Skip Navigation LinksHome > August 2014 - Volume 20 - Issue 8 > Role of Intestinal Bacteria in the Pathogenesis of Pouchitis
Inflammatory Bowel Diseases:
doi: 10.1097/MIB.0000000000000055
Basic Review Articles

Role of Intestinal Bacteria in the Pathogenesis of Pouchitis

Batista, Daisy MD; Raffals, Laura MD, MS

Free Access
Article Outline
Collapse Box

Author Information

Division of Gastroenterology and Hepatology, Mayo Clinic, Rochester, Minnesota.

Reprints: Laura Raffals, MD, MS, Division of Gastroenterology and Hepatology, Mayo Clinic, 200 First Street SW, Mayo East 9, Rochester, MN 55905 (e-mail: raffals.laura@mayo.edu).

The authors have no conflicts of interest to disclose.

Received February 28, 2014

Accepted March 24, 2014

Collapse Box

Abstract

Pouchitis is a common complication seen in patients with ulcerative colitis who undergo total proctocolectomy with ileal pouch anal anastomosis. Bacteria seem to play an important role in the development of pouchitis, although this role is not well defined. Because technology has advanced, we are able to apply molecular techniques to describe the structure and function of the pouch microbial community. In recent years, several studies have been performed comparing the pouch microbiota in patients with ulcerative colitis with healthy pouches and pouchitis. Many of these studies have suggested that pouchitis is characterized by dysbiosis and/or decreased microbial diversity. There has not been a clear pattern identifying a pathogenic organism or a group of organisms responsible for pouchitis. This review summarizes recent studies exploring the pouch microbiota in health and disease, the relationship of bacterial metabolites and pouchitis, and the role of antibiotics and probiotics for the treatment and prevention of pouchitis.

The prevalence of ulcerative colitis (UC) has been reported to be as high as 249 per 100,000 persons in North America.1 Approximately, 30% of patients with UC go on to require total proctocolectomy after 15 years of disease.2 The indication for surgery is often medically refractory disease or the development of dysplasia or colorectal cancer. The surgical treatment of choice for many patients is total proctocolectomy with creation of an ileal pouch anal anastomosis (IPAA) (Fig. 1). This procedure has greatly enhanced the quality of life for many patients and allows patients to avoid the need for a permanent ileostomy. This same procedure is also performed for patients with familial adenomatous polyposis (FAP).

FIGURE 1
FIGURE 1
Image Tools

Pouchitis, a term referring to inflammation of the pouch, is the most common complication occurring in patients with IPAA with rates as high as 46%.4–7 Approximately, 50% of patients with pouchitis may experience recurrent inflammation, and an unfortunate 5% to 10% suffer from chronic pouchitis requiring immunomodulator or biologic therapy or even the removal of the pouch.8–10 Interestingly, patients with FAP who undergo IPAA rarely develop pouchitis, which suggests there may be an important genetic contribution to the development of pouchitis.

The symptoms of pouchitis can include increased bowel frequency, bleeding, urgency, abdominal cramping, and tenesmus. The diagnosis of pouchitis is based on the presence of symptoms as well as endoscopic and histological evidence of inflammation of the pouch. Pouchoscopy can help identify other conditions affecting the pouch, such as pouch ischemia, Crohn's disease of the pouch, cuffitis, infections such as cytomegalovirus, or less commonly autoimmune enteropathy. In cases where there is a suspicion for mechanical complications such as a fistula or stricture, abdominal imaging with contrast pouchogram, computed tomography, or magnetic resonance enterography can be helpful to delineate the anatomy of the pouch, identify complications involving the pouch, and determine the extent and nature of inflammation.

Although we do not know the cause of pouchitis, we do believe that the intestinal microbial community plays an important role in maintaining pouch health or driving pouch inflammation. In support of this assumption, it is observed that pouchitis only occurs after restoration of the fecal stream through the pouch.11,12 Second, antibiotics are our most effective treatment of pouchitis.13,14 Probiotics, such as VSL#3 have also been shown to prevent pouchitis.15–17 In addition, several genes associated with the innate immune response and microbial sensing and recognition have been associated with an increased risk for pouchitis including the NOD2/CARD15 gene,18,19 Il-1 receptor antagonist gene,20 and Toll-like receptor genes.21

The intestinal microbiota may influence the health of the gastrointestinal tract through its interactions with the host immune system. Intestinal epithelial cells provide a mechanical barrier preventing the penetration of luminal bacteria and toxins. There is a mucus layer immediately adjacent to the epithelium. Bacteria reside in this mucus and are believed to instruct and regulate the host immune system at the level of the gut epithelium through numerous complex mechanisms.22,23

Although all of these observations implicate the enteric microbiota for the maintenance of pouch health and development of disease, we have much to learn about the mechanisms involved. In this article, we will discuss our understanding of the relationship between the pouch microbiome and pouchitis, and the therapeutic roles of antibiotics and probiotics.

