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Anaerobes: Antibiotic resistance, clinical significance, and the role of susceptibility testing
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Anaerobe 12 (2006) 115–121 www.elsevier.com/locate/anaerobe
Mini-review
Anaerobes: Antibiotic resistance, clinical significance, and the role of susceptibility testing David W. Hecht Hines VA Hospital, Loyola University Medical Center, 2160 S. First Avenue, Maywood, IL 60153, USA Received 25 October 2005; accepted 27 October 2005 Available online 6 December 2005
1. Introduction Antimicrobial resistance among many pathogenic anaerobic bacteria has increased significantly over the past two–three decades, paralleling similar trends among nonanaerobic pathogens. Awareness of these disturbing findings is limited among both clinicians and microbiologists due to a number of confounding factors, including lack of isolation and susceptibility testing at most clinical laboratories, non-standardization of testing methods and reporting, and lack of correlation of clinical outcome with resistance. Standardization of testing methods and recommendations for additional susceptibility testing by the Clinical Laboratories Standards Institute (CLSI) were most recently published in 2004, allowing for better comparison among published studies [1]. Correlation of clinical failures with known antibiotic resistance and the discovery and characterization of resistance determinants and their spread has resulted in both increased recognition of the problem and has even resulted in changes to published treatment standards for infections involving anaerobes. 2. Patterns of antibiotic resistance Antimicrobial susceptibility testing of anaerobes is still rarely performed at most medical centers [2]. Thus, tracking of antibiotic resistance patterns is often limited to published reference laboratory studies that include either single site or multi-hospital data that typically report susceptibility of the Bacteroides fragilis group [3–6]. Among reported studies using approved standardized Tel.: +1 708 216 3232; fax: +1 708 216 8198.
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methods, significant variation in susceptibility is seen among different countries, geographic areas within countries, and even medical centers within the same city [4,7–11]. Despite these variable results, some general trends have emerged that have had a direct effect on the recommendations for empiric antibiotic therapy. In particular, clindamycin, cephalosporin, and cephamycin resistance among the B. fragilis group has grown sufficiently large that clinicians and laboratories can no longer assume susceptibility to these agents without testing, with neither national nor local data from other institutions sufficient to predict susceptibility of anaerobes at one’s own medical center [3–5]. A brief overview of current resistance patterns for anaerobic bacteria and known mechanisms of resistance and resistance transfer is provided below [12]. 3. Gram-negative bacilli 3.1. B. fragilis group Although B. fragilis is the most frequently isolated and generally the most susceptible species within the group, resistance to penicillins among all members of the group is high. More than 95% of all species are resistant to penicillin G and ampicillin, o50% susceptible to ticarcillin [6], and only 70% susceptible to piperacillin [3,6,13]. The common mechanism of resistance to penicillin and ampicillin is a chromosomally encoded functional class 2e cephalosporinase, which has not been shown to be transferable [14,15]. Fortunately, this nearly ubiquitous Bacteroides b-lactamase is inhibited by all three of the parenteral b-lactam–b-lactamase inhibitor combinations including ampicillin/sulbactam, ticarcillin/clavulanate, and piperacillin/tazobactam, with less than 2% resistance to these agents reported in most surveys [3].
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Of available cephalosporins and cephamycins with reported efficacy against B. fragilis group infections, cefoxitin remains the most active, with 80–90% of isolates susceptible, followed by cefotetan, although the latter is much less active against non-B. fragilis members of the group [5]. Generally, first and third generation cephalosporins have poor in vitro activity against nearly all members of the B. fragilis group, inhibiting o50% of isolates in most studies, although ceftizoxime has been shown to be clinically efficacious [16]. Resistance to cefoxitin and cefotaxime is encoded by cepA and cfxA blactamase genes, which are transferable via a plasmid or a mobilizable transposon [17]. Carbapenem (imipenem, ertapenem, meropenem) resistance fortunately remains rare, accounting for less than 1% of all B. fragilis group strains worldwide [3,6]. This broad spectrum b-lactam resistance is encoded by one of two nearly identical genes, cfiA and ccrA, expressing a class B metallo-b-lactamase that confers resistance to all b-lactam antibiotics including the three b-lactamase inhibitor combinations [18,19]. Interestingly, DNA homology II subgroup of B. fragilis (5%) are harbingers of cfiA (ccrA), although typically not expressed at a sufficient level to detect in vitro resistance due to the absence of a promoter for the gene [20]. However, high-level expression of these genes can be selected in vitro following a single step mutation under antibiotic pressure. Such selected strains typically are found to have a promoter containing insertion sequence (IS) inserted immediately upstream of the cfiA (ccrA) gene resulting in increased levels of expression of the metallo-b-lactamase [21–23]. Two alternative non-b-lactamase but relatively uncommon mechanisms of resistance to b-lactam agents include either alterations in penicillin-binding-proteins PBP1 and PBP2 or porin mutations [24,25]. These mechanisms result in either a decreased affinity or outer membrane changes that decrease transport for the b-lactam drug, respectively [26]. Clindamycin resistance among B. fragilis group has become widely prevalent over the last two decades, with resistance ranging between 10% and 40% in many surveys worldwide [3,4,7,27]. High-level resistance to clindamycin is mediated by an MLS-type mechanism via 23S rRNA methylases ErmF, ErmFS, ErmG or ErmB [12,28–30]. The corresponding determinants are frequently located on one of several transferable plasmids or conjugative transposons, which likely accounts for their rapid dissemination along with the tetracycline resistant conjugative transposons [31–33]. 5-nitro-imidazole (metronidazole) resistance remains rare among Bacteroides sp., despite its widespread use since 1960. Resistance to metronidazole has been reported from several Western European countries, the Middle East, Asia, Africa, and Canada, while only one strain with an MIC exceeding 16 mg/mL has been reported in the US [3,27,34–36]. Resistance to metronidazole is most commonly expressed by one of seven known nim genes (A–G)
