Prevalence and association of macrolide–lincosamide– streptogramin B (MLSB) resistance with resistance to moxifloxacin in Clostridium difficile

Grit Ackermann1,*, Angelika Degner1, Stuart H. Cohen2, Joseph Silva Jr2 and Arne C. Rodloff1

1 Institute for Medical Microbiology and Epidemiology of Infectious Diseases, University of Leipzig, Liebigstrasse 24, 04103 Leipzig, Germany; 2 Department of Internal Medicine, Division of Infectious Diseases and Immunologic Diseases, University of California–Davis, Medical Centre, Sacramento, CA, USA

Received 9 August 2002; returned 10 October 2002; revised 25 November 2002; accepted 4 December 2002


    Abstract
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Clostridium difficile remains the leading cause of nosocomially acquired diarrhoea. C. difficile usually exhibits resistance against ß-lactam antibiotics, whereas susceptibility to other drugs may vary. This study investigated the antimicrobial susceptibility of C. difficile to different antibiotics over a period of time and characterizes molecular mechanisms for resistance. One hundred and seventy-three toxigenic and 19 non-toxigenic C. difficile strains, recovered from patients in two university hospitals in Germany between 1986 and 2001, were investigated for their susceptibility to erythromycin, clindamycin, moxifloxacin, vancomycin and metronidazole employing the Etest. The genetic background for resistance was analysed using PCR and DNA sequencing. All strains were susceptible to vancomycin and metronidazole. Resistance to erythromycin, clindamycin and moxifloxacin was found in 27%, 36% and 12% of the tested strains, respectively. High-level resistance (MIC > 128 mg/L) against erythromycin and clindamycin was detected in 25% of the strains tested. Thirty-four of the macrolide–lincosamide–streptogramin B (MLSB)-resistant strains carried the erythromycin resistance methylase gene. The results indicate an increase in the prevalence of resistance to MLSB and fluoroquinolone antibiotics in C. difficile. Fluoroquinolone resistance is associated with resistance to MLSB antimicrobials.

Keywords: Clostridium difficile, antimicrobials, susceptibility


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Clostridium difficile is the most common nosocomial enteric pathogen causing pseudomembranous colitis, antibiotic-associated colitis and antibiotic-associated diarrhoea. During hospitalization ~20% of patients acquire the organism, and depending on known risk factors like antibiotic treatment, older age and prolonged hospital stay they develop C. difficile-associated diarrhoea (CDAD).1,2 In many patients, colonizing microorganisms in the gastrointestinal tract are exposed to different chemotherapeutic and antimicrobial agents. Development of resistance and spreading of resistance mechanisms within these organisms is of growing interest. A clindamycin-resistant C. difficile strain was found to be responsible for a large outbreak of diarrhoea in four hospitals in the USA. Colonization with clindamycin-resistant strains increased the risk of CDAD.3

Vancomycin and metronidazole are still first-line therapeutics for the treatment of severe CDAD. Several studies have reported elevated MICs for C. difficile and metronidazole.47 However, metronidazole resistance in C. difficile has no clinical relevance and is not responsible for treatment failure. Impaired susceptibility of C. difficile to vancomycin has been reported by Peláez et al.7 Reduced susceptibility against moxifloxacin, a new fluoroquinolone, was shown in clonal and non-clonal isolates recovered from various patients.8,9 A mutation in one of the target enzymes of fluoroquinolones, gyrase A (encoded by the gyrA gene), was found to be associated with this phenotypic resistance.8

Post-transcriptional methylation of 23S ribosomal rRNA confers resistance to macrolide–lincosamide–streptogramin B (MLSB)-type antibiotics.10 Target modification occurs at the level of the ribosomes through the Erm N-methyltransferase that is encoded by the erythromycin resistance methylase gene (erm). Several classes of erm gene have been described.11,12 Mutations in domain V of 23S rRNA also alter the target and confer resistance to MLSB antimicrobials.12 Additionally, enzymes that inactivate macrolides and the presence of multi-component macrolide efflux pumps were found to contribute to erythromycin resistance.10,11

Resistance to clindamycin, erythromycin and tetracycline in C. difficile was reported to be transferred without plasmid-DNA involvement.13 MLSB resistance in C. difficile is encoded by the erythromycin ribosomal methylase gene B [erm(B)] for which a location on a mobilizable non-conjugative element Tn5398 in the chromosome was shown.14,15 Two studies reported the detection of two copies of the erm(B) gene in C. difficile.16,17 High-level resistance against MLSB antimicrobials usually requires an erm gene, whereas efflux-mediated resistance confers low-level resistance.6

Correlation between resistance of C. difficile to antimicrobial agents for which different molecular mechanisms of resistance exist, or the association of antimicrobial resistance with virulence properties, has not been investigated. In the study presented here, we observed increased resistance of clinical strains of C. difficile to different antimicrobial classes. The genotypes of these resistant strains were analysed.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
C. difficile strains

One hundred and ninety-two C. difficile isolates recovered from patients from two university hospitals in Germany were investigated (hospital 1: n = 69; hospital 2: n = 123). Only one isolate per patient was included. The isolates were obtained from patients with symptoms of CDAD (I: 1986–1995, n = 50; II: 1996–2000, n = 82; III: 2001, n = 60).

