JAC Advance Access originally published online on October 17, 2006
Journal of Antimicrobial Chemotherapy 2006 58(6):1160-1167; doi:10.1093/jac/dkl420
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Clindamycin-induced enrichment and long-term persistence of resistant Bacteroides spp. and resistance genes
1 Department of Laboratory Medicine, Karolinska Institute SE-141 86 Stockholm, Sweden 2 Section for Natural Sciences, Södertörn University College SE-141 89 Huddinge, Sweden 3 Department of Microbiology, Swedish University of Agricultural Sciences, SE-750 07 Uppsala, Sweden 4 Medical Products Agency SE-751 03 Uppsala, Sweden
*Corresponding author. Tel: +46-18-17-48-39; Fax: +46-87-11-39-18; E-mail: charlotta.edlund{at}mpa.se
Received 30 March 2006; returned 29 April 2006; revised 25 August 2006; accepted 22 September 2006
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Abstract |
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Objectives: The aim was to study the long-term consequences of 1 week clindamycin administration regarding selection and persistence of resistance, resistance determinants and diversity of the Bacteroides spp. in the intestinal microflora.
Methods: A total of 1306 Bacteroides isolates were collected from constitutively cultured faecal samples during a 2 year period from eight healthy volunteers. The strains were identified by biochemical and genotyping methods. MIC values were determined by the agar dilution method and presence of resistance genes was screened by real-time PCR.
Results: Ecological changes in the intestinal microflora persisting up to 24 months were recorded after a 7 day clindamycin administration to four healthy volunteers. Compared to a control group, not exposed to clindamycin, an enrichment and stabilization of resistant Bacteroides strains and resistance determinants were discovered up to 2 years after clindamycin exposure.
Conclusions: The results indicate that even a short-term antibiotic administration can cause long-term alterations in the commensal microbiota of individual subjects, detectable 2 years after dosing. The recorded selection and persistence of resistant strains and resistance genes, illustrates the importance of increasing our knowledge of the role of the abundant intestinal microbial community as a reservoir for spread of resistance.
Keywords: antimicrobial resistance , intestinal microflora , clindamycin
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Introduction |
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The dynamic and complex intestinal ecosystem is often affected by low and sub-inhibitory drug concentrations when a patient undergoes an antibiotic treatment, regardless of the infectious cause. Such disturbances can create an ideal environment for direct selection of resistant bacteria and enrichment of already present resistance determinants and resistant strains. The intestine may thus act as a reservoir of resistance genes, which can spread via both intra- and inter-species lateral gene transfer to transient and already present strains.1 Clindamycin administration has previously been shown to markedly suppress the anaerobic microbial community2–4 including members of the Bacteroides group, which account for
30% of the intestinal bacterial flora.5 Some bacteria belonging to the Bacteroides genus are opportunists that can cause severe infections and often appear in mixed infections. An increasing rate of resistance to a wide range of agents, including clindamycin, has been demonstrated among Bacteroides spp.6,7 Mechanisms of resistance to clindamycin in anaerobes include mutations in the binding site of the ribosomal target, efflux pump mediated resistance and the erythromycin resistance methylases (erm) genes. Among Bacteroides spp., the most commonly occurring erm genes are erm(F), erm(G) and erm(B), although other genes within the group have been detected in Bacteroides spp. more recently.8,9 Resistance determinants, including the erm genes, are often carried on transmissible elements such as conjugative transposons and plasmids contributing to their spread.8,10,11 Linkage of erm(F) and the tetracycline resistance gene tet(Q) on conjugative transposons has been described, and both genes are frequently found in clinical Bacteroides isolates.8,10 Resistance rates to different antibiotics are increasing in clinical isolates within the Bacteroides group,6,12,13 which limits the therapeutic alternatives in the treatment of anaerobic and mixed infections. Resistance rates of up to 70% against tetracycline and up to 100% against ampicillin are reported.6,14,15 Acquisition of tetracycline resistance is often mediated by the uptake of new resistance genes encoding ribosomal protection proteins, although other mechanisms also are of importance.9,16 Resistance to ß-lactam antibiotics is most frequently mediated by ß-lactamases, commonly encoded by cepA in clinical Bacteroides fragilis isolates, or infrequently by cfxA, which both contribute to high-level resistance.17 Several other ß-lactamases have also been described among the Bacteroides spp.18
The main aim of the present investigation was to study potential ecological alterations in the intestinal microbiota up to 24 months after 1 week clindamycin administration with emphasis on emergence and persistence of resistant strains and resistance genes in intestinal Bacteroides spp.