that encode nitroimidazole reductases [37]. Nitroimidazole reductases likely compete with the normal reduction of the 5-nitro-imidazoles prodrugs by ferredoxin. nim gene encoded nitroimidazole reductase transforms the nitro group of the prodrug into an amine derivative resulting in a non-toxic form of the drug [38]. nim genes have been identified on transferable plasmids, including those that closely resemble a clindamycin resistance plasmid, increasing concern about their rapid dissemination [39]. Non-nim gene associated imidazole resistance has also been reported, typically following extensive use of metronidazole in an individual patient. The exact mechanism of resistance is unknown, although strains can be generated in vitro by exposure to metronidazole [40,41]. Among fluoroquinolones, the Food and Drug Administration recently approved moxifloxacin for complicated skin and skin structure infections, that can contain anaerobes. Moxifloxacin exhibits very good but incomplete in vitro activity against a broad range of anaerobes. In fact, in vitro resistance to moxifloxacin and trovafloxacin among B. fragilis group has increased from 30% in 1998 to 43% in 2001 [42]. Resistance among Bacteroides sp. is associated with mutations in gyrA (gyrase) and gyrB and/ or by increased expression of efflux pumps [43–45]. The precise interactions of these different mechanisms and the role of other possible pumps, is not completely worked out. However, mutations that occur in both gyrA, gyrB, and efflux pumps pose potentially significant hurdles to this class of agents among Bacteroides [46]. Transferable fluoroquinolone resistance has not been described among anaerobic bacteria. Tigecycline is the latest FDA approved agent in a new class of antimicrobials, the glycylcyclines, with activity against anaerobes. This compound is a t-butylglyclamido derivative of minocycline that has excellent in vitro activity against all members of the B. fragilis group and most other anaerobes, and is approved for mixed infections of skin and soft tissue and intra-abdominal origins [47]. Resistance to this agent is rare among Bacteroides, and mechanisms of resistance are not known at this time. 3.2. Prevotella, Porphyromonas, and Fusobacterium In general, data on the susceptibility of these organisms (mostly former Bacteroides species) is more limited when compared to the B. fragilis group. Overall, these organisms are more susceptible than the B. fragilis group. Currently, about 50% of Prevotella sp. are resistant to penicillin and ampicillin due to b-lactamase production either of the CfxA-type, or other b-lactamases approaching those of Bacteroides. However, in contrast to the B. fragilis group, many retain susceptibility to piperacillin, cefoxitin, cefotetan, and ceftizoxime at 70–90% [6,7,48–50]. For Porphyromonas sp., resistance to penicillin is much lower ranging between 8% and 17%. The mechanism of resistance is also a cfxA-encoded b-lactamase [51]. Both genera are nearly uniformly susceptible to carbapenems, metronidazole, and
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chloramphenicol, and the b-lactam/b-lactamase inhibitor combinations. However, clindamycin resistance has been observed among both Prevotella sp. (0–11%) and Porphyromonas sp. (0–35%) [6,7]. Clindamycin resistance among Prevotella and Porphyromonas is associated with the presence of either an ermF or ermG gene [52]. Tetracycline resistance among Prevotella sp. have been reported as high as 50%, associated with the presence of tet(Q), tet(M), and tet(W) via a ribosomal protection mechanism, and tet(Q) has been identified in Porphyromonas sp. [53–56]. Transfer of tetracycline resistance among Prevotella has been described via conjugative transposons similar to that of Bacteroides, while transfer of other resistance determinants is unknown [57]. Among Fusobacterium sp., penicillin resistance remains uncommon [58], with B-lactamases the readily identified mechanism of resistance when it is found [53]. In general, 490% of Fusobacterium spp. are susceptible to cephalosporins and cephamycins, including cefoxitin, cefotetan, and ceftizoxime [58,59], while tetracycline resistance has also reported with increasing frequency. Both tet (W) and tet (M) have been identified in tetracycline resistant F. nucleatum [54,60]. 3.3. Other Gram-negative bacilli When looked for, Bilophila wadsworthia is a common organism of the gastrointestinal tract, that frequently produces b-lactamase, and is resistant to penicillin and ampicillin with high MIC90 values also when testing piperacillin and ceftizoxime. However, this organism is susceptible to clindamycin, cefoxitin, b-lactam/b-lactamase inhibitor combinations, carbapenems, and metronidazole [53,61,62]. Campylobacter gracilis (formerly Bacteroides gracilis) is susceptible to most agents tested, including blactam/b-lactamase inhibitor combinations, cefoxitin, ceftizoxime, ceftriaxone, and clindamycin [63]. On the other hand, Sutterella wadsworthensis, often isolated from the same samples and identified as C. gracilis, is far more resistant including clindamycin, ceftizoxime, piperacillin, and/or metronidazole [63]. Campylobacter rectus and C. curvus (formerly Wolinella) vary in their susceptibility to blactams, but remain very susceptible to chloramphenicol, metronidazole, and clindamycin [64]. The mechanisms of resistance and resistance transfer among these three groups of organisms are unknown. 4. Gram-positive bacilli and cocci 4.1. Non-spore forming Gram-positive bacilli The Eubacterium group, Actinomyces, Propionibacterium, and Bifidobacterium are usually susceptible to blactam agents, including the penicillins, cephalosporins and cephamycins, carbapenems, and b-lactam/b-lactamase inhibitor combinations. Lactobacillus spp. are variably susceptible to cephalosporins, and may be inhibited
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effectively only by penicillin [65], while most non-spore forming Gram-positive anaerobes are resistant to metronidazole [66]. 4.2. Spore forming Gram-positive bacilli Clostridium perfringens is generally very susceptible to most anti-anaerobic agents including penicillin [67]. However, non-perfringens Clostridium sp. and C. difficile have highly variable in vitro susceptibility [49,68,69]. Resistance among non-perfringens species includes clindamycin and blactams including penicillin, cephalosporins, and imipenem [70]. Resistance to clindamycin has been attributed to ermQ and ermP in C. perfringens and ermB in C. difficile [71–73]. The genes corresponding to b-lactam resistance have not been identified. Tetracycline resistance is high among C. perfringens 10–76%, with tetA and tetB encoding an efflux protein and a ribosome protection-type protein, respectively [74,75]. C. clostridioforme has been recently been identified as a mixture of three different species, with a majority of strains resistant to penicillin via a blactamase. There is also variable resistance to clindamycin and moxifloxacin [76]. Among C. difficile, clindamycin resistance is widespread, with 490% of strains resistant in one study. Resistance is mediated by the erm gene family of which these determinants are transferable [72,77]. Tetracycline resistance in C. difficile is encoded by tet(M), and may be decreasing in prevalence [77,78]. However, fluoroquinolone resistance has emerged as a new potentially major problem among C. difficile. Outbreaks of C. difficile have been reported with the use of gatifloxacin and levofloxacin, with high MICs reported for many of the outbreak strains [79]. The mechanism of fluoroquinolone resistance appears to be associated with gyrA and gyrB mutations in C. perfringens and C. difficile, with nucleotide substitutions found in topoisomerase IV genes in C. perfringens [5,80,81]. 