Clostridium difficile infection in diabetes

diabetes research and clinical practice 105 (2014) 285–294

Contents available at ScienceDirect

Diabetes Research and Clinical Practice journ al h ome pa ge : www .elsevier.co m/lo cate/diabres

Review

Clostridium difficile infection in diabetes Hui-Qi Qu a,*, Zhi-Dong Jiang b a

Human Genetics Center, The University of Texas School of Public Health, Houston, TX, USA Center for Infectious Diseases, Division of Epidemiology, Human Genetics and Environmental Sciences, The University of Texas School of Public Health, Houston, TX, USA b

article info

abstract

Article history:

Diabetes-related hospitalization and hospital utilization is a serious challenge to the health

Received 12 August 2013

care system, a situation which may be further aggravated by nosocomial Clostridium difficile

Received in revised form

(C. difficile) infection (CDI). Studies have demonstrated that diabetes increases the risk of

26 January 2014

recurrent CDI with OR (95% CI) 2.99 (1.88, 4.76). C. difficile is a gram-positive, spore-forming

Accepted 13 June 2014

anaerobic bacterium which is widely distributed in the environment. Up to 7% of healthy

Available online 21 June 2014

adults and up to 45% of infants may have asymptomatic intestinal carriage of C. difficile. A

Keywords:

based molecular typing methods are available for typing C. difficile isolates. C. difficile

Clostridium difficile

virulence evolved independently in the highly epidemic lineages, associated with the

large number of strains of C. difficile have been identified. A number of PCR or sequence-

Diabetes

expression of toxin genes and other virulence factors. This article briefly reviews recent

Host immunity

progresses in the bateriology of C. difficile and highlights the limited knowledge of potential

Gut microbiota

mechanisms for the increased risk of CDI in diabetes which warrants further research. # 2014 Elsevier Ireland Ltd. All rights reserved.

Contents 1. 2. 3.

4.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . Clostridium difficile infection and diabetes . . . . . Bateriology of Clostridium difficile . . . . . . . . . . . . 3.1. Molecular typing of C. difficile . . . . . . . . . 3.2. Molecular pathogenicity of C. difficile . . . 3.2.1. tcdA and tcdB . . . . . . . . . . . . . . . . 3.2.2. tcdR, tcdE, and tcdC in PaLoc . . . . 3.2.3. cdtA and cdtB . . . . . . . . . . . . . . . . 3.2.4. Other virulence factors . . . . . . . . Mechanistic study on the increased CDI risk in Acknowledgements . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

........ ........ ........ ........ ........ ........ ........ ........ ........ diabetes . ........ ........

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* Corresponding author. Tel.: +1 713 500 9950; fax: +1 713 500 0900. E-mail address: [email protected] (H.-Q. Qu). http://dx.doi.org/10.1016/j.diabres.2014.06.002 0168-8227/# 2014 Elsevier Ireland Ltd. All rights reserved.

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1.

diabetes research and clinical practice 105 (2014) 285–294

Introduction

Clostridium difficile (C. difficile) infection (CDI) is a major nosocomial infection [1]. The infection mostly occurs after the gut microbiota is disrupted by the administration of antibiotics, which impairs colonization resistance against C. difficile [2]. CDI accounts for approximately 15% to 25% of antibiotic-associated diarrhea [3]. In addition to antibiotic use, advanced age (>65 years old), impaired immune function, contact with CDI patients or their health care providers, and the use of acid-suppressive therapy, particularly proton pump inhibitors, are common factors increasing the risk of CDI [4–6]. The clinical symptoms include diarrhea, abdominal pain and fever. Sigmoidoscopic examination may reveal pseudomembranes colitis. In rare case, CDI may cause fulminant colitis in which patients may develop septic shock, toxic megacolon, and intestinal perforation [7,8]. Stool culture is the most sensitive test for laboratory diagnosis of CDI and essential for epidemiological studies, whereas it is not clinical practical because of its slow turnaround time [9,10]. Stool culture takes as long as 9 days for results, i.e. 2 to 5 days for culture, 2 days for enrichment, and 1 to 2 days for the isolate toxin cell culture cytotoxicity neutralization assay (CCNA) [10]. Instead, C. difficile glutamate dehydrogenase (GDH) test of stool specimens by enzyme immunoasssays (EIAs) followed by tests for toxins is recommended for CDI diagnosis in suspected patients [11]. The toxin testing by a direct stool CCNA requires only 1 to 2 days [10]. In cases that GDH testing is positive but toxin testing is negative, stool culture with isolate toxin testing is most useful and may detect additional 23% toxigenic C. difficile [10]. Stool culture followed by identification of a toxigenic isolate thus provides the standard against which other clinical test results should be compared, and the availability of an isolate also allows for strain typing and antimicrobial susceptibility testing [9,10]. Oral metronidazole or vancomycin are effective in most cases. However, 15–20% patients may have relapsing infection. Lipiarmycin (also known as clostomicin, tiacumicin, diffimicin, PAR-101, OPT-80, fidaxomicin, and Dificid), an 18membered macrocyclic-lactone antibiotic produced by Actinomycete species, is a novel antibiotic for CDI by inhibition of the switch region of bacterial RNA polymerase (RNAP) [12]. Lipiarmycin has high in vitro activity against C. difficile but low activity against the typical intestinal flora, and is barely absorbed systemically and achieves high stool concentrations [13]. Recurrence is less frequent with lipiarmycin than with vancomycin [14]. Fecal bacteriotherapy to restore disordered gut microbiota has been shown to be highly effective for recurrent CDI patients with disease resolution in 92% patients [15]. Fecal bacteriotherapy involves infusing intestinal microorganisms in a liquid suspension of stool from a healthy donor to restore the intestinal microbiota of a diseased individual [15,16]. To date, fecal bacteriotherapy still presents significant scientific and regulatory challenges. Although donor material is widely available, the complex in stool composition raises concerns about its safety and acceptability, and prevents the procedure from becoming a standard therapeutic option [17]. Further randomized placebo-controlled study on its efficacy and

safety are still underway [18]. Monoclonal antibodies against C. difficile toxins are also under development [19]. Endospores are pivotal to C. difficile transmission. C. difficile spores can survive up to 5 months in environment [20]. C. difficile spores are resistant to prolonged exposure to high temperatures and 70% ethanol, but are effectively inactivated by sporicidal detergents [21]. The proper practice of hand hygiene to prevent C. difficile transmission is to wash hands with soap and water to remove any spores because alcohol-based hand sanitizers cannot kill the spores [9]. CDI is linked to 14,000 deaths in the US each year (http:// www.cdc.gov/vitalsigns/hai/). Due to the emerging hypervirulent strains of C. difficile, this number may still increase [22]. CDI has formed a serious economic burden to the health care system. For the US health care system, the conservative estimation of direct cost for management of CDI is $3.4 billion per year [23].

2.