Back to Top | Article Outline

RELATIONSHIP OF THE POUCH MICROBIOTA IN HEALTH AND DISEASE

Research exploring the role of the microbial community in inflammatory bowel disease (IBD) has exploded in recent years. The great advances in technology and bioinformatics allowing investigators to perform high throughput genetic sequencing of the bacterial communities and synthesize the data in meaningful ways have fostered this explosion. Despite this recent boon in the field of microbial ecology, investigators have been interested in the role of the pouch microbiota for decades. Initial studies were performed using culture methods, biasing results to include only those organisms cultivatable (estimated to be <30%–40% of gut bacterial species).24 The first reports examining the pouch microbiota date back to the 1960s, when Gorbach et al25 published their comparisons of the intestinal microflora in ileostomy effluent compared with microflora in the ileal pouch. Before this study, it was recognized that anaerobic bacteria dominate the bacterial composition of stool in healthy individuals. Gorbach's study found that ileal effluent was mostly composed of aerobic bacteria, and chronically inflamed pouches had a relatively equal proportion of anaerobic and aerobic organisms. Subsequent studies noted an increased ratio of anaerobes/aerobes in stool collected from ileal pouches compared with ileostomy effluent.26 This compositional difference may be explained by the function of the pouch, which is a reservoir for stool, promoting fecal stasis and subsequent shifts in the structure of the pouch microbiome.

Most current studies of the pouch microbiota use culture-independent molecular techniques to characterize the microbial community. One recent study explored the evolution of the mucosal microbiota of the pouch after takedown of the protective ileostomy.3 In this study, Young et al described the development of the pouch microbiota in 4 patients. Pouchoscopy and collection of mucosal and luminal samples were performed before ileostomy takedown and at several time points after restoration of the fecal stream through the pouch. There was a significant shift in the microbial community in all 4 individuals after ileostomy closure. Overall diversity increased in 3 of the 4 subjects, and the overall density of microorganisms in the pouch increased in all individuals. There was a clear shift from facultative to more obligate anaerobes in the pouch after restoration of the fecal stream, consistent with previous studies of the pouch microbial community. The microbial community was distinct in each individual. One individual's microbial community was more similar to that seen in the healthy colon. This individual continued to remain healthy for 2 years of follow-up, whereas the other 3 patients whose communities were not as similar to the healthy colon went on to develop pouchitis. Falk et al27 also performed a longitudinal study of pouch microbial communities in 2 patients with IPAA. These patients were followed for 1 year after ileostomy closure. The composition of the microbial community shifted during the 12 months of follow-up. Similar to Young's study, each subject had a unique mucosal-associated microbial community; however, both of these subjects' microbial communities became “colon-like” and neither subject developed pouchitis. Although the number of patients studied in these 2 studies is small, their findings suggest that the development of the pouch microbial community into a diverse community similar to that found in the healthy colon promotes pouch health.

Studies have explored the pouch microbiota in patients with and without pouchitis. Komanduri et al28 performed the first study of this kind using modern molecular techniques to describe the pouch microbiota. They pooled ileal/pouch mucosal samples from patients with no history of pouchitis, a history of pouchitis, and non-IBD controls. They found an increase in Fusobacter (Proteobacteria) and a decrease in Streptococci (phylum Firmicutes) in the group with pouchitis. Members of the Clostridium group (Clostridium paraputrificum), Enterics (Escherichia coli/Shigella sp), and the Streptococcus group accounted for a greater proportion of the microbial community in the control pouch group compared with the pouchitis group.

