Chronic inflammatory bowel diseases (IBD) are the ailments of the gastrointestinal (GI) tract of unknown etiology. Within IBD, there are 3 distinct disease entities that show different clinical symptoms and affect different parts of the GI tract: Crohn disease (CD), ulcerative colitis (UC), and an indirect form—nonspecific indeterminate colitis. Undoubtedly, these are inflammatory conditions, which can be confirmed by histological examination of the tissue samples collected from the GI tract of patients, yet the cause of these changes remains unknown. Apart from the activity of the gut-associated lymphoid tissue (GALT), the genetic predisposition and the influence of the commensal bacteria have been postulated.
The commensal microbiota of the GI tract plays an important role in the process of digestion and absorption of nutrients and displays the protective function against the invasion of pathogenic microorganisms by both creating the resistance to colonization and affecting the immune system of the host (1). The number of bacterial cells living in the human intestines equals approximately 1014, that is, 100 trillion. In 1 mL of intestinal content, there are about 1011 bacterial cells (2). The microbiota of the GI tract mainly consists of anaerobic bacteria (Clostridium, Eubacterium, Bacteroides, and Bifidobacterium) and, in smaller amounts, species of aerobic and facultative anaerobic bacteria (Enterobacteriaceae, Streptococcus, Staphylococcus, and Lactobacillus) (3). So far, owing to the 16S rRNA sequence analysis, >400 bacterial species in the intestinal ecosystem have been identified, the majority of which is noncultivable (2,4). Some authors, such as Frank et al and Vitali et al (5,6), point out that the number of bacteria in the human GI tract can reach >1000 various species. This intestinal ecosystem is called a microbiome (2,5). It should be emphasized that the intestinal microbiota in the fecal content varies quantitatively considering the specific types and species of bacteria from the microbiota adhering to the mucosa and intestinal crypts (4,7). The passage of food takes place in the GI tract and, consequently, part of the microbiota called planktonic (or transient) material, is quickly excreted with the fecal mass. As a result, the quantitative and qualitative composition of the planktonic flora is significantly varied and dependent mostly on the type of diet. In a healthy person, intestinal bacteria do not have direct contact with the epithelium of the intestinal wall because they are separated from it with the mucus layer (8,9). We assume that in the GI tract, the composition of the microbiota is not homogeneous in the transverse section of the large intestine, and it is determined as it gets closer to the intestinal mucosa. In the literature, apart from the studies by Swidsinski et al (10) examining the so-called biostructure of the intestinal microbiota, there have been no reports concerning this subject (11). Therefore, the question arises whether there is intestinal microbiota diversity in the intestinal transverse section in adolescents with IBD as compared with healthy people, which was the main objective of this work. Three consecutive stool fractions obtained in the subsequent phases of preparing patients for colonoscopy and colon tissue samples were examined for this purpose. In addition, the results were compared with the use of quantitative cultures as well as fluorescent in situ hybridization (FISH). Moreover, the influence of bacterial flora on the mucin degradation in fraction III and attached to the mucosal layer was checked.
PATIENTS AND METHODS
The study included 61 patients ages 1 to 18 years (Table 1), hospitalized at the Department of Pediatrics, Gastroenterology, and Nutrition of the Jagiellonian University Medical College in Krakow. Clinical characteristics of patients with IBD are described in Table 2. The inclusion criteria were bloody diarrhea, the presence of fecal occult blood, chronic diarrhea, abdominal pain and/or poor weight gain, and histopathological diagnosis of CD or UC. The exclusion criteria included children younger than 1 year, patients treated with antibiotics up to 30 days before enrollment in the study, confirmed infectious background of the disease, confirmed malabsorption or maldigestion syndromes, primary secretory enteropathies, solely breast-feeding, immunodeficiencies, and children whose parents or legal guardians did not give written consent for their children to participate in the study. The control group comprised the patients with functional disorders of the GI tract and excluded IBD.
All of the adolescents enrolled in the study were fed with a normal oral diet, adequate for age, in keeping with protein and energy daily requirements. They did not confirm drinking alcohol and smoking. We assessed the nutritional status of the patients using the Cole index (12).