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Materials and methods |
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Experimental set-up and sampling
Eight healthy volunteers, who had not received any antimicrobial agents in the previous 12 months, were included in the study. Four of the subjects, A–D (3 females and 1 male, aged 31–49 years), received 150 mg of clindamycin capsules (Dalacin; Pharmacia, Stockholm, Sweden) orally four times daily for 7 days. Four subjects, E–H (3 females and 1 male, aged 33–58 years), who did not receive any antibiotics during the study period, were included as a control group, in order to follow the normal variation of the intestinal microflora over time. No specific dietary restrictions were made. None of the volunteers in the control group received any antibiotics during the entire study period of 2 years. Two of the subjects in the clindamycin group were treated for upper respiratory tract infections during the investigation period; Subject A received penicillin for 7 days after 13 months, and Subject D was treated with doxycycline for 7 days at 10 months post-clindamycin administration, i.e. 5 and 8 weeks prior to the subsequent sampling occasions, respectively. The samples from these subjects were closely monitored for any potential impact of this unplanned antibiotic exposure. The study was approved by the regional ethics committee of Karolinska University Hospital Huddinge, Stockholm, Sweden.
Faecal samples were collected prior to (day 0) and at the last day of clindamycin administration (day 7), after 2 weeks (day 21), subsequently every third month for 1 year, and then every sixth month up to 2 years post-administration (3, 6, 9, 12, 18 and 24 months). For the control group, faecal sampling was performed at corresponding intervals. The samples were stored at –70°C until analysis.
Assay of clindamycin concentrations
Concentrations of clindamycin in faeces (mg/kg) were measured in samples from days 0, 7 and 21, by the agar diffusion method using Antibiotic medium No 1 (Difco, Detroit, MI, USA) and Micrococcus luteus ATCC 9341 was included as a control strain.
Culture techniques
The faecal samples were diluted 10-fold to 10–7 and inoculated on kanamycin-vancomycin-blood agar, selective for Bacteroides spp., and on non-selective blood agar media. The plates were incubated anaerobically (BBL GasPak Anaerobic System, Cockeysville, MD, USA) at 37°C for 48 h. Presumed Bacteroides colonies were enumerated and, if possible, 20 colonies of Bacteroides spp. were picked from each sample and stored at –70°C until assayed. In order to get representative isolates since only 20 colonies per sample were analysed, one experienced technician performed all microbiological analyses, and the colonies were chosen to mirror the different morphotypes with respect to their abundance in the cultured sample.
Identification of Bacteroides by phenotypic and genotypic methods
The Bacteroides isolates were morphologically identified to genus level and to species level by biochemical tests.19,20 As controls, the reference strains B. fragilis ATCC 25285, Bacteroides distasonis ATCC 8503, Bacteroides ovatus ATCC 8483, Bacteroides thetaiotaomicron ATCC 29741, Bacteroides uniformis ATCC 8492 and Bacteroides vulgatus ATCC 8482 were included. The clonal identity of clindamycin-resistant Bacteroides isolates (MIC 
8 mg/L) was studied by rep-PCR. This method has previously been shown to be able to distinguish between strains of Bacteroides species.21 The overall results of these analyses will be reported in a separate publication.
