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Prevalence and association of PCR ribotypes of Clostridium difficile isolated from symptomatic patients from Warsaw with macrolide-lincosamide-streptogramin B (MLS_B) type resistance

Hanna Pituch¹, Jon S. Brazier², Piotr Obuch-Woszczatyski¹, Dorota Wultaska¹, Felicja Meisel-Mikoajczyk¹ and Mirosaw uczak¹

¹ Department of Medical Microbiology, Medical University of Warsaw, 5 Chaubinski Street, 02-004 Warsaw, Poland

² Anaerobe Reference Laboratory, National Public Health Service Microbiology Cardiff, University Hospital of Wales, Cardiff, UK

Correspondence
Hanna Pituch
hanna.pituch{at}ib.amwaw.edu.pl

Received 24 June 2005
Accepted 9 October 2005

Abbreviations: CDAD, Clostridium difficile-associated diarrhoea; CPE, cytopathic effect; MLS_B, macrolide-lincosamide-streptogramin B.

	INTRODUCTION

TOP INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

 
Clostridium difficile is the main aetiological agent of antibiotic-associated diarrhoea, antibiotic-associated colitis and pseudomembranous colitis. The pathogenicity of this bacterium is determined by the production of two major toxins: enterotoxin A (TcdA) (A) and cytotoxin (TcdB) (B) (Borriello, 1998). However, C. difficile strains producing only toxin B (A^–B⁺) can be isolated from cases of C. difficile-associated diarrhoea (CDAD) throughout the world (Brazier et al., 1999; Pituch et al., 2001, 2003; Kuijper et al., 2001; Alfa et al., 2000; Barbut et al., 2002; Johnson et al., 2003).

The application of different typing methods has revealed that C. difficile is a heterogeneous species. Workers at the Anaerobe Reference Laboratory (ARL) in Cardiff developed a modification of the PCR-ribotyping method, based on the polymorphism in the 16S–23S rDNA intergenic region spacer, for the routine typing of C. difficile (O'Neill et al., 1996). Every bacterial strain contains several rRNA operons, and there is a strain-dependent variation in the size and number of the 16S–23S intergenic spacer regions. Macrolide-lincosamide-streptogramin B (MLS_B)-resistant C. difficile strains have been demonstrated to be the cause of epidemics of CDAD in different countries (Johnson et al., 1999; Kuijper et al., 2001; Pituch et al., 2003). High-level resistance to clindamycin and erythromycin in C. difficile is encoded by the ermB gene (the erythromycin ribosomal methylase gene B), and this determinant is located on a conjugative transposon called Tn5398 (Wust & Hardegger, 1993; Mullany et al., 1995; Farrow et al., 2001). The ermB MLS_B-resistance determinant from C. difficile 630 contains two copies of an erm (B) gene (Farrow et al., 2000). Spigaglia & Mastrantonio (2004) described five phenotypic classes (EC-a to EC-e), with characteristic susceptibility/resistance patterns to erythromycin and clindamycin, among the C. difficile strains.

The aim of this study was to determine the distribution of PCR ribotypes of C. difficile circulating in the University Hospital in Warsaw, Poland and to establish the relationship with MLS_B-type resistance.

	METHODS

TOP INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

 
C. difficile isolates. A total of 785 liquid stool samples was collected from patients for whom clinicians specifically requested investigations for C. difficile toxin, between January 2002 and December 2003. The samples originated from patients hospitalized in different wards of University Hospital in Warsaw. From these faecal samples, 79 C. difficile isolates were obtained from different wards: transplant (n=36), surgery (n=16), internal medicine (n=16), orthopaedics (n=6), intensive care (n=2), dermatology (n=1), gynaecology (n=1), urology (n=1). Three reference strains were included in this study as controls, namely, toxigenic C. difficile VPI 10463 (A⁺B⁺), a non-toxigenic C. difficile NIHBRRIGS 8050 (A^–B^–) and A^–B⁺ C. difficile GAI 95 601, isolated by H. Kato (Institute of Anaerobic Bacteriology, Gifu University School of Medicine, Gifu, Japan), were used as an internal control for detecting repeating sequences in the tcdA gene.