4.3. Gram-positive cocci At present, only Peptococcus niger remains in this genus, and several of the species in the genus Peptostreptococcus have been reclassified into other genera including Anaerococcus, Finegoldia, Micromonas, Peptoniphilus, and Peptostreptococcus [82]. In general, these organisms have variable resistance to pencillins (7–10%) clindamycin (7–20%), and metronidazole (5–10%), while retaining a much greater susceptibility to b-lactam/b-lactamase inhibitors, cephalosporins, carbapenems, and chloramphenicol. [6,66,83–85]. Fluoroquinolone resistance among Peptostreptococcus has been reported with 14% and 27% resistant to ciprofloxacin and levofloxacin, respectively [6,7]. PBP alterations appear to account for most b-lactam resistance, while the nimB gene has been demonstrated in two highly metronidazole resistant strains of Finegoldia magna. Interestingly, 19 of 21 strains of other Gram-positive cocci that did not demonstrate resistance were also found to have nimB genes, but
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the isolates were not resistant by susceptibility testing [86,87].
clinicians can rely upon in the same manner as for nonanaerobic pathogen testing.
5. Susceptibility testing methods
5.1. Clinical relevance
Although still rarely performed in the US, susceptibility testing of anaerobic bacteria is becoming recommended more frequently for both surveillance and individual isolate purposes (see Ref. [88] for the history of susceptibility testing methods for anaerobes). The most recent rigorously standardized methods for susceptibility testing of anaerobic bacteria were published by CLSI in 2004 [1]. Two methods are approved by CLSI, with a third method approved by the FDA [89]. The reference method remains agar dilution, a highly reproducible standard for testing all anaerobes, and to which other methods have now been correlated. This method, although labor intensive, is appropriate for testing batches of isolates, such as for surveillance purposes, as recommended by CLSI. Currently, CLSI recommends testing X100 anaerobic isolates on an annual basis at individual hospitals using the agar dilution method to monitor for resistance trends. Two alternative methods have also been correlated with the reference standard. The broth micro-dilution method is approved by CLSI, and correlates well with the agar dilution standard when testing all members of the B. fragilis group. This method provides a convenient userfriendly approach, uses the identical broth medium as agar dilution, and allows the user to determine susceptibility for a single isolate to several antibiotics simultaneously. Unfortunately, correlation for non-Bacteroides anaerobes has not proven satisfactory due to poor growth of many of these organisms in broth. CLSI currently does not recommend this method for non-Bacteroides anaerobes, although theoretically a laboratory could validate broth micro-dilution results against the standard themselves for use with other organisms. The FDA approved E-test (AB Biodisk) provides a simple gradient method that has also been validated against the agar dilution standard. This convenient method is best suited for single isolate/ antibiotic testing. Both non-agar dilution methods are well suited for testing individual isolates when indicated, particularly when selection of therapy for positive blood cultures, for persistent infection, for prolonged therapy, or for known resistance of a particular anaerobic species. CLSI recommends anaerobe susceptibility testing for isolates known to be highly virulent, and for which susceptibility cannot be predicted including Bacteroides sp., Prevotella sp., Fusobacterium sp., Clostridium sp., Bilophila wadsworthia, and Sutterella wadworthensis. Other testing methods such as broth disk elution and disk diffusion have failed to consistently correlate with standard methods, and are not recommended for testing. Current and future adoption of the standardized methods worldwide will allow much better comparisons among reported studies, as well as susceptibility results that
Both the in vitro resistance trends and correlation with acquisition of resistance genes gives support to the concern about treatment failures. Few studies have definitively associated resistance with clinical failure. Clinical studies of anaerobic bacteria are confounded by several factors that affect outcome. Most infections involving anaerobes are of the mixed type that also include aerobes and or facultative anaerobes and are typically tissue destructive, requiring debridement. Thus, the role of a resistant anaerobe is sometimes difficult to determine in this setting. While several retrospective studies have correlated clinical failure with antibiotic resistance [90–92], the prospective observational study of Bacteroides bacteremia clearly demonstrates the association of clinical failure with resistant pathogens when a single pathogen can be identified as associated with infection [93]. The results and impact of resistance trends associated with clinical failures has resulted in recent changes in the recommendations for empiric treatment of intra-abdominal infections. Both cefoxitin and cefotetan are now discouraged as first line therapy, and clindamycin is no longer listed as first line therapy [94]. Among maxillofacial infections, a consensus group in Spain lists amoxicillin/ clavulanate as first line therapy over that of penicillin [95]. As additional data and studies demonstrate the rise in resistance to some agents, and association with clinical failures, other recommendations are likely. 6. Conclusions Antibiotic resistance among anaerobes continues to rise, which is really not surprising given the parallel observations among aerobes over the last several decades. Awareness of this problem through more surveillance testing using standardized methods, and appropriate use of individual isolate testing that directly affects antibiotic choice will hopefully result in better patient outcomes, although additional studies in this area are sorely needed. Acknowledgements DH is supported by National Institutes of Health RO1AI050122 and The Office of Research and Development, Medical Research Services, Department of Veterans Affairs. References [1] National Committee for Clinical Laboratory Standards. Methods for antimicrobial susceptibility testing of anaerobic bacteria. 6th ed. M11-A6, 2004.