Antimicrobial susceptibility testing
The MICs of clindamycin, tetracycline and ampicillin were determined by the agar dilution method as recommended by CLSI, formerly NCCLS.22 Breakpoints for Bacteroides spp. for clindamycin, tetracycline and ampicillin were R 8, R 16 and R 2 mg/L, respectively, as recommended by CLSI.
Real-time PCR
One microlitre of a bacterial colony was suspended in 100 µL of milliQ water, heated at 95°C for 15 min to lyse the cells and centrifuged (10 000 g) for 7 min. The supernatant, free from cellular debris, was stored at –20°C until used.
All isolates were screened for the presence of resistance genes [erm(B), erm(F), erm(G), tet(Q), cepA and cfxA] using real-time PCR. Isolates expressing phenotypic clindamycin resistance, but that were negative for the specific erm genes tested, were also screened by universal erm gene primers E1/E2. Primers were designed using Beacon Designer 4.0 (PREMIER Biosoft International, Palo Alto, CA, USA). PCR amplification was performed using a Light Cycler (Roche Diagnostics, Mannheim, Germany) and Light Cycler-FastStart DNA Master SYBR green I kit (Roche Diagnostics) with 3 or 4 mM of MgCl2 and 0.5 µM of each gene specific primer. All primer sequences used for the real-time PCR are listed in Table 1. PCR programs were as follows: initial denaturation for 600 s, amplification for 25–30 cycles with denaturation at 95°C for 5 s, annealing at 65 or 68°C for 8 s and extension at 72°C for 5 s. The transition rate was 20°C/s. Melting curve analysis was conducted in three segments: 95°C for 60 s, 67°C for 60 s and then increasing to 95°C. The transition rate was 20°C/s for the first and second segment and 0.1°C/s for the last segment. These programs were used for all genes. In addition, for erm(F) an initial profile using six short cycles of amplification, each with a descending annealing temperature (70°C–60°C), was used in order to get the amplified product.
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PCR products representing each of the genes erm(B), erm(F), erm(G), tet(Q), cepA and cfxA, generated by conventional PCR, were sequenced in order to verify the identities of the amplified genes. Prior to sequencing, the PCR products were purified using the QIA quick PCR purification kit (QIAGEN GmbH, Hilden, Germany) to remove free primers and nucleotides. Sequences were obtained using an ABI 310 Genetic Analyser according to the manufacturer (Perkin Elmer, Foster City, CA, USA).
Assay of efflux mediated resistance
Low-level clindamycin-resistant isolates (MIC 8 mg/L) lacking erm genes were tested for the presence of innate efflux pump-mediated resistance mechanisms by performing susceptibility testing on agar media with and without the efflux pump inhibitor reserpine (Sigma Chemical Co., St Louis, MO, USA), at a concentration of 20 mg/L. B. thetaiotaomicron ATCC 29741 and ATCC 29148 were used as control strains.
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Results |
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Clindamycin concentration in faeces
The median clindamycin concentration on day 7 (the last day of administration) was 133 mg/kg (ranging from 89 to 185 mg/kg), while clindamycin concentrations were under the detection limit in all samples collected on days 0 and 21.
Quantitative alterations in B. fragilis group species over time
In the clindamycin-exposed group, median decrease in numbers of Bacteroides cfu was 3 log at day 7, the last day of antibiotic administration, while at day 21 the numbers were normalized to pre-treatment levels. In the control group the levels were more stable and Bacteroides isolates were continuously found at high numbers (median 108 cfu/g faeces) during the entire study period.