Before the planned colonoscopy procedure, the patients underwent the standard preparation. For 2 days before colonoscopy, all of the patients received a liquid diet and lactulose, followed by cleansing of the GI tract using the normal saline enema (0.9% NaCl 200–500 mL, depending on the size of the child) repeated 4 times and oral administration of the laxative sodium phosphate (0.6–0.8 mL/kg, maximal dose 45 mL, given in 2 doses). During this time, the patients were encouraged to drink as much as possible to improve the cleansing process. Normal saline enema was administered in the supine or left-side position and the children were encouraged to hold the fluid for 30 minutes. After each preparatory phase, the stool sample was collected to obtain the 3 fractions: I (before first enema), II (after first dose of sodium phosphate and second enema), and III (after the last saline enema). Colon biopsies from the mucosa were taken during colonoscopy. The aim of the examination of the intestinal content of each fraction was to determine whether the qualitative and quantitative composition of the microbiota present in the collected samples was similar to the mucosa of the biopsy samples, as well as whether it changed during the following washing. The stool samples in sterile foil bags were immediately frozen on dry ice. The biopsy samples were transferred directly into Schaedler anaerobic broth medium (Difco, BD, Franklin Lakes, NJ) with 10% glycerol. All of the procedures were performed as fast as possible, using sterile instruments and ensuring the integrity of the intestinal tissue. The colon tissue samples were immediately frozen on dry ice. Then the samples were transported to the laboratory of the Chair of Microbiology of the Jagiellonian University Medical College, where classical microbiological diagnostic tests and examination with the use of quantitative FISH were conducted.
To determine the quantitative composition of the bacterial microbiota adhering to the intestinal mucosa, the quantitative culture method was applied. All of the procedures were performed in an anaerobic chamber (MACS-MG 500 Work Station, DW Scientific, Shipley, UK) in N2 (85%) + H2 (10%) + CO2 (5%) atmosphere. Approximately 0.5 g of the stool samples and 0.02 to 0.05 g of tissue samples were precisely weighed and subsequently homogenized in 1 mL of Schaedler's medium (Oxoid, Hampshire, UK). The homogenate was serially diluted in Schaedler's medium (100 μL of homogenate + 900 μL of Schaedler's medium) and plated onto the solid media differentiating for aerobic and anaerobic bacteria: blood agar (Oxoid) for Streptococcus and the rest were aerobic bacteria, McConkey agar (Oxoid) for Enterobacteriaceae, bile esculin azid agar (Biocorp, Warsaw, Poland) for Enterococcus, MRS agar (Oxoid) for lactic acid bacteria, Wilkins Chalgren agar (Oxoid) for Bacteroides, and agar for Bifidobacterium(13). Aerobic bacteria were cultured in the aerobic conditions at a temperature of 37°C for 24 hours, whereas cultures of anaerobes were grown in an anaerobic chamber for 72 hours. The cultivated colonies underwent the process of routine identification to determine the genus or species using Gram staining and API tests (bioMérieux, Warsaw, Poland) and afterwards they were counted. The results of the numbers of cells of the specific bacterial group in the weighed sample of the fecal and tissue mass were converted into 1 g of the mass to make the quantitative comparisons among the patients.