Clostridium difficile infection and diabetes

Diabetes-related hospitalization and hospital utilization is a serious challenge to health care system [24], a situation which may be further aggravated by nosocomial CDI. According to the American Diabetes Association, nearly 26 million children and adults in the United States have diabetes; another 79 million Americans have prediabetes and are at risk for developing type 2 diabetes. Recent estimates project that as many as 1 in 3 American adults will have diabetes in 2050 unless we take steps to stop diabetes [25]. Diabetes impairs host immunity, and increase the risk of a number of infectious diseases [26–28]. With the international pandemic of type 2 diabetes, it is becoming a major driving force of the epidemic of infectious diseases. The infectious diseases with increased risk in diabetes include CDI. Studies have demonstrated that diabetes increases the risk of recurrent CDI [29,30]. The combined effect of the two studies [29,30] has OR (95% CI) = 2.99(1.88, 4.76) for the increased risk of recurrent CDI in diabetes. In addition, studies showed that diabetes increases the risk of metronidazole treatment failure [31], and is independently associated with C. difficile positivity (CDP) status [32]. To understand the pathogenesis mechanisms of diabetes-increased risk of infectious diseases is a critical approach to gain knowledge for the control of these situations. Also diabetes can serve as a model to understand the interaction between host and pathogens, as demonstrated by our study on the molecular mechanisms of increased risk of tuberculosis in diabetes [27,28]. To discuss these issues, we reviewed recent progress of the molecular bateriology of C. difficile and highlight the limited knowledge of some possible mechanisms in the next sections.

3.

Bateriology of Clostridium difficile

C. difficile is a gram-positive, spore-forming anaerobic bacterium. It was first isolated in meconium samples of normal newborn infants as a component of the normal intestinal flora by Hall and O’Toole [33]. Since the 1970s, C. difficile was recognized as the cause of antibiotic-associated pseudomembranous colitis [34–37]. C. difficile is widely distributed in the

diabetes research and clinical practice 105 (2014) 285–294

environment, e.g. soil, river and sea water, and won’t cause disease normally [38]. Up to 7% healthy adults [39] and up to 45% infants [40] may have asymptomatic intestinal carriage of C. difficile. The C. difficile genome consists of a circular chromosome of 4,290,252 bp encoding 3,776 predicted coding sequences (CDS), and a plasmid of 7881 bp encoding 11 CDSs [41]. For the encoded genes, 19.7% genes are conserved across different C. difficile strains [42].

3.1.

Molecular typing of C. difficile

To date, a large number of strains of C. difficile have been identified. A number of PCR or sequence-based molecular typing methods are available for typing C. difficile isolates, e.g. restriction endonuclease analysis (REA), pulsed-field gel electrophoresis (PFGE), PCR-ribotyping, multilocus variablenumber tandem-repeat analysis (MLVA), amplified fragment length polymorphism (AFLP), surface layer protein A gene sequence typing (slpAST), and multilocus sequence typing (MLST) [43,44]. Three of these methods have been most commonly used to type the strains of C. difficile, i.e. REA, PFGE, and PCR ribotyping [44]. REA and PFGE are restriction enzyme based methods. In the REA typing, the chromosomal DNA of C. difficile is usually digested with HindIII restriction enzyme, which has numerous restriction sites in the C. difficile genome [45]. Because of producing numerous DNA fragments after HindIII digestion, the banding pattern of REA electrophoresis is complicated to read, which limits the clinical application of REA. In the PFGE typing, the chromosomal DNA of C. difficile is usually digested with SmaI restriction enzyme, which gives 7– 15 restriction fragments ranging from 10 to 1100 kbp [46]. Compared with conventional electrophoresis which only separate DNA fragments <30 kbp, PFGE is able to separate large DNA molecules up to 2 Mbp by employing alternately pulsed, perpendicularly oriented electrical fields [47]. Intact genome DNA without degradation is important for PFGE typing, especially for the bands of the large DNA fragments. In North America, a PFGE strain type is designed as NAP (North American Pulsotype) and numerical number based on the banding patterns of electrophoresis, e.g. the epidemic strain NAP1. Compared with PCR ribotyping, the PFGE typing is more complicated, and has low throughput and high consumable costs. In addition, approaches to avoid DNA degradation during PFGE typing are important [48]. Ribotyping is the taxonomic typing method based on evolutionary neutral polymorphisms in the housekeeping genes flanking the ribosomal operons and the copy number difference of ribosomal operons among bacterial species [49]. Conventional ribotyping is based on restriction enzyme digestion of genomic DNA and southern blot visualization of band patterns of DNA fragments [50]. PCR ribotyping enables more expedite and sensitive analysis of ribotypes by using specific primers to amplify the intergenic spacer region (ISR) between the 16S and 23S rRNA genes. Length polymorphisms of the PCR amplicons are used to distinguish different bacterial strains [51]. The application of PCR ribotyping in C. difficile typing has been well documented [52,53]. The PCR ribotyping of C. difficile follows the ECDC (European Centre for Disease Prevention and Control)—Cardiff nomenclature. Like the REA and PFGE methods, the PCR ribotyping is also a

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fingerprint image analysis by electrophoresis banding patterns, and the availability of reference library of fully characterized PCR ribotypes is essential [52]. The restriction enzyme based methods or the PCR ribotyping for C. difficile typing are essentially based on independent reference systems, and it is difficult to correlate the nomenclatures from different typing methods. With the rapid progress of DNA genotyping microarrays and genomic sequencing technology, the unambiguous typing of C. difficile strains by genome-wide genotyping or sequencing, with the genome sequences of different strains as a unified reference system, has a great potential to become practical in the near future [54]. Such a genome-wide genotyping or sequencing based system may need to be updated continuously with evolving of C. difficile and emergence of new strains. Before the available of this kind of typing systems, the profiling of major toxins or other important functional genes of C. difficile are critical for the epidemic monitoring and control of CDI.

3.2.

Molecular pathogenicity of C. difficile

The major genetic locus that mediates the pathogenicity of C. difficile is PaLoc (pathogenicity locus), which is 19.6 kb in length and contains five genes, i.e. in the sequence of physical positions tcdR (previously known as tcdD), tcdB, tcdE, tcdA, and tcdC [55,56]. Compared with the strain typing, the toxin testing is more important clinically as the strain typing described above does not reflect the existence of the toxin genes. As shown by recent study, C. difficile virulence evolved independently in the highly epidemic lineages [57].

3.2.1.

tcdA and tcdB

tcdA and tcdB encodes two principle virulence factors of C. difficile, toxin A (TcdA) and toxin B (TcdB). TcdA and TcdB are structurally and functionally homologous with 63% amino acid (aa) homology [58]. The two toxins are the prototypes of the family of large clostridial toxins (LCTs), a family of glycosyltransferases modifying small GTP-binding proteins [59]. The toxins of the LCT family catalyze the transfer of a glucosyl moiety from UDP-glucose to the intracellular small GTPases, and render the small GTPases functionally inactive [60]. LCT toxins contain four distinct structural domains, i.e. the Nterminal glucosyltransferase domain for the biological activity, the cysteine protease domain for autocatalytic toxin processing, the translocation domain for pore formation and delivery of the catalytic domain into the cytosol, and the C-terminal receptor–binding domain (containing the C-terminal repeat region) for endocytosis [56,61,62]. These functional domains mediate the action of TcdA and TcdB in four major steps: (1) The C-terminal receptor–binding domain binds the surface of the target cell and the toxins enter the cell by endocytosis [63]; (2) low pH of the endosome induces conformational changes for translocation of the N terminus across the membrane [64,65]; (3) the cysteine protease domain catalyzes the autocleavage reaction and release the glucosyltransferase [66]; (4) The glucosyltransferase catalyzes the covalent binding of glucose to small GTPases and inactivates small GTPases [67]. Both TcdA and TcdB glucosylate Rho family substrates, while TcdA has a wider substrate spectrum [68]. TcdA is also capable of modifying Rap family GTPases [69], while Rap

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GTPases have important roles in cell adhesion and cell junction formation [70]. Initially, TcdA was taken as the enterotoxin causing disease, and the pathogenic effect of TcdB was assumed depending on TcdA [71]. Later, TcdB was found of 1000-fold higher cytopathic potency toward cultured cells [68]. Since TcdA-negative and TcdB-positive CDI outbreaks were reported, the pathogenic role of TcdB in CDI was further emphasized [72,73]. By homologous recombination, Lyras et al. found, in the hamster C. difficile-associated diseases (CDAD) model, tcdA mutant (A B+) retained a wild-type virulence phenotype, but tcdB mutant (A+B ) had a significantly attenuated virulence phenotype [74], thereby further highlighted TcdB as essential for the virulence of C. difficile. Using a different hamster model of CDI, Kuehne at al. reestablished the importance of both toxin A and toxin B by showing that isogenic mutants of C. difficile producing either toxin A or toxin B alone can cause fulminant disease in the hamster model of infection [75].