McLaughlin et al29 examined the mucosal-adherent microbiota in ileal pouch biopsy samples of 16 patients with UC (8 of whom had pouchitis) and 8 with FAP. Through cloning and sequencing 3184 full-length bacterial 16S rRNA genes, they found a significant increase in Proteobacteria (P = 0.019) and a significant decrease in Bacteroidetes (P = 0.001) and Faecalibacterium prausnitzii (P = 0.029) in the patients with UC compared with FAP. In contrast to Komanduri's study, they did not observe a significant dysbiosis in the patients with pouchitis compared with those without pouchitis. Bacterial diversity was significantly greater in patients with FAP compared with patients with UC without pouchitis (P = 0.019), whereas diversity was greater in patients with UC without pouchitis compared with patients with UC with pouchitis (P = 0.009).

A similar study using terminal restriction fragment length polymorphism and DNA sequencing compared the pouch fecal and mucosal-associated microbiota in patients with UC with and without pouchitis and patients with FAP with pouchitis.30 Similar to the study by McLaughlin et al, they noted that the microbial community in healthy UC pouches was unique to the microbial community in FAP pouches. There were also notable differences in the inflamed pouch compared with both the healthy UC pouch and the FAP pouch. Patients with UC with pouchitis had more terminal restriction fragment length polymorphism peaks consistent with Clostridium and Eubacterium genera compared with healthy UC and FAP pouches. UC pouches also had fewer peaks of Lactobacillus and Streptococcus genera compared with FAP pouches. Sequencing of pooled fecal samples revealed a greater abundance of Firmicutes and Verrucomicrobia and fewer Bacteroidetes and Proteobacteria in patients with pouchitis compared with FAP.

One of the most recently published studies characterized mucosal-associated microbial communities present in the pouch and afferent limb of patients with IPAA through pyrosequencing the 16S rRNA V1-V3 hypervariable region followed by quantitative PCR for targeted bacteria.31 This study included patients with UC with and without pouchitis, FAP pouches, and pouches with Crohn's disease–like features. They found decreased bacterial diversity in patients with pouchitis compared with those with noninflamed pouches, including FAP individuals. In addition, there were statistically significant differences in patients with pouchitis with Bacteroidetes detected less frequently and Proteobacteria detected more frequently.

Antibiotic treatment seems to influence the composition of the pouch microbial community. Tannock et al32 examined stool from patients with chronic pouchitis, healthy pouches, and FAP pouches. Patients on antibiotic therapy for chronic pouchitis had a less diverse microbial community compared with those with chronic pouchitis off antibiotic treatment (P < 0.0001). FISH analysis showed that patients with chronic pouchitis on antibiotics had a reduced phylogenic gap compared with patients with chronic pouchitis off antibiotics and patients with healthy pouches. Patients with pouchitis off antibiotic treatment had fewer Firmicutes and more Proteobacteria.

It is challenging to identify a consistent theme among the many cross-sectional studies examining the structure of the pouch microbiota. These challenges stem from differences in methodology (molecular techniques used to analyze the microbial communities) and differences in the type of samples processed. Microbial communities present in luminal (or fecal) samples are likely to differ from those in mucosal biopsies. Some of the studies pooled samples for patient cohorts, which may also affect results because pooling of samples may not detect important differences in individual patients. Nonetheless, studies clearly demonstrate a difference between pouch microbial communities in patients with UC compared with patients with FAP. There also seems to be a change in the microbial community during pouchitis, although these changes are not consistent across studies.

Back to Top | Article Outline

BACTERIAL METABOLITES—SHORT CHAIN FATTY ACIDS

To understand the role of the microbiota in pouchitis, we must expand our understanding of the microbiota beyond the phylotypic description of microbial communities. The inconsistencies among studies characterizing the structure of the pouch microbial communities in health and disease may reflect the importance of the function of the microbial community rather than which microbial species are present or absent.

