The Skin Research Centre, Division of Microbiology, School of Biochemistry and Molecular Biology, University of Leeds, Leeds LS2 9JT, UK
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Abstract |
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Introduction |
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The aim of this study was to determine the prevalence of different erythromycin and tetracycline resistance phenotypes in populations of aerobic coryneform bacteria and FURECs from the human axilla and fourth toe cleft. These moist skin sites are the major habitats of these organisms.9 One reason for selecting these particular antibiotics is that we already know that cutaneous populations of staphylococci are commonly resistant to them, as are propionibacteria from treated acne patients.1012 Thus it seemed likely that the other major resident skin bacteria would demonstrate similar resistance. A second reason for looking at erythromycin resistance was to detect novel phenotypes that were incompatible with the possession of either macrolidelincosamidestreptogramin type B (MLS) or macrolidestreptogramin type B (MS) resistance determinants (erm genes or msrA).
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Materials and methods |
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Aerobic coryneform bacteria and FURECs were isolated from either the fourth toe cleft of the right foot or from both axillae of 28 healthy volunteers (22 male, six female; age range 2146 years), none of whom were receiving antibiotic therapy at the time of sampling. Only three volunteers were sampled from both sites, to minimize the chance of re-isolating the same strains. Samples from the fourth toe cleft were taken, using sterile cotton-tipped swabs moistened in full-strength wash fluid, from the base of the cleft and along the sides of the toes forming the cleft.13 Axillary bacteria were collected using the detergent scrub technique of Williamson & Kligman.13 Volunteers had been instructed to stop using deodorants for 1 week before sample collection.
Growth media
Total viable counts were obtained by plating 0.1 mL aliquots of decimal dilutions of wash fluid on to coryneform agar as described by Leeming et al.,14 with the inclusion of furazolidone 6 mg/L to inhibit the growth of staphylococci.15 The same medium with the addition of either erythromycin 5 mg/L or tetracycline 10 mg/L was used to obtain differential counts of resistant bacteria.11 Plates were examined after 48 h incubation at 37°C followed by 48 h at room temperature to enhance pigmentation and make it easier to discriminate colony types. All distinct colony types from the selective media were Gram-stained to differentiate Gram-positive cocci and Gram-positive pleomorphic rods. Isolates of different morphology were excluded. Following primary isolation, strains were purified and maintained on either Direct Sensitest Agar (DST) for Gram-positive cocci or DST plus 5% sterile defibrinated horse blood (E&O Laboratories, Bonnybridge, UK) and 0.1% Tween-80 (DST-BT) for coryneforms. Strains were stored in liquid nitrogen.
Gram-positive cocci were typed using API ID 32 Staph strips (bioMérieux, Basingstoke, UK) and coryneforms using API Coryne strips (bioMérieux). In addition all isolates were screened for resistance to mupirocin using 5 µg discs.
Antibiotic susceptibility and MICs
The antibiogram of each isolate was determined by disc testing on DST or DST-BT as appropriate using 20 mL volumes of medium in 90 mm diameter Petri dishes. The antibiotics and concentrations used are shown in Table I. Micrococcus luteus NCTC 9341 and Corynebacterium glutamicum R163 were used as controls. Erythromycin-resistant strains were further tested for erythromycin-inducible resistance to pristinamycin IA, spiramycin and clindamycin. Blunting of the inhibition zone proximal to the erythromycin disc was taken as evidence of inducibility. A minority of isolates of both FURECs and coryneforms which gave small zone diameters (
15 mm) or grew up to penicillin discs were qualitatively screened for ß-lactamase production using the chromogenic substrate, nitrocefin.
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Clindamycin and spiramycin were dissolved and diluted in distilled water. Erythromycin and azithromycin were dissolved in ethanol and diluted in distilled water. Pristinamycin was dissolved in dimethylsulphoxide, and furazolidone in acetone.
Media components, chemicals and testing kits
All growth media and nitrocefin were obtained from Oxoid, Basingstoke, UK; all other chemicals and antibiotics were from SigmaAldrich, Poole, UK with the exceptions of pristinamycin IA and azithromycin which were gifts from RhônePoulenc Rorer (De Vitry-Alfortville, France) and Pfizer (Sandwich, Kent), respectively. Antibiotic-impregnated discs were from Mast Laboratories, Bootle, UK.