Culture and identification of C. difficile isolates. Isolation of C. difficile was performed on selective Columbia Agar supplemented with cycloserine-cefoxitin and amphotericin B (CCCA medium; bioMerieux), as described previously (Pituch et al., 2001). Plates were incubated in an anaerobic chamber (Forma Scientific) at 37 °C for 4 days. The isolates were identified as C. difficile by the characteristic morphology of the colonies and horse odour, green-yellow fluorescence under UV light (365 nm), Gram staining and the API 20A biochemical test (bioMérieux).

Toxin detection. A single colony was transferred into brain heart infusion broth (BHI) (Difco) and grown for 48 h. Supernatants were collected by centrifugation at 3000 g for 15 min. TcdA was detected by an immunochromatography assay using anti-toxin A antibody labelled with latex: the C. difficile toxin A test (Oxoid). Additionally, the immunoenzymic assay C. difficile TOX A/B test (TechLab) was used for detection of either TcdA or TcdB toxins, or both. The procedures were carried out according to the manufacturer's instructions. TcdB was detected by a cytotoxicity assay on the McCoy cell line. Tenfold serial dilutions of culture filtrate were added in duplicate to McCoy cells and incubated for 24 h. C. difficile strain VPI 10463 was used as a positive control. The cytopathic effect (CPE) was observed by inverse microscopy. If this CPE could be neutralized by polyclonal antiserum to C. difficile (C. difficile TOX-B Test; TechLab), the test was considered positive.

PCR assays for detection of tcdA and tcdB genes. Crude template DNA was prepared using genomic DNA PREP-PLUS (A & A Biotechnology) according to the manufacturer's instructions. A 630 bp fragment of the tcdA gene and 399 bp fragment of tcdB gene were amplified using specific primer pairs YT28-YT29 and YT17-YT18, respectively, as described previously (Pituch et al., 2003). The cycling conditions for both PCRs were: one predenaturation cycle at 94 °C for 45 s, and 55 °C for 30 s and 70 °C for 45 s, for 35 cycles, as described previously. Deletion in repeating regions of the tcdA gene was detected with the NK9-NKV011 primer pairs, as described previously (Kato et al., 1999). The PCR cycling conditions were 95 °C for 20 s, 60 °C for 2 min for 40 cycles.

PCR assay for detection of binary-toxin genes (cdtA and cdtB). Primers described by Stubbs et al. (1999) were used for amplification of the binary-toxin genes cdtA and cdtB, as described previously (Pituch et al., 2005).

PCR ribotyping. All isolates were typed by the PCR-ribotyping method described by O'Neill et al. (1996) and Stubbs et al. (1999). The oligonucleotide primers used were P3 (5'-CTG GGG TGA AGT CGT AAC AAG G-3') and P4 (5'-GCG CCC TTT GTA GCT TGA CC-3'). Banding patterns were compared with those of the library of PCR ribotypes at the ARL, Cardiff.

Determination of antibiotic susceptibility and ermB gene PCR. MICs of clindamycin and erythromycin were determined by E-test (AB Biodisc) according to the manufacturer's instructions. Cultures were adjusted to an OD₉₅₀ 1 (using a bioMérieux ATB 1550) on the McFarland scale and were streaked to confluence on the surface of Brucella agar plates. Plastic strips with antibiotics were applied and the plates were incubated anaerobically at 37 °C for 48 h. The MIC was measured at the intercept of the inhibition ellipses. According to NCCLS (National Committee for Clinical Laboratory Standards) recommendations, resistance was defined as follows: 8 mg clindamycin l^–1 and 8 mg erythromycin l^–1. A 688 bp fragment of ermB was amplified using specific primer pairs 2980 (5'-AAT AAG TAA ACA GGT AAC GTT-3') and 2981 (5'-GCT CCT TGG AAG C TG TCA GTA G-3') (Johnson et al., 1999). The PCR cycling conditions included 30 cycles of 60 s at 95 °C, 120 s at 55 °C and 180 s at 72 °C.

