate that target DNA represented too small proportion of the total to be detected with our PCR assay.

A drawback in this study was the high dropout rate of the participants (6/27 IBS patients and 7/22 controls). This could be partly explained by the long duration (6 months) of the study. The reasons for withdrawal of the IBS patients were as follows: illness or hospitalization due to reasons other than IBS (3 subjects), pregnancy (2 subjects), and noncompliance (1 subject). Consequently, analysis results of the microbiota over the 6-months period cannot be directly compared with the results of samples taken at the beginning of the study. During the 6-month follow-up period, the IBS patients ingested a placebo capsule containing cellulose daily (see Materials and Methods). Although cellulose could affect the gut microbiota, the daily amount of cellulose in the capsule was negligible (<200 mg) and therefore it is unlikely that the placebo treatment could have caused the differences between analyses of pooled samples and samples taken at the starting point. Another issue concerning the analysis of microbiota arises from the fact that 10 out of 27 IBS patients had regular IBS medication. The great majority of the medications were commercial fiber analogues, but there was also one subject taking laxatives, one taking antidiarrheals, and one taking spasmolytics. All patients were already on medication prior to the study and continued it throughout the study. Hence, it is assumed that the microbiota was stable throughout the trial. No statistically significant differences were observed between the microbiota of IBS patients receiving medication and IBS patients not on medication when these two groups were compared (data not shown). It must be pointed out, though, that the patients received different medications and therefore, the possible effects on the microbiota are most likely dissimilar.

Multiple testing (20 different real-time PCR assays and subsequent statistical analyses) was performed in this work. Considering the heterogeneous nature of the test group (age, gender, medication) and small patient numbers, this may have lead into getting statistically significant results by chance. However, it seems to us more likely that small subject numbers and the subject-to-subject variation hindered observation of more precise patterns of the microbial content in the feces of IBS patients. Indeed, further studies are needed to confirm this study. Technically, however, the real-time PCR provides the so far the most precise method for analysis of GI microbiota.

The possible implications of the observed microbial differences between IBS patients and the healthy subjects, or between different IBS subgroups, remain unclear. Little is yet known about the relevance of individual microbial species or strains on the GI health. An imbalanced microbiota has been associated with some diseases, but the ability of certain bacterial species, or strains to impair, or enhance GI health is poorly known. This complicates the interpretation of the clinical significance. It is, nonetheless, known that an imbalanced microbiota during for instance antibiotic treatment can give rise to GI symptoms. The aim of the study was to compare the microbiota in IBS patients and healthy controls, but the importance of individual species or strains of the normal microbiota in giving rise to symptoms requires further studies. In conclusion, this study supports the earlier suggestions of the changed GI microbiota in IBS. Fecal microbiota of the IBS subgroups seem to differ from each other, which is reasonable considering the different kinds of symptoms experienced by these patients. However, we are aware that most intestinal bacteria are still uncharacterized, and may therefore have escaped the PCR analyses. To overcome this, we have extended our analyses in sequencing of 16S ribosomal RNA gene libraries, generated from group-wise pooled fecal DNA samples of the four subject groups. We expect to uncover more details regarding the intestinal microbiota and IBS in the future.

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Acknowledgements

This study was supported by the Finnish Technology Centre (TEKES). We thank Sinikka Ahonen for excellent technical assistance. We also thank Department of Microbiology, National Public Health Institute, Helsinki, Finland for kindly providing bacterial strains.