Species diversity
The most prevalent species in both groups was B. thetaiotaomicron, although the composition differed between the groups during the study period, and species such as B. vulgatus, B. uniformis, B. ovatus and B. fragilis were also found (Table 2). A decrease in species diversity was recorded after clindamycin administration; while seven disparate species were identified at day 0, only two different species were detected at day 7. In contrast, high species diversity was seen over time in the control group, with at least five different Bacteroides species identified at each sampling occasion, in each subject. The median numbers of species in the subjects over time in the exposed group and control group were 4 (range 2–7) and 6 (range 5–7), respectively. After clindamycin exposure, clindamycin-resistant strains of B. thetaiotaomicron dominated the Bacteroides flora up to 24 months. While clindamycin-resistant B. thetaiotaomicron strains comprised 3% of the isolates pre-exposure, these strains made up 76% of the analysed isolates after 7 days. In the following samples, up to 18 months, the levels of B. thetaiotaomicron isolates resistant to clindamycin remained very high (mean 59%, range 40–98%). After 2 years, 34% of the analysed Bacteroides isolates in the exposed group were still clindamycin-resistant B. thetaiotaomicron. The species composition over time in the non-exposed group is shown in Table 2.
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Resistance levels to clindamycin, tetracycline and ampicillin
The levels of clindamycin resistance over time for both groups are illustrated in Figure 1. In the control group, 700 of the 701 Bacteroides isolates examined were susceptible to clindamycin. In contrast, a distinct increase in resistance to clindamycin was detected, from 7% pre-exposure to 95% resistant isolates directly after clindamycin administration. This was mainly due to selection of single resistant strains in each exposed subject. The overall results of the clonal analyses will be reported in a separate publication. The frequency of clindamycin resistance remained at an increased level at 75% 2 years after clindamycin intake. In three of the four exposed subjects (A, B and D) a similar pattern was revealed with 100% clindamycin-resistant isolates recovered from all samples collected at the end of administration and until 9 months. Notably, from these three exposed individuals, clindamycin-resistant genotypes, selected during the administration, could be identified up to 18 and 24 months post-administration. From two of these subjects, A and B, one clone each, identified by rep-PCR, that were susceptible to clindamycin at day 0 gained resistance within 2 weeks after the administration by acquiring erm genes, which were maintained over the entire study period. In Subject D, a single low-level clindamycin-resistant, erm-negative, B. thetaiotaomicron strain (MIC 8 mg/L), which constituted 5% of the analysed colonies from day 0, came to dominate the Bacteroides community in all but one of the subsequent samples. According to the reserpine assay, a weak efflux pump mediated resistance might be contributing to the resistance in the low-level resistant strain, the MIC decreased by two dilutions in the presence of reserpine, compared to one dilution step in the control strains.
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Resistance to tetracycline and ampicillin was compared for the treated and untreated groups over time. The results showed moderate to high levels of tetracycline resistance among the Bacteroides spp. in both groups at levels of 77% and 47% resistant isolates in the exposed and the control group, respectively (Figure 2). Thus, tetracycline resistance was present both with and without coincident clindamycin resistance, although tetracycline resistance increased with the enrichment of clindamycin resistance, as compared to the control group where the clindamycin resistance was low, <1%, during the study period (Figure 2). Resistance to ampicillin was high in both groups, close to 100% at all time points (range 84–100%).
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Unfortunately, two of the individuals in the clindamycin group received additional antibiotic treatment during the study period. The effects of these 7 day exposures to doxycycline and penicillin V, respectively, were carefully analysed regarding their potential impacts on resistance levels and presence of resistance genes associated with clindamycin, tetracycline and ampicillin resistance. In none of these subjects did the pattern observed in the samples preceding the treatment change significantly in these respects (Table 3).
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Presence of resistance determinants
A selection of isolates carrying erm(F) and erm(G) genes following clindamycin administration was apparent among the exposed subjects, while no isolates carrying the erm(B) gene were detected (Table 3). The erm(F) gene was detected in samples from two exposed subjects and erm(G) in isolates from another subject. None of the clindamycin resistance determinants screened for were found in the control group. The total presence of erm genes in analysed isolates in relation to phenotypic resistance to clindamycin is shown in Figure 2. While high-level clindamycin resistance (MIC > 64 mg/L) in isolates recovered from Subjects A–C with few exceptions was associated with the presence of an erm gene, no erm gene was identified in the low-level resistant isolates (MIC 8–16 mg/L) collected over time from Subject D, even when using universal primers for the erm genes.