	RESULTS

TOP INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

 
Toxigenicity of clinical isolates

During the study period, a total of 785 stool samples was investigated for C. difficile. All of these faecal samples were from symptomatic adults hospitalized in different units of the University Hospital in Warsaw. The overall incidence of toxin-positive C. difficile-positive symptomatic patients among those with suspected CDAD was 66 % (526), with individual incidence in the units as follows: transplant 64 % (156), surgery 23 % (22) and internal medicine 57 % (72). From toxin-positive patients, 79 C. difficile isolates were obtained: 36 from transplant patients, 16 from patients hospitalized in the internal medicine unit, 16 from patients hospitalized in the surgery unit, 6 from orthopaedic patients, 2 from patients hospitalized in the intensive care unit, 1 from a urology patient, 1 from a gynaecology patient and 1 from a patient hospitalized in the dermatology unit. These 79 C. difficile isolates were analysed for TcdA and/or TcdB toxins. Among these, 36 were A⁺B⁺, as demonstrated by the C. difficile toxin A test, the TOX A/B test and the TcdB cytotoxicity testing on McCoy cells. Thirty-six isolates were A^–B⁺, because TcdA could not be detected using the commercial latex test for TcdA, but a CPE on the McCoy cell line was observed. The TOX A/B tests gave positive results for all 36 A^–B⁺ strains. The remaining seven C. difficile isolates were A^–B^– because all tests gave negative results. For the 72 isolates designated A⁺B⁺ or A^–B⁺, PCR amplification with primer pairs YT28-YT29 and YT17-YT18 generated products of 630 bp and 399 bp for the tcdA and tcdB genes, respectively. For the seven A^–B^– isolates, PCR with the above primer pairs did not generate a product. PCR for detecting repeating sequences in toxin A with NK9-NKV011 primer pairs for 36 A^–B⁺ strains generated a 700 bp product similar to that obtained for the Japanese GAI 95 601 C. difficile strain, and similar to that obtained for prevalent group of A^–B⁺ strains (serogroup F, toxinotype VIII, ribotype 017). The presence of the cdtA and cdtB genes was tested by PCR for the same set of strains. Both cdtA and cdtB genes were identified in A⁺B⁺ C. difficile isolate no. 2145.

PCR ribotyping

By PCR ribotyping, we distinguished 21 different types among 79 C. difficile isolates as shown in Table 1. Of the 35 C. difficile A⁺B⁺ isolates 15 could be classified into visually distinct ribotypes. One distinct A⁺B⁺CDT⁺ isolate belonged to ribotype 033. All isolates producing only toxin B (A^–B⁺) belonged to ribotype 017. Seven non-toxigenic (A^–B^–) isolates were classified into four ribotypes: three isolates belonged to ribotype 010, two to ribotype 128, one to ribotype 031 and one to ribotype 114. Interestingly, among C. difficile strains isolated from patients hospitalized in the internal unit, 4 A⁺B⁺ isolates belonged to 3 ribotypes, ribotype 023 (1), ribotype 046 (1) and ribotype 070 (2), but 12 A^–B⁺ strains belonged to ribotype 017. Among strains isolated in the surgery unit the 4 A⁺B⁺ isolates belonged to 3 ribotypes, ribotype 046 (1), ribotype 090 (1), ribotype 094 (2), but 11 A^–B⁺ strains belonged to ribotype 017. One non-toxigenic (A^–B^–) strain belonged to ribotype 010.

Three phenotypic classes of patterns of susceptibility/resistance to erythromycin/clindamycin were identified in all strains studied (Table 2). In our laboratory, the EC-a class was characterized by susceptibility to both erythromycin (MIC range 0·023–1·5 mg l^–1) and clindamycin (MIC range 0·023–3·0 mg l^–1), the EC-b class by high level resistance to both erythromycin and clindamycin (MIC256 mg l^–1), and the new class designated by us as EC-f by decreased susceptibility to clindamycin (MIC 4 mg l^–1) and susceptibility to erythromycin (MIC range 0·5–0·75 mg l^–1). The phenotypic EC-a class was composed of 36 C. difficile isolates, the EC-b class of 39 isolates and the EC-f class of 4 isolates.