3.2.2.

tcdR, tcdE, and tcdC in PaLoc

Besides tcdB and tcdA, there are three other genes tcdR, tcdE, and tcdC in PaLoc with important roles in CDAD (Fig. 1a) [41,76]. tcdR maps to the upstream of tcdB in PaLoc, and encodes an alternative sigma factor that directs C. difficile RNA polymerase to recognize the promoters of tcdB and tcdA [77]. tcdR serves as the major positive regulator of tcdB and tcdA expression and mediates environmental response [78]. tcdE is located between tcdB and tcdA, and encodes a holin-like pore-forming protein, whose pore-forming activity may allow the specific release of TcdA and TcdB to the extracellular environment [79,80]. tcdC at downstream of tcdA encodes a putative negative regulator of toxin gene expression, which may destabilize the TcdR holoenzyme to prevent transcription of the PaLoc [81]. A nonsense mutation causing a pre-termination codon and the production of a truncated protein of tcdC may contribute to the hypervirulence of the BI/NAP1/027 strain [82,83]. However, the roles of tcdE in toxin release and tcdC in toxin production are still contentious and lack of empirical evidence [84–86].

3.2.3.

cdtA and cdtB

Another important locus involved in CDAD is the C. difficile transferase locus (CdtLoc). This locus encodes a binary toxin

which was first identified in the strain CD196 [87]. Three genes are located in this locus, including cdtR, cdtA and cdtB (Fig. 1b). cdtA encodes the ADP-ribosyltransferase subunit CdtA (the enzymatic component of the binary toxin), and cdtB encodes the ADP-ribosyltransferase binding protein (the binding component of the binary toxin). Either protein alone is noncytotoxic, but acquires cytotoxicity by forming the binary toxin CDT [88]. cdtR encodes a LytTR family response regulator, which plays a key role in the regulatory control of the binary toxin production [89]. The CDT binary toxin is an actin-specific ADP-ribosyltransferase. Through ADP-ribosylation of monomeric G-actin, the CDT binary toxin prevents actin polymerization, promotes depolymerization, and disrupts the cytoskeleton of the cell [90,91]. Recent study by Schwan et al. shows that the CDT binary toxin inhibits F-actin formation and causes microfilament depolymerization. Without grossly changing the cell morphology, this effect induces cellular protrusions in epithelial cells and leads to increased adherence of bacteria [92]. The role of the CDT binary toxin in CDAD is still unclear. The production of the binary toxin has no necessary correlation with the C. difficile ribotypes [93]. The binary toxin genes were detected in approximately 6–15% C. difficile Strains [94,95]. The CDT binary toxin may play an adjunctive role to toxins A and B in the severity of CDAD while itself may not be sufficient to cause disease [96,97]. However, a case of bacteremia with a LCT-negative, binary toxin-positive strain of C. difficile has been reported [98].

3.2.4.

Other virulence factors

Numerous virulence factors of C. difficile are still to be recognized. Antimicrobial resistance genes mediated through conjugative transposons are commonly seen in C. difficile Strains, e.g. tetW and tetM encoding ribosomal protection proteins conferring resistance to tetracycline, and ermB encoding rRNA adenine N-6-methyltransferase conferring resistance to erythromycin [41,99]. Stress response proteins, including rubrerythrins, tellurium resistance proteins, bacterioferritin, catalase, and superoxide dismutase may contribute to the protection of spores from oxidative stress during germination [21]. Many adhesion factors including surface layer proteins (SLPs) contribute to the gut colonization of

Fig. 1 – The two pathogenicity loci of the C. difficile genome. (a) The PaLoc of C. difficile strain CD630. This figure is based on the NCBI reference sequence: NC_009089.1 [133]. (b) The Cdt locus of C. difficile strain CD196. The CDT toxin was first identified in the strain CD196 [87]. This figure is based on the NCBI reference sequence: NC_013315.1 [134].

diabetes research and clinical practice 105 (2014) 285–294

C. difficile [100]. Cwp84, a cysteine protease has been characterized, which is associated with the S-layer proteins and contributes to the degradation of the host tissue integrity [101]. Detection of virulence factors is important for the control of CDI epidemic and for clinical decisions of individualized treatment. Enzyme immunoassay (EIA) testing have been used for the detection of C. difficile toxin A and B in stool specimens [9,102]. Toxinotyping based on the restriction fragment length polymorphisms (RFLPs) of the tcdB and tcdA genes remains to be a practical approach to monitor the epidemic strains [103,104]. Direct amplification of virulence genes including tcdA, tcdB, cdtA, and cdtB, etc. by polymerase chain reaction (PCR) is being widely adopted [94,105,106]. Futhuremore, rapid progress of genomic technologies enables transcriptome analysis of C. difficile, which is rapidly increasing and unprecedentedly sophisticating our knowledge on important molecules involved in CDI, e.g. host adaptation [107], nutrient acquisition [108], and heat resistance [109]. The application of high-throughput whole genome sequencing technologies is discovering novel genes and molecular mechanisms within the evolving C. difficile [57].

4. Mechanistic study on the increased CDI risk in diabetes Diabetes impairs host immunity, and increases the risk of a number of infectious diseases [26]. In CDI, the normal function of macrophages may be important to contain the infection of C. difficile spores [110,111]. After phagocytosis of C. difficile spores by macrophages, spores can remain dormant and be able to survive and produce cytotoxic effects on macrophages [110]. Our previous study showed decreased expression of ATPase, H+ transporting, lysosomal 50/57 kDa, V1 subunit H in diabetes patients by a longitudinal investigation [27]. The principle function of V-ATPase is to acidify intracellular compartments [112,113]. V-ATPase is involved in endocytosis through the acidification of the phagosome [114]—a critical process in intracellular microbial killing [115], especially in specialized phagocytic cells such as macrophages. Among 376 genes related to human tuberculosis [116], we also observed decreased expression of the HK2 gene in diabetes [28]. The HK2 gene encodes hexokinase 2, which is a critical mediator of aerobic glycolysis [117]. Aerobic glycolysis is the unique energy source for macrophages [118]. Decreased expression of HK2 may impair macrophage function, thus increasing the risk of tuberculosis. In addition, host genes involved in the activation of macrophages may also be involved in the CDI risk in diabetes. MyD88, encoded by the myeloid differentiation primary response 88 gene, is a key downstream adapter for most Toll-like receptors (TLRs) and interleukin-1 receptors (IL1Rs) and plays a central role in the innate and adaptive immune response [119,120]. C. difficile spore-mediated transmission to mice deficient in MyD88 leads to a severe intestinal disease that is often fatal, while transmission to control mice treated with antibiotics results in self-limiting mucosal inflammation of the large intestine [121]. MyD88-deficiency also increases risk of metabolic syndrome and diabetes in mice [122]. Among the TLRs, Ryan et al. showed that toll-like receptor 4 (TLR4) mediates the recognition of the surface layer