Microbially-derived short chain fatty acids (SCFA) are the most extensively studied bacterial metabolite in the context of IBD. In the colon, anaerobic bacteria ferment undigested polysaccharides producing SCFA, such as acetic, propionic, and butyric acid. SCFA provide energy to colonocytes, possess anti-inflammatory properties, and may regulate host gene expression.33 Butyrate is the preferred energy source for the intestinal epithelium and has generated a lot of interest in the research community. Bacteria that produce butyrate are phylogenetically diverse, including Eubacterium spp. and Roseburia spp. of the Clostridium cluster XIVa and F. prausnitzii of the Clostridium cluster IV.34

Vital et al35 described the establishment of butyrate-producing communities in the ileal pouch in patients with an IPAA. This study was performed from luminal samples collected from the same patients described in the previously discussed study by Young et al. Using 454 pyrotag sequencing and quantitative polymerase chain reaction, they targeted genes coding for the butyrate-synthesizing enzymes. All 4 patients developed abundant butyrate-producing communities by 2 months after restoration of the fecal stream through the pouch, although the structure of these communities varied among the individuals. Only 1 individual developed a community comparable with communities seen in the healthy controls, and this individual was the only 1 of the 4 studied who did not go on to develop pouchitis in the 2-year follow-up.

SCFA seem to be reduced in the fecal contents of patients with pouchitis.36 This reduction in butyrate is likely a result of a shift in the microbial community composition. Investigators have described an overabundance of sulfate-reducing bacteria in patients with pouches for the treatment of UC but not in patients with pouches for FAP.37 Other studies have described increased sulfate-reducing bacteria in the stool of patients with pouchitis compared with the stool of patients with healthy pouches.38 The overgrowth of sulfate-reducing bacteria can lead to increased hydrogen sulfide production that blocks the utilization of butyrate by colonocytes.39,40 Although we cannot definitively explain the significance of decreased butyrate in pouchitis, simply replacing butyrate in this patient population does not seem to reverse inflammation. Several small studies evaluating the use of exogenous SCFA in the form of suppositories or enemas have failed to show significant improvement.41 Nonetheless, it is not known if a reduction of butyrate is responsible for the development of pouchitis or an epiphenomenon related to changes in the microbial community during inflammation. Butyrate's role in pouchitis still needs to be elucidated.

Back to Top | Article Outline

PROBIOTICS

Probiotics are nonpathogenic living organisms that belong to the natural host flora and provide important health benefits to the host.6,42 Experimental studies in animal models of colitis, such as the IL-10 knockout mice, have suggested that probiotics might attenuate the development and the severity of colitis.43,44 These findings have not played out in most IBD clinical trials. However, probiotics do seem to have some benefits in patients with IPAA.

VSL#3 is a probiotic compound containing 4 strains of Lactobacillus, 3 strains of Bifidobacterium, and 1 strain of Streptococcus. In a randomized, double-blinded placebo-controlled trial, Gionchetti et al16 evaluated the effectiveness of VSL#3 in the maintenance of remission of chronic pouchitis after treatment with antibiotics. Forty patients with clinical and endoscopic remission at the time of study entry were randomized to placebo or 6 g/d of VSL#3. Three patients (15%) in the VSL#3 group had pouchitis relapse within the 9-month follow-up period, compared with 20 (100%) of patients in the placebo group. Interestingly, 100% of the patients who maintained remission with VSL#3 therapy experienced a relapse of pouchitis within 4 months of stopping probiotic therapy. VSL#3 is also effective at preventing pouchitis in patients with a newly created IPAA. Another study by Gionchetti et al15 randomized 40 patients to receive VSL#3 or placebo after fecal continuity was restored through the IPAA. Patients receiving VSL#3 were less likely to develop pouchitis (2/20 versus 8/20, P < 0.05) within the first year after takedown of their protective ileostomy.

Other probiotic formulations such as Lactobacillus rhamnosus GG have been studied in pouchitis with less consistent results.45,46 A small randomized, double-blinded placebo-controlled study by Kuisma et al46 evaluated the effectiveness of L. rhamnosus GG as a primary therapy for pouchitis in 20 patients with endoscopically confirmed active pouchitis. There were no differences observed between treatment groups measured by clinical or endoscopic response, deeming this probiotic ineffective as a primary therapy for pouchitis. However, the same probiotic formulation seemed to be beneficial as a prophylactic agent for pouchitis.45 Thirty-nine patients underwent IPAA from 1996 to 2001 received L. rhamnosus immediately after IPAA, and 78 patients who underwent IPAA between 1989 and 1996 did not receive any prophylactic treatment. Subjects who received treatment with L. rhamnosus had a lower cumulative risk of pouchitis at 3 years (7%) compared with 29% in the group who did not receive L. rhamnosus (P = 0.011).