Of the C. difficile isolates of class EC-a, all were ermB negative. Out of the 37 strains showing the EC-b phenotype, 35 harboured the ermB gene, but 2 strains did not. All four C. difficile strains of class EC-f were ermB gene negative.

	DISCUSSION

TOP INTRODUCTION METHODS RESULTS DISCUSSION REFERENCES

 
The present study describes a comparative analysis of C. difficile strains isolated from symptomatic patients hospitalized in the University Hospital in Warsaw between 2002 and 2003. Our investigations focused on toxigenicity studies, comparative analysis of ribotypes isolated in Poland with the UK ARL PCR-ribotype library, and MLS_B-type resistance. We analysed 79 C. difficile strains isolated over a 2 year period from a large group of patients with antibiotic-associated diarrhoea. Among the strains investigated in this study 44·3 % (35/79) were A⁺B⁺, 1·3 % (1/79) were A⁺B⁺CDT⁺, 45·5 % (36/79) were A^–B⁺ and 8·9 % (7/79) were A^–B^–. Among 33 strains isolated in the period 1999–2001 from patients with CDAD in our hospital, 45 % were A⁺B⁺ but 55 % were A^–B⁺. The findings of the present study confirmed the high prevalence of A^–B⁺ isolates in our institution, as was described previously (for 1999–2001) (Pituch et al., 2003).

The data generated from the present study showed that all 79 C. difficile strains isolated from patients with antibiotic-associated diarrhoea were typable by the PCR-ribotyping method. Interestingly, the 35 C. difficile-positive patients harboured 15 highly diverse PCR ribotypes of A⁺B⁺ strains.

Two distinct toxigenic (A⁺B⁺) clones were found to circulate our hospital (PCR ribotypes 014 and 046). Rotimi et al. (2003) described three different clones belonging to PCR ribotypes 097, 078 and 039, among isolates from patients in hospitals in Kuwait. The most predominant ribotype in a Hungarian survey was PCR ribotype 087 (A⁺B⁺), which accounted for 39 % of all isolates (Urban et al., 2001). In Warsaw, we did not observe these ribotypes that were present in Hungary and Kuwait. Among clinical strains isolated in the University Hospital in Warsaw only one isolate was found that belonged to ribotype 001, which is dominant in the United Kingdom (Stubbs et al., 1999). The most predominant PCR ribotype in the University Hospital in Warsaw (between 2002 and 2003), from a survey of 79 isolates, was ribotype 017 (A^–B⁺), which accounted for 45·5 % of all isolates. During the 24 month period, an outbreak of CDAD cases caused by C. difficile strains ribotype 017 occurred among 12 patients at the internal unit versus 4 patients with CDAD caused by A⁺B⁺ C. difficile strains (ribotypes 023, 046 and 070), and among 16 patients at the surgery unit versus 4 patients with CDAD caused by toxigenic strains (ribotypes 046, 090, 094). C. difficile A^–B⁺ strains isolated between 1999 and 2001 belonged to the Polish ribotype designated ribotype A (Pituch et al., 2003). One C. difficile binary positive strain (no. 2145) belonged to ribotype 033. In our earlier study, the same isolate belonged to Polish ribotype D (Pituch et al., 2005).

In our hospital we observed MLS_B-type resistant variant toxin C. difficile strains that were responsible for many cases of CDAD. In our earlier study we found MLS_B resistance among all analysed C. difficile A^–B⁺ strains and we concluded that C. difficile strains harbouring the ermB gene are significantly associated with CDAD (Pituch et al., 2001, 2003). Van den Berg et al. (2004) analysed 39 C. difficile strains that did not produce toxin A but produced toxin B (A^–B⁺), originating from Canada, the United States, Poland, the United Kingdom, France, Japan and The Netherlands. All Polish A^–B⁺ isolates, designated earlier as Polish ribotype A, belonged to ribotype 017. Clindamycin resistance encoded by the ermB gene was found in 33 of the 39 C. difficile strains. They concluded that clindamycin-resistant C. difficile A^–B⁺ strains of PCR ribotype 017 have a clonal worldwide spread. MLS_B-type resistance is frequently found in C. difficile strains that are resistant to other antibiotics (Ackermann et al., 2003; Barbut et al., 2002). It is interesting because Delmee & Avesani (1988) did not find resistance to clindamycin in toxin variant C. difficile strains (A^–B⁺).