Simultaneously, the samples were subjected to quantitative FISH analysis. The stool sample was preliminarily homogenized using a stomacher device (Interscience, Markham, Canada) for 3 minutes. The content of the foil bag was transferred under sterile conditions to sterile Falcon 15-mL test tubes (Greiner Bio-One, Wemmel, Belgium). The samples were centrifuged at 3000 rpm for 10 minutes to remove saline applied in the enema solution before the colonoscopy procedure. The supernatant fluid was discarded and the sediment was precisely weighed. The amount of 0.5 g of the fecal sample was used for FISH analysis. It was suspended in 4500 μL of the sterile phosphate buffered saline (PBS). Four to 5 sterile glass beads of the diameter of 5 mm were added to the suspension and the whole was intensively vortexed using the laboratory mixer for 3 minutes. The test tube with the homogenate was centrifuged at 1500 rpm for 1 minute. Each tissue sample was precisely weighed. Then the tissue samples were homogenized in 1 mL of saline using a FastPrep machine (Qbiogene, Heidelberg, Germany) at 4.0 m/s for 1 minute. The fluid containing the bacteria was mixed with 4% paraformaldehyde solution (Sigma-Aldrich, Carlsbad, CA) in the proportions 1:3 to be fixed (ie, 300 μL of the supernatant fluid to 900 μL of paraformaldehyde). The process of sample fixation was carried out for 15 minutes at a temperature of 4°C. Afterward, 10 μL of the fixed suspension was placed onto a SuperFrostPlus (MENZEL-GLÄSER, Braunschweig, Germany) microscopic slide over an area of 1 cm2 with the use of sterile disposable loops and the whole was dried. After that, the slide was washed with PBS and 1 to 2 mL of 96% methanol (POCH, Gliwice, Poland) was poured over it. The whole was subsequently incubated under cover for 30 minutes at a temperature of −20°C. On completion of the fixation process, methanol was washed out with warm (50°C) PBS and 50 μL of the diluted solutions of lysozyme (1 mg/mL) and lysostaphin (0.05 mg/mL, Sigma-Aldrich) were placed into the preparation. The whole was incubated for 5 minutes at 37°C and rinsed with warm PBS. The hybridization was performed using the following probes (GENOMED, Warsaw, Poland): STREP—Streptococcus genus (14), 5′-GGT ATT AGC AYC TGT TTC CA-3′; Lab158—Lactobacillus and Enterococcus genera (15), 5′-GGT ATT AGC AYC TGT TTC CA-3′; Bif164—Bifidobacterium genus (16), 5′-CAT CCG GCA TTA CCA CCC-3′; BAC303—Bacteroides genus (17), 5′-CCA ATG TGG GGG ACC TT-3′; ECOLI—Escherichia coli(14), 5′-GCA AAG GTA TTA ACT TTA CTC CC-3′; Erec482—Clostridium coccoides (Blautia coccoides) (18,19), 5′-GCT TCT TAG TCA RGT ACC G-3′. To conduct the process of hybridization, 5 μL of the examined probe solution (50 ng/μL) was mixed with 45 μL of the hybridization buffer (20 mmol/L Tris HCl; 0.9 mol/L NaCl; 0.1% sodium dodecyl sulfate; pH 7.2) heated to 50°C. The prepared hybridization solution (50 μL) was transferred onto the preparation using an automatic pipette, and afterward, the slides were placed in the humid chamber covered with aluminum foil at 50°C for 3 hours. Having completed the hybridization process, the preparation was rinsed with warm washing buffer (of the same composition as the hybridization buffer except for sodium dodecyl sulfate). The preparation was stained with 4′,6′-diamino-2-phenylindole (Sigma-Aldrich) for 5 minutes. After that, it was thoroughly washed with sterile distilled water and dried in dark conditions. The specimen was analyzed using BX51 fluorescence microscope (Olympus, Warsaw, Poland) with the application of AnalySIS software (Soft Imaging, Olympus). To calculate the bacterial content in 1 g of the sample mass, a mathematical formula with the assumption that 0.5 g of the centrifuged fecal mass corresponds to the volume of 0.5 mL was applied:
Equation (Uncited)Image Tools
where NB is the number of bacteria in 1 g of the fecal mass; a, average number of bacteria calculated within 5 fields of view; fm, fecal mass in milligrams; fM, fecal mass in grams; 17217, number of fields of view within the area of 1 cm2; 120, conversion of the number of bacteria in the volume of fecal fixed samples (300 μL); 300, volume of the fixed sample (microliter); 100, conversion of the number of bacteria in the colon tissue sample in the volume of 1 mL of saline; 4500, volume of PBS buffer, which was used for fecal sample suspension (microliter); and cfu, colony-forming units.