Phenotypic tetracycline resistance was associated with tet(Q) genes in 99% of isolates from the exposed group and in 59% of isolates from the control group (Figure 2). The level of isolates carrying tet(Q) rose directly after the clindamycin exposure from 42% to 74% (Table 3). During the following sampling occasions (day 21 up to 24 months) a mean of 80% of the isolates were positive for the gene, thus the levels were still very high after 2 years when tet(Q) was detected in 84% of the isolates. In the control group, the mean total frequency of isolates positive for tet(Q) was 28% during the study period, the levels varied randomly between different sampling occasions (range 5–51%). A co-selection of tet(Q) and erm(F) genes was detected in Bacteroides isolates from Subjects B and C in the exposed group. At day 0 no isolates harboured both genes, 8% harboured erm(F) and 56% harboured tet(Q). At day 7, 55% of the Bacteroides isolates were found to carry both genes [91% were positive for erm(F) and 64% were positive for tet(Q)] and after 9 months 95% were positive for both genes, 100% of the isolates were positive for erm(F) and 95% were positive for tet(Q).
The ampicillin resistance determinant, cepA, was detected at low levels (6–8%) in the Bacteroides isolates, equally distributed over time in both groups, and no co-selection of the cepA gene was seen after clindamycin exposure (Table 3). The gene was exclusively found in B. fragilis species. The cephalosporinase encoding gene, cfxA, was recovered from 5% of the isolates in the exposed group in high-level ampicillin-resistant B. ovatus and B. vulgatus strains (>256 mg/L), mainly originating from one subject.
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Discussion |
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In the present study, faecal specimens from healthy persons were repeatedly sampled for 2 years, and by analysing the predominant Bacteroides species, using different microbiological and molecular techniques, the influence of 7 days of clindamycin administration was monitored regarding resistance, resistance genes and species diversity. While the numbers of Bacteroides cfu returned to pre-treatment levels within a few weeks after clindamycin exposure, dramatic qualitative alterations regarding species diversity and enrichment of resistant strains and resistance genes remained up to 2 years. There was also a decline in the variety of different Bacteroides species isolated, from seven different species pre-exposure to two species detectable at day 7. Resistant B. thetaiotaomicron dominated markedly forming up to 98% of the Bacteroides community in post-exposure samples. These findings were in contrast to the control group, representing a healthy gut flora, where a greater diversity of species was detected throughout the investigation period, which is in line with previous reports,23 and clindamycin resistance levels were low. This apparent prolonged effect of clindamycin on Bacteroides diversity may have an impact on the maintenance of a healthy intestinal homeostasis. It is well known that clindamycin treatment is associated with an increased burden of Clostridium difficile infections and colonization by vancomycin-resistant enterococci, features which are both consequences of disturbed colonization resistance.24–26 In spite of the short-term administration, a significantly increased prevalence of clindamycin- and tetracycline-resistant isolates persisted for at least 2 years. Resistance levels for clindamycin among analysed isolates exceeded 50% in all post-exposure samples, compared to 0–1% in samples from untreated subjects. This finding was unexpected and discloses the huge and deleterious consequences that may result from a common course of antibiotic treatment, routinely prescribed for uncomplicated conditions.
Enriched resistant strains were stably persistent in three of the exposed subjects for 18–24 months, indicating an adaptation and unaffected or restored fitness, although these strains all harboured an erm gene, either erm(F) or erm(G). The erm(F) and erm(G) genes are often associated with the Bacteroides genera and the frequency of clinical erm-positive strains have increased over the last few decades.8 While a significant co-selection between erm(F) and tet(Q) was recorded in the present study, no correlation could be detected between the levels of cepA or cfxA and the other resistance genes studied.