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proteins (SLPs) of C. difficile and activates innate and adaptive immunity [123]. However, the roles of MyD88 and TLR4 in the increased CDI risk in diabetes is unclear. Dasu et al. found that subjects with type 2 diabetes had increased expression of MyD88 and TLR4 [124]. Although impaired host immunity in diabetes [26,28] may contribute to the susceptibility of CDI in diabetes, we must acknowledge that impaired colonization resistance against C. difficile is the major cause of CDI [2]. A plausible hypothesis of the colonization resistance against CDI is the niche exclusion hypothesis, i.e. exclusion of toxigenic C. difficile by nontoxigenic C. difficile [2,125,126]. Pre-colonization of the intestinal tract with non-toxigenic C. difficile may exclude toxigenic C. difficile by the ability of the non-toxigenic C. difficile strain to outcompete toxigenic C. difficile for a limiting nutrient [125,126]. Chang et al. showed that patients with RCDI had decreased species of Bacteroidetes and Firmicutes in their stool as compared to patients with just single episode of CDI [127]. On the other hand, changed ratio of Bacteroidetes and Firmicutes in diabetes gut microbiota has been suggested by recent studies [128,129]. According to the study by Larsen et al., the ratio of Bacteroidetes to Firmicutes increased in diabetes [128]. In contrast, obese mice and humans without diabetes had a reduction in the abundance of Bacteroidetes and a proportional increase in Firmicutes [130]. Most of the components of Firmicutes belong to the class Clostridia. The proportion of Clostridia in diabetes is significantly lower than that in controls [128]. This change of gut microbiota may explain some of the increased risk of CDI in diabetes through weakened niche exclusion against C. difficile. In addition, virulence of pathogenic C. difficile may also vary depending on the community structure of the flora [131]. The known success of fecal transplantation in the treatment of refractory or recurrent CDI [15], as well as the research enthusiasm of fecal transplantation in the treatment diabetes [17,132] (ClinicalTrials.gov Identifier: of NCT01790711), is transmitting positive message about fecal bacteriotherapy. However, fecal bacteriotherapy presents significantly scientific and regulatory challenges on the Food Drug Administration (FDA) regulatory policies of novel live biologics to assure the safety and rights of subjects. This issue highlights the importance of mechanistic study on the microbiota change associated with CDI in diabetes. Through further study, a true and deeper understanding of the impact of diabetes on the gut microbiota, as well as a better knowledge of probiotic and prebiotic mechanisms with clinical indication may enable successful future implementation of effective bacteriotherapy for CDI in diabetes using microbial or polymicrobial consortium.

Conflict of interest statement None declared.

Acknowledgments H.Q.Q. is supported by intramural funding from the University of Texas School of Public Health. The funder had no role in decision to publish or preparation of the manuscript.

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references

[1] Archibald LK, Banerjee SN, Jarvis WR. Secular trends in hospital-acquired Clostridium difficile disease in the United States 1987–2001. J Infect Dis 2004;189:1585–9. [2] Britton RA, Young VB. Interaction between the intestinal microbiota and host in Clostridium difficile colonization resistance. Trends Microbiol 2012;20:313–9. [3] Bartlett JG. Clostridium difficile: clinical considerations. Rev Infect Dis 1990;12(Suppl 2):S243–51. [4] Rupnik M, Wilcox MH, Gerding DN. Clostridium difficile infection: new developments in epidemiology and pathogenesis. Nat Rev Microbiol 2009;7:526–36. [5] McFarland LV, Mulligan ME, Kwok RY, Stamm WE. Nosocomial acquisition of Clostridium difficile infection. N Engl J Med 1989;320:204–10. [6] Dial S, Delaney JA, Barkun AN, Suissa S. Use of gastric acid-suppressive agents and the risk of communityacquired Clostridium difficile-associated disease. JAMA 2005;294:2989–95. [7] Kelly CP, LaMont JT. Clostridium difficile infection. Annu Rev Med 1998;49:375–90. [8] Deneve C, Janoir C, Poilane I, Fantinato C, Collignon A. New trends in Clostridium difficile virulence and pathogenesis. Int J Antimicrob Agents 2009;33(Suppl 1):S24–8. [9] Cohen Stuart H, Gerding Dale N, Stuart Johnson, Kelly Ciaran P, Loo Vivian G, McDonald LC, Pepin J, Wilcox Mark H. Clinical practice guidelines for Clostridium difficile infection in adults: 2010 update by the Society for Healthcare Epidemiology of America (SHEA) and the Infectious Diseases Society of America (IDSA). Infect Control Hosp Epidemiol 2010;31:431–55. [10] Reller ME, Lema CA, Perl TM, Cai M, Ross TL, Speck KA, Carroll KC. Yield of stool culture with isolate toxin testing versus a two-step algorithm including stool toxin testing for detection of toxigenic Clostridium difficile. J Clin Microbiol 2007;45:3601–5. [11] Crobach MJT, Dekkers OM, Wilcox MH, Kuijper EJ. European Society of Clinical Microbiology and Infectious Diseases (ESCMID): data review and recommendations for diagnosing Clostridium difficile-infection (CDI). Clin Microbiol Infect 2009;15:1053–66. [12] Srivastava A, Talaue M, Liu S, Degen D, Ebright RY, Sineva E, Chakraborty A, Druzhinin SY, Chatterjee S, Mukhopadhyay J, Ebright YW, Zozula A, Shen J, Sengupta S, Niedfeldt RR, Xin C, Kaneko T, Irschik H, Jansen R, Donadio S, Connell N, Ebright RH. New target for inhibition of bacterial RNA polymerase: ‘switch region’. Curr Opin Microbiol 2011;14:532–43. [13] Shue YK, Sears PS, Shangle S, Walsh RB, Lee C, Gorbach SL, Okumu F, Preston RA. Safety, tolerance, and pharmacokinetic studies of OPT-80 in healthy volunteers following single and multiple oral doses. Antimicrob Agents Chemother 2008;52:1391–5. [14] Drekonja DM, Butler M, MacDonald R, Bliss D, Filice GA, Rector TS, Wilt TJ. Comparative effectiveness of Clostridium difficile treatments: a systematic review. Ann Intern Med 2011;155:839–47. [15] Gough E, Shaikh H, Manges AR. Systematic review of intestinal microbiota transplantation (fecal bacteriotherapy) for recurrent Clostridium difficile infection. Clin Infect Dis 2011;53:994–1002. [16] Surawicz CM, Alexander J. Treatment of refractory and recurrent Clostridium difficile infection. Nat Rev Gastroenterol Hepatol 2011;8:330–9.