Spigaglia & Mastrantonio (2004) performed a comparative analysis of C. difficile strains belonging to distinct genetic lineages, focusing on PaLoc (pathogenicity locus) analysis and antibiotic resistance. In this survey, five classes of patterns of susceptibility/resistance (EC-a to EC-e) to erythromycin and clindamycin were identified. Most of the recent isolates belonged to EC-d and EC-e classes, although erythromycin resistant in vitro, they did not harbour the ermB gene, and two strains of the EC-d class were resistant to clindamycin only after induction with a subinhibitory concentration of the antibiotic. In our study, the main phenotypic classes of patterns of susceptibility/resistance to erythromycin/clindamycin, for all Polish strains, were identified: EC-a, EC-b (described by Spigaglia & Mastrantonio, 2004) and a new class designated by us as EC-f. We did not observe classes EC-c, EC-d and EC-e in our collection. The analysis of the genetic background of the resistance to clindamycin and erythromycin using PCR, showed the presence of the erythromycin-resistance methylase gene (ermB) in 36 C. difficile strains belonging to the EC-b₁ phenotypic subclass. Interestingly, two MLS_B-resistant C. difficile strains belonging to PCR ribotype 046, from the EC-b₂ phenotypic subclass, were ermB negative. Resistance in those strains could possibly be due to mutations within the target site in the 23S rRNA, or a new mechanism of high resistance. Spigaglia & Mastrantonio (2004) showed that the EC-b phenotypic class in their Italian collection was always associated with the presence of an ermB gene, but Ackermann et al. (2003) have described ermB-negative C. difficile strains with high level resistance to clindamycin and/or erythromycin. Ackermann et al. (2003) suggest it is probable that other erm genes are responsible for this high-level resistance to clindamycin and erythromycin.

In streptococci, as well as in many other Gram-positive bacteria, target-site modification is a common resistance mechanism (Weisblum, 1995). Genes belonging to the ermAM (ermB) gene class, which was founded in Streptococcus pyogenes, are linked with MLS_B resistance. In addition, Seppala et al. (1998) found in S. pyogenes a gene other than ermB that mediated MLS_B resistance called ermTR.

In our collection of C. difficile strains, we identified phenotype EC-f by the decreased susceptibility to clindamycin (MIC 4 mg l^–1) and susceptibility to erythromycin (MIC range 0·5–0·75 mg l^–1), and they were always ermB-gene negative. Spigaglia & Mastrantonio (2004) described that only C. difficile strains with susceptibility or decreased susceptibility to clindamycin were ermB-negative. Efflux-mediated resistance confers only a low-level resistance to antimicrobial agents. Interestingly, among C. difficile strains belonging to ribotype 017 we observed three type clindamycin/erythromycin resistance patterns: EC-a/ermB negative, EC-b₁/ermB positive, EC-f/ermB negative. MLS_B type resistance was always associated with the ermB gene. Clindamycin-resistant C. difficile strains were found to be responsible for a large outbreak of CDAD in four hospitals in the USA. High-level resistance to clindamycin (MIC256 mg l^–1) was present in all 85 epidemic-strain isolates, but in only 7 out of 46 non-epidemic strain isolates. The epidemic strains were also highly resistant to erythromycin (MIC256 mg l^–1) (Johnson et al., 1999). In summary, resistance against clindamycin and erythromycin among Polish A^–B⁺ (ribotype 017) C. difficile strains was very frequent (94 %), but among A⁺B⁺ and A^–B^– strains it was very rare (11 and 3 %, respectively). However, clindamycin resistance among C. difficile strain isolates is not new (Gerding & Johnson, 2001), but further work is needed to elucidate the association of MLS_B resistance with epidemic-spreading toxin variant C. difficile strains.

	ACKNOWLEDGEMENTS

 
This work was supported by Ministry of Scientific Research and Information Technology, grant no. 2 P05D 074 27

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