All strains were grown anaerobically on a selective medium (cycloserine–cefoxitin–fructose agar) at 37°C for 48 h.18 For antimicrobial susceptibility testing, strains were grown on Columbia agar supplemented with sheep erythrocytes, haemin and vitamin K. Strains were maintained in cooked meat broth (Becton Dickinson Microbiology Systems Europe, Heidelberg, Germany).

Toxin detection

Diagnostic procedures in hospital 2 encompassed both toxin detection in stool samples employing toxin A/B-ELISA (r-biopharm, Darmstadt, Germany) and culture. From hospital 1 only the C. difficile strains that were cultured from stool samples were available.

All strains were analysed by PCR for tcdA/tcdB gene sequences as described elsewhere.19

Antimicrobial susceptibility testing

MICs of erythromycin, clindamycin, moxifloxacin, vancomycin and metronidazole were determined by Etest (AB Biodisk, Solna, Sweden). The Etest was carried out by inoculating the surface of pre-reduced Columbia agar plates containing vitamin K1, haemin and 5% defibrinated sheep red blood cells with a 1 McFarland standard-matched inoculum. The inoculation was performed with cotton-tipped swabs that were streaked three times, rotating the plate ~90 degrees each time to ensure an even distribution of inoculum. Etest strips were used according to the manufacturer’s instructions. Bacteroides fragilis ATCC 25285 and Staphylococcus aureus ATCC 29123 were used as reference strains.

According to DIN recommendations,20 resistance was defined as follows: erythromycin >= 8 mg/L; clindamycin >= 8 mg/L; vancomycin >= 16 mg/L; metronidazole >= 8 mg/L; moxifloxacin >= 4 mg/L.

DNA extraction techniques

An isolated colony from each strain was transferred with an inoculating loop into a 0.6 mL tube containing 100 µL of sterile water, and was boiled at 100°C for 10 min followed by centrifugation at low speed (3000g) to remove cell debris. The DNA in the supernatant was used for amplification reactions. Highly pure DNA was prepared with the QiaAmp DNA Mini Kit (Qiagen, Hilden, Germany).

For PCR ribotyping, template nucleic acid was prepared by re-suspension of cells in a 5% solution of Chelex 100 (Bio-Rad, Hercules, CA, USA) and boiling for 12 min. Cellular debris was removed by centrifugation (15 000g for 10 min) and the supernatant used for PCR.21

PCR methods

Toxin A and B gene fragments were amplified as published elsewhere.19 A 247 bp fragment of the gyrA gene was amplified using specific primers CdgaV (5'-TTTAAAGCCAGTTCATAG-3') and CdgaR (5'-GAACCAAAGTTACCATG-3').8 Thirty cycles of the following PCR profile were run: 30 s at 95°C, 30 s at 48°C and 60 s at 72°C. The resulting DNA fragments were purified with Amicon Microcon-PCR Centrifugal Filter Devices (Millipore Corporation, Bedford, MA, USA). Complementary strands were sequenced on an ABI310 sequencer (Perkin Elmer Applied Biosystems, Foster City, CA, USA) using either PCR primer. erm(B) sequences were amplified using primers 2980 (5'-AATAAGTAAACAGGTAACGTT-3') and 2981 (5'-GCTCCTTGGAAGCTGTCAGTAG-3').3

PCR ribotyping

PCR ribotyping was carried out using two primers (16S: 5'-CTGGGGTGAAGTCGTAACAAGG-3' and 23S: 5'-GCGCCCTTTGTAGCTTGACC-3').21 Amplification products were concentrated to a final volume of 50 µL using a Speedvac. Electrophoresis (400 mA, 80 V) was run in 1.5% Metaphor agarose (BMA, Rockland, ME, USA) for ~4 h. Products were visualized by staining the gel in ethidium bromide.


    Results
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Toxigenicity of clinical isolates

Ninety per cent (173/192) of the C. difficile strains investigated in this study were toxigenic, as determined either by toxin A/B-ELISA or tcdA/tcdB gene PCR. Four strains were found to be toxin A gene-positive/toxin B-gene negative. PCR for toxin genes was carried out for gene fragments only, thus the possibility of deletions in the toxin gene is not known.

Antimicrobial susceptibility

MICs of erythromycin, clindamycin, moxifloxacin, vancomycin and metronidazole for the 192 C. difficile strains are shown in Table 1. All strains were susceptible to metronidazole and vancomycin. Resistance against erythromycin, clindamycin and moxifloxacin was found in 27%, 36% and 12% of the strains tested, respectively. Considering specific time frames, an increasing number of resistant strains was observed (Figure 1). High-level resistance (MIC > 128 mg/L) against erythromycin and clindamycin was found in 26% and 25% of the strains tested, respectively. Moxifloxacin resistance was found almost always (in 87%) in strains resistant to erythromycin and clindamycin (95% confidence interval 68–97%).