[17] Borody TJ, Khoruts A. Fecal microbiota transplantation and emerging applications. Nat Rev Gastroenterol Hepatol 2012;9:88–96. [18] van Nood E, Vrieze A, Nieuwdorp M, Fuentes S, Zoetendal EG, de Vos WM, Visser CE, Kuijper EJ, Bartelsman JFWM, Tijssen JGP, Speelman P, Dijkgraaf MGW, Keller JJ. Duodenal infusion of donor feces for recurrent Clostridium difficile. N Engl J Med 2013;368:407–15. [19] Lowy I, Molrine DC, Leav BA, Blair BM, Baxter R, Gerding DN, Nichol G, Thomas Jr WD, Leney M, Sloan S, Hay CA, Ambrosino DM. Treatment with monoclonal antibodies against Clostridium difficile toxins. N Engl J Med 2010;362:197–205. [20] Kim KH, Fekety R, Batts DH, Brown D, Cudmore M, Silva Jr J, Waters D. Isolation of Clostridium difficile from the environment and contacts of patients with antibioticassociated colitis. J Infect Dis 1981;143:42–50. [21] Lawley TD, Croucher NJ, Yu L, Clare S, Sebaihia M, Goulding D, Pickard DJ, Parkhill J, Choudhary J, Dougan G. Proteomic and genomic characterization of highly infectious Clostridium difficile 630 spores. J Bacteriol 2009;191:5377–86. [22] Cookson B. Hypervirulent strains of Clostridium difficile. Postgrad Med J 2007;83:291–5. [23] Dubberke Erik R, Wertheimer Albert I. Review of current literature on the economic burden of Clostridium difficile infection. Infect Control Hosp Epidemiol 2009;30:57–66. [24] Aubert RE, Geiss LS, Ballard DJ, Cocanougher B, Herman WH. Diabetes-related hospitalization and hospital utilization. Diabetes Am 1995;2:553–69. [25] Boyle JP, Thompson TJ, Gregg EW, Barker LE, Williamson DF. Projection of the year 2050 burden of diabetes in the US adult population: dynamic modeling of incidence, mortality, and prediabetes prevalence. Popul Health Metrics 2010;8:29. [26] Geerlings SE, Hoepelman AI. Immune dysfunction in patients with diabetes mellitus (DM). FEMS Immunol Med Microbiol 1999;26:259–65. [27] Molina MF, Qu HQ, Rentfro AR, Nair S, Lu Y, Hanis CL, McCormick JB, Fisher-Hoch SP. Decreased expression of ATP6V1H in type 2 diabetes: a pilot report on the diabetes risk study in Mexican Americans. Biochem Biophys Res Commun 2011;412:728–31. [28] Qu HQ, Rentfro AR, Lu Y, Nair S, Hanis CL, McCormick JB, Fisher-Hoch SP. Host susceptibility to tuberculosis: insights from a longitudinal study of gene expression in diabetes. Int J Tuberc Lung Dis 2012;16:370–2. [29] Shakov R, Salazar RS, Kagunye SK, Baddoura WJ, DeBari VA. Diabetes mellitus as a risk factor for recurrence of Clostridium difficile infection in the acute care hospital setting. Am J Infect Control 2011;39:194–8. [30] Dial S, Alrasadi K, Manoukian C, Huang A, Menzies D. Risk of Clostridium difficile diarrhea among hospital inpatients prescribed proton pump inhibitors: cohort and casecontrol studies. Can Med Assoc J 2004;171:33–8. [31] Jung KS, Park JJ, Chon YE, Jung ES, Lee HJ, Jang HW, Lee KJ, Lee SH, Moon CM, Lee JH, Shin JK, Jeon SM, Hong SP, Kim TI, Kim WH, Cheon JH. Risk factors for treatment failure and recurrence after metronidazole treatment for Clostridium difficile-associated diarrhea. Gut Liver 2010;4:332–7. [32] Rodrigues MA, Brady RR, Rodrigues J, Graham C, Gibb AP. Clostridium difficile infection in general surgery patients; identification of high-risk populations. Int J Surg 2010;8:368–72. [33] Hall IC, O’Toole E. Intestinal flora in new-born infants: with a description of a new pathogenic anaerobe, bacillus difficilis. Am J Dis Child 1935;49:390–402.

diabetes research and clinical practice 105 (2014) 285–294

[34] Tedesco FJ, Barton RW, Alpers DH. Clindamycinassociated colitis. A prospective study. Ann Intern Med 1974;81:429–33. [35] Bartlett JG. Historical perspectives on studies of Clostridium difficile and C. difficile infection. Clin Infect Dis 2008;46:S4– 11. [36] Bartlett JG, Onderdonk AB, Cisneros RL, Kasper DL. Clindamycin-associated colitis due to a toxin-producing species of Clostridium in hamsters. J Infect Dis 1977;136:701–5. [37] Bartlett JG, Chang TW, Gurwith M, Gorbach SL, Onderdonk AB. Antibiotic-associated pseudomembranous colitis due to toxin-producing clostridia. N Engl J Med 1978;298:531–4. [38] Al Saif N, BRAZIER JS. The distribution of Clostridium difficile in the environment of South Wales. J Med Microbiol 1996;45:133–7. [39] Kato H, Kita H, Karasawa T, Maegawa T, Koino Y, Takakuwa H, Saikai T, Kobayashi K, Yamagishi T, Nakamura S. Colonisation and transmission of Clostridium difficile in healthy individuals examined by PCR ribotyping and pulsed-field gel electrophoresis. J Med Microbiol 2001;50:720–7. [40] Rousseau C, Poilane I, De Pontual L, Maherault A-C, Le Monnier A, Collignon A. Clostridium difficile carriage in healthy infants in the community: a potential reservoir for pathogenic strains. Clin Infect Dis 2012;55:1209–15. [41] Sebaihia M, Wren BW, Mullany P, Fairweather NF, Minton N, Stabler R, Thomson NR, Roberts AP, Cerdeno-Tarraga AM, Wang H, Holden MTG, Wright A, Churcher C, Quail MA, Baker S, Bason N, Brooks K, Chillingworth T, Cronin A, Davis P, Dowd L, Fraser A, Feltwell T, Hance Z, Holroyd S, Jagels K, Moule S, Mungall K, Price C, Rabbinowitsch E, Sharp S, Simmonds M, Stevens K, Unwin L, Whithead S, Dupuy B, Dougan G, Barrell B, Parkhill J. The multidrug-resistant human pathogen Clostridium difficile has a highly mobile, mosaic genome. Nat Genet 2006;38:779–86. [42] Stabler RA, Gerding DN, Songer JG, Drudy D, Brazier JS, Trinh HT, Witney AA, Hinds J, Wren BW. Comparative phylogenomics of Clostridium difficile reveals clade specificity and microevolution of hypervirulent strains. J Bacteriol 2006;188:7297–305. [43] Killgore G, Thompson A, Johnson S, Brazier J, Kuijper E, Pepin J, Frost EH, Savelkoul P, Nicholson B, van den Berg RJ, Kato H, Sambol SP, Zukowski W, Woods C, Limbago B, Gerding DN, McDonald LC. Comparison of seven techniques for typing international epidemic strains of Clostridium difficile: restriction endonuclease analysis, pulsed-field gel electrophoresis, PCR-ribotyping, multilocus sequence typing, multilocus variable-number tandem-repeat analysis, amplified fragment length polymorphism, and surface layer protein A gene sequence typing. J Clin Microbiol 2008;46:431–7. [44] Janezic S, Rupnik M. Molecular typing methods for Clostridium difficile: pulsed-field gel electrophoresis and PCR ribotyping. Methods Mol Biol 2010;646:55–65. [45] Clabots CR, Johnson S, Bettin KM, Mathie PA, Mulligan ME, Schaberg DR, Peterson LR, Gerding DN. Development of a rapid and efficient restriction endonuclease analysis typing system for Clostridium difficile and correlation with other typing systems. J Clin Microbiol 1993;31:1870–5. [46] Bidet P, Lalande V, Salauze B, Burghoffer B, Avesani V, Delmee M, Rossier A, Barbut F, Petit JC. Comparison of PCR-ribotyping, arbitrarily primed PCR, and pulsed-field gel electrophoresis for typing Clostridium difficile. J Clin Microbiol 2000;38:2484–7. [47] Schwartz DC, Cantor CR. Separation of yeast chromosome-sized DNAs by pulsed field gradient gel electrophoresis. Cell 1984;37:67–75.