Mucin Degradation Assay in a Petri Dish
To verify the ability of the fecal and mucosal flora to degrade mucin, the Zhou et al (20) method was used. Only the fecal fraction III and the mucosal tissue flora were examined in this assay because they were located nearest to the colon epithelium. The homogenized fecal and tissue samples were placed on the agar-containing medium B without glucose. After culture and staining, the diameter of mucin lysis zone (discolored halo) around the colonies of studied samples was measured.
The comparisons were made using the χ2 test for description of the total distribution of the studied bacterial taxons in the large intestinal content in the fecal fractions. The Steel test was used for analysis of differences between the groups of patients with CD and UC and controls for the absolute numbers of bacteria. The Mann-Whitney U test was used for the analysis of differences in the ability to degrade mucin. All of the analyses were conducted using SAS 9.1 package and SAS Enterprise Guide 3.0 (SAS Institute Inc, Cary, NC).
The study was approved by the Jagiellonian University bioethical committee (KBET/236/B/2002). Informed consent was obtained from all of the patients’ legal guardians and patients older than 16 years of age included in the study.
Using the culture method and the FISH technique, 3 consecutive stool fractions obtained during the preparation of the patients for colonoscopy and the colon tissue samples were examined. In the tissue samples, results were obtained using the culture method only. The FISH method has not allowed us to show the presence of bacterial cells in the samples. In further analyses, the patients experiencing indeterminate colitis were omitted because of the insignificant number of such cases (n = 3).
The analysis of the results obtained from the examination of the samples from patients with CD and UC proved that the quantitative composition of the bacterial microbiota changed in the consecutive fecal fractions and the tissue samples of the examined groups (Fig. 1A, B, D, E). In the patients in the control group who did not experience IBD, the composition of the bacterial microbiota of the consecutive fecal fractions and the tissue samples was similar (Fig. 1C, F). The described relation was demonstrated by both culture method and the FISH technique. The statistical analyses performed with the use of the χ2 test showed that the total distribution of the bacterial content of the examined taxons in all 3 fecal fractions and in the tissue samples in the specific disease entity and in the control group were characteristic for the examined group of patients and differed significantly (P < 0.0001) (Figure 1A–F).
In general, in the results obtained using the culture method, the largest differences in the total proportion of bacteria were visible in the Bifidobacterium genus, whose number declines with the consecutive fecal fractions and tissue samples (in the CD group 79%-27%-44%-5%; in the UC group 46%-52%-22%-5%), whereas in the control group, it remained at a high invariable level of >90%. The above-mentioned differences were also confirmed with the use of the FISH technique; however, in the hybridization, the percentages of Bifidobacterium in the examined groups were not as high: in the CD group 38%-16%-47%; in the UC group 74%-59%-27%; and in the control group 45%-57%-80%. In the patients experiencing CD, the percentage of bacteria from Streptococcus genus in the subsequent fecal fractions and in the tissue samples increased in the culture method (4%-18%-2%-79%) in comparison with the control group (1%-2%-2%-3%); this tendency was confirmed using the FISH method (6%-2%-15% and 9%-1%-0%) (Fig. 1). In patients with UC, a growing percentage of lactobacilli was observed using culture methods (29%-27%-49%-90%) in the consecutive fecal fractions and the tissue, whereas a minimal increase (2%-3%-8%-6%) was noted in the control group. The same analysis performed with the application of the FISH technique provided similar results (0%-2%-13% and 3%-5%-1%). The application of the culture method enabled us to confirm the increase in the percentage of Enterobacteriaceae in the consecutive fecal fractions and the tissue in the group of patients experiencing CD (1%-1%-31%-13%) in comparison with the control group (0%-1%-0%-11%). In the group of patients with UC, the percentage of Enterobacteriaceae was 25%-19%-18%-1%. The results for E coli (representative species of Enterobacteriaceae) using the FISH technique were less diversified among the specific fractions and studied groups: in the CD group (7%-3%-11%), in the UC group (7%-2%-2%), and in the control group (6%-1%-3%) (Fig. 1). Bacteroides spp were detected in significant amounts using the FISH technique. In the patients with CD, their percentage dropped in the consecutive fecal fractions (36%-15%-17%), whereas in the patients experiencing UC, it increased (5%-34%-44%). In the control group, the number of Bacteroides spp in the consecutive fecal fractions fell (22%-15%-9%). There were trace elements of Bacteroides spp detected with the use of the culture technique in tissue samples: 2% in the UC and 1% in the control group. Enterococci were identified by means of the culture method and it was shown that in the groups of the patients with CD and UC, the percentage of these bacteria was variable in the consecutive fecal fractions and in the colon tissue (11%-2%-21%-1% and 0%-2%-3%-2%, respectively), in comparison with the control group, in which the percentages did not exceed 1% (Fig. 1). For species of Clostridium coccoides (Blautia coccoides), the identification was performed only with the use of the FISH technique. Their quantities were diversified depending on the fecal fraction and the examined group of patients and they did not display significant regularities as the remaining examined bacterial taxons: CD (13%-58%-8%), UC (5%-1%-14%), and control (15%-21%-7%) (Fig. 1).