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Table 1.  Cumulative percentage of C. difficile strains (n = 192) inhibited by five antimicrobials
 


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Figure 1. Resistance against clindamycin, erythromycin and moxifloxacin determined in 192 C. difficile strains isolated from 1986 to 2001. According to DIN recommendations, resistance was defined as follows: erythromycin >= 8 mg/L; clindamycin >= 8 mg/L; moxifloxacin >= 4 mg/L;20 95% confidence intervals are shown.

 
Genotyping

The erm(B) gene was detected in 34 of 83 MLSB-resistant strains (41%); all of the erm(B)-carrying organisms were high-level MLSB resistant. The percentage of erm(B)-negative strains was found to be very high. Thus, PCR from erm(B)-negative strains was repeated using high quality DNA. In order to identify erm gene sequences harbouring sequence changes, Southern blotting followed by hybridization using digoxigenin-labelled probes was carried out. With both methods, no erm gene sequences were found. Nineteen of the 24 moxifloxacin-resistant strains harboured a gyrA mutation and five isolates were found to have the wild-type sequence (Table 2).


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Table 2.  Genotype of C. difficile strains resistant to moxifloxacin (MXF-R) and erythromycin/clindamycin (ERY/CLI-R)
 
Using PCR ribotyping, ~80 ribotypes were identified. A PCR ribotype was defined as the existence of clearly discernible, reproducible differences in PCR ribotype pattern from those of the other existing types.21 There was no association between PCR ribotyping group and antimicrobial susceptibility. Antimicrobial resistance was not caused by clonal expression.


    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
In 34 of 83 MLSB-resistant strains, an erm(B) gene could be detected as the basis for the resistance to erythromycin and/or clindamycin. Resistance to erythromycin and clindamycin increased in parallel from 1986 to 2001 (30->50% and 18->35%, respectively), as could be expected since resistance to both antibiotics is mediated by erm(B). The increase is not statistically significant (P value comparing numbers of periods I and II: 0.07, and periods II and III: 0.051), but shows a trend. Moxifloxacin resistance was almost always detected together with resistance to erythromycin and/or clindamycin. Of 24 C. difficile isolates resistant to moxifloxacin, only three were susceptible to erythromycin and clindamycin (12.5%). One of these 21 strains was found to be resistant to moxifloxacin and clindamycin/erythromycin and was gyrA-/erm(B)-negative.

Forty-nine of 83 MLSB-resistant C. difficile strains were erm(B) gene-negative. Resistance in those strains could be due to mutations within the target sequence in the 23S rRNA, could be efflux mediated or caused by inactivating enzymes. Mutations in domain V of 23S rRNA have not yet been described for C. difficile. Low-level MLSB resistance is known to be efflux mediated. A multicomponent efflux pump used for transportation of different classes of chemotherapeutics into and out of the cell could be involved in the aetiology of MLSB and fluoroquinolone resistance. However, 15 of the 49 erm(B)-negative C. difficile strains showed high-level resistance to clindamycin and/or erythromycin. Efflux genes or enzymes that inactivate antibiotics have not yet been described for C. difficile. erm genes have been isolated and characterized from diverse sources.12

Probably other erm genes confer MLSB resistance in C. difficile, which are not detectable with the primers used in this study. It is also possible that modifications of the antimicrobial target due to mutations could have caused resistance to erythromycin and clindamycin. Efflux mechanisms may also play an important role in resistance to fluoroquinolones. Resistance of C. difficile against moxifloxacin was rare 7 years ago (2%), but has reached alarming numbers in recent years (20%). In five of the 24 moxifloxacin-resistant strains, no mutation in the topoisomerase II gene (gyrA) was detected. Neither the sequence of topoisomerase IV, the second target enzyme for fluoroquinolone antimicrobials, nor efflux-mediated resistance mechanisms of C. difficile have been described in the literature. As shown in earlier studies, in vitro selected moxifloxacin-resistant strains and clinical isolates, high-level resistant to moxifloxacin, can contain the wild-type sequence.8

erm(B) class genes have been detected in a wide variety of bacterial genera, indicating their potential for intergenic transfer.12,14 Mutations in topoisomerase genes leading to resistance have been described for topoisomerases II and IV of Gram-positive and -negative bacteria.22,23 Multiple drug resistance is a well known phenomenon in bacteria such as Mycobacterium tuberculosis, methicillin-resistant S. aureus and vancomycin-resistant enterococci.

C. difficile colonizes the gastrointestinal tract of humans. Antimicrobials excreted via the gastrointestinal tract exert selective pressure on the colonizing microflora. Transfer of resistance genes within a pool of multiple drug-resistant bacteria seems to be a likely process. The donors of antimicrobial resistance genes in the gut may rarely be identified because of their lack of pathogenicity. The acceptance and accumulation of multiple drug resistance mechanisms by pathogenic bacteria limit therapeutic options and management of severe infections. Restrictive and intelligent use of antimicrobial agents can enhance their lifetime as well as delay the development of bacterial resistance.


    Acknowledgements
 
We thank Yajarayma J. Tang-Feldman for critical reading of the manuscript.


    Footnotes
 
* Corresponding author. Tel: +49-341-971-5200; Fax: +49-341-971-5209; E-mail: ackermg{at}medizin.uni-leipzig.de Back


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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
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