291

[48] Gal M, Northey G, Brazier JS. A modified pulsed-field gel electrophoresis (PFGE) protocol for subtyping previously non-PFGE typeable isolates of Clostridium difficile polymerase chain reaction ribotype 001. J Hosp Infect 2005;61:231–6. [49] Bouchet V, Huot H, Goldstein R. Molecular genetic basis of ribotyping. Clin Microbiol Rev 2008;21:262–73. [50] Grimont F, Grimont PAD. Ribosomal ribonucleic acid gene restriction patterns as potential taxonomic tools. Ann Inst Pasteur/Microbiol 1986;137:165–75. [51] Kostman JR, Alden MB, Mair M, Edlind TD, LiPuma JJ, Stull TL. A universal approach to bacterial molecular epidemiology by polymerase chain reaction ribotyping. J Infect Dis 1995;171:204–8. [52] Stubbs SLJ, Brazier JS, O’Neill GL, Duerden BI. PCR targeted to the 16S–23S rRNA gene intergenic spacer region of Clostridium difficile and construction of a library consisting of 116 different PCR ribotypes. J Clin Microbiol 1999;37:461– 3. [53] Bidet P, Barbut F, Lalande V, Burghoffer B, Petit JC. Development of a new PCR-ribotyping method for Clostridium difficile based on ribosomal RNA gene sequencing. FEMS Microbiol Lett 1999;175:261–6. [54] Didelot X, Eyre D, Cule M, Ip C, Ansari A, Griffiths D, Vaughan A, O’Connor L, Golubchik T, Batty E, Piazza P, Wilson D, Bowden R, Donnelly P, Dingle K, Wilcox M, Walker S, Crook D, Peto T, Harding R. Microevolutionary analysis of Clostridium difficile genomes to investigate transmission. Genome Biol 2012;13:R118. [55] Rupnik M, Dupuy B, Fairweather NF, Gerding DN, Johnson S, Just I, Lyerly DM, Popoff MR, Rood JI, Sonenshein AL, Thelestam M, Wren BW, Wilkins TD, von Eichel-Streiber C. Revised nomenclature of Clostridium difficile toxins and associated genes. J Med Microbiol 2005;54:113–7. [56] Voth DE, Ballard JD. Clostridium difficile toxins: mechanism of action and role in disease. Clin Microbiol Rev 2005;18:247–63. [57] He M, Sebaihia M, Lawley TD, Stabler RA, Dawson LF, Martin MJ, Holt KE, Seth-Smith HM, Quail MA, Rance R, Brooks K, Churcher C, Harris D, Bentley SD, Burrows C, Clark L, Corton C, Murray V, Rose G, Thurston S, van Tonder A, Walker D, Wren BW, Dougan G, Parkhill J. Evolutionary dynamics of Clostridium difficile over short and long time scales. Proc Nat Acad Sci USA 2010;107:7527–32. [58] von Eichel-Streiber C, Laufenberg-Feldmann R, Sartingen S, Schulze J, Sauerborn M. Comparative sequence analysis of the Clostridium difficile toxins A and B. Mol Gen Genet 1992;233:260–8. [59] von Eichel-Streiber C, Boquet P, Sauerborn M, Thelestam M. Large clostridial cytotoxins—a family of glycosyltransferases modifying small GTP-binding proteins. Trends Microbiol 1996;4:375–82. [60] Just I, Gerhard R. Large clostridial cytotoxins. Rev Physiol Biochem Pharmacol 2004;152:23–47. [61] Jank T, Aktories K. Structure and mode of action of clostridial glucosylating toxins: the ABCD model. Trends Microbiol 2008;16:222–9. [62] Albesa-Jove´ D, Bertrand T, Carpenter EP, Swain GV, Lim J, Zhang J, Haire LF, Vasisht N, Braun V, Lange A, von EichelStreiber C, Svergun DI, Fairweather NF, Brown KA. Four distinct structural domains in Clostridium difficile toxin B visualized using SAXS. J Mol Biol 2010;396:1260–70. [63] Frisch C, Gerhard R, Aktories K, Hofmann F, Just I. The complete receptor–binding domain of Clostridium difficile toxin A is required for endocytosis. Biochem Biophys Res Commun 2003;300:706–11. [64] Pruitt RN, Chambers MG, Ng KK-S, Ohi MD, Lacy DB. Structural organization of the functional domains of

292

[65]

[66]

[67]

[68]

[69]

[70] [71]

[72]

[73]

[74]

[75]

[76]

[77]

[78]

[79]

[80]

[81]

[82]

diabetes research and clinical practice 105 (2014) 285–294

Clostridium difficile toxins A and B. Proc Nat Acad Sci 2010;107:13467–72. Qa’Dan M, Spyres LM, Ballard JD. pH-Induced Conformational Changes in Clostridium difficile toxin B. Infect Immun 2000;68:2470–4. Egerer M, Giesemann T, Jank T, Satchell KJF, Aktories K. Auto-catalytic cleavage of Clostridium difficile toxins A and B depends on cysteine protease activity. J Biol Chem 2007;282:25314–21. Just I, Selzer J, Wilm M, Eichel-Streiber Cv, Mann M, Aktories K. Glucosylation of Rho proteins by Clostridium difficile toxin B. Nature 1995;375:500–3. Chaves-Olarte E, Weidmann M, Eichel-Streiber C, Thelestam M. Toxins A and B from Clostridium difficile differ with respect to enzymatic potencies, cellular substrate specificities, and surface binding to cultured cells. J Clin Invest 1997;100:1734–41. Pruitt RN, Chumbler NM, Rutherford SA, Farrow MA, Friedman DB, Spiller B, Lacy DB. Structural determinants of Clostridium difficile toxin A glucosyltransferase activity. J Biol Chem 2012;287:8013–20. Raaijmakers JH, Bos JL. Specificity in Ras and Rap signaling. J Biol Chem 2009;284:10995–9. Lyerly DM, Saum KE, MacDonald DK, Wilkins TD. Effects of Clostridium difficile toxins given intragastrically to animals. Infect Immun 1985;47:349–52. al-Barrak A, Embil J, Dyck B, Olekson K, Nicoll D, Alfa M, Kabani A. An outbreak of toxin A negative, toxin B positive Clostridium difficile-associated diarrhea in a Canadian tertiary-care hospital. Can Commun Dis Rep 1999;25:65–9. Alfa MJ, Kabani A, Lyerly D, Moncrief S, Neville LM, AlBarrak A, Harding GK, Dyck B, Olekson K, Embil JM. Characterization of a toxin A-negative, toxin B-positive strain of Clostridium difficile responsible for a nosocomial outbreak of Clostridium difficile-associated diarrhea. J Clin Microbiol 2000;38:2706–14. Lyras D, O’Connor JR, Howarth PM, Sambol SP, Carter GP, Phumoonna T, Poon R, Adams V, Vedantam G, Johnson S, Gerding DN, Rood JI. Toxin B is essential for virulence of Clostridium difficile. Nature 2009;458:1176–9. Kuehne SA, Cartman ST, Heap JT, Kelly ML, Cockayne A, Minton NP. The role of toxin A and toxin B in Clostridium difficile infection. Nature 2010;467:711–3. Karen CC, John GB. Biology of Clostridium difficile: implications for epidemiology and diagnosis. Annu Rev Microbiol 2011;65:501–21. Mani N, Dupuy B. Regulation of toxin synthesis in Clostridium difficile by an alternative RNA polymerase sigma factor. Proc Nat Acad Sci 2001;98:5844–9. Mani N, Lyras D, Barroso L, Howarth P, Wilkins T, Rood JI, Sonenshein AL, Dupuy B. Environmental response and autoregulation of Clostridium difficile TxeR, a sigma factor for toxin gene expression. J Bacteriol 2002;184:5971–8. Govind R, Dupuy B. Secretion of Clostridium difficile toxins A and B requires the holin-like protein TcdE. PLoS Pathog 2012;8:e1002727. Tan KS, Wee BY, Song KP. Evidence for holin function of tcdE gene in the pathogenicity of Clostridium difficile. J Med Microbiol 2001;50:613–9. Matamouros S, England P, Dupuy B. Clostridium difficile toxin expression is inhibited by the novel regulator TcdC. Mol Microbiol 2007;64:1274–88. Carter GP, Douce GR, Govind R, Howarth PM, Mackin KE, Spencer J, Buckley AM, Antunes A, Kotsanas D, Jenkin GA, Dupuy B, Rood JI, Lyras D. The anti-sigma factor TcdC modulates hypervirulence in an epidemic BI/NAP1/027 clinical isolate of Clostridium difficile. PLoS Pathog 2011;7:e1002317.