The unplanned administration of penicillin and doxycycline did not seem to cause any detectable impact on the intestinal microflora of these patients. According to several studies penicillin is associated with no or only minor disturbances in the normal microflora, while doxycycline, despite not significantly suppressing the numbers of intestinal bacteria, has been shown to cause short-term selection of tetracycline-resistant strains.27,28
To our knowledge, this is the first study where the dynamics in the intestinal microflora of antibiotic-exposed subjects have been closely monitored up to 24 months regarding species diversity and resistance development, in relation to the natural fluctuations recorded in a control group. A persistent enrichment of resistant strains and erm genes was observed for up to 2 years after clindamycin administration, which emphasizes the importance of the intestinal flora as a reservoir of resistance genes.
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None to declare.
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Acknowledgements |
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This work was supported by grants from the Microbes and Man research programme; Swedish Foundation of Strategic Research.
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References |
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1 Salyers AA, Gupta A, Wang Y. (2004) Human intestinal bacteria as reservoirs for antibiotic resistance genes. Trends Microbiol 12:412–16.[CrossRef][Web of Science][Medline]
2
Sullivan A, Barkholt L, Nord CE. (2003) Lactobacillus acidophilus, Bifidobacterium lactis and Lactobacillus F19 prevent antibiotic-associated ecological disturbances of Bacteroides fragilis in the intestine. J Antimicrob Chemother 52:308–11.
3
Leigh DA and Simmons K. (1978) Effect of clindamycin and lincomycin therapy on faecal flora. J Clin Pathol 31:439–43.
4
Jernberg C, Sullivan A, Edlund C, et al. (2005) Monitoring of antibiotic-induced alterations in the human intestinal microflora and detection of probiotic strains by use of terminal restriction fragment length polymorphism. Appl Environ Microbiol 71:501–6.
5
Franks AH, Harmsen HJ, Raangs GC, et al. (1998) Variations of bacterial populations in human feces measured by fluorescent in situ hybridization with group-specific 16S rRNA-targeted oligonucleotide probes. Appl Environ Microbiol 64:3336–45.
6 Hedberg M and Nord CE. (2003) Antimicrobial susceptibility of Bacteroides fragilis group isolates in Europe. Clin Microbiol Infect 9:475–88.[CrossRef][Web of Science][Medline]
7
Aldridge KE, Ashcraft D, Cambre K, et al. (2001) Multicenter survey of the changing in vitro antimicrobial susceptibilities of clinical isolates of Bacteroides fragilis group, Prevotella, Fusobacterium, Porphyromonas, and Peptostreptococcus species. Antimicrob Agents Chemother 45:1238–43.
8
Shoemaker NB, Vlamakis H, Hayes K, et al. (2001) Evidence for extensive resistance gene transfer among Bacteroides spp. and among Bacteroides and other genera in the human colon. Appl Environ Microbiol 67:561–8.