A recent study by Persborn et al47 evaluated the effects of probiotics on the mucosal barrier function and permeability in patients with pouchitis. After completing a 4-week course of antibiotics for active pouchitis, 16 patients with UC were enrolled to receive Ecologic 825 for 8 weeks. Ecologic 825 contains 9 viable probiotic strains of Bifidobacterium, Lactobacillus, and Lactococcus. Pouch biopsies were obtained during active pouchitis, after 4 weeks of antibiotic treatment, and after 8 weeks of probiotic treatment. Transmucosal permeability of mucosal biopsies was assessed using Ussing chambers. Active pouchitis was associated with an increase passage of E. coli K12, which did not correct after treatment with antibiotics, despite significant decreases in pouch disease activity index. However, after probiotic therapy bacterial passage significantly decreased suggesting that a restoration of mucosal permeability comparable with that of healthy pouches. Horseradish peroxidase was used to measure transcellular permeability. After probiotic therapy, horseradish peroxidase flux returned to the same levels seen in patients with healthy pouches. Paracellular permeability assessed by 51Creatinine-labelled ethylenediaminetetraacetic acid (51CrEDTA) probe did not seem affected in pouchitis. In contrast to previous studies where the pouch microbiota was altered by probiotic treatment,16 this study did not show a difference in the composition of the pouch microbiota after 8 weeks of probiotic therapy.

A meta-analysis by Shen et al48 included randomized controlled trials evaluating the therapeutic effect of probiotics in IBD and pouchitis. Subgroup analysis did not find a significant benefit with probiotic treatment for maintenance therapy of pouchitis. When trials using VSL#3 were looked at separately, there did seem to be a benefit in preventing relapse in patients with pouchitis, (P < 0.00001, risk ratio = 0.20) with little heterogeneity (P = 0.88, I2 = 0%).

Although several studies have shown the benefits of probiotics in the prevention and maintenance of remission in pouchitis, the exact mechanisms by which they exert their benefits are complex and incompletely understood. Furthermore, it is not yet clear which bacterial species or combinations are optimal to modulate pouch inflammation. Further studies in this area are needed to better understand and optimize the role of probiotics in controlling pouch inflammation.

Back to Top | Article Outline

ANTIBIOTICS FOR TREATMENT OF POUCHITIS

Antibiotics are the first-line therapy for patients with pouchitis, although there are little data to support their efficacy. In a small randomized clinical trial of 16 patients, ciprofloxacin (1000 mg/d) and metronidazole (20 mg·kg−1·d−1) for 14 days resulted in a significant reduction in the pouch disease activity index.49 Treatment with ciprofloxacin resulted in a greater reduction in pouch disease activity index and symptoms compared with metronidazole. Ciprofloxacin was also better tolerated than metronidazole.

A small randomized, placebo-controlled pilot trial evaluated the efficacy of rifaximin for the treatment of pouchitis.50 Eight patients were randomized to rifaximin, and 10 patients received placebo. At week 4, 25% of patients treated with rifaximin were in clinical remission compared with 0% in the placebo group, although this difference was not statistically significant (P = 0.2059). An open-label study of rifaximin suggested a benefit for maintenance therapy with variable doses of rifaximin.51 In this study, 51 patients with chronic pouchitis were treated with a 2-week course of antibiotics and then transitioned to rifaximin for maintenance therapy. At 3 months, 33 (65%) patients maintained remission. At 1 year, 19 (58%) patients continued to maintain their remission.

Despite the lack of large studies examining the role of antibiotics, ciprofloxacin or metronidazole are the first-line treatments for acute pouchitis. Because ciprofloxacin is well tolerated, it is often the first choice for treatment. Rifaximin, amoxicillin–clavulanic acid, tetracycline, and erythromycin are other antibiotics commonly used in practice.

Patients who do not respond to a course of a single antibiotic may benefit from a combination of antibiotics. Several studies have examined the efficacy of antibiotic combinations. In the first study, patients who did not achieve a remission with a course of a single antibiotic were randomized to ciprofloxacin (1 g/d) along with rifaximin (2 g/d) for 15 days.14 Sixteen of the 18 patients in this study improved or achieved a remission. Combinations of ciprofloxacin (1 g/d) along with metronidazole (1 g/d) for 4 weeks,52 and ciprofloxacin (1 g/d) along with tinidazole (15 mg·kg−1·d−1) for 4 weeks53 have also shown efficacy in treating refractory pouchitis.