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[63] Molitoris E, Wexler HM, Finegold SM. Sources and antimicrobial susceptibilities of Campylobacter gracilis and Sutterella wadsworthensis. Clin Infect Dis 1997;25(Suppl. 2):S264–5. [64] Wexler HM, Molitoris D, St John S, Vu A, Read EK, Finegold SM. In vitro activities of faropenem against 579 strains of anaerobic bacteria. Antimicrob Agents Chemother 2002;46:3669–75. [65] Goldstein EJ, Citron DM, Vreni MC, Tyrrell K, Warren Y. Activities of gemifloxacin (SB 265805, LB20304) compared to those of other oral antimicrobial agents against unusual anaerobes. Antimicrob Agents Chemother 1999;43:2726–30. [66] Pelak BA, Citron DM, Motyl M, Goldstein EJ, Woods GL, Teppler H. Comparative in vitro activities of ertapenem against bacterial pathogens from patients with acute pelvic infection. J Antimicrob Chemother 2002;50:735–41. [67] Hecht DW, Wexler HM. In vitro susceptibility of anaerobes to quinolones in the United States. Clin Infect Dis 1996;23(Suppl. 1):S2–8. [68] Goldstein EJ, Citron DM, Warren Y, Tyrrell K, Merriam CV. In vitro activity of gemifloxacin (SB 265805) against anaerobes. Antimicrob Agents Chemother 1999;43:2231–5. [69] Goldstein EJ, Citron DM, Vreni MC, Tyrrell K, Warren Y. Activities of gemifloxacin (SB 265805, LB20304) compared to those of other oral antimicrobial agents against unusual anaerobes. Antimicrob Agents Chemother 1999;43:2726–30. [70] Jamal WY, Mokaddas EM, Verghese TL, Rotimi VO. In vitro activity of 15 antimicrobial agents against clinical isolates of Clostridium difficile in Kuwait. Int J Antimicrob Agents 2002;20:270–4. [71] Ackermann G, Degner A, Cohen SH, Silva Jr J, Rodloff AC. Prevalence and association of macrolide–lincosamide–streptogramin B (MLS(B)) resistance with resistance to moxifloxacin in Clostridium difficile. J Antimicrob Chemother 2003;51:599–603. [72] Farrow KA, Lyras D, Rood JI. Genomic analysis of the erythromycin resistance element Tn5398 from Clostridium difficile. Microbiology 2001;147:2717–28. [73] Farrow KA, Lyras D, Polekhina G, Koutsis K, Parker MW, Rood JI. Identification of essential residues in the Erm(B) rRNA methyltransferase of Clostridium perfringens. Antimicrob Agents Chemother 2002;46:1253–61. [74] Johansson A, Greko C, Engstrom BE, Karlsson M. Antimicrobial susceptibility of Swedish, Norwegian and Danish isolates of Clostridium perfringens from poultry, and distribution of tetracycline resistance genes. Vet Microbiol 2004;99:251–7. [75] Sloan J, McMurry LM, Lyras D, Levy SB, Rood JI. The Clostridium perfringens Tet P determinant comprises two overlapping genes: tetA(P), which mediates active tetracycline efflux, and tetB(P), which is related to the ribosomal protection family of tetracycline-resistance determinants. Mol Microbiol 1994;11:403–15. [76] Finegold SM, Song Y, Liu C, Hecht DW, Summanen P, Kononen E, et al. Clostridium clostridioforme: a mixture of three clinically important species. Eur J Clin Microbiol Infect Dis 2005;24: 319–24. [77] Bendle JS, James PA, Bennett PM, Avison MB, Macgowan AP, Al Shafi KM. Resistance determinants in strains of Clostridium difficile from two geographically distinct populations. Int J Antimicrob Agents 2004;24:619–21. [78] Adams V, Lyras D, Farrow KA, Rood JI. The clostridial mobilisable transposons. Cell Mol Life Sci 2002;59:2033–43. [79] Changela U, Cannon JP, Aneziokoro C, Shah PS, Thottapurathu L, Lentino J. Risk factors and mortality associated with Clostridium difficile-associated diarrhoea at a VA hospital. Int J Antimicrob Agents 2004;24:562–6. [80] Dridi L, Tankovic J, Burghoffer B, Barbut F, Petit JC. gyrA and gyrB mutations are implicated in cross-resistance to Ciprofloxacin and moxifloxacin in Clostridium difficile. Antimicrob Agents Chemother 2002;46:3418–21. [81] Rafii F, Park M, Novak JS. Alterations in DNA gyrase and topoisomerase IV in resistant mutants of Clostridium perfringens found after in vitro treatment with fluoroquinolones. Antimicrob Agents Chemother 2005;49:488–92.