The collective presentation of the quantities of bacteria in the consecutive fecal fractions and in the tissue samples is shown in Table 3, where the absolute values of the bacterial numbers and their percentages in relation to the total number of bacteria in the examined fecal fraction and in the colon tissue samples are displayed. The Steel test was applied to analyze the differences between the groups of the patients experiencing CD, UC, and the control group for absolute values of the bacterial numbers. The significant differences (P < 0.05) were detected only between fractions II and III. Using the culture method, the sole significant difference was found in fecal fraction II for Enterobacteriaceae in comparison with the control group in the group of patients experiencing UC. Owing to the application of the FISH technique in the group of patients experiencing CD, the significant differences were merely confirmed in relation to the group of patients experiencing UC for the following bacteria: Bifidobacterium (fraction II), Streptococcus (fraction II), and Lactobacillus (fraction III) (Table 3). In the group of patients experiencing UC, the considerable differences were presented in relation to the control group for the following bacteria: Bacteroides (fraction II), Bifidobacterium (fraction II), E coli (fraction II), Streptococcus (fraction II), and Lactobacillus (fraction III) (Table 3).
When studying the ability of bacterial flora attached to the mucous layer to degrade mucin, it was found that the measurements between the UC group (19.8 ± 0.94 mm diameter of mucin lysis zone) and the control group (15.3 ± 1.78 mm) were significantly different (P = 0.045; Mann-Whitney U test). There was no difference between the CD group (18.3 ± 1.25 mm) and the control group and between the CD and UC groups. Also, the Mann-Whitney U test analysis showed no significance between the CD group (14.3 ± 0.94 mm) and the UC group (17.3 ± 3.30 mm) in comparison with the control group (16.7 ± 2.87) and between CD and UC when fraction III samples were tested.
The objective of the present study was to examine whether and how the composition of the bacterial microbiota in the intestinal cross-section of the patients experiencing IBD changed in comparison with the healthy individuals. Consequently, 3 consecutive fecal fractions obtained during the preparation of the patient for colonoscopy procedure and the colon tissue samples were analyzed. Fraction I probably contained the planktonic flora. Fraction II consisted of the intestinal content in our opinion situated closer to the intestinal wall. Fraction III was the intestinal content, which was in direct contact with the intestinal mucosa layer. Subsequently, examination of the tissue samples enabled us to evaluate the composition of bacterial flora attached to the colon mucus layer.
The examination of the bacterial microbiota obtained from the above-mentioned 3 fractions and the tissue samples enabled us to demonstrate the diversified quantitative composition between the main bacterial taxons colonizing the colon in the groups of patients experiencing CD and UC. In the control group, the similarity between specific fractions and the colon tissue was confirmed. The statistical analysis demonstrated that in fecal fraction I, there was no significance in the bacterial groups examined using the culture method and the FISH technique. The same results were obtained for the colon tissue samples, but the examination was carried out only by culture technique. The significant results were obtained in fractions II and III in the specific bacterial groups.