9 Roberts MC. (2004) Resistance to macrolide, lincosamide, streptogramin, ketolide, and oxazolidinone antibiotics. Mol Biotechnol 28:47–62.[CrossRef][Web of Science][Medline]
10 Whittle G, Shoemaker NB, Salyers AA. (2002) The role of Bacteroides conjugative transposons in the dissemination of antibiotic resistance genes. Cell Mol Life Sci 59:2044–54.[CrossRef][Web of Science][Medline]
11 Salyers AA and Amabile-Cuevas CF. (1997) Why are antibiotic resistance genes so resistant to elimination? Antimicrob Agents Chemother 41:2321–5.[Web of Science][Medline]
12 Vedantam G and Hecht DW. (2003) Antibiotics and anaerobes of gut origin. Curr Opin Microbiol 6:457–61.[CrossRef][Web of Science][Medline]
13 Phillips I, King A, Nord CE, et al. (1992) Antibiotic sensitivity of the Bacteroides fragilis group in Europe. European Study Group. Eur J Clin Microbiol Infect Dis 11:292–304.[CrossRef][Web of Science][Medline]
14 Hecht DW. (2004) Prevalence of antibiotic resistance in anaerobic bacteria: worrisome developments. Clin Infect Dis 39:92–7.[CrossRef][Web of Science][Medline]
15 Ulger (Toprak) N, Celik C, Cakici O, et al. (2004) Antimicrobial susceptibilities of Bacteroides fragilis and Bacteroides thetaiotaomicron strains isolated from clinical specimens and human intestinal microbiota. Anaerobe 10:255–9.[CrossRef][Web of Science][Medline]
16 Roberts MC. (2005) Update on acquired tetracycline resistance genes. FEMS Microbiol Lett 245:195–203.[CrossRef][Web of Science][Medline]
17 Paula GR, Falcao LS, Antunes EN, et al. (2004) Determinants of resistance in Bacteroides fragilis strains according to recent Brazilian profiles of antimicrobial susceptibility. Int J Antimicrob Agents 24:53–8.[Web of Science][Medline]
18 Rasmussen BA, Bush K, Tally FP. (1997) Antimicrobial resistance in anaerobes. Clin Infect Dis 24:Suppl 1, S110–20.
19
Morgan JR, Liu PY, Smith JA. (1976) Semi-microtechnique for the biochemical characterization of anaerobic bacteria. J Clin Microbiol 4:315–8.
20 Jouisimies-Somer HR, Summanen P, Citron D, et al. (2002) Wadsworth-KTL Anaerobic Bacteriology Manual 6th edn (Star publishing company, Belmont, CA).
21
Moraes SR, Goncalves RB, Mouton C, et al. (2000) Use of rep-PCR to define genetic relatedness among Bacteroides fragilis strains. J Med Microbiol 49:279–84.
22 National Committee for Clinical Laboratory Standards. (2004) Methods for Antimicrobial Susceptibility Testing of Anaerobic Bacteria—Sixth Edition: Approved Standards M11-A6.(NCCLS, Wayne, PA, USA).
23
Duerden BI. (1980) The isolation and identification of Bacteroides spp. from the normal human faecal flora. J Med Microbiol 13:69–78.
24 Job ML and Jacobs NF Jr. (1997) Drug-induced Clostridium difficile-associated disease. Drug Saf 17:37–46.[Web of Science][Medline]
25 Sullivan A, Edlund C, Nord CE. (2001) Effect of antimicrobial agents on the ecological balance of human microflora. Lancet Infect Dis 1:101–14.[CrossRef][Medline]
26 Lautenbach E, LaRosa LA, Marr AM, et al. (2003) Changes in the prevalence of vancomycin-resistant enterococci in response to antimicrobial formulary interventions: impact of progressive restrictions on use of vancomycin and third-generation cephalosporins. Clin Infect Dis 36:440–6.[CrossRef][Web of Science][Medline]
27 Nord CE and Heimdahl A. (1988) Impact of different antimicrobial agents on the colonisation resistance in the intestinal tract with special reference to doxycycline. Scand J Infect Dis Suppl 53:50–8.[Medline]
28 Edlund C and Nord CE. (1993) Ecological impact of antimicrobial agents on human intestinal microflora. Alpe Adria Microbiology J 3:137–64.
29
Arthur M, Molinas C, Mabilat C, et al. (1990) Detection of erythromycin resistance by the polymerase chain reaction using primers in conserved regions of erm rRNA methylase genes. Antimicrob Agents Chemother 34:2024–6.
30
Ono T, Shiota S, Hirota K, et al. (2000) Susceptibilities of oral and nasal isolates of Streptococcus mitis and Streptococcus oralis to macrolides and PCR detection of resistance genes. Antimicrob Agents Chemother 44:1078–80.
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