ARTICLE IN PRESS D.W. Hecht / Anaerobe 12 (2006) 115–121 [82] Song Y, Liu C, McTeague M, Finegold SM. 16S ribosomal DNA sequence-based analysis of clinically significant Gram-positive anaerobic cocci. J Clin Microbiol 2003;41:1363–9. [83] Hoellman DB, Kelly LM, Jacobs MR, Appelbaum PC. Comparative antianaerobic activity of BMS 284756. Antimicrob Agents Chemother 2001;45:589–92. [84] Brazier JS, Hall V, Morris TE, Gal M, Duerden BI. Antibiotic susceptibilities of Gram-positive anaerobic cocci: results of a sentinel study in England and Wales. J Antimicrob Chemother 2003;52: 224–8. [85] Baquero F, Reig M. Resistance of anaerobic bacteria to antimicrobial agents in Spain. Eur J Clin Microbiol Infect Dis 1992;11:1016–20. [86] Handal T, Olsen I, Walker CB, Caugant DA. Detection and characterization of beta-lactamase genes in subgingival bacteria from patients with refractory periodontitis. FEMS Microbiol Lett 2005;242:319–24. [87] Theron MM, Janse Van Rensburg MN, Chalkley LJ. Nitroimidazole resistance genes (nimB) in anaerobic Gram-positive cocci (previously Peptostreptococcus spp.). J Antimicrob Chemother 2004;54:240–2. [88] Hecht DW. Evolution of anaerobe susceptibility testing in the United States. Clin Infect Dis 2002;35:S28–35. [89] Rosenblatt JE, Gustafson DR. Evaluation of the Etest for susceptibility testing of anaerobic bacteria. Diagn Microbiol Infect Dis 1995;22:279–84.
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[90] Bieluch VM, Cuchural GJ, Snydman DR, Gorbach SL, Tally FP. Clinical importance of cefoxitin-resistant Bacteroides fragilis isolates. Diagn Microbiol Infect Dis 1987;7:119–26. [91] Snydman DR, Cuchural Jr GJ, McDermott L, Gill M. Correlation of various in vitro testing methods with clinical outcomes in patients with Bacteroides fragilis group infections treated with cefoxitin: a retrospective analysis. Antimicrob Agents Chemother 1992;36:540–4. [92] Dalmau D, Cayouette M, Lamothe F, Vincelette J, Lachance N, Bourgault AM, et al. Clindamycin resistance in the Bacteroides fragilis group: association with hospital-acquired infections. Clin Infect Dis 1997;24:874–7. [93] Nguyen MH, Yu VL, Morris AJ, McDermott L, Wagener MW, Harrell L, et al. Antimicrobial resistance and clinical outcome of Bacteroides bacteremia: findings of a multicenter prospective observational trial. Clin Infect Dis 2000;30:870–6. [94] Solomkin JS, Mazuski JE, Baron EJ, Sawyer RG, Nathens AB, DiPiro JT, et al. Guidelines for the selection of anti-infective agents for complicated intra-abdominal infections. Clin Infect Dis 2003;37:997–1005. [95] Bascones MA, Aguirre Urizar JM, Bermejo FA, Blanco CA, GayEscoda C, Gonzalez-Moles MA, et al. Consensus statement on antimicrobial treatment of odontogenic bacterial infections. Med Oral Patol Oral Cir Bucal 2004;9:369–76.
Anaerobe 12 (2006) 115–121 www.elsevier.com/locate/anaerobe
Mini-review
Anaerobes: Antibiotic resistance, clinical significance, and the role of susceptibility testing David W. Hecht Hines VA Hospital, Loyola University Medical Center, 2160 S. First Avenue, Maywood, IL 60153, USA Received 25 October 2005; accepted 27 October 2005 Available online 6 December 2005
1. Introduction Antimicrobial resistance among many pathogenic anaerobic bacteria has increased significantly over the past two–three decades, paralleling similar trends among nonanaerobic pathogens. Awareness of these disturbing findings is limited among both clinicians and microbiologists due to a number of confounding factors, including lack of isolation and susceptibility testing at most clinical laboratories, non-standardization of testing methods and reporting, and lack of correlation of clinical outcome with resistance. Standardization of testing methods and recommendations for additional susceptibility testing by the Clinical Laboratories Standards Institute (CLSI) were most recently published in 2004, allowing for better comparison among published studies [1]. Correlation of clinical failures with known antibiotic resistance and the discovery and characterization of resistance determinants and their spread has resulted in both increased recognition of the problem and has even resulted in changes to published treatment standards for infections involving anaerobes. 2. Patterns of antibiotic resistance Antimicrobial susceptibility testing of anaerobes is still rarely performed at most medical centers [2]. Thus, tracking of antibiotic resistance patterns is often limited to published reference laboratory studies that include either single site or multi-hospital data that typically report susceptibility of the Bacteroides fragilis group [3–6]. Among reported studies using approved standardized Tel.: +1 708 216 3232; fax: +1 708 216 8198.