The thesis concerning the layered structure of the fecal bacterial flora in the colon is not reflected in the literature; nevertheless, our results may prove its validity. What is more, Swidsinski et al (10) describe the phenomenon of the spatial organization of the intestinal bacterial flora, applying the term “the intestinal microbiota biostructure” (10,11). In our research, the gradual assimilation of the composition of the successive fecal fractions to the bacterial flora present on the surface of the mucus membrane of the colonial epithelial cells has been presented. This can be observed in the UC group and in the control group. In the CD group, this tendency is less noticeable. Interestingly enough, in the control group examined with the application of both microbiological culture and the FISH method, a significant microbiological similarity of the subsequent fecal fractions and the colonic mucsa is observed. This can be related to the fact that the thickness of the mucus layer separating the colonic epithelial cells from the bacterial flora located in the colon light is considerably larger in comparison with the patients experiencing CD and UC, which has been proved in the previous thesis (8). In addition, the analysis of the capacity of the bacterial flora present on the surface of the intestinal mucosa to degrade the mucin has confirmed that only in the UC group as compared with the control group can the bacteria significantly degrade the mucin. Therefore, it cannot be excluded that damage to the mucus and simultaneous easier access to the epithelial cells affect the composition of the bacterial flora as a result of the accessibility of new receptors to the bacterial cells (21). This can constitute the cause of variability in the quantitative composition of the examined bacterial taxons in the subsequent fecal fractions—the closer to the intestinal wall, the higher the probability of interaction with the epithelial cells and with the GALT.
In the CD group, in the intestinal tissue, the quantitative composition of specific bacterial taxons rapidly changes in relation to its successive fecal fractions differently from the UC group. The reason can be that a decrease in the thickness of the intestinal mucosa in this disease entity is mainly connected with the mutations of the mucin-encoding genes (22,23), not with the influence of the bacterial flora itself on the stability of this protein. The results obtained by us, where the considerable effect of the bacterial flora on the process of degrading the mucin has not been presented, confirm this. Furthermore, Ardesjö et al (24) proved that the autoimmune reaction against the goblet cells located in the intestinal epithelium and responsible for production and secretion of the intestinal mucosa, which may result in the atrophy of the mucus layer, was present in 84% of patients with CD. Obviously, in the case of UC, we cannot exclude the influence of other factors, except for the bacterial flora, on the atrophy of the mucus barrier, such as the Muc gene mutations, which is pointed out by Myerscough et al (25). Consequently, it is possible that in the case of CD, the composition of the colonic bacterial flora is less related to the condition of the intestinal mucous membrane.
When analyzing the data obtained in this work, it should be taken into consideration that inducing acute diarrhea while preparing patients for colonoscopy may disturb the spatial organization of the colonic bacterial flora and simultaneously hinder the proper interpretation of the results.
The largest diversification in the quantitative composition was detected with Bifidobacterium. The application of both the culture method and the FISH technique allowed us to demonstrate that the presence of these bacteria in patients with IBD in all of the fecal fractions and the tissue samples was lower as compared with the control group of patients. The decreased colonization of the intestinal mucosa by Bifidobacterium was frequently described in the scientific literature (9,26,27). This fact emphasizes the important role of Bifidobacterium species in the process of preserving the ecological homeostasis in child colons. These microorganisms, apart from the lactic acid bacteria, are listed as a reservoir of the strains with probiotic properties. As a result, the possibility of checking and using these properties in the treatment of IBD appears (28,29).
Also surprising was the fact of a large increase in lactobacilli populations in adolescents with UC. Some bacteria belonging to this genus are considered probiotic, meaning beneficial to the host. It is vital to remember that the genus Lactobacillus is a broad group of bacteria, encompassing many species with different characteristics and requirements. It is probable that the chronic inflammatory state in the course of UC somehow promotes the propagation of some selected species of lactobacilli. In the present study, only the quantitative aspects of the presence of these bacteria in bowel flora of adolescents with IBD were analyzed, omitting their speciation, which was beyond the scope and objectives of this work. It has been demonstrated by us, however, that populations of bacteria producing hydrogen peroxide including peroxide-positive Lactobacillus spp are elevated in bowel biopsies from adolescents with IBD, and they may contribute to perpetuation of the inflammation (9).