Data analysis
Within-group differences in median bacterial counts were determined using Wilcoxon's matched pairs. Between-group differences in the proportion of resistant bacteria were determined using the MannWhitney U test. All computations were performed on release 8.0 of Minitab.
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Results |
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Discussion |
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Although breakpoint data are not given for cutaneous aerobic coryneforms and FURECs in the recent NCCLS guidelines,17 comparison of data given for other organisms, e.g. Staphylococcus spp., Streptococcus spp. and Enterococcus spp., with the zone sizes used in this study to assign co-resistance shows that the criteria used to define resistance were rigorous (see Table V). Analysis of co-resistance adds an extra dimension of complexity to the picture outlined above. Although all the antibiotic resistance identified could be traced either to agents used topically or to antibiotics that reach intact skin, the resistance profiles were highly dependent upon bacterial type. For example, the most common co-resistance in the cutaneous aerobic coryneforms was to fusidic acid and chloramphenicol, antibiotics that are used topically. The resistance pattern in the FURECs differed from that in both the CNS10 and the cutaneous propionibacteria,16 with a higher prevalence of resistance to chloramphenicol and trimethoprim. Notably, the most common co-resistance in FURECs was to the fluoroquinolone, norfloxacin. In the other groups of skin bacteria, the aerobic Propionibacterium spp. and the CNS, there is a paucity of information on the prevalence of fluoroquinolone resistance in untreated subjects. Further studies are also required to quantify the effects of fluoroquinolone treatment on the acquisition of fluoroquinolone resistance by propionibacteria, aerobic coryneforms and FURECs, although it has been shown that the cutaneous CNS acquire resistance to ciprofloxacin rapidly as a result of excretion of the drug in sweat.18 This route of delivery of quinolones to the skin surface may be significant in that FURECs preferentially colonize moist skin surface sites.9 Unfortunately, it was not possible to discriminate between sensitivity and resistance to penicillins in either cutaneous aerobic coryneforms or FURECs. Measurement of zone sizes of all the sensitive and resistant strains included in this study failed to reveal a bimodal distribution to distinguish the two populations. A lack of breakpoint data for coryneform suseptibility to ß-lactams has also been noted by other workers19 and this, together with the observation that none of the isolates with smaller zones of inhibition produced penicillinase, may point to an alternative resistance mechanism in these isolates. One possibility could be a succession of mutations leading to steadily increasing resistance with no clear distinction between sensitive and resistant strains.
The six tetracycline-resistant FURECs (MICs 864 mg/L) were sensitive to the other tetracyclines, doxycycline and minocycline. In contrast, the aerobic cutaneous coryneforms fell into two groups: those that were resistant to tetracycline alone (modal MIC 16 mg/L), and those (comprising nine strains) that were resistant to all three tetracyclines, for which the modal MIC of tetracycline was much higher (256 mg/L). Resistance to tetracycline alone would suggest carriage of either determinants encoding tetracycline efflux systems of the major facilitator type20 or, as recently reported in Corynebacterium striatum, an ABC transporter mediating drug efflux encoded by tetAB.21 Alternatively, coryneform resistance to all three tetracyclines appears to mirror the phenotype identified in staphylococci mediated via ribosomal protection in which TetO/TetM encodes a homologue of elongation factor G.22
Analysis of the MLS resistance phenotypes suggested the presence of several different antibiotic resistance mechanisms, some of which may be novel. Most cutaneous aerobic coryneforms and a number of FURECs showed classic MLS resistance, easily explained either by expression of inducible/constitutive erythromycin ribosomal methylase (erm) genes23 or by a mutation at base 2058 in the 23S ribosomal RNA.24 However, some isolates exhibited unusual phenotypes which could not be placed in these categories and are not compatible with other known erythromycin resistance mechanisms. Examples of these included (i) the group A FURECs which were erythromycin resistant, sensitive to clindamycin and pristinamycin IA and weakly inducibly resistant to spiramycin (the majority of erythromycin-resistant FURECs fell into this group); (ii) the group B1 aerobic cutaneous coryneforms which expressed low-level resistance to erythromycin and azithromycin, but were sensitive to pristinamycin IA, clindamycin and/or spiramycin; and (iii) the group B2 aerobic cutaneous coryneforms which expressed low-level resistance to erythromycin, spiramycin and clindamycin and usually to azithromycin and pristinamycin IA. Resistance to the MLS antibiotics in the latter two groups was non-inducible and, because of the low level of some of the resistance identified, it was found that MIC determinations provided a more reliable indicator of resistance phenotype than the use of disc tests. It is likely that genetic analysis of these strains will identify previously unreported antibiotic resistance genes and that this approach would be facilitated by the development of genetic technology using C. glutamicum.21