Back to Top | Article Outline

CONCLUSIONS

Pouchitis is a common complication experienced by patients with a history of UC who undergo colectomy with IPAA. Pouchitis is not commonly seen in patients with FAP who undergo the same surgery. There is strong evidence to suggest that bacteria are important to the development of pouchitis, although this relationship is not well understood. The pouch microbial community evolves over the first months to year after fecal continuity through the pouch is restored. Some individuals develop a pouch microbiota similar to that of the healthy colon. The development of a “colon-like” microbiota may promote a healthy pouch. Cross-sectional studies describing the structure of the pouch microbial community in healthy and inflamed pouches have failed to identify a pathogenic organism or a group of organisms responsible for pouchitis. It is possible that the function of the pouch microbial community is more important than the phylogenic structure of the microbial community. Longitudinal studies examining the pouch microbiota and the functional characteristics of the pouch are needed to better understand the role the pouch microbiota that plays in the development of pouchitis.

Back to Top | Article Outline

REFERENCES

1. Molodecky NA, Soon IS, Rabi DM, et al. Increasing incidence and prevalence of the inflammatory bowel diseases with time, based on systematic review. Gastroenterology. 2012; 142:46–54.

e42; quiz e30


2. Gionchetti P, Amadini C, Rizzello F, et al. Diagnosis and treatment of pouchitis. Best practice and research. Best Pract Res Clin Gastroenterol. 2003; 17:75–87.

3. Young VB, Raffals LH, Huse SM, et al. Multiphasic analysis of the temporal development of the distal gut microbiota in patients following ileal pouch anal anastomosis. Microbiome. 2013; 1:9

4. Penna C, Dozois R, Tremaine W, et al. Pouchitis after ileal pouch-anal anastomosis for ulcerative colitis occurs with increased frequency in patients with associated primary sclerosing cholangitis. Gut. 1996; 38:234–239.

5. Fazio VW, Ziv Y, Church JM, et al. Ileal pouch-anal anastomoses complications and function in 1005 patients. Ann Surg. 1995; 222:120–127.

6. Ferrante M, Declerck S, De Hertogh G, et al. Outcome after proctocolectomy with ileal pouch-anal anastomosis for ulcerative colitis. Inflamm Bowel Dis. 2008; 14:20–28.

7. Fleshner P, Ippoliti A, Dubinsky M, et al. Both preoperative perinuclear antineutrophil cytoplasmic antibody and anti-CBir1 expression in ulcerative colitis patients influence pouchitis development after ileal pouch-anal anastomosis. Clin Gastroenterol Hepatol. 2008; 6:561–568.

8. Simchuk EJ, Thirlby RC. Risk factors and true incidence of pouchitis in patients after ileal pouch-anal anastomoses. World J Surg. 2000; 24:851–856.

9. Stahlberg D, Gullberg K, Liljeqvist L, et al. Pouchitis following pelvic pouch operation for ulcerative colitis: incidence, cumulative risk, and risk factors. Dis Colon Rectum. 1996; 39:1012–1018.

10. Salemans JM, Nagengast FM, Lubbers EJ, et al. Postoperative and long-term results of ileal pouch-anal anastomosis for ulcerative colitis and familial polyposis coli. Dig Dis Sci. 1992; 37:1882–1889.

11. Nicholls RJ, Belliveau P, Neill M, et al. Restorative proctocolectomy with ileal reservoir: a pathophysiological assessment. Gut. 1981; 22:462–468.

12. Santavirta J, Mattila J, Kokki M, et al. Mucosal morphology and fecal bacteriology after ileoanal anastomosis. Int J Colorectal Dis. 1991; 6:38–41.

13. Holubar SD, Cima RR, Sandborn WJ, et al. Treatment and prevention of pouchitis after ileal pouch-anal anastomosis for chronic ulcerative colitis. Cochrane Database Syst Rev. 2010;

CD001176


14. Gionchetti P, Rizzello F, Venturi A, et al. Antibiotic combination therapy in patients with chronic, treatment-resistant pouchitis. Aliment Pharmacol Ther. 1999; 13:713–718.

15. Gionchetti P, Rizzello F, Helwig U, et al. Prophylaxis of pouchitis onset with probiotic therapy: a double-blind, placebo-controlled trial. Gastroenterology. 2003; 124:1202–1209.