Using the culture method, it was demonstrated that in the group of patients with CD larger quantities of Enterobacteriaceae (31%) appeared in the III fraction and 13% in the tissue samples, despite the fact that in the previous fecal fractions, the percentage fluctuated at approximately with 1%. Also, in the group of children with UC, the presence of Enterobacteriaceae was detected in all of the fecal fractions. In the control group in all 3 fractions, this percentage did not exceed 1% with the exception of tissue samples, where the percentage was 8%. The increasing percentage of Enterobacteriaceae in the stools of the patients with IBD was confirmed by other authors (26,30). Mostly, it is the proinflammatory influence of the bacterial LPS liberated by Gram-negative bacteria and the increase in the number of the adherent-invasive Escherichia coli strains adhering to the intestinal epithelium and having the ability to produce the hemolysins destroying this tissue that is postulated (31). Sasaki et al (32) and Darfeuille-Michaud et al (33) proved that the adherent-invasive Escherichia coli strains displayed a higher incidence in patients with CD than in the patients with UC and healthy individuals (32,33). This may be reflected by the increasing percentage of Enterobacteriaceae in the III fecal fraction and the colon tissue in patients with CD, which was shown in the examinations performed. The application of the FISH technique enabled us to observe an insignificant increase in the number of the species of E coli in all of the examined fecal fractions in children experiencing CD in relation to the control group, yet the most significant difference was identified in the III fecal fraction. The percentage of Enterobacteriaceae in this fraction as compared with the control group, detected by the use of the culture method, was considerably higher and equaled approximately 31%. The ECOLI probe specific solely to E coli was applied in the FISH technique, which did not allow us to detect the remaining species of Enterobacteriaceae spp, which presumably influenced the detection of the lower percentage of this bacterial group.
Some authors depict the increasing populations of Bacteroides spp in the fecal microbiota as well as adhering to the intestinal mucosa of the patients with IBD (34,35). In the following work, this regularity for the fecal samples examined with the use of the FISH technique was confirmed. In patients with UC, the percentage of Bacteroides spp increased with the consecutive fecal fractions as opposed to the control group and patients with CD, in which the described relation was reversed. The higher colonization of the colon by Bacteroides spp in the group of patients with UC may have been the result of occupying the ecological niche abandoned by other anaerobic bacteria (eg, Bifidobacterium) living in the GI tract in children who do not have IBD. Consequently, the higher concentration of Bacteroides endotoxins may lead to the increased activity of GALT and exacerbation of the inflammatory condition as it is observed for Enterobacteriaceae spp. The application of the culture method to detect the Bacteroides genus allowed us to identify the trace elements of these bacteria.
The use of both the culture method and the FISH technique was aimed at obtaining more complete results and comparing both research methods. The culture method possesses specific limitations concerning, among other things, the difficulties in detecting anaerobic bacteria, such as Bifidobacterium, Bacteroides, or Clostridium, because of the toxic effect of oxygen present in the atmosphere. This was also proved in this work. Owing to the application of the FISH technique, the presence of the above-mentioned bacteria was demonstrated in the quantity of 1010 cfu/mL, whereas for Bacteroides, the highest number was 103 cfu/mL using the culture method (Table 3).
We have carried out our investigation using fecal samples, which were stored at −70°C during the transport from the department to the laboratory. According to Achá (36) and Dan et al (37), freezing does not influence the viability of the fecal samples.
The significance was demonstrated solely in 1 case with the use of the culture method; the FISH technique allowed us to demonstrate it in 8 cases (Table 3). The method based on the detection of the nucleic acids, as the FISH is, enables us to identify the dead bacteria as well as the noncultivable (for other reason) ones. As a result, we can conclude that the FISH molecular technique applied in microbiological fecal examinations allowed us to obtain more reliable results than the culture method. Unfortunately, the results of the examination of tissue sections could not be obtained using the FISH method. This is because it is a low-sensitivity technique that does not allow the detection of bacteria <3 × 103 cfu/mL(g) (8,38). Tissue weights were low (0.02–0.05 g), and the sample homogenates were diluted by paraformaldehyde solution. In this case, the method that can be recommended is quantitative polymerase chain reaction.
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