E-mail address: [email protected]. 1075-9964/$ - see front matter Published by Elsevier Ltd. doi:10.1016/j.anaerobe.2005.10.004
methods, significant variation in susceptibility is seen among different countries, geographic areas within countries, and even medical centers within the same city [4,7–11]. Despite these variable results, some general trends have emerged that have had a direct effect on the recommendations for empiric antibiotic therapy. In particular, clindamycin, cephalosporin, and cephamycin resistance among the B. fragilis group has grown sufficiently large that clinicians and laboratories can no longer assume susceptibility to these agents without testing, with neither national nor local data from other institutions sufficient to predict susceptibility of anaerobes at one’s own medical center [3–5]. A brief overview of current resistance patterns for anaerobic bacteria and known mechanisms of resistance and resistance transfer is provided below [12]. 3. Gram-negative bacilli 3.1. B. fragilis group Although B. fragilis is the most frequently isolated and generally the most susceptible species within the group, resistance to penicillins among all members of the group is high. More than 95% of all species are resistant to penicillin G and ampicillin, o50% susceptible to ticarcillin [6], and only 70% susceptible to piperacillin [3,6,13]. The common mechanism of resistance to penicillin and ampicillin is a chromosomally encoded functional class 2e cephalosporinase, which has not been shown to be transferable [14,15]. Fortunately, this nearly ubiquitous Bacteroides b-lactamase is inhibited by all three of the parenteral b-lactam–b-lactamase inhibitor combinations including ampicillin/sulbactam, ticarcillin/clavulanate, and piperacillin/tazobactam, with less than 2% resistance to these agents reported in most surveys [3].
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Of available cephalosporins and cephamycins with reported efficacy against B. fragilis group infections, cefoxitin remains the most active, with 80–90% of isolates susceptible, followed by cefotetan, although the latter is much less active against non-B. fragilis members of the group [5]. Generally, first and third generation cephalosporins have poor in vitro activity against nearly all members of the B. fragilis group, inhibiting o50% of isolates in most studies, although ceftizoxime has been shown to be clinically efficacious [16]. Resistance to cefoxitin and cefotaxime is encoded by cepA and cfxA blactamase genes, which are transferable via a plasmid or a mobilizable transposon [17]. Carbapenem (imipenem, ertapenem, meropenem) resistance fortunately remains rare, accounting for less than 1% of all B. fragilis group strains worldwide [3,6]. This broad spectrum b-lactam resistance is encoded by one of two nearly identical genes, cfiA and ccrA, expressing a class B metallo-b-lactamase that confers resistance to all b-lactam antibiotics including the three b-lactamase inhibitor combinations [18,19]. Interestingly, DNA homology II subgroup of B. fragilis (5%) are harbingers of cfiA (ccrA), although typically not expressed at a sufficient level to detect in vitro resistance due to the absence of a promoter for the gene [20]. However, high-level expression of these genes can be selected in vitro following a single step mutation under antibiotic pressure. Such selected strains typically are found to have a promoter containing insertion sequence (IS) inserted immediately upstream of the cfiA (ccrA) gene resulting in increased levels of expression of the metallo-b-lactamase [21–23]. Two alternative non-b-lactamase but relatively uncommon mechanisms of resistance to b-lactam agents include either alterations in penicillin-binding-proteins PBP1 and PBP2 or porin mutations [24,25]. These mechanisms result in either a decreased affinity or outer membrane changes that decrease transport for the b-lactam drug, respectively [26]. Clindamycin resistance among B. fragilis group has become widely prevalent over the last two decades, with resistance ranging between 10% and 40% in many surveys worldwide [3,4,7,27]. High-level resistance to clindamycin is mediated by an MLS-type mechanism via 23S rRNA methylases ErmF, ErmFS, ErmG or ErmB [12,28–30]. The corresponding determinants are frequently located on one of several transferable plasmids or conjugative transposons, which likely accounts for their rapid dissemination along with the tetracycline resistant conjugative transposons [31–33]. 5-nitro-imidazole (metronidazole) resistance remains rare among Bacteroides sp., despite its widespread use since 1960. Resistance to metronidazole has been reported from several Western European countries, the Middle East, Asia, Africa, and Canada, while only one strain with an MIC exceeding 16 mg/mL has been reported in the US [3,27,34–36]. Resistance to metronidazole is most commonly expressed by one of seven known nim genes (A–G)
that encode nitroimidazole reductases [37]. Nitroimidazole reductases likely compete with the normal reduction of the 5-nitro-imidazoles prodrugs by ferredoxin. nim gene encoded nitroimidazole reductase transforms the nitro group of the prodrug into an amine derivative resulting in a non-toxic form of the drug [38]. nim genes have been identified on transferable plasmids, including those that closely resemble a clindamycin resistance plasmid, increasing concern about their rapid dissemination [39]. Non-nim gene associated imidazole resistance has also been reported, typically following extensive use of metronidazole in an individual patient. The exact mechanism of resistance is unknown, although strains can be generated in vitro by exposure to metronidazole [40,41]. Among fluoroquinolones, the Food and Drug Administration recently approved moxifloxacin for complicated skin and skin structure infections, that can contain anaerobes. Moxifloxacin exhibits very good but incomplete in vitro activity against a broad range of anaerobes. In fact, in vitro resistance to moxifloxacin and trovafloxacin among B. fragilis group has increased from 30% in 1998 to 43% in 2001 [42]. Resistance among Bacteroides sp. is associated with mutations in gyrA (gyrase) and gyrB and/ or by increased expression of efflux pumps [43–45]. The precise interactions of these different mechanisms and the role of other possible pumps, is not completely worked out. However, mutations that occur in both gyrA, gyrB, and efflux pumps pose potentially significant hurdles to this class of agents among Bacteroides [46]. Transferable fluoroquinolone resistance has not been described among anaerobic bacteria. Tigecycline is the latest FDA approved agent in a new class of antimicrobials, the glycylcyclines, with activity against anaerobes. This compound is a t-butylglyclamido derivative of minocycline that has excellent in vitro activity against all members of the B. fragilis group and most other anaerobes, and