There was strong evidence for a link between tetracycline and erythromycin resistance in the aerobic cutaneous coryneforms. In both axilla and toe cleft the vast majority of these bacteria were sensitive to erythromycin. Erythromycin-resistant strains represented <10% of the median count in the toe cleft and <1% in the axilla (Table II). In the absence of linkage, a majority of tetracycline-resistant strains would, therefore, be expected to be erythromycin sensitive. Only seven isolates were tetracycline resistant without also being erythromycin resistant (Table IV
). This undoubtedly reflects both antibiotic usage as well as genetic linkage of the resistance determinants themselves. For example, in C. striatum, the plasmid pTP10, which carries the tetracycline resistance determinant tetAB, also carries genes encoding resistance to kanamycin, erythromycin and chloramphenicol.21 There are clearly many examples of linked antibiotic resistance genes in cutaneous CNS.25 Interestingly, mobile genetic elements have yet to be identified in the cutaneous propionibacteria and resistance to both erythromycin and tetracycline has been shown to be caused by mutations in genes encoding ribosomal RNA.24,26
In addition to the short-term use of antibiotics for the treatment of acute infections, the long-term use of antibiotics in dermatology, especially for the treatment of acne vulgaris, represents a significant source of selective pressure for the evolution of antibiotic-resistant organisms.16 The results of this study show that the axilla and toe cleft of untreated subjects harbour a significant reservoir of G+C-rich antibiotic-resistant bacteria. In addition many of these express novel antibiotic resistance phenotypes. It is certain that during antibiotic treatment the proportion of antibiotic-resistant cutaneous aerobic coryneforms and FURECs will increase significantly. Clearly they share a common niche with the G+C-rich cutaneous propionibacteria and the A+T-rich CNS. Thus it is likely that, under conditions of prolonged selection, gene exchange between these groups of bacteria will provide a significant impetus for the further evolution and spread of antibiotic resistance among cutaneous bacteria.
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Acknowledgments |
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Notes |
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References |
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2 . Larson, E. L., McGinley, K. J., Leyden, J. J., Cooley, M. E. & Talbot, G. H. (1986). Skin colonisation with antibiotic-resistant (JK group) and antibiotic-sensitive lipophilic diphtheroids in hospitalised and normal adults. Journal of Infectious Diseases 153, 7016.[ISI][Medline]
3 . Leyden, J. J. & McGinley, K. J. (1993). Coryneform bacteria. In The Skin Microflora and Microbial Skin Disease, (Noble, W. C., Ed.), pp. 10217. Cambridge University Press.
4 . Roth, R. R. & James, W. D. (1989). Microbiology of the skin: resident flora, ecology, infection. Journal of the American Academy of Dermatology 20, 36790.[ISI][Medline]
5 . Stackebrandt, E., Koch, C., Gvozdiak, O. & Schumann, P. (1995). Taxonomic dissection of the genus Micrococcus: Kocuria gen. nov., Nesterenkonia gen. nov., Kytococcus gen. nov., Dermacoccus gen. nov., and Micrococcus Cohn 1872 gen. emend. International Journal of Systematic Bacteriology 45, 68292.[Abstract]
6 . Roberts, M. C., Leonard, R. B., Briselden, A., Schoenknecht, F. D. & Coyle, M. B. (1992). Characterisation of antibiotic-resistant Corynebacterium striatum strains. Journal of Antimicrobial Chemotherapy 30, 46374.[Abstract]
7 . Soriano, F., Zapardiel J. & Nieto, E. (1995). Antimicrobial susceptibilities of Corynebacterium species and other non-spore-forming gram-positive bacilli to 18 antimicrobial agents. Antimicrobial Agents and Chemotherapy 39, 20814.[Abstract]
8 . von Eiff, C., Herrmann, M. & Peters, G. (1995). Antimicrobial susceptibilities of Stomatococcus mucilaginosus and of Micrococcus spp. Antimicrobial Agents and Chemotherapy 39, 26870.[Abstract]
9 . Noble, W. C. (1981). In Microbiology of Human Skin, (Rook, A., Ed.). Lloyd Luke, London.