[83] Warny M, Pepin J, Fang A, Killgore G, Thompson A, Brazier J, Frost E, McDonald LC. Toxin production by an emerging strain of Clostridium difficile associated with outbreaks of severe disease in North America and Europe. Lancet 2005;366:1079–84. [84] Murray R, Boyd D, Levett PN, Mulvey MR, Alfa MJ. Truncation in the tcdC region of the Clostridium difficile PathLoc of clinical isolates does not predict increased biological activity of Toxin B or Toxin A. BMC Infect Dis 2009;9:103. [85] Cartman ST, Kelly ML, Heeg D, Heap JT, Minton NP. Precise manipulation of the Clostridium difficile chromosome reveals a lack of association between the tcdC genotype and toxin production. Appl Environ Microbiol 2012;78:4683–90. [86] Olling A, Seehase S, Minton NP, Tatge H, Schro¨ter S, Kohlscheen S, Pich A, Just I, Gerhard R. Release of TcdA and TcdB from Clostridium difficile cdi 630 is not affected by functional inactivation of the tcdE gene. Microb Pathog 2012;52:92–100. [87] Popoff MR, Rubin EJ, Gill DM, Boquet P. Actin-specific ADPribosyltransferase produced by a Clostridium difficile strain. Infect Immun 1988;56:2299–306. [88] Sundriyal A, Roberts AK, Ling R, McGlashan J, Shone CC, Acharya KR. Expression, purification and cell cytotoxicity of actin-modifying binary toxin from Clostridium difficile. Protein Expression Purif 2010;74:42–8. [89] Carter GP, Lyras D, Allen DL, Mackin KE, Howarth PM, O’Connor JR, Rood JI. Binary toxin production in Clostridium difficile is regulated by CdtR, a LytTR family response regulator. J Bacteriol 2007;189:7290–301. [90] Considine RV, Simpson LL. Cellular and molecular actions of binary toxins possessing ADP-ribosyltransferase activity. Toxicon 1991;29:913–36. [91] Perelle S, Gibert M, Bourlioux P, Corthier G, Popoff MR. Production of a complete binary toxin (actin-specific ADPribosyltransferase) by Clostridium difficile CD196. Infect Immun 1997;65:1402–7. [92] Schwan C, Stecher B, Tzivelekidis T, van Ham M, Rohde M, Hardt WD, Wehland J, Aktories K. Clostridium difficile toxin CDT induces formation of microtubule-based protrusions and increases adherence of bacteria. PLoS Pathog 2009;5:e1000626. [93] Poilane I, Fantinato C, Cruaud P, Collignon A. [Epidemiological study of Clostridium difficile strains isolated in Jean-Verdier-Rene-Muret hospitals from 2001 to 2007]. Pathol Biol (Paris) 2008;56:412–6. [94] Goncalves C, Decre D, Barbut F, Burghoffer B, Petit JC. Prevalence and characterization of a binary toxin (actinspecific ADP-ribosyltransferase) from Clostridium difficile. J Clin Microbiol 2004;42:1933–9. [95] Geric B, Johnson S, Gerding DN, Grabnar M, Rupnik M. Frequency of binary toxin genes among Clostridium difficile strains that do not produce large clostridial toxins. J Clin Microbiol 2003;41:5227–32. [96] Geric B, Carman RJ, Rupnik M, Genheimer CW, Sambol SP, Lyerly DM, Gerding DN, Johnson S. Binary toxin– producing, large clostridial toxin–negative Clostridium difficile strains are enterotoxic but do not cause disease in hamsters. J Infect Dis 2006;193:1143–50. [97] Barbut F, Decre´ D, Lalande V, Burghoffer B, Noussair L, Gigandon A, Espinasse F, Raskine L, Robert J, Mangeol A, Branger C, Petit J-C. Clinical features of Clostridium difficileassociated diarrhoea due to binary toxin (actin-specific ADP-ribosyltransferase)-producing strains. J Med Microbiol 2005;54:181–5. [98] Elliott B, Reed R, Chang BJ, Riley TV. Bacteremia with a large clostridial toxin-negative, binary toxin-positive strain of Clostridium difficile. Anaerobe 2009;15:249–51.