is approved for mixed infections of skin and soft tissue and intra-abdominal origins [47]. Resistance to this agent is rare among Bacteroides, and mechanisms of resistance are not known at this time. 3.2. Prevotella, Porphyromonas, and Fusobacterium In general, data on the susceptibility of these organisms (mostly former Bacteroides species) is more limited when compared to the B. fragilis group. Overall, these organisms are more susceptible than the B. fragilis group. Currently, about 50% of Prevotella sp. are resistant to penicillin and ampicillin due to b-lactamase production either of the CfxA-type, or other b-lactamases approaching those of Bacteroides. However, in contrast to the B. fragilis group, many retain susceptibility to piperacillin, cefoxitin, cefotetan, and ceftizoxime at 70–90% [6,7,48–50]. For Porphyromonas sp., resistance to penicillin is much lower ranging between 8% and 17%. The mechanism of resistance is also a cfxA-encoded b-lactamase [51]. Both genera are nearly uniformly susceptible to carbapenems, metronidazole, and
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chloramphenicol, and the b-lactam/b-lactamase inhibitor combinations. However, clindamycin resistance has been observed among both Prevotella sp. (0–11%) and Porphyromonas sp. (0–35%) [6,7]. Clindamycin resistance among Prevotella and Porphyromonas is associated with the presence of either an ermF or ermG gene [52]. Tetracycline resistance among Prevotella sp. have been reported as high as 50%, associated with the presence of tet(Q), tet(M), and tet(W) via a ribosomal protection mechanism, and tet(Q) has been identified in Porphyromonas sp. [53–56]. Transfer of tetracycline resistance among Prevotella has been described via conjugative transposons similar to that of Bacteroides, while transfer of other resistance determinants is unknown [57]. Among Fusobacterium sp., penicillin resistance remains uncommon [58], with B-lactamases the readily identified mechanism of resistance when it is found [53]. In general, 490% of Fusobacterium spp. are susceptible to cephalosporins and cephamycins, including cefoxitin, cefotetan, and ceftizoxime [58,59], while tetracycline resistance has also reported with increasing frequency. Both tet (W) and tet (M) have been identified in tetracycline resistant F. nucleatum [54,60]. 3.3. Other Gram-negative bacilli When looked for, Bilophila wadsworthia is a common organism of the gastrointestinal tract, that frequently produces b-lactamase, and is resistant to penicillin and ampicillin with high MIC90 values also when testing piperacillin and ceftizoxime. However, this organism is susceptible to clindamycin, cefoxitin, b-lactam/b-lactamase inhibitor combinations, carbapenems, and metronidazole [53,61,62]. Campylobacter gracilis (formerly Bacteroides gracilis) is susceptible to most agents tested, including blactam/b-lactamase inhibitor combinations, cefoxitin, ceftizoxime, ceftriaxone, and clindamycin [63]. On the other hand, Sutterella wadsworthensis, often isolated from the same samples and identified as C. gracilis, is far more resistant including clindamycin, ceftizoxime, piperacillin, and/or metronidazole [63]. Campylobacter rectus and C. curvus (formerly Wolinella) vary in their susceptibility to blactams, but remain very susceptible to chloramphenicol, metronidazole, and clindamycin [64]. The mechanisms of resistance and resistance transfer among these three groups of organisms are unknown. 4. Gram-positive bacilli and cocci 4.1. Non-spore forming Gram-positive bacilli The Eubacterium group, Actinomyces, Propionibacterium, and Bifidobacterium are usually susceptible to blactam agents, including the penicillins, cephalosporins and cephamycins, carbapenems, and b-lactam/b-lactamase inhibitor combinations. Lactobacillus spp. are variably susceptible to cephalosporins, and may be inhibited
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effectively only by penicillin [65], while most non-spore forming Gram-positive anaerobes are resistant to metronidazole [66]. 4.2. Spore forming Gram-positive bacilli Clostridium perfringens is generally very susceptible to most anti-anaerobic agents including penicillin [67]. However, non-perfringens Clostridium sp. and C. difficile have highly variable in vitro susceptibility [49,68,69]. Resistance among non-perfringens species includes clindamycin and blactams including penicillin, cephalosporins, and imipenem [70]. Resistance to clindamycin has been attributed to ermQ and ermP in C. perfringens and ermB in C. difficile [71–73]. The genes corresponding to b-lactam resistance have not been identified. Tetracycline resistance is high among C. perfringens 10–76%, with tetA and tetB encoding an efflux protein and a ribosome protection-type protein, respectively [74,75]. C. clostridioforme has been recently been identified as a mixture of three different species, with a majority of strains resistant to penicillin via a blactamase. There is also variable resistance to clindamycin and moxifloxacin [76]. Among C. difficile, clindamycin resistance is widespread, with 490% of strains resistant in one study. Resistance is mediated by the erm gene family of which these determinants are transferable [72,77]. Tetracycline resistance in C. difficile is encoded by tet(M), and may be decreasing in prevalence [77,78]. However, fluoroquinolone resistance has emerged as a new potentially major problem among C. difficile. Outbreaks of C. difficile have been reported with the use of gatifloxacin and levofloxacin, with high MICs reported for many of the outbreak strains [79]. The mechanism of fluoroquinolone resistance appears to be associated with gyrA and gyrB mutations in C. perfringens and C. difficile, with nucleotide substitutions found in topoisomerase IV genes in C. perfringens [5,80,81]. 4.3. Gram-positive cocci At present, only Peptococcus niger remains in this genus, and several of the species in the genus Peptostreptococcus have been reclassified into other genera including Anaerococcus, Finegoldia, Micromonas, Peptoniphilus, and Peptostreptococcus [82]. In general, these organisms have variable resistance to pencillins (7–10%) clindamycin (7–20%), and metronidazole (5–10%), while retaining a much greater susceptibility to b-lactam/b-lactamase inhibitors, cephalosporins, carbapenems, and chloramphenicol. [6,66,83–85]. Fluoroquinolone resistance among Peptostreptococcus has been reported with 14% and 27% resistant to ciprofloxacin and levofloxacin, respectively [6,7]. PBP alterations appear to account for most b-lactam resistance, while the nimB gene has been demonstrated in two highly metronidazole resistant strains of Finegoldia magna. Interestingly, 19 of 21 strains of other Gram-positive cocci that did not demonstrate resistance were also found to have nimB genes, but
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the isolates were not resistant by susceptibility testing [86,87].