10 . Cove, J. H., Eady, E. A. & Cunliffe, W. J. (1990). Skin carriage of antibiotic-resistant coagulase-negative staphylococci in untreated subjects. Journal of Antimicrobial Chemotherapy 25, 45969.[Abstract]
11 . Eady, E. A., Cove, J. H., Blake, J., Holland, K. T. & Cunliffe, W. J. (1988). Recalcitrant acne vulgaris. Clinical, biochemical and microbiological investigation of patients not responding to antibiotic treatment. British Journal of Dermatology 118, 41523.[ISI][Medline]
12 . Leyden, J. J., McGinley, K. J., Cavalieri, S., Webster, G. F., Mills, O. H. & Kligman, A. M. (1983). Propionibacterium acnes resistance to antibiotics in acne patients. Journal of the American Academy of Dermatology 8, 415.[ISI][Medline]
13 . Williamson, P. & Kligman, A. M. (1965). A new method for the quantitative investigation of cutaneous bacteria. Journal of Investigative Dermatology 45, 498503.[ISI][Medline]
14 . Leeming, J. P., Holland, K. T. & Cunliffe, W. J. (1984). The microbial ecology of pilosebaceous units isolated from human skin. Journal of General Microbiology 130, 8037.[ISI][Medline]
15 . Marshall, J. (1988). The microbial ecology of the human foot. Ph.D. Thesis, Department of Microbiology, University of Leeds, UK.
16 . Eady, E. A. (1998). Bacterial resistance in acne. Dermatology 196, 5966.[ISI][Medline]
17 . National Committee for Clinical Laboratory Standards. (1997). Performance Standards for Antimicrobial Susceptibility TestingEighth Informational Supplement: Approved Standard M100-S8. NCCLS, Wayne, PA.
18 . Høiby, N., Jarlov, J. O., Kemp, M., Tvede, M., Bangsborg, J. M., Kjerulf, A. et al.(1997). Excretion of ciprofloxacin in sweat and multiresistant Staphylococcus epidermidis. Lancet 349, 1679.[ISI][Medline]
19 . Funke, G., Pünter, V. & von Graevenitz, A. (1996). Antimicrobial susceptibility patterns of some recently established coryneform bacteria. Antimicrobial Agents and Chemotherapy 40, 28748.[Abstract]
20 . Levy, S. B. (1992). Active efflux mechanisms for antimicrobial resistance. Antimicrobial Agents and Chemotherapy 36, 695703.[ISI][Medline]
21 . Tauch, A., Krieft, S., Pühler, A. & Kalinowski, J. (1999). The tetAB genes of the Corynebacterium striatum R-plasmid pTP10 encode an ABC transporter and confer tetracycline, oxytetracycline and oxacillin resistance in Corynebacterium glutamicum. FEMS Microbiology Letters 173, 2039.[ISI][Medline]
22 . Burdett, V. (1996). Tet(M)-promoted release of tetracycline from ribosomes is GTP dependent. Journal of Bacteriology 178, 324651.[Abstract]
23 . Weisblum, B. (1985). Inducible resistance to macrolides, lincosamides and streptogramin type B antibiotics: the resistance phenotype, its biological diversity, and structural elements that regulate expressiona review. Journal of Antimicrobial Chemotherapy 16, Suppl. A, 6390.[ISI][Medline]
24 . Ross, J. I., Eady, E. A., Cove, J. H., Jones, C. E., Ratyal, A. H., Miller, Y. W. et al. (1997). Clinical resistance to erythromycin and clindamycin in cutaneous propionibacteria isolated from acne patients is associated with mutations in 23S rRNA. Antimicrobial Agents and Chemotherapy 41, 11625.[Abstract]
25 . Skurray, R. A. & Firth, N. (1997). Molecular evolution of multiply-resistant staphylococci. In Antibiotic Resistance: Origins, Evolution, Selection and Spread, (Chadwick, D. J. & Goode, J., Eds). Ciba Foundation Symposium Vol. 207, pp. 16783. John Wiley, Chichester, UK.
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Ross, J. I., Eady, E. A., Cove, J. H. & Cunliffe, W. J. (1998). 16S rRNA mutation associated with tetracycline resistance in a gram-positive bacterium. Antimicrobial Agents and Chemotherapy 42, 17025.
Received 3 August 1999; returned 20 December 1999; revised 31 January 2000; accepted 24 February 2000