16. Gionchetti P, Rizzello F, Venturi A, et al. Oral bacteriotherapy as maintenance treatment in patients with chronic pouchitis: a double-blind, placebo-controlled trial. Gastroenterology. 2000; 119:305–309.

17. Gionchetti P, Rizzello F, Morselli C, et al. High-dose probiotics for the treatment of active pouchitis. Dis Colon Rectum. 2007; 50:2075–2082.

18. Meier CB, Hegazi RA, Aisenberg J, et al. Innate immune receptor genetic polymorphisms in pouchitis: is CARD15 a susceptibility factor? Inflamm Bowel Dis. 2005; 11:965–971.

19. Tyler AD, Milgrom R, Stempak JM, et al. The NOD2insC polymorphism is associated with worse outcome following ileal pouch-anal anastomosis for ulcerative colitis. Gut. 2013; 62:1433–1439.

20. Carter MJ, Di Giovine FS, Cox A, et al. The interleukin 1 receptor antagonist gene allele 2 as a predictor of pouchitis following colectomy and IPAA in ulcerative colitis. Gastroenterology. 2001; 121:805–811.

21. Lammers KM, Ouburg S, Morre SA, et al. Combined carriership of TLR9-1237C and CD14-260T alleles enhances the risk of developing chronic relapsing pouchitis. World J Gastroenterol. 2005; 11:7323–7329.

22. Duerkop BA, Vaishnava S, Hooper LV. Immune responses to the microbiota at the intestinal mucosal surface. Immunity. 2009; 31:368–376.

23. Berkes J, Viswanathan VK, Savkovic SD, et al. Intestinal epithelial responses to enteric pathogens: effects on the tight junction barrier, ion transport, and inflammation. Gut. 2003; 52:439–451.

24. Hayashi H, Sakamoto M, Benno Y. Phylogenetic analysis of the human gut microbiota using 16S rDNA clone libraries and strictly anaerobic culture-based methods. Microbiol Immunol. 2002; 46:535–548.

25. Gorbach SL, Plaut AG, Nahas L, et al. Studies of intestinal microflora. II. Microorganisms of the small intestine and their relations to oral and fecal flora. Gastroenterology. 1967; 53:856–867.

26. Nasmyth DG, Godwin PG, Dixon MF, et al. Ileal ecology after pouch-anal anastomosis or ileostomy. A study of mucosal morphology, fecal bacteriology, fecal volatile fatty acids, and their interrelationship. Gastroenterology. 1989; 96:817–824.

27. Falk A, Olsson C, Ahrne S, et al. Ileal pelvic pouch microbiota from two former ulcerative colitis patients, analysed by DNA-based methods, were unstable over time and showed the presence of Clostridium perfringens. Scand J Gastroenterol. 2007; 42:973–985.

28. Komanduri S, Gillevet PM, Sikaroodi M, et al. Dysbiosis in pouchitis: evidence of unique microfloral patterns in pouch inflammation. Clin Gastroenterol Hepatol. 2007; 5:352–360.

29. McLaughlin SD, Walker AW, Churcher C, et al. The bacteriology of pouchitis: a molecular phylogenetic analysis using 16S rRNA gene cloning and sequencing. Ann Surg. 2010; 252:90–98.

30. Zella GC, Hait EJ, Glavan T, et al. Distinct microbiome in pouchitis compared to healthy pouches in ulcerative colitis and familial adenomatous polyposis. Inflamm Bowel Dis. 2011; 17:1092–1100.

31. Tyler AD, Knox N, Kabakchiev B, et al. Characterization of the gut-associated microbiome in inflammatory pouch complications following ileal pouch-anal anastomosis. PLoS One. 2013; 8:e66934

32. Tannock GW, Lawley B, Munro K, et al. Comprehensive analysis of the bacterial content of stool from patients with chronic pouchitis, normal pouches, or familial adenomatous polyposis pouches. Inflamm Bowel Dis. 2012; 18:925–934.

33. Hamer HM, Jonkers D, Venema K, et al. Review article: the role of butyrate on colonic function. Aliment Pharmacol Ther. 2008; 27:104–119.