clinicians can rely upon in the same manner as for nonanaerobic pathogen testing.
5. Susceptibility testing methods
5.1. Clinical relevance
Although still rarely performed in the US, susceptibility testing of anaerobic bacteria is becoming recommended more frequently for both surveillance and individual isolate purposes (see Ref. [88] for the history of susceptibility testing methods for anaerobes). The most recent rigorously standardized methods for susceptibility testing of anaerobic bacteria were published by CLSI in 2004 [1]. Two methods are approved by CLSI, with a third method approved by the FDA [89]. The reference method remains agar dilution, a highly reproducible standard for testing all anaerobes, and to which other methods have now been correlated. This method, although labor intensive, is appropriate for testing batches of isolates, such as for surveillance purposes, as recommended by CLSI. Currently, CLSI recommends testing X100 anaerobic isolates on an annual basis at individual hospitals using the agar dilution method to monitor for resistance trends. Two alternative methods have also been correlated with the reference standard. The broth micro-dilution method is approved by CLSI, and correlates well with the agar dilution standard when testing all members of the B. fragilis group. This method provides a convenient userfriendly approach, uses the identical broth medium as agar dilution, and allows the user to determine susceptibility for a single isolate to several antibiotics simultaneously. Unfortunately, correlation for non-Bacteroides anaerobes has not proven satisfactory due to poor growth of many of these organisms in broth. CLSI currently does not recommend this method for non-Bacteroides anaerobes, although theoretically a laboratory could validate broth micro-dilution results against the standard themselves for use with other organisms. The FDA approved E-test (AB Biodisk) provides a simple gradient method that has also been validated against the agar dilution standard. This convenient method is best suited for single isolate/ antibiotic testing. Both non-agar dilution methods are well suited for testing individual isolates when indicated, particularly when selection of therapy for positive blood cultures, for persistent infection, for prolonged therapy, or for known resistance of a particular anaerobic species. CLSI recommends anaerobe susceptibility testing for isolates known to be highly virulent, and for which susceptibility cannot be predicted including Bacteroides sp., Prevotella sp., Fusobacterium sp., Clostridium sp., Bilophila wadsworthia, and Sutterella wadworthensis. Other testing methods such as broth disk elution and disk diffusion have failed to consistently correlate with standard methods, and are not recommended for testing. Current and future adoption of the standardized methods worldwide will allow much better comparisons among reported studies, as well as susceptibility results that
Both the in vitro resistance trends and correlation with acquisition of resistance genes gives support to the concern about treatment failures. Few studies have definitively associated resistance with clinical failure. Clinical studies of anaerobic bacteria are confounded by several factors that affect outcome. Most infections involving anaerobes are of the mixed type that also include aerobes and or facultative anaerobes and are typically tissue destructive, requiring debridement. Thus, the role of a resistant anaerobe is sometimes difficult to determine in this setting. While several retrospective studies have correlated clinical failure with antibiotic resistance [90–92], the prospective observational study of Bacteroides bacteremia clearly demonstrates the association of clinical failure with resistant pathogens when a single pathogen can be identified as associated with infection [93]. The results and impact of resistance trends associated with clinical failures has resulted in recent changes in the recommendations for empiric treatment of intra-abdominal infections. Both cefoxitin and cefotetan are now discouraged as first line therapy, and clindamycin is no longer listed as first line therapy [94]. Among maxillofacial infections, a consensus group in Spain lists amoxicillin/ clavulanate as first line therapy over that of penicillin [95]. As additional data and studies demonstrate the rise in resistance to some agents, and association with clinical failures, other recommendations are likely. 6. Conclusions Antibiotic resistance among anaerobes continues to rise, which is really not surprising given the parallel observations among aerobes over the last several decades. Awareness of this problem through more surveillance testing using standardized methods, and appropriate use of individual isolate testing that directly affects antibiotic choice will hopefully result in better patient outcomes, although additional studies in this area are sorely needed. Acknowledgements DH is supported by National Institutes of Health RO1AI050122 and The Office of Research and Development, Medical Research Services, Department of Veterans Affairs. References [1] National Committee for Clinical Laboratory Standards. Methods for antimicrobial susceptibility testing of anaerobic bacteria. 6th ed. M11-A6, 2004.
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