diabetes research and clinical practice 105 (2014) 285–294

[99] Fry PR, Thakur S, Abley M, Gebreyes WA. Antimicrobial resistance, toxinotype, and genotypic profiling of Clostridium difficile isolates of swine origin. J Clin Microbiol 2012;50:2366–72. [100] Calabi E, Calabi F, Phillips AD, Fairweather NF. Binding of Clostridium difficile surface layer proteins to gastrointestinal tissues. Infect Immun 2002;70:5770–8. [101] Janoir C, Pe´chine´ S, Grosdidier C, Collignon A. Cwp84, a surface-associated protein of Clostridium difficile is a cysteine protease with degrading activity on extracellular matrix proteins. J Bacteriol 2007;189:7174–80. [102] Barbut F, Kajzer C, Planas N, Petit JC. Comparison of three enzyme immunoassays, a cytotoxicity assay, and toxigenic culture for diagnosis of Clostridium difficileassociated diarrhea,. J Clin Microbiol 1993;31:963–7. [103] Rupnik M, Avesani V, Janc M, von Eichel-Streiber C, Delme´e M. A novel toxinotyping scheme and correlation of toxinotypes with serogroups of Clostridium difficile isolates. J Clin Microbiol 1998;36:2240–7. [104] Rupnik M. Clostridium difficile toxinotyping. Methods Mol Biol 2010;646:67–76. [105] Stubbs S, Rupnik M, Gibert M, Brazier J, Duerden B, Popoff M. Production of actin-specific ADP-ribosyltransferase (binary toxin) by strains of Clostridium difficile. FEMS Microbiol Lett 2000;186:307–12. [106] Persson S, Torpdahl M, Olsen KE. New multiplex PCR method for the detection of Clostridium difficile toxin A (tcdA) and toxin B (tcdB) and the binary toxin (cdtA/cdtB) genes applied to a Danish strain collection. Clin Microbiol Infect 2008;14:1057–64. [107] Janvilisri T, Scaria J, Thompson AD, Nicholson A, Limbago BM, Arroyo LG, Songer JG, Grohn YT, Chang YF. Microarray identification of Clostridium difficile core components and divergent regions associated with host origin. J Bacteriol 2009;191:3881–91. [108] Antunes A, Camiade E, Monot M, Courtois E, Barbut F, Sernova NV, Rodionov DA, Martin-Verstraete I, Dupuy B. Global transcriptional control by glucose and carbon regulator CcpA in Clostridium difficile. Nucleic Acids Res 2012;40:10701–18. [109] Ternan NG, Jain S, Srivastava M, McMullan G. Comparative transcriptional analysis of clinically relevant heat stress response in Clostridium difficile strain 630. PLoS One 2012;7:e42410. [110] Paredes-Sabja D, Cofre-Araneda G, Brito-Silva C, PizarroGuajardo M, Sarker MR. Clostridium difficile @@spore– macrophage interactions: spore survival. PLoS One 2012;7:e43635. [111] Paredes-Sabja D, Sarker MR. Interactions between Clostridium perfringens spores and Raw 264.7 macrophages. Anaerobe 2012;18:148–56. [112] Stevens TH, Forgac M. Structure, function and regulation of the vacuolar (H+)-ATPase. Annu Rev Cell Dev Biol 1997;13:779–808. [113] Geyer M, Fackler OT, Peterlin BM. Subunit H of the VATPase involved in endocytosis shows homology to betaadaptins. Mol Biol Cell 2002;13:2045–56. [114] Lee BY, Jethwaney D, Schilling B, Clemens DL, Gibson BW, Horwitz MA. The Mycobacterium bovis bacille Calmette– Guerin phagosome proteome. Mol Cell Proteomics (United States) 2010;9:32–53. [115] Ip WK, Sokolovska A, Charriere GM, Boyer L, Dejardin S, Cappillino MP, Yantosca LM, Takahashi K, Moore KJ, LacyHulbert A, Stuart LM. Phagocytosis and phagosome acidification are required for pathogen processing and MyD88-dependent responses to Staphylococcus aureus. J Immunol (United States) 2010;184:7071–81. [116] Berry MP, Graham CM, McNab FW, Xu Z, Bloch SA, Oni T, Wilkinson KA, Banchereau R, Skinner J, Wilkinson RJ. An

[117]

[118]

[119]

[120]

[121]

[122]

[123]

[124]

[125]

[126]

[127]

[128]

[129]

[130]

[131]

[132]

293

interferon-inducible neutrophil-driven blood transcriptional signature in human tuberculosis. Nature 2010;466:973–7. Wolf A, Agnihotri S, Micallef J, Mukherjee J, Sabha N, Cairns R, Hawkins C, Guha A. Hexokinase 2 is a key mediator of aerobic glycolysis and promotes tumor growth in human glioblastoma multiforme. J Exp Med 2011;208:313–26. Rist R, Naftalin RJ. Dexamethasone inhibits the hexose monophosphate shunt in activated rat peritoneal macrophages by reducing hexokinase-dependent sugar uptake. Biochem J 1991;278:129–35. von Bernuth H, Picard C, Jin Z, Pankla R, Xiao H, Ku C-L, Chrabieh M, Mustapha IB, Ghandil P, Camcioglu Y. Pyogenic bacterial infections in humans with MyD88 deficiency. Science 2008;321:691–6. Warner N, Nu´n˜ez G. MyD88: a critical adaptor protein in innate immunity signal transduction. J Immunol 2013;190:3–4. Lawley TD, Clare S, Walker AW, Goulding D, Stabler RA, Croucher N, Mastroeni P, Scott P, Raisen C, Mottram L. Antibiotic treatment of Clostridium difficile carrier mice triggers a supershedder state, spore-mediated transmission, and severe disease in immunocompromised hosts. Infect Immun 2009;77:3661– 3669. Hosoi T, Yokoyama S, Matsuo S, Akira S, Ozawa K. Myeloid differentiation factor 88 (MyD88)-deficiency increases risk of diabetes in mice. PLoS One 2010;5:e12537. Ryan A, Lynch M, Smith SM, Amu S, Nel HJ, McCoy CE, Dowling JK, Draper E, O’Reilly V, McCarthy C. A role for TLR4 in Clostridium difficile infection and the recognition of surface layer proteins. PLoS Pathog 2011;7:e1002076. Dasu MR, Devaraj S, Park S, Jialal I. Increased toll-like receptor (TLR) activation and TLR ligands in recently diagnosed type 2 diabetic subjects. Diabetes Care 2010;33:861–8. Merrigan MM, Sambol SP, Johnson S, Gerding DN. Prevention of fatal Clostridium difficile—associated disease during continuous administration of clindamycin in hamsters. J Infect Dis 2003;188:1922–7. Sambol SP, Merrigan MM, Tang JK, Johnson S, Gerding DN. Colonization for the prevention of Clostridium difficile disease in hamsters. J Infect Dis 2002;186:1781–9. Chang JY, Antonopoulos DA, Kalra A, Tonelli A, Khalife WT, Schmidt TM, Young VB. Decreased diversity of the fecal microbiome in recurrent Clostridium difficileassociated diarrhea. J Infect Dis 2008;197:435–8. Larsen N, Vogensen FK, van den Berg FWJ, Nielsen DS, Andreasen AS, Pedersen BK, Al-Soud WA, Sørensen SJ, Hansen LH, Jakobsen M. Gut microbiota in human adults with type 2 diabetes differs from non-diabetic adults. PLoS One 2010;5:e9085. Wu X, Ma C, Han L, Nawaz M, Gao F, Zhang X, Yu P, Zhao C, Li L, Zhou A, Wang J, Moore JE, Millar BC, Xu J. Molecular characterisation of the faecal microbiota in patients with type II diabetes. Curr Microbiol 2010;61:69–78. Ley RE, Ba¨ckhed F, Turnbaugh P, Lozupone CA, Knight RD, Gordon JI. Obesity alters gut microbial ecology. Proc Nat Acad Sci USA 2005;102:11070–5. Ananthakrishnan AN. Clostridium difficile infection: epidemiology, risk factors and management. Nat Rev Gastroenterol Hepatol 2010;8:17–26. Vrieze A, Van Nood E, Holleman F, Salojarvi J, Kootte RS, Bartelsman JF, Dallinga-Thie GM, Ackermans MT, Serlie MJ, Oozeer R, Derrien M, Druesne A, Van Hylckama Vlieg JE, Bloks VW, Groen AK, Heilig HG, Zoetendal EG, Stroes ES, de Vos WM, Hoekstra JB, Nieuwdorp M. Transfer of

294

diabetes research and clinical practice 105 (2014) 285–294

intestinal microbiota from lean donors increases insulin sensitivity in individuals with metabolic syndrome. Gastroenterology 2012;143:913–6. e917. [133] Monot M, Boursaux-Eude C, Thibonnier M, Vallenet D, Moszer I, Medigue C, Martin-Verstraete I, Dupuy B. Reannotation of the genome sequence of Clostridium difficile strain 630. J Med Microbiol 2011;60:1193–9.

[134] Stabler RA, He M, Dawson L, Martin M, Valiente E, Corton C, Lawley TD, Sebaihia M, Quail MA, Rose G, Gerding DN, Gibert M, Popoff MR, Parkhill J, Dougan G, Wren BW. Comparative genome and phenotypic analysis of Clostridium difficile 027 strains provides insight into the evolution of a hypervirulent bacterium. Genome Biol 2009;10:R102.