34. Louis P, Young P, Holtrop G, et al. Diversity of human colonic butyrate-producing bacteria revealed by analysis of the butyryl-CoA:acetate CoA-transferase gene. Environ Microbiol. 2010; 12:304–314.

35. Vital M, Penton CR, Wang Q, et al. A gene-targeted approach to investigate the intestinal butyrate-producing bacterial community. Microbiome. 2013; 1:8

36. Clausen MR, Tvede M, Mortensen PB. Short-chain fatty acids in pouch contents from patients with and without pouchitis after ileal pouch-anal anastomosis. Gastroenterology. 1992; 103:1144–1153.

37. Duffy M, O'Mahony L, Coffey JC, et al. Sulfate-reducing bacteria colonize pouches formed for ulcerative colitis but not for familial adenomatous polyposis. Dis Colon Rectum. 2002; 45:384–388.

38. Coffey JC, Rowan F, Burke J, et al. Pathogenesis of and unifying hypothesis for idiopathic pouchitis. Am J Gastroenterol. 2009; 104:1013–1023.

39. Smith FM, Coffey JC, Kell MR, et al. A characterization of anaerobic colonization and associated mucosal adaptations in the undiseased ileal pouch. Colorectal Dis. 2005; 7:563–570.

40. Roediger WE, Duncan A, Kapaniris O, et al. Reducing sulfur compounds of the colon impair colonocyte nutrition: implications for ulcerative colitis. Gastroenterology. 1993; 104:802–809.

41. Wischmeyer P, Pemberton JH, Phillips SF. Chronic pouchitis after ileal pouch-anal anastomosis: responses to butyrate and glutamine suppositories in a pilot study. Mayo Clin Proc. 1993; 68:978–981.

42. Fuller R. Probiotics in human medicine. Gut. 1991; 32:439–442.

43. Madsen KL, Doyle JS, Jewell LD, et al. Lactobacillus species prevents colitis in interleukin 10 gene-deficient mice. Gastroenterology. 1999; 116:1107–1114.

44. Schultz M, Veltkamp C, Dieleman LA, et al. Continuous feeding of lactobacillus plantarum attenuates established colitis in interleukin-10 deficient mice. Gastroenterology. 1998; 114:A1081

45. Gosselink MP, Schouten WR, van Lieshout LM, et al. Delay of the first onset of pouchitis by oral intake of the probiotic strain lactobacillus rhamnosus GG. Dis Colon Rectum. 2004; 47:876–884.

46. Kuisma J, Mentula S, Jarvinen H, et al. Effect of lactobacillus rhamnosus GG on ileal pouch inflammation and microbial flora. Aliment Pharmacol Ther. 2003; 17:509–515.

47. Persborn M, Gerritsen J, Wallon C, et al. The effects of probiotics on barrier function and mucosal pouch microbiota during maintenance treatment for severe pouchitis in patients with ulcerative colitis. Aliment Pharmacol Ther. 2013; 38:772–783.

48. Shen J, Zuo ZX, Mao AP. Effect of probiotics on inducing remission and maintaining therapy in ulcerative colitis, Crohn's disease, and pouchitis: meta-analysis of randomized controlled trials. Inflamm Bowel Dis. 2014; 20:21–35.

49. Shen B, Achkar JP, Lashner BA, et al. A randomized clinical trial of ciprofloxacin and metronidazole to treat acute pouchitis. Inflamm Bowel Dis. 2001; 7:301–305.

50. Isaacs KL, Sandler RS, Abreu M, et al. Rifaximin for the treatment of active pouchitis: a randomized, double-blind, placebo-controlled pilot study. Inflamm Bowel Dis. 2007; 13:1250–1255.

51. Shen B, Remzi FH, Lopez AR, et al. Rifaximin for maintenance therapy in antibiotic-dependent pouchitis. BMC Gastroenterol. 2008; 8:26

52. Mimura T, Rizzello F, Helwig U, et al. Four-week open-label trial of metronidazole and ciprofloxacin for the treatment of recurrent or refractory pouchitis. Aliment Pharmacol Ther. 2002; 16:909–917.

53. Shen B, Fazio VW, Remzi FH, et al. Combined ciprofloxacin and tinidazole therapy in the treatment of chronic refractory pouchitis. Dis Colon Rectum. 2007; 50:498–508.

Copyright © 2014 Crohn's & Colitis Foundation of